diff --git a/.Rprofile b/.Rprofile index 5b36bc35..81b960f5 100644 --- a/.Rprofile +++ b/.Rprofile @@ -1,26 +1 @@ source("renv/activate.R") - -# Set the repos using the renv.lock file -renv_json <- jsonlite::read_json("renv.lock") -renv_r_repos <- renv_json$R$Repositories - -# Extract the names -repo_names <- purrr::flatten_chr( - purrr::map(renv_r_repos, - ~ .x$Name) -) - -# Extract the URLs -repo_urls <- purrr::flatten_chr( - purrr::map(renv_r_repos, - ~ .x$URL) -) - -# Set the repo names -names(repo_urls) <- repo_names - -# Set the options -options(repos = repo_urls) - -# Remove all these objects -rm(renv_json, renv_r_repos, repo_names, repo_urls) diff --git a/.github/workflows/build-docker.yml b/.github/workflows/build-docker.yml index 13cfb828..22feafa8 100644 --- a/.github/workflows/build-docker.yml +++ b/.github/workflows/build-docker.yml @@ -19,16 +19,27 @@ on: - renv.lock - requirements.txt - current-modules.json + - .github/workflows/build-docker.yml -# A workflow run is made up of one or more jobs that can run sequentially or in parallel +env: + REGISTRY_IMAGE: ccdl/training_rstudio jobs: - # This workflow contains a single job called "build" build: - # The type of runner that the job will run on - runs-on: ubuntu-latest + strategy: + fail-fast: false + matrix: + include: + - platform: linux/amd64 + runner: ubuntu-latest + - platform: linux/arm64 + runner: ubuntu-24.04-arm + runs-on: ${{ matrix.runner }} - # Steps represent a sequence of tasks that will be executed as part of the job steps: + - name: Prepare env variables + run: | + platform=${{ matrix.platform }} + echo "PLATFORM_PAIR=${platform//\//-}" >> $GITHUB_ENV - name: Check out the repo uses: actions/checkout@v5 @@ -36,7 +47,7 @@ jobs: if: startsWith(github.ref, 'refs/tags/') run: | GIT_TAG=${GITHUB_REF#refs/tags/} - MODULES_TAG=$(jq -r '.release-tag' current-modules.json) + MODULES_TAG=$(jq -r '."release-tag"' current-modules.json) if [ "$GIT_TAG" != "$MODULES_TAG" ]; then echo "Error: current-modules.json release-tag ($MODULES_TAG) does not match git tag ($GIT_TAG)" echo "Please update release-tag in current-modules.json to match the git tag (and modules, if needed), then update the GitHub release accordingly." @@ -44,23 +55,22 @@ jobs: fi - name: Load 1Password secrets - uses: 1password/load-secrets-action@v2 + uses: 1password/load-secrets-action@v3 with: export-env: true env: OP_SERVICE_ACCOUNT_TOKEN: ${{ secrets.TRAINING_OP_SERVICE_ACCOUNT_TOKEN }} DOCKER_USER: ${{ secrets.OP_DOCKER_USERNAME }} DOCKER_PASSWORD: ${{ secrets.OP_DOCKER_PASSWORD }} - ACTION_MONITORING_SLACK: ${{ secrets.OP_ACTION_MONITORING_SLACK }} - # Login to Dockerhub - name: Login to DockerHub uses: docker/login-action@v3 with: username: ${{ env.DOCKER_USER }} password: ${{ env.DOCKER_PASSWORD }} - # set up Docker build + - name: Set up QEMU + uses: docker/setup-qemu-action@v3 - name: Set up Docker Buildx uses: docker/setup-buildx-action@v3 @@ -68,7 +78,74 @@ jobs: id: meta uses: docker/metadata-action@v5 with: - images: ccdl/training_rstudio + images: ${{ env.REGISTRY_IMAGE }} + + # Build Docker image, push only on push events + - name: Build Docker image + id: build + uses: docker/build-push-action@v6 + with: + push: ${{ github.event_name == 'push' }} + platforms: ${{ matrix.platform }} + outputs: type=image,push-by-digest=true,name-canonical=true + labels: ${{ steps.meta.outputs.labels }} + tags: ${{ env.REGISTRY_IMAGE }} + cache-from: type=registry,ref=${{ env.REGISTRY_IMAGE }}:buildcache-${{ env.PLATFORM_PAIR }} + cache-to: type=registry,ref=${{ env.REGISTRY_IMAGE }}:buildcache-${{ env.PLATFORM_PAIR }},mode=max + + - name: Export digest + if: github.event_name == 'push' + run: | + mkdir -p ${{ runner.temp }}/digests + digest="${{ steps.build.outputs.digest }}" + touch "${{ runner.temp }}/digests/${digest#sha256:}" + + - name: Upload digest + if: github.event_name == 'push' + uses: actions/upload-artifact@v4 + with: + name: digests-${{ env.PLATFORM_PAIR }} + path: ${{ runner.temp }}/digests/* + if-no-files-found: error + retention-days: 1 + + merge: + runs-on: ubuntu-latest + needs: + - build + # only merge the manifests and push on push events + if: github.event_name == 'push' + steps: + - name: Load 1Password secrets + uses: 1password/load-secrets-action@v3 + with: + export-env: true + env: + OP_SERVICE_ACCOUNT_TOKEN: ${{ secrets.TRAINING_OP_SERVICE_ACCOUNT_TOKEN }} + DOCKER_USER: ${{ secrets.OP_DOCKER_USERNAME }} + DOCKER_PASSWORD: ${{ secrets.OP_DOCKER_PASSWORD }} + ACTION_MONITORING_SLACK: ${{ secrets.OP_ACTION_MONITORING_SLACK }} + + - name: Download digests + uses: actions/download-artifact@v4 + with: + path: ${{ runner.temp }}/digests + pattern: digests-* + merge-multiple: true + - name: Login to DockerHub + uses: docker/login-action@v3 + with: + username: ${{ env.DOCKER_USER }} + password: ${{ env.DOCKER_PASSWORD }} + + - name: Set up Docker Buildx + uses: docker/setup-buildx-action@v3 + + - name: Docker meta + id: meta + uses: docker/metadata-action@v5 + with: + images: ${{ env.REGISTRY_IMAGE }} # each github tag will create a matching tag on dockerhub, # with the most recent given the "latest" tag # the most recent push to master will get an "edge" tag @@ -76,18 +153,19 @@ jobs: type=ref,event=tag type=edge,branch=master - # Build Docker image, push only on push events - - name: Build Docker image - uses: docker/build-push-action@v5 - with: - push: ${{ github.event_name == 'push' }} - tags: ${{ steps.meta.outputs.tags }} - cache-from: type=registry,ref=ccdl/training_rstudio:buildcache - cache-to: type=registry,ref=ccdl/training_rstudio:buildcache,mode=max + - name: Create manifest list and push + working-directory: ${{ runner.temp }}/digests + run: | + docker buildx imagetools create $(jq -cr '.tags | map("-t " + .) | join(" ")' <<< "$DOCKER_METADATA_OUTPUT_JSON") \ + $(printf '${{ env.REGISTRY_IMAGE }}@sha256:%s ' *) + + - name: Inspect image + run: | + docker buildx imagetools inspect ${{ env.REGISTRY_IMAGE }}:${{ steps.meta.outputs.version }} # If we have a failure, Slack us - name: Report failure to Slack - if: ${{ github.event_name == 'push' }} + if: ${{ failure() }} uses: ravsamhq/notify-slack-action@v2 with: status: ${{ job.status }} diff --git a/.github/workflows/make-live.yml b/.github/workflows/make-live.yml index d10d0a2c..c37b4f6f 100644 --- a/.github/workflows/make-live.yml +++ b/.github/workflows/make-live.yml @@ -14,20 +14,22 @@ jobs: make-live: # The type of runner that the job will run on runs-on: ubuntu-latest - container: - image: ccdl/training_rstudio:edge steps: + - name: Free disk space + run: | + sudo rm -rf /usr/local/lib/android + sudo rm -rf /usr/share/dotnet + sudo rm -rf /opt/ghc + sudo rm -rf /usr/local/share/boost + sudo rm -rf "$AGENT_TOOLSDIRECTORY" + # Print free disk space + df -h + # Checks-out your repository under $GITHUB_WORKSPACE, so your job can access it - name: Checkout code uses: actions/checkout@v4 - - name: Configure git - run: | - git config --global --add safe.directory "$GITHUB_WORKSPACE" - git config --local user.email "actions@github.com" - git config --local user.name "GitHub Actions" - - name: Load 1Password secrets uses: 1password/load-secrets-action@v2 with: @@ -43,15 +45,26 @@ jobs: env: AWS_DEFAULT_REGION: us-east-1 run: | - aws s3 sync s3://ccdl-training-data/training-modules/ . + aws s3 sync s3://ccdl-training-data/training-modules/ . --no-progress + + - name: Pull latest Docker image + run: | + docker pull ccdl/training_rstudio:edge - name: Render notebooks env: RENDER_RMD: ${{ github.event.inputs.rendering }} - run: bash scripts/render-live.sh + run: | + docker run --rm \ + --mount type=bind,source="$GITHUB_WORKSPACE",target=/training-modules \ + -w /training-modules \ + -e RENDER_RMD \ + ccdl/training_rstudio:edge \ + bash scripts/render-live.sh # Make changes to pull request here - name: Create PR with rendered notebooks + id: cpr uses: peter-evans/create-pull-request@v6 with: token: ${{ env.DOCS_BOT_GITHUB_TOKEN }} diff --git a/.github/workflows/render-rmds.yml b/.github/workflows/render-rmds.yml index 3b3cd56c..14271387 100644 --- a/.github/workflows/render-rmds.yml +++ b/.github/workflows/render-rmds.yml @@ -6,20 +6,29 @@ on: branches: - master paths: - - '**.Rmd' - - '!**-live.Rmd' # don't trigger for live-only changes - - '!**/exercise*.Rmd' # or exercise notebooks - - '!**/setup/**.Rmd' # or setup notebooks - - 'scripts/make-live.R' - - 'scripts/render-live.sh' + - ".github/workflows/render-rmds.yml" + - "**.Rmd" + - "!**-live.Rmd" # don't trigger for live-only changes + - "!**/exercise*.Rmd" # or exercise notebooks + - "!**/setup/**.Rmd" # or setup notebooks + - "scripts/make-live.R" + - "scripts/render-live.sh" jobs: test-render: runs-on: ubuntu-latest - container: - image: ccdl/training_rstudio:edge steps: + - name: Free disk space + run: | + sudo rm -rf /usr/local/lib/android + sudo rm -rf /usr/share/dotnet + sudo rm -rf /opt/ghc + sudo rm -rf /usr/local/share/boost + sudo rm -rf "$AGENT_TOOLSDIRECTORY" + # Print free disk space + df -h + - name: Checkout code uses: actions/checkout@v4 @@ -36,8 +45,16 @@ jobs: env: AWS_DEFAULT_REGION: us-east-1 run: | - aws s3 sync s3://ccdl-training-data/training-modules/ . + aws s3 sync s3://ccdl-training-data/training-modules/ . --no-progress + - name: Pull latest Docker image + run: | + docker pull ccdl/training_rstudio:edge - name: Render notebooks - run: bash scripts/render-live.sh + run: | + docker run --rm \ + --mount type=bind,source="$GITHUB_WORKSPACE",target=/training-modules \ + -w /training-modules \ + ccdl/training_rstudio:edge \ + bash scripts/render-live.sh diff --git a/CONTRIBUTING.md b/CONTRIBUTING.md index 15b2b996..34b25a16 100644 --- a/CONTRIBUTING.md +++ b/CONTRIBUTING.md @@ -215,7 +215,7 @@ In practice, this means that you will not need to add individual R packages to t To use the Docker image for development, pull from Docker Hub with: ``` -docker pull --platform linux/amd64 ccdl/training_rstudio:edge +docker pull ccdl/training_rstudio:edge ``` To run the container and mount a local volume, use the following from the root of this repository: diff --git a/Dockerfile b/Dockerfile index e551636f..9d08dc91 100644 --- a/Dockerfile +++ b/Dockerfile @@ -1,5 +1,6 @@ # Build salmon from source in a separate image -FROM ubuntu:22.04 AS build +# matching base image from https://github.com/rocker-org/rocker-versioned2/blob/master/dockerfiles/r-ver_4.5.2.Dockerfile +FROM docker.io/library/ubuntu:noble AS build # Build dependencies RUN apt-get update -qq @@ -15,6 +16,7 @@ RUN DEBIAN_FRONTEND=noninteractive apt-get install -y --no-install-recommends \ libdeflate-dev \ libisal-dev \ liblzma-dev \ + libzstd-dev \ make \ pkg-config \ unzip \ @@ -29,7 +31,7 @@ RUN unzip awscliv2.zip RUN ./aws/install # Build salmon -ARG SALMON_VERSION=1.10.1 +ARG SALMON_VERSION=1.10.3 RUN curl -LO https://github.com/COMBINE-lab/salmon/archive/refs/tags/v${SALMON_VERSION}.tar.gz RUN tar xzf v${SALMON_VERSION}.tar.gz RUN mkdir salmon-${SALMON_VERSION}/build @@ -45,12 +47,12 @@ RUN cd fastp-${FASTP_VERSION} && \ make && make install # Main image with Biocconductor and other tools -FROM bioconductor/bioconductor_docker:3.19 AS final +FROM bioconductor/bioconductor_docker:3.22 AS final LABEL maintainer="ccdl@alexslemonade.org" WORKDIR /rocker-build/ -# Additonal dependencies for AWS runtime +# Additional dependencies for AWS runtime RUN apt-get update -qq RUN DEBIAN_FRONTEND=noninteractive apt-get install -y --no-install-recommends \ glibc-source \ @@ -66,21 +68,19 @@ RUN DEBIAN_FRONTEND=noninteractive apt-get install -y --no-install-recommends \ # Python packages COPY requirements.txt requirements.txt -RUN pip install -r requirements.txt +RUN pip install -r requirements.txt --break-system-packages # Use renv for R packages WORKDIR /usr/local/renv ENV RENV_CONFIG_CACHE_ENABLED=FALSE RUN Rscript -e "install.packages('renv')" -# Temporary fix for broken(?) RSamtools package -RUN Rscript -e "install.packages('BiocManager'); BiocManager::install('Rsamtools')" COPY renv.lock renv.lock -RUN Rscript -e "renv::restore()" \ - rm -rf ~/.cache/R/renv && \ - rm -rf /tmp/downloaded_packages && \ - rm -rf /tmp/Rtmp* +RUN Rscript -e "options(pkgType='binary'); renv::restore(repos = c(CRAN = 'https://packagemanager.posit.co/cran/__linux__/noble/latest'))" \ + && rm -rf ~/.cache/R/renv \ + && rm -rf /tmp/downloaded_packages \ + && rm -rf /tmp/Rtmp* # copy aws, salmon, and fastp binaries from the build image COPY --from=build /usr/local/aws-cli/ /usr/local/aws-cli/ diff --git a/RNA-seq/01-qc_trim_quant.Rmd b/RNA-seq/01-qc_trim_quant.Rmd index aa6ca7ac..f5299c1c 100644 --- a/RNA-seq/01-qc_trim_quant.Rmd +++ b/RNA-seq/01-qc_trim_quant.Rmd @@ -247,7 +247,7 @@ The index we used was built with `-k 23` and can be found here: index/Homo_sapiens/short_index ``` -Using a smaller value for _k_ than the default (_k_ = 31) is appropriate for shorter reads and may improve sensitivity when using `--validateMappings` according to the [Salmon documentation](https://salmon.readthedocs.io/en/latest/salmon.html#preparing-transcriptome-indices-mapping-based-mode). +Using a smaller value for _k_ than the default (_k_ = 31) is appropriate for shorter reads and may improve sensitivity when using `--validateMappings` according to the [Salmon documentation](https://salmon.readthedocs.io/en/latest/salmon.html#salmon). #### `-l` @@ -285,5 +285,5 @@ The `--threads` argument controls the number of threads that are available to Sa This in essence controls how much of the mapping can occur in parallel. If you had access to a computer with many cores, you could increase the number of threads to make quantification go faster. -**Navigate to** `data/gastric-cancer/salmon_quant/SRR585571/aux_info` **and open** `meta_info.json`**. +**Navigate to** `data/gastric-cancer/salmon_quant/SRR585570/aux_info` **and open** `meta_info.json`**. Look for a field called** `percent_mapped` **-- what value does this sample have?** diff --git a/RNA-seq/02-gastric_cancer_tximeta.Rmd b/RNA-seq/02-gastric_cancer_tximeta.Rmd index 0738a6c5..a8fe752e 100644 --- a/RNA-seq/02-gastric_cancer_tximeta.Rmd +++ b/RNA-seq/02-gastric_cancer_tximeta.Rmd @@ -120,24 +120,24 @@ sample_meta_df <- readr::read_tsv(meta_file) sample_meta_df ``` -We'll want this information to be added to the `coldata`, which we can do by using a join function to match up the rows between the two data frames and combine them. +We'll want this information to be added to the `coldata`, which we can do by using a join function to match up the rows between the two data frames and combine them to create a new data frame `coldata_joined`. ```{r join-sample_meta_df} -coldata <- coldata |> +coldata_joined <- coldata |> dplyr::inner_join(sample_meta_df, by = c("names" = "srr_accession")) -coldata +coldata_joined ``` ## Import expression data with `tximeta` -Using the `coldata` data frame that we set up, we can now run the `tximeta()` to import our expression data while automatically finding and associating the transcript annotations that were used when we performed the quantification. +Using the `coldata_joined` data frame that we set up, we can now run the `tximeta()` to import our expression data while automatically finding and associating the transcript annotations that were used when we performed the quantification. The first time you run `tximeta()` you may get a message about storing downloaded transcriptome data in a cache directory so that it can retrieve the data more quickly the next time. We recommend you use the cache, and accept the default location. ```{r tximeta, live = TRUE} -txi_data <- tximeta(coldata) +txi_data <- tximeta(coldata_joined) ``` *tximeta currently works easily for most human and mouse datasets, but requires a [few more steps for other species](https://bioconductor.org/packages/release/bioc/vignettes/tximeta/inst/doc/tximeta.html#What_if_checksum_isn%E2%80%99t_known). diff --git a/RNA-seq/02-gastric_cancer_tximeta.nb.html b/RNA-seq/02-gastric_cancer_tximeta.nb.html index 99d9795d..ce6fa77e 100644 --- a/RNA-seq/02-gastric_cancer_tximeta.nb.html +++ b/RNA-seq/02-gastric_cancer_tximeta.nb.html @@ -3264,8 +3264,8 @@

Summarize to gene

# Summarize to the gene level
 gene_summarized <- summarizeToGene(txi_data)
- -
loading existing EnsDb created: 2025-07-16 21:08:01
+ +
loading existing EnsDb created: 2025-11-20 20:30:47
obtaining transcript-to-gene mapping from database
diff --git a/components/dependencies.R b/components/dependencies.R index fe381a4e..358fe6d8 100644 --- a/components/dependencies.R +++ b/components/dependencies.R @@ -36,3 +36,6 @@ library(scDblFinder) # Loom file format functions for Single Cell data library(LoomExperiment) + +# Needed for SingleR to run de with wilcox, for cell type exercises +library(scrapper) diff --git a/components/dictionary.txt b/components/dictionary.txt index b6fadbe6..0387b616 100644 --- a/components/dictionary.txt +++ b/components/dictionary.txt @@ -1,5 +1,6 @@ ʹ ⚠️ +µm Adelie ADT ADTs @@ -15,6 +16,7 @@ AllCells ALSF Amezquita AML +anaplastic Anders Angerer AnnotationDbi @@ -46,6 +48,8 @@ biomolecular bioRxiv biospecimen Blasi +blastema +blastemal BMC Bonferroni bp @@ -110,6 +114,7 @@ dplyr dropdown DropSeq dumpFeatures +ECM effector eigengene embeddings @@ -145,6 +150,7 @@ FCS FDR fgsea fibrotic +fiducials Figshare FLI FN @@ -191,6 +197,7 @@ hematopoietic Hemberg hexamer Hippen +histologic histologies Hm Homebrew @@ -314,6 +321,7 @@ Novia NPC NRAS NRASG +num octothorps OkabeIto oligodendrocyte @@ -343,6 +351,7 @@ pDC PDX ped permalink +permeabilized phenotypes Phred Picelli @@ -390,6 +399,7 @@ rmd Rmd RMS roadmap +roxygen RPKMs rRNA Rscript @@ -490,6 +500,7 @@ vectorization vectorized versicolor virginica +Visium vitro VST Wattenberg diff --git a/current-modules.json b/current-modules.json index bb8d68be..fe8c34c7 100644 --- a/current-modules.json +++ b/current-modules.json @@ -1,5 +1,5 @@ { - "release-tag": "2025-dev", + "release-tag": "2025-december", "modules": ["scRNA-seq-advanced"], "reference-modules": ["scRNA-seq"] } diff --git a/intro-to-R-tidyverse/01-intro_to_base_R.Rmd b/intro-to-R-tidyverse/01-intro_to_base_R.Rmd index 79a9ec9d..5240f597 100644 --- a/intro-to-R-tidyverse/01-intro_to_base_R.Rmd +++ b/intro-to-R-tidyverse/01-intro_to_base_R.Rmd @@ -418,13 +418,9 @@ question_values %in% values_1_to_20 _Data frames are one of the most useful tools for data analysis in R._ They are tables which consist of rows and columns, much like a _spreadsheet_. Each column is a variable which behaves as a _vector_, and each row is an observation. -We will begin our exploration with dataset of measurements from three penguin species measured, which we can find in the [`palmerpenguins` package](https://allisonhorst.github.io/palmerpenguins/). -We'll talk more about packages soon! -To use this dataset, we will load it from the `palmerpenguins` package using a `::` (more on this later) and assign it to a variable named `penguins` in our current environment. - -```{r penguin-library} -penguins <- palmerpenguins::penguins -``` +We will begin our exploration with dataset of measurements from three penguin species measured using the built-in R dataset `penguins`. +(This dataset was added as a built-in dataset to R version `4.5.0`; +if you are working with an earlier version of R, it is available via the [`palmerpenguins` package](https://allisonhorst.github.io/palmerpenguins/) - we'll talk more about packages soon!) ![drawings of penguin species](diagrams/lter_penguins.png) Artwork by [@allison_horst](https://twitter.com/allison_horst) @@ -456,32 +452,33 @@ This provides a short view of the **str**ucture and contents of the data frame. str(penguins) ``` -You'll notice that the column `species` is a _factor_: This is a special type of character variable that represents distinct categories known as "levels". -We have learned here that there are three levels in the `species` column: Adelie, Chinstrap, and Gentoo. +You'll notice that the columns `species`, `island`, and `sex` are labelled as _factor_: This is a special type of character variable that represents distinct categories known as "levels". +Other columns are labelled as _num_ for numeric (with decimals), or _int_ for integer (numeric without decimals). + We might want to explore individual columns of the data frame more in-depth. We can examine individual columns using the dollar sign `$` to select one by name: ```{r penguins-subset} -# Extract bill_length_mm as a vector -penguins$bill_length_mm +# Extract bill_len as a vector +penguins$bill_len # indexing operators can be used on these vectors too -penguins$bill_length_mm[1:10] +penguins$bill_len[1:10] ``` We can perform our regular vector operations on columns directly. ```{r penguins-col-mean, live = TRUE} -# calculate the mean of the bill_length_mm column -mean(penguins$bill_length_mm, +# calculate the mean of the bill_len column +mean(penguins$bill_len, na.rm = TRUE) # remove missing values before calculating the mean ``` We can also calculate the full summary statistics for a single column directly. ```{r penguins-col-summary, live = TRUE} -# show a summary of the bill_length_mm column -summary(penguins$bill_length_mm) +# show a summary of the bill_len column +summary(penguins$bill_len) ``` Extract `species` as a vector and subset it to see a preview. diff --git a/intro-to-R-tidyverse/02-intro_to_ggplot2.Rmd b/intro-to-R-tidyverse/02-intro_to_ggplot2.Rmd index fbbd2ae0..0cb33754 100644 --- a/intro-to-R-tidyverse/02-intro_to_ggplot2.Rmd +++ b/intro-to-R-tidyverse/02-intro_to_ggplot2.Rmd @@ -32,8 +32,8 @@ We performed three sets of contrasts: **More ggplot2 resources:** - [ggplot2 website](https://ggplot2.tidyverse.org/) +- [ggplot2 book](https://ggplot2-book.org/) - [Handy cheatsheet for ggplot2 (pdf)](https://github.com/rstudio/cheatsheets/raw/main/data-visualization.pdf) -- [_Data Visualization, A practical introduction_](https://socviz.co/) - [Data visualization chapter of _R for Data Science_](https://r4ds.hadley.nz/data-visualize.html) - [ggplot2 online tutorial](http://r-statistics.co/Complete-Ggplot2-Tutorial-Part1-With-R-Code.html) diff --git a/intro-to-R-tidyverse/03-intro_to_tidyverse.Rmd b/intro-to-R-tidyverse/03-intro_to_tidyverse.Rmd index b3d3d915..bc58746c 100644 --- a/intro-to-R-tidyverse/03-intro_to_tidyverse.Rmd +++ b/intro-to-R-tidyverse/03-intro_to_tidyverse.Rmd @@ -52,7 +52,8 @@ library(tidyverse) Note that if we had not imported the tidyverse set of packages using `library()` like above, and we wanted to use a tidyverse function like `read_tsv()`, we would need to tell R what package to find this function in. To do this, we would use `::` to tell R to load in this function from the `readr` package by using `readr::read_tsv()`. You will see this `::` method of referencing libraries within packages throughout the course. -We like to use it in part to remove any ambiguity in which version of a function we are using; it is not too uncommon for different packages to use the same name for very different functions! +We like to use it in part to remove any ambiguity in which version of a function we are using, even if the package was imported; it is not too uncommon for different packages to use the same name for very different functions! +We'll use `::` syntax for external functions throughout this notebook. ## Managing directories @@ -92,6 +93,7 @@ But we've also seen that this function will throw an error if you try to create A different option is to use the [`fs`](https://fs.r-lib.org/) package, which provides functions for you to interact with your computer's file system with a more consistent behavior than the base R functions. One function from this package is `fs::dir_create()` (note that it has an _underscore_, not a period), and much like the base R `dir.create()`, it creates directories. It has some other helpful features too: + - It will simply do nothing if that directory already exists; no errors, and nothing will get overwritten - It allows creating _nested_ directories by default, i.e. in one call make directories inside of other directories @@ -142,16 +144,15 @@ data_dir <- "data" ``` Although base R has functions to read in data files, the functions in the `readr` package (part of the tidyverse) are faster and more straightforward to use so we are going to use those here. -Because the file we are reading in is a TSV (tab separated values) file we will be using the `read_tsv` function. -There are analogous functions for CSV (comma separated values) files (`read_csv()`) and other files types. +Because the file we are reading in is a TSV (tab separated values) file we will be using the `readr::read_tsv()` function. +There are analogous functions for CSV (comma separated values) files (`readr::read_csv()`) and other files types. ## Read in the differential expression analysis results file ```{r read-results} stats_df <- readr::read_tsv( - file.path(data_dir, - "gene_results_GSE44971.tsv") - ) + file.path(data_dir, "gene_results_GSE44971.tsv") +) ``` Following the template of the previous chunk, use this chunk to read in the file `GSE44971.tsv` that is in the `data` folder and save it in the variable `gene_df`. @@ -159,9 +160,8 @@ Following the template of the previous chunk, use this chunk to read in the file ```{r read-expr, live = TRUE} # Use this chunk to read in data from the file `GSE44971.tsv` gene_df <- readr::read_tsv( - file.path(data_dir, - "GSE44971.tsv") - ) + file.path(data_dir, "GSE44971.tsv") +) ``` Use this chunk to explore what `gene_df` looks like. @@ -182,16 +182,18 @@ _Note:_ If you are using a version of `R` prior to 4.1 (or looking at older code That pipe was the inspiration for the native R pipe we are using here. While there are some minor differences, you can mostly treat them interchangeably as long as you load the `magrittr` package or `dplyr`, which also loads that version of the pipe. -For example, the output from this: +We'll being our journey into pipes using some core functions from the `dplyr` package (a core tidyverse package that offers "pliers" for your data). + +As our first example, the output from this: ```{r filter} -filter(stats_df, contrast == "male_female") +dplyr::filter(stats_df, contrast == "male_female") ``` ...is the same as the output from this: ```{r filter-pipe} -stats_df |> filter(contrast == "male_female") +stats_df |> dplyr::filter(contrast == "male_female") ``` This can make your code cleaner and easier to follow a series of related commands. @@ -200,9 +202,9 @@ Let's look at an example with our stats of of how the same functions look with o *Example 1:* without pipes: ```{r steps-nopipe} -stats_arranged <- arrange(stats_df, t_statistic) -stats_filtered <- filter(stats_arranged, avg_expression > 50) -stats_nopipe <- select(stats_filtered, contrast, log_fold_change, p_value) +stats_arranged <- dplyr::arrange(stats_df, t_statistic) +stats_filtered <- dplyr::filter(stats_arranged, avg_expression > 50) +stats_nopipe <- dplyr::select(stats_filtered, contrast, log_fold_change, p_value) ``` UGH, we have to keep track of all of those different intermediate data frames and type their names so many times here! @@ -213,15 +215,15 @@ It's annoying and makes it harder for people to read. ```{r steps-pipe, live = TRUE} # Example of the same modifications as above but with pipes! -stats_pipe <- stats_df |> - arrange(t_statistic) |> - filter(avg_expression > 50) |> - select(contrast, log_fold_change, p_value) +stats_pipe <- stats_df |> + dplyr::arrange(t_statistic) |> + dplyr::filter(avg_expression > 50) |> + dplyr::select(contrast, log_fold_change, p_value) ``` What the `|>` (pipe) is doing here is feeding the result of the expression on its left into the first argument of the next function (to its right, or on the next line here). We can then skip that first argument (the data in these cases), and move right on to the part we care about at that step: what we are arranging, filtering, or selecting in this case. -The key insight that makes the pipe work here is to recognize that each of these functions (`arrange`, `filter`, and `select`) are fundamental `dplyr` (a tidyverse package) functions which work as "data in, data out." +The key insight that makes the pipe work here is to recognize that each of these functions (`dplyr::arrange`, `dplyr::filter`, and `dplyr::select`) are fundamental `dplyr` functions which work as "data in, data out." In other words, these functions operate on data frames, and return data frames; you give them a data frame, and they give you back a data frame. Because these functions all follow a "data in, data out" framework, we can chain them together with pipe and send data all the way through the...pipeline! @@ -239,12 +241,12 @@ Now that hopefully you are convinced that the tidyverse can help you make your c ## Common tidyverse functions Let's say we wanted to filter this gene expression dataset to particular sample groups. -In order to do this, we would use the function `filter()` as well as a logic statement (usually one that refers to a column or columns in the data frame). +In order to do this, we would use the function `dplyr::filter()` as well as a logic statement (usually one that refers to a column or columns in the data frame). ```{r filter-gene} # Here let's filter stats_df to only keep the gene_symbol "SNCA" stats_df |> - filter(gene_symbol == "SNCA") + dplyr::filter(gene_symbol == "SNCA") ``` We can use `filter()` similarly for numeric statements. @@ -252,7 +254,7 @@ We can use `filter()` similarly for numeric statements. ```{r filter-numeric, live = TRUE} # Here let's filter the data to rows with average expression values above 50 stats_df |> - filter(avg_expression > 50) + dplyr::filter(avg_expression > 50) ``` We can apply multiple filters at once, which will require all of them to be satisfied for every row in the results: @@ -260,8 +262,10 @@ We can apply multiple filters at once, which will require all of them to be sati ```{r filter-2, live = TRUE} # filter to highly expressed genes with contrast "male_female" stats_df |> - filter(contrast == "male_female", - avg_expression > 50) + dplyr::filter( + contrast == "male_female", + avg_expression > 50 + ) ``` When we are filtering, the `%in%` operator can come in handy if we have multiple items we would like to match. @@ -278,7 +282,7 @@ stats_df$gene_symbol %in% genes_of_interest ```{r filter-in, live = TRUE} # filter to keep only genes of interest stats_df |> - filter(gene_symbol %in% c("SNCA", "CDKN1A")) + dplyr::filter(gene_symbol %in% c("SNCA", "CDKN1A")) ``` Let's return to our first `filter()` and build on to it. @@ -290,9 +294,11 @@ Let's also save this as a new data frame called `stats_filtered_df`. # filter to highly expressed "male_female" # and select gene_symbol, log_fold_change and t_statistic stats_filtered_df <- stats_df |> - filter(contrast == "male_female", - avg_expression > 50) |> - select(log_fold_change, t_statistic) + dplyr::filter( + contrast == "male_female", + avg_expression > 50 + ) |> + dplyr::select(log_fold_change, t_statistic) ``` Let's say we wanted to arrange this dataset so that the genes are arranged by the smallest p values to the largest. @@ -300,7 +306,7 @@ In order to do this, we would use the function `arrange()` as well as the column ```{r arrange} stats_df |> - arrange(p_value) + dplyr::arrange(p_value) ``` What if we want to sort from largest to smallest? @@ -310,15 +316,15 @@ We can use the same function, but instead use the `desc()` function and now we a ```{r arrange-desc} # arrange descending by avg_expression stats_df |> - arrange(desc(avg_expression)) + dplyr::arrange(desc(avg_expression)) ``` What if we would like to create a new column of values? -For that we use `mutate()` function. +For that we use `dplyr::mutate()` function. ```{r mutate} stats_df |> - mutate(log10_p_value = -log10(p_value)) + dplyr::mutate(log10_p_value = -log10(p_value)) ``` What if we want to obtain summary statistics for a column or columns? @@ -327,19 +333,26 @@ Here we will use summarize to calculate two summary statistics of log-fold chang ```{r summarize} stats_df |> - summarize(mean(log_fold_change), - sd(log_fold_change)) + dplyr::summarize( + # name the columns mean_lfc an sd_lfc + mean_lfc = mean(log_fold_change), + sd_lfc = sd(log_fold_change) + ) ``` What if we'd like to obtain a summary statistics but have them for various groups? Conveniently named, there's a function called `group_by()` that seamlessly allows us to do this. Also note that `group_by()` allows us to group by multiple variables at a time if you want to. +We'll use this function to create a grouped summary data frame and save it to a variable `stats_summary_df`. + ```{r summarize-groups, live = TRUE} stats_summary_df <- stats_df |> - group_by(contrast) |> - summarize(mean(log_fold_change), - sd(log_fold_change)) + dplyr::group_by(contrast) |> + dplyr::summarize( + mean_lfc = mean(log_fold_change), + sd_lfc = sd(log_fold_change) + ) ``` Let's look at a preview of what we made: @@ -454,7 +467,7 @@ In this case we want to match the gene information between the two, so we will s stats_df |> # Join based on their shared column # Called ensembl_id in stats_df and called Gene in gene_df - inner_join(gene_df, by = c('ensembl_id' = 'Gene')) + dplyr::inner_join(gene_df, by = c('ensembl_id' = 'Gene')) ``` ## Save data to files diff --git a/pathway-analysis/01-overrepresentation_analysis.Rmd b/pathway-analysis/01-overrepresentation_analysis.Rmd index 8e7a62f1..afc79f52 100644 --- a/pathway-analysis/01-overrepresentation_analysis.Rmd +++ b/pathway-analysis/01-overrepresentation_analysis.Rmd @@ -59,9 +59,7 @@ library(org.Mm.eg.db) # We'll create a directory to specifically hold the ORA results if it doesn't # exist yet results_dir <- file.path("results", "leukemia") -if (!dir.exists(results_dir)) { - dir.create(results_dir, recursive = TRUE) -} +fs::dir_create(results_dir) ``` #### Input files @@ -107,7 +105,7 @@ The results we're interested in here come from mouse samples, so we can obtain j mm_msigdb_df <- msigdbr(species = "Mus musculus") ``` -MSigDB contains 8 different gene set collections. +MSigDB contains 9 different gene set collections. H: hallmark gene sets C1: positional gene sets @@ -117,23 +115,26 @@ MSigDB contains 8 different gene set collections. C5: GO gene sets C6: oncogenic signatures C7: immunologic signatures + C8: cell type signatures In this example, we will use canonical pathways which are ([ref](https://www.gsea-msigdb.org/gsea/msigdb/collections.jsp)): > Gene sets from pathway databases. Usually, these gene sets are canonical representations of a biological process compiled by domain experts. And are a subset of `C2: curated gene sets`. -Specifically, we will use the [KEGG (Kyoto Encyclopedia of Genes and Genomes)](https://www.genome.jp/kegg/) pathways. +Specifically, we will use the [KEGG (Kyoto Encyclopedia of Genes and Genomes)](https://www.genome.jp/kegg/) pathways, which is denoted as `CP:KEGG_MEDICUS` in this version of `msigdbr`. ```{r filter_to_kegg} # Filter the mouse data frame to the KEGG pathways that are included in the # curated gene sets mm_kegg_df <- mm_msigdb_df |> - dplyr::filter(gs_cat == "C2", # curated gene sets - gs_subcat == "CP:KEGG") # KEGG pathways + dplyr::filter( + gs_collection == "C2", # curated gene set + gs_subcollection == "CP:KEGG_MEDICUS" # KEGG pathways + ) ``` -*Note: We could specified that we wanted the KEGG gene sets using the `category` and `subcategory` arguments of `msigdbr()`, but we're going for general steps!* +*Note: We could specified that we wanted the KEGG gene sets using the `collection` and `subcollection` arguments of `msigdbr()`, but we're going for general steps!* ```{r mm_kegg_columns} colnames(mm_kegg_df) @@ -179,18 +180,20 @@ The function we will use to map from Ensembl gene IDs to gene symbols is called ```{r map_to_symbol} # This returns a named vector which we can convert to a data frame, where # the keys (Ensembl IDs) are the names -symbols_vector <- mapIds(org.Mm.eg.db, # Specify the annotation package - # The vector of gene identifiers we want to - # map - keys = vs_low_df$Gene, - # What type of gene identifiers we're starting - # with - keytype = "ENSEMBL", - # The type of gene identifier we want returned - column = "SYMBOL", - # In the case of 1:many mappings, return the - # first one. This is default behavior! - multiVals = "first") +symbols_vector <- mapIds( + org.Mm.eg.db, # Specify the annotation package + # The vector of gene identifiers we want to + # map + keys = vs_low_df$Gene, + # What type of gene identifiers we're starting + # with + keytype = "ENSEMBL", + # The type of gene identifier we want returned + column = "SYMBOL", + # In the case of 1:many mappings, return the + # first one. This is default behavior! + multiVals = "first" +) # We would like a data frame we can join to the DGE results symbols_df <- data.frame( @@ -210,10 +213,12 @@ Let's do this first for the comparison to the low stem cell capacity population. ```{r add_symbols, live = TRUE} vs_low_df <- symbols_df |> # An *inner* join will only return rows that are in both data frames - dplyr::inner_join(vs_low_df, - # The name of the column that contains the Ensembl gene IDs - # in the left data frame and right data frame - by = c("ensembl_id" = "Gene")) + dplyr::inner_join( + vs_low_df, + # The name of the column that contains the Ensembl gene IDs + # in the left data frame and right data frame + by = c("ensembl_id" = "Gene") + ) ``` ### Drop `NA` values @@ -226,7 +231,7 @@ This will also drop genes that have an Ensembl gene identifier but no gene symbo ```{r complete_cases} # Remove rows that are not complete (e.g., contain NAs) by filtering to only # complete rows -vs_low_df <- vs_low_df |> +vs_low_complete_df <- vs_low_df |> tidyr::drop_na() ``` @@ -290,10 +295,12 @@ We'll start with the high stem cell capacity vs. low stem cell capacity populati Genes with positive log2 fold-changes (LFC) will be more highly expressed in the high stem cell capacity cells based on how we set up the analysis. ```{r high_capacity_genes} -vs_low_genes <- vs_low_df |> +vs_low_genes <- vs_low_complete_df |> # Filter to the positive LFC and filter based on significance too (padj) - dplyr::filter(log2FoldChange > 0, - padj < 0.05) |> + dplyr::filter( + log2FoldChange > 0, + padj < 0.05 + ) |> # Return a vector of gene symbols dplyr::pull(gene_symbol) ``` @@ -306,8 +313,10 @@ Now, we'll take the same steps for our other results. ```{r unsorted_genes_to_remove, live = TRUE} vs_unsorted_genes <- vs_unsorted_df |> - dplyr::filter(log2FoldChange > 0, - padj < 0.05) |> + dplyr::filter( + log2FoldChange > 0, + padj < 0.05 + ) |> dplyr::pull(gene_symbol) ``` @@ -335,8 +344,10 @@ We can use another function for set operations, `intersect()`, to get our backgr ```{r get_background_set} # intersect() will return the genes in both sets - we are using the entire data # frame here (complete cases), not just the significant genes -background_set <- intersect(vs_low_df$gene_symbol, - vs_unsorted_df$gene_symbol) +background_set <- intersect( + vs_low_complete_df$gene_symbol, + vs_unsorted_df$gene_symbol +) # Remove anything that couldn't reliably be measured/assessed in both from the # genes of interest list - using intersect() will drop anything in the first set @@ -381,13 +392,13 @@ kegg_result_df <- data.frame(kegg_ora_results@result) We can use a dot plot to visualize our significant enrichment results. -```{r dotplot, live = TRUE} +```{r dotplot, fig.width = 10, live = TRUE} enrichplot::dotplot(kegg_ora_results) ``` We can use an [UpSet plot](https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4720993/) to visualize the **overlap** between the gene sets that were returned as significant. -```{r upsetplot, live = TRUE} +```{r upsetplot, fig.width = 10, live = TRUE} enrichplot::upsetplot(kegg_ora_results) ``` diff --git a/pathway-analysis/02-gene_set_enrichment_analysis.Rmd b/pathway-analysis/02-gene_set_enrichment_analysis.Rmd index 3c79f68b..1d6dd544 100644 --- a/pathway-analysis/02-gene_set_enrichment_analysis.Rmd +++ b/pathway-analysis/02-gene_set_enrichment_analysis.Rmd @@ -52,6 +52,10 @@ library(clusterProfiler) library(msigdbr) ``` +```{r set seed} +set.seed(2020) +``` + ### Directories and Files #### Directories @@ -63,9 +67,7 @@ input_dir <- file.path("..", "RNA-seq", "results", "NB-cell") # We will create a directory to specifically hold our GSEA results if it does # not yet exist output_dir <- file.path("results", "NB-cell") -if (!dir.exists(output_dir)) { - dir.create(output_dir, recursive = TRUE) -} +fs::dir_create(output_dir) ``` #### Input files @@ -119,11 +121,13 @@ We'll use the Hallmark collection for GSEA. Notably, there are only 50 gene sets included in this collection. The fewer gene sets we test, the lower our multiple hypothesis testing burden. -We can retrieve only the Hallmark gene sets by specifying `category = "H"` to the `msigdbr()` function. +We can retrieve only the Hallmark gene sets by specifying `collection = "H"` to the `msigdbr()` function. ```{r hallmark_sets, live = TRUE} -hs_hallmark_df <- msigdbr(species = "Homo sapiens", - category = "H") +hs_hallmark_df <- msigdbr( + species = "Homo sapiens", + collection = "H" +) ``` ## Differential gene expression results @@ -238,14 +242,18 @@ Now for the analysis! We can use the `GSEA()` function to perform GSEA with any generic set of gene sets, but there are several functions for using specific, commonly used gene sets (e.g., `gseKEGG()`). ```{r run_gsea} -gsea_results <- GSEA(geneList = lfc_vector, # ordered ranked gene list - minGSSize = 25, # minimum gene set size - maxGSSize = 500, # maximum gene set set - pvalueCutoff = 0.05, - pAdjustMethod = "BH", # correction for multiple hypothesis testing - TERM2GENE = dplyr::select(hs_hallmark_df, - gs_name, - gene_symbol)) +gsea_results <- GSEA( + geneList = lfc_vector, # ordered ranked gene list + minGSSize = 25, # minimum gene set size + maxGSSize = 500, # maximum gene set set + pvalueCutoff = 0.05, + pAdjustMethod = "BH", # correction for multiple hypothesis testing + TERM2GENE = dplyr::select( + hs_hallmark_df, + gs_name, + gene_symbol + ) +) ``` The warning about ties means that there are multiple genes that have the same log2 fold change value. diff --git a/pathway-analysis/03-gene_set_variation_analysis.Rmd b/pathway-analysis/03-gene_set_variation_analysis.Rmd index abcbb998..6e8862c1 100644 --- a/pathway-analysis/03-gene_set_variation_analysis.Rmd +++ b/pathway-analysis/03-gene_set_variation_analysis.Rmd @@ -57,9 +57,7 @@ input_dir <- file.path("data", "open-pbta") # Create a directory specifically for the results using this dataset output_dir <- file.path("results", "open-pbta") -if (!dir.exists(output_dir)) { - dir.create(output_dir, recursive = TRUE) -} +fs::dir_create(output_dir) ``` #### Input @@ -164,9 +162,7 @@ gsva_results <- gsva( kcdf = "Gaussian", # enrichment score is the difference between largest positive and negative maxDiff = TRUE - ), - # Use 4 cores for multiprocessing - BPPARAM = BiocParallel::MulticoreParam(4) + ) ) ``` @@ -211,8 +207,9 @@ random_gene_sets <- rep(gene_set_size, nreps) |> # Repeat gene sizes so we run purrr::map( # Sample the vector of all genes, choosing the number of items specified # in the element of gene set size - ~ base::sample(x = all_genes, - size = .x) + \(geneset_size) { + base::sample(x = all_genes, size = geneset_size) + } ) ``` @@ -224,9 +221,13 @@ The Hallmarks list we used earlier stored the gene set names as the name of the # number lengths_vector <- random_gene_sets |> # Get the length of each gene set (number of genes) - purrr::map(~ length(.x)) |> + purrr::map(length) |> # Make it "pathway_" - purrr::map(~ paste0("pathway_", .x)) |> + purrr::map( + \(geneset_size) { + paste0("pathway_", geneset_size) + } + ) |> # Return a vector purrr::flatten_chr() @@ -247,9 +248,7 @@ random_gsva_results <- gsva( maxSize = 500, kcdf = "Gaussian", maxDiff = TRUE - ), - # Use 4 cores for multiprocessing - BPPARAM = BiocParallel::MulticoreParam(4) + ) ) ``` diff --git a/renv.lock b/renv.lock index 752ed02d..d6d0ab6b 100644 --- a/renv.lock +++ b/renv.lock @@ -1,2128 +1,5955 @@ { "R": { - "Version": "4.4.0", + "Version": "4.5.2", "Repositories": [ { "Name": "BioCsoft", - "URL": "https://bioconductor.org/packages/3.19/bioc" + "URL": "https://bioconductor.org/packages/3.22/bioc" }, { "Name": "BioCann", - "URL": "https://bioconductor.org/packages/3.19/data/annotation" + "URL": "https://bioconductor.org/packages/3.22/data/annotation" }, { "Name": "BioCexp", - "URL": "https://bioconductor.org/packages/3.19/data/experiment" + "URL": "https://bioconductor.org/packages/3.22/data/experiment" }, { "Name": "BioCworkflows", - "URL": "https://bioconductor.org/packages/3.19/workflows" + "URL": "https://bioconductor.org/packages/3.22/workflows" }, { "Name": "BioCbooks", - "URL": "https://bioconductor.org/packages/3.19/books" + "URL": "https://bioconductor.org/packages/3.22/books" }, { "Name": "CRAN", "URL": "https://p3m.dev/cran/latest" + }, + { + "Name": "BioCcontainers", + "URL": "https://bioconductor.org/packages/3.22/container-binaries/bioconductor_docker" } ] }, "Bioconductor": { - "Version": "3.19" + "Version": "3.22" }, "Packages": { "ALL": { "Package": "ALL", - "Version": "1.46.0", + "Version": "1.52.0", "Source": "Bioconductor", - "Repository": "Bioconductor 3.19", - "Requirements": [ - "Biobase", - "R" - ], - "Hash": "dd00b008c7f784712d11015bbd59bd4c" + "Title": "A data package", + "Date": "2009-07-22", + "Author": "Xiaochun Li", + "Description": "Data of T- and B-cell Acute Lymphocytic Leukemia from the Ritz Laboratory at the DFCI (includes Apr 2004 versions)", + "Maintainer": "Robert Gentleman ", + "License": "Artistic-2.0", + "Depends": [ + "R (>= 2.10)", + "Biobase (>= 2.5.5)" + ], + "Suggests": [ + "rpart" + ], + "biocViews": "ExperimentData, CancerData, LeukemiaCancerData", + "git_url": "https://git.bioconductor.org/packages/ALL", + "git_branch": "RELEASE_3_22", + "git_last_commit": "0bc53bb", + "git_last_commit_date": "2025-10-29", + "Repository": "Bioconductor 3.22", + "NeedsCompilation": "no" }, "AUCell": { "Package": "AUCell", - "Version": "1.26.0", + "Version": "1.32.0", "Source": "Bioconductor", - 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Laboratory of Computational Biology. VIB-KU Leuven Center for Brain & Disease Research. Leuven, Belgium.", + "Maintainer": "Gert Hulselmans ", + "Description": "AUCell allows to identify cells with active gene sets (e.g. signatures, gene modules...) in single-cell RNA-seq data. AUCell uses the \"Area Under the Curve\" (AUC) to calculate whether a critical subset of the input gene set is enriched within the expressed genes for each cell. The distribution of AUC scores across all the cells allows exploring the relative expression of the signature. Since the scoring method is ranking-based, AUCell is independent of the gene expression units and the normalization procedure. 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This package provides an R interface by relying on the 'Rcpp' package, exposing the same interface as the original Python wrapper to 'Annoy'. See for more on 'Annoy'. 'Annoy' is released under Version 2.0 of the Apache License. Also included is a small Windows port of 'mmap' which is released under the MIT license.", + "License": "GPL (>= 2)", + "Depends": [ + "R (>= 3.1)" + ], + "Imports": [ + "methods", + "Rcpp" + ], + "LinkingTo": [ + "Rcpp" + ], + "Suggests": [ + "tinytest" ], - "Hash": "f6baa1e06fb6c3724f601a764266cb0d" + "URL": "https://github.com/eddelbuettel/rcppannoy, https://dirk.eddelbuettel.com/code/rcpp.annoy.html", + "BugReports": "https://github.com/eddelbuettel/rcppannoy/issues", + "NeedsCompilation": "yes", + "RoxygenNote": "7.3.2", + "Encoding": "UTF-8", + "VignetteBuilder": "Rcpp", + "Author": "Dirk Eddelbuettel [aut, cre] (ORCID: ), Erik Bernhardsson [aut] (Principal author of Annoy)", + "Maintainer": "Dirk Eddelbuettel ", + "Repository": "CRAN" }, "RcppArmadillo": { "Package": "RcppArmadillo", - "Version": "0.12.8.2.1", - "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", - "Rcpp", - "methods", + "Version": "15.2.3-1", + "Source": "Repository", + "Type": "Package", + "Title": "'Rcpp' Integration for the 'Armadillo' Templated Linear Algebra Library", + "Date": "2025-12-16", + "Authors@R": "c(person(\"Dirk\", \"Eddelbuettel\", role = c(\"aut\", \"cre\"), email = \"edd@debian.org\", comment = c(ORCID = \"0000-0001-6419-907X\")), person(\"Romain\", \"Francois\", role = \"aut\", comment = c(ORCID = \"0000-0002-2444-4226\")), person(\"Doug\", \"Bates\", role = \"aut\", comment = c(ORCID = \"0000-0001-8316-9503\")), person(\"Binxiang\", \"Ni\", role = \"aut\"), person(\"Conrad\", \"Sanderson\", role = \"aut\", comment = c(ORCID = \"0000-0002-0049-4501\")))", + "Description": "'Armadillo' is a templated C++ linear algebra library aiming towards a good balance between speed and ease of use. 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Several utilities are also provided to compare and evaluate clustering results.", + "Authors@R": "c( person(\"Aaron\", \"Lun\", role = c(\"aut\", \"cre\"), email = \"infinite.monkeys.with.keyboards@gmail.com\"), person(\"Stephanie\", \"Hicks\", role=\"ctb\"), person(\"Basil\", \"Courbayre\", role=\"ctb\"), person(\"Tuomas\", \"Borman\", role=\"ctb\"), person(\"Leo\", \"Lahti\", role=\"ctb\") )", + "Imports": [ + "stats", + "methods", + "utils", + "cluster", "Matrix", "Rcpp", - "S4Vectors", - "cluster", "igraph", - "methods", - "stats", - "utils" + "S4Vectors", + "BiocParallel", + "BiocNeighbors" ], - "Hash": "ed9597168d850071aa9abbbef7be7204" + "Suggests": [ + "knitr", + "rmarkdown", + "testthat", + "BiocStyle", + "dynamicTreeCut", + "scRNAseq", + "scuttle", + "scater", + "scran", + "pheatmap", + "viridis", + "mbkmeans", + "kohonen", + "apcluster", + "DirichletMultinomial", + "vegan", + "fastcluster" + ], + "biocViews": "ImmunoOncology, Software, GeneExpression, Transcriptomics, SingleCell, Clustering", + "LinkingTo": [ + "Rcpp", + "assorthead" + ], + "Collate": "AllClasses.R AllGenerics.R AgnesParam.R approxSilhouette.R bluster-package.R DbscanParam.R DianaParam.R AffinityParam.R BlusterParam.R bootstrapStability.R ClaraParam.R clusterRMSD.R clusterSweep.R compareClusterings.R DmmParam.R FixedNumberParam.R HclustParam.R HierarchicalParam.R KmeansParam.R linkClusters.R makeSNNGraph.R MbkmeansParam.R mergeCommunities.R neighborPurity.R nestedClusters.R NNGraphParam.R pairwiseModularity.R pairwiseRand.R PamParam.R RcppExports.R SomParam.R TwoStepParam.R utils.R", + "License": "GPL-3", + "NeedsCompilation": "yes", + "VignetteBuilder": "knitr", + "SystemRequirements": "C++17", + "RoxygenNote": "7.3.3", + "Encoding": "UTF-8", + "git_url": "https://git.bioconductor.org/packages/bluster", + "git_branch": "RELEASE_3_22", + "git_last_commit": "b47a2df", + "git_last_commit_date": "2025-10-29", + "Repository": "Bioconductor 3.22", + "Author": "Aaron Lun [aut, cre], 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Broom provides three verbs that each provide different types of information about a model. tidy() summarizes information about model components such as coefficients of a regression. glance() reports information about an entire model, such as goodness of fit measures like AIC and BIC. augment() adds information about individual observations to a dataset, such as fitted values or influence measures.", + "License": "MIT + file LICENSE", + "URL": "https://broom.tidymodels.org/, https://github.com/tidymodels/broom", + "BugReports": "https://github.com/tidymodels/broom/issues", + "Depends": [ + "R (>= 4.1)" + ], + "Imports": [ "backports", - "dplyr", - "ellipsis", - "generics", - "glue", - "lifecycle", - "purrr", - "rlang", - "stringr", - "tibble", - "tidyr" - ], - "Hash": "fd25391c3c4f6ecf0fa95f1e6d15378c" - }, - "broom.helpers": { - "Package": "broom.helpers", - "Version": "1.15.0", - "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", - "broom", "cli", - "dplyr", - 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It provides a universal interface for gene functional annotation from a variety of sources and thus can be applied in diverse scenarios. It provides a tidy interface to access, manipulate, and visualize enrichment results to help users achieve efficient data interpretation. Datasets obtained from multiple treatments and time points can be analyzed and compared in a single run, easily revealing functional consensus and differences among distinct gene clusters.", + "Depends": [ + "R (>= 4.2.0)" + ], + "Imports": [ "AnnotationDbi", - "DOSE", - "GO.db", - "GOSemSim", - "R", - "downloader", + "DOSE (>= 3.23.2)", "dplyr", - "enrichplot", - "gson", + "enrichplot (>= 1.9.3)", + "GO.db", + "GOSemSim (>= 2.27.2)", + "gson (>= 0.0.7)", "httr", "igraph", "magrittr", @@ -2133,1002 +5960,2847 @@ "stats", "tidyr", "utils", - "yulab.utils" + "yulab.utils (>= 0.2.3)" ], - "Hash": "4dcafdc7266ccabdde011cbab04b1730" + "Suggests": [ + "AnnotationHub", + "BiocManager", + "jsonlite", + "readr", + "org.Hs.eg.db", + "quarto", + "testthat" + ], + "VignetteBuilder": "quarto", + "ByteCompile": "true", + "License": "Artistic-2.0", + "URL": "https://yulab-smu.top/contribution-knowledge-mining/", + "BugReports": "https://github.com/YuLab-SMU/clusterProfiler/issues", + "biocViews": "Annotation, Clustering, GeneSetEnrichment, GO, KEGG, MultipleComparison, Pathways, Reactome, Visualization", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.3", + "git_url": "https://git.bioconductor.org/packages/clusterProfiler", + "git_branch": "RELEASE_3_22", + "git_last_commit": "16e6517", + "git_last_commit_date": "2025-12-15", + "Repository": "Bioconductor 3.22", + "NeedsCompilation": "no", + "Author": "Guangchuang Yu [aut, cre, cph] (ORCID: ), Li-Gen Wang [ctb], Xiao Luo [ctb], Meijun Chen [ctb], Giovanni Dall'Olio [ctb], Wanqian Wei [ctb], Chun-Hui Gao [ctb] (ORCID: )" }, "coda": { "Package": "coda", "Version": "0.19-4.1", "Source": "Repository", - 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Qualitative, sequential, and diverging color palettes based on HCL colors are provided along with corresponding ggplot2 color scales. Color palette choice is aided by an interactive app (with either a Tcl/Tk or a shiny graphical user interface) and shiny apps with an HCL color picker and a color vision deficiency emulator. Plotting functions for displaying and assessing palettes include color swatches, visualizations of the HCL space, and trajectories in HCL and/or RGB spectrum. Color manipulation functions include: desaturation, lightening/darkening, mixing, and simulation of color vision deficiencies (deutanomaly, protanomaly, tritanomaly). Details can be found on the project web page at and in the accompanying scientific paper: Zeileis et al. 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This can make it hard to detect conflicts, particularly when they arise because a package update creates ambiguity that did not previously exist. 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It has also been used extensively in the book Fundamentals of Data Visualization.", + "URL": "https://wilkelab.org/cowplot/", + "BugReports": "https://github.com/wilkelab/cowplot/issues", + "Depends": [ + "R (>= 3.5.0)" + ], + "Imports": [ + "ggplot2 (>= 3.5.2)", "grid", "gtable", + "grDevices", "methods", "rlang", "scales" ], - "Hash": "8ef2084dd7d28847b374e55440e4f8cb" + "License": "GPL-2", + "Suggests": [ + "Cairo", + "covr", + "dplyr", + "forcats", + "gridGraphics (>= 0.4-0)", + "knitr", + "lattice", + "magick", + "maps", + "PASWR", + "patchwork", + "rmarkdown", + "ragg", + "testthat (>= 1.0.0)", + "tidyr", + "vdiffr (>= 0.3.0)", + "VennDiagram" + ], + "VignetteBuilder": "knitr", + "Collate": "'add_sub.R' 'align_plots.R' 'as_grob.R' 'as_gtable.R' 'axis_canvas.R' 'cowplot.R' 'draw.R' 'get_plot_component.R' 'get_axes.R' 'get_titles.R' 'get_legend.R' 'get_panel.R' 'gtable.R' 'key_glyph.R' 'plot_grid.R' 'save.R' 'set_null_device.R' 'setup.R' 'stamp.R' 'themes.R' 'utils_ggplot2.R'", + "RoxygenNote": "7.3.2", + "Encoding": "UTF-8", + "NeedsCompilation": "no", + "Author": "Claus O. 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Compared to other approaches 'cpp11' strives to be safe against long jumps from the C API as well as C++ exceptions, conform to normal R function semantics and supports interaction with 'ALTREP' vectors.", + "License": "MIT + file LICENSE", + "URL": "https://cpp11.r-lib.org, https://github.com/r-lib/cpp11", + "BugReports": "https://github.com/r-lib/cpp11/issues", + "Depends": [ + "R (>= 4.0.0)" + ], + "Suggests": [ + "bench", + "brio", + "callr", + "cli", + "covr", + "decor", + "desc", + "ggplot2", + "glue", + "knitr", + "lobstr", + "mockery", + "progress", + "rmarkdown", + "scales", + "Rcpp", + "testthat (>= 3.2.0)", + "tibble", + "utils", + "vctrs", + "withr" ], - "Hash": "5a295d7d963cc5035284dcdbaf334f4e" + "VignetteBuilder": "knitr", + "Config/Needs/website": "tidyverse/tidytemplate", + "Config/testthat/edition": "3", + "Config/Needs/cpp11/cpp_register": "brio, cli, decor, desc, glue, tibble, vctrs", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.2", + "NeedsCompilation": "no", + "Author": "Davis Vaughan [aut, cre] (ORCID: ), Jim Hester [aut] (ORCID: ), Romain François [aut] (ORCID: ), Benjamin Kietzman [ctb], Posit Software, PBC [cph, fnd]", + "Maintainer": "Davis Vaughan ", + "Repository": "CRAN" }, "crayon": { "Package": "crayon", - "Version": "1.5.2", - "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ + "Version": "1.5.3", + "Source": "Repository", + "Title": "Colored Terminal Output", + "Authors@R": "c( person(\"Gábor\", \"Csárdi\", , \"csardi.gabor@gmail.com\", role = c(\"aut\", \"cre\")), person(\"Brodie\", \"Gaslam\", , \"brodie.gaslam@yahoo.com\", role = \"ctb\"), person(\"Posit Software, PBC\", role = c(\"cph\", \"fnd\")) )", + "Description": "The crayon package is now superseded. Please use the 'cli' package for new projects. Colored terminal output on terminals that support 'ANSI' color and highlight codes. It also works in 'Emacs' 'ESS'. 'ANSI' color support is automatically detected. Colors and highlighting can be combined and nested. New styles can also be created easily. 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Some knowledge of 'libcurl' is recommended; for a more-user-friendly web client see the 'httr2' package which builds on this package with http specific tools and logic.", + "License": "MIT + file LICENSE", + "SystemRequirements": "libcurl (>= 7.73): libcurl-devel (rpm) or libcurl4-openssl-dev (deb)", + "URL": "https://jeroen.r-universe.dev/curl", + "BugReports": "https://github.com/jeroen/curl/issues", + "Suggests": [ + "spelling", + "testthat (>= 1.0.0)", + "knitr", + "jsonlite", + "later", + "rmarkdown", + "httpuv (>= 1.4.4)", + "webutils" ], - "Hash": "411ca2c03b1ce5f548345d2fc2685f7a" + "VignetteBuilder": "knitr", + "Depends": [ + "R (>= 3.0.0)" + ], + "RoxygenNote": "7.3.2", + "Encoding": "UTF-8", + "Language": "en-US", + "NeedsCompilation": "yes", + "Author": "Jeroen Ooms [aut, cre] (ORCID: ), Hadley Wickham [ctb], Posit Software, PBC [cph]", + "Maintainer": "Jeroen Ooms ", + "Repository": "CRAN" }, "data.table": { "Package": "data.table", - "Version": "1.15.4", + "Version": "1.18.2.1", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", + "Title": "Extension of `data.frame`", + "Depends": [ + "R (>= 3.4.0)" + ], + "Imports": [ "methods" ], - "Hash": "8ee9ac56ef633d0c7cab8b2ca87d683e" + "Suggests": [ + "bit64 (>= 4.0.0)", + "bit (>= 4.0.4)", + "R.utils (>= 2.13.0)", + "xts", + "zoo (>= 1.8-1)", + "yaml", + "knitr", + "markdown" + ], + "Description": "Fast aggregation of large data (e.g. 100GB in RAM), fast ordered joins, fast add/modify/delete of columns by group using no copies at all, list columns, friendly and fast character-separated-value read/write. 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\"Girlich\", role = \"aut\"), person(\"Edgar\", \"Ruiz\", role = \"aut\"), person(\"Posit Software, PBC\", role = c(\"cph\", \"fnd\")) )", + "Description": "A 'dplyr' back end for databases that allows you to work with remote database tables as if they are in-memory data frames. Basic features works with any database that has a 'DBI' back end; more advanced features require 'SQL' translation to be provided by the package author.", + "License": "MIT + file LICENSE", + "URL": "https://dbplyr.tidyverse.org/, https://github.com/tidyverse/dbplyr", + "BugReports": "https://github.com/tidyverse/dbplyr/issues", + "Depends": [ + "R (>= 3.6)" + ], + "Imports": [ + "blob (>= 1.2.0)", + "cli (>= 3.6.1)", + "DBI (>= 1.1.3)", + "dplyr (>= 1.1.2)", + "glue (>= 1.6.2)", + "lifecycle (>= 1.0.3)", "magrittr", "methods", - "pillar", - "purrr", - "rlang", - "tibble", - "tidyr", - "tidyselect", + "pillar (>= 1.9.0)", + "purrr (>= 1.0.1)", + "R6 (>= 2.2.2)", + "rlang (>= 1.1.1)", + "tibble (>= 3.2.1)", + "tidyr (>= 1.3.0)", + "tidyselect (>= 1.2.1)", "utils", - "vctrs", - "withr" + "vctrs (>= 0.6.3)", + "withr (>= 2.5.0)" ], - "Hash": "39b2e002522bfd258039ee4e889e0fd1" + "Suggests": [ + "bit64", + "covr", + "knitr", + "Lahman", + "nycflights13", + "odbc (>= 1.4.2)", + "RMariaDB (>= 1.2.2)", + "rmarkdown", + "RPostgres (>= 1.4.5)", + "RPostgreSQL", + "RSQLite (>= 2.3.8)", + "testthat (>= 3.1.10)" + ], + "VignetteBuilder": "knitr", + "Config/Needs/website": "tidyverse/tidytemplate", + "Config/testthat/edition": "3", + "Config/testthat/parallel": "TRUE", + "Encoding": "UTF-8", + "Language": "en-gb", + "RoxygenNote": "7.3.3", + "Collate": "'db-sql.R' 'utils-check.R' 'import-standalone-types-check.R' 'import-standalone-obj-type.R' 'utils.R' 'sql.R' 'escape.R' 'translate-sql-cut.R' 'translate-sql-quantile.R' 'translate-sql-string.R' 'translate-sql-paste.R' 'translate-sql-helpers.R' 'translate-sql-window.R' 'translate-sql-conditional.R' 'backend-.R' 'backend-access.R' 'backend-hana.R' 'backend-hive.R' 'backend-impala.R' 'verb-copy-to.R' 'backend-mssql.R' 'backend-mysql.R' 'backend-odbc.R' 'backend-oracle.R' 'backend-postgres.R' 'backend-postgres-old.R' 'backend-redshift.R' 'backend-snowflake.R' 'backend-spark-sql.R' 'backend-sqlite.R' 'backend-teradata.R' 'build-sql.R' 'data-cache.R' 'data-lahman.R' 'data-nycflights13.R' 'db-escape.R' 'db-io.R' 'db.R' 'dbplyr.R' 'explain.R' 'ident.R' 'import-standalone-s3-register.R' 'join-by-compat.R' 'join-cols-compat.R' 'lazy-join-query.R' 'lazy-ops.R' 'lazy-query.R' 'lazy-select-query.R' 'lazy-set-op-query.R' 'memdb.R' 'optimise-utils.R' 'pillar.R' 'progress.R' 'sql-build.R' 'query-join.R' 'query-select.R' 'query-semi-join.R' 'query-set-op.R' 'query.R' 'reexport.R' 'remote.R' 'rows.R' 'schema.R' 'simulate.R' 'sql-clause.R' 'sql-expr.R' 'src-sql.R' 'src_dbi.R' 'table-name.R' 'tbl-lazy.R' 'tbl-sql.R' 'test-frame.R' 'testthat.R' 'tidyeval-across.R' 'tidyeval.R' 'translate-sql.R' 'utils-format.R' 'verb-arrange.R' 'verb-compute.R' 'verb-count.R' 'verb-distinct.R' 'verb-do-query.R' 'verb-do.R' 'verb-expand.R' 'verb-fill.R' 'verb-filter.R' 'verb-group_by.R' 'verb-head.R' 'verb-joins.R' 'verb-mutate.R' 'verb-pivot-longer.R' 'verb-pivot-wider.R' 'verb-pull.R' 'verb-select.R' 'verb-set-ops.R' 'verb-slice.R' 'verb-summarise.R' 'verb-uncount.R' 'verb-window.R' 'zzz.R'", + "NeedsCompilation": "no", + "Author": "Hadley Wickham [aut, cre], Maximilian Girlich [aut], Edgar Ruiz [aut], Posit Software, PBC [cph, fnd]", + "Maintainer": "Hadley Wickham ", + "Repository": "CRAN" }, "digest": { "Package": "digest", - 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Please note that this package is not meant to be deployed for cryptographic purposes for which more comprehensive (and widely tested) libraries such as 'OpenSSL' should be used.", + "URL": "https://github.com/eddelbuettel/digest, https://eddelbuettel.github.io/digest/, https://dirk.eddelbuettel.com/code/digest.html", + "BugReports": "https://github.com/eddelbuettel/digest/issues", + "Depends": [ + "R (>= 3.3.0)" + ], + "Imports": [ "utils" ], - "Hash": "698ece7ba5a4fa4559e3d537e7ec3d31" + "License": "GPL (>= 2)", + "Suggests": [ + "tinytest", + "simplermarkdown", + "rbenchmark" + ], + "VignetteBuilder": "simplermarkdown", + "Encoding": "UTF-8", + "NeedsCompilation": "yes", + "Author": "Dirk Eddelbuettel [aut, cre] (ORCID: ), Antoine Lucas [ctb] (ORCID: ), Jarek Tuszynski [ctb], Henrik Bengtsson [ctb] (ORCID: ), Simon Urbanek [ctb] (ORCID: ), Mario Frasca [ctb], Bryan Lewis [ctb], Murray Stokely [ctb], Hannes Muehleisen [ctb] (ORCID: ), Duncan Murdoch [ctb], Jim Hester [ctb] (ORCID: ), Wush Wu [ctb] (ORCID: ), Qiang Kou [ctb] (ORCID: ), Thierry Onkelinx [ctb] (ORCID: ), Michel Lang [ctb] (ORCID: ), Viliam Simko [ctb], Kurt Hornik [ctb] (ORCID: ), Radford Neal [ctb] (ORCID: ), Kendon Bell [ctb] (ORCID: ), Matthew de Queljoe [ctb], Dmitry Selivanov [ctb] (ORCID: ), Ion Suruceanu [ctb] (ORCID: ), Bill Denney [ctb] (ORCID: ), Dirk Schumacher [ctb], András Svraka [ctb] (ORCID: ), Sergey Fedorov [ctb] (ORCID: ), Will Landau [ctb] (ORCID: ), Floris Vanderhaeghe [ctb] (ORCID: ), Kevin Tappe [ctb], Harris McGehee [ctb], Tim Mastny [ctb], Aaron Peikert [ctb] (ORCID: ), Mark van der Loo [ctb] (ORCID: ), Chris Muir [ctb] (ORCID: ), Moritz Beller [ctb] (ORCID: ), Sebastian Campbell [ctb] (ORCID: ), Winston Chang [ctb] (ORCID: ), Dean Attali [ctb] (ORCID: ), Michael Chirico [ctb] (ORCID: ), Kevin Ushey [ctb] (ORCID: ), Carl Pearson [ctb] (ORCID: )", + "Maintainer": "Dirk Eddelbuettel ", + "Repository": "CRAN" + }, + "dir.expiry": { + "Package": "dir.expiry", + "Version": "1.18.0", + "Source": "Bioconductor", + "Date": "2024-10-17", + "Title": "Managing Expiration for Cache Directories", + "Description": "Implements an expiration system for access to versioned directories. Directories that have not been accessed by a registered function within a certain time frame are deleted. This aims to reduce disk usage by eliminating obsolete caches generated by old versions of packages.", + "Authors@R": "person(\"Aaron\", \"Lun\", role=c(\"aut\", \"cre\"), email=\"infinite.monkeys.with.keyboards@gmail.com\")", + "License": "GPL-3", + "Imports": [ + "utils", + "filelock" + ], + "Suggests": [ + "rmarkdown", + "knitr", + "testthat", + "BiocStyle" + ], + "biocViews": "Software, Infrastructure", + "VignetteBuilder": "knitr", + "RoxygenNote": "7.3.2", + "Encoding": "UTF-8", + "git_url": "https://git.bioconductor.org/packages/dir.expiry", + "git_branch": "RELEASE_3_22", + "git_last_commit": "6d768b3", + "git_last_commit_date": "2025-10-29", + "Repository": "Bioconductor 3.22", + "NeedsCompilation": "no", + "Author": "Aaron Lun [aut, cre]", + "Maintainer": "Aaron Lun " }, "doParallel": { "Package": "doParallel", "Version": "1.0.17", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", - "foreach", - "iterators", + "Type": "Package", + "Title": "Foreach Parallel Adaptor for the 'parallel' Package", + "Authors@R": "c(person(\"Folashade\", \"Daniel\", role=\"cre\", email=\"fdaniel@microsoft.com\"), person(\"Microsoft\", \"Corporation\", role=c(\"aut\", \"cph\")), person(\"Steve\", \"Weston\", role=\"aut\"), person(\"Dan\", \"Tenenbaum\", role=\"ctb\"))", + "Description": "Provides a parallel backend for the %dopar% function using the parallel package.", + "Depends": [ + "R (>= 2.14.0)", + "foreach (>= 1.2.0)", + "iterators (>= 1.0.0)", "parallel", "utils" ], - 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In addition fast functions for generating random numbers according to a uniform, normal and exponential distribution are included. The latter two use the Ziggurat algorithm originally proposed by Marsaglia and Tsang (2000, ). The fast sampling methods support unweighted sampling both with and without replacement. 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The goal of 'dtplyr' is to allow you to write 'dplyr' code that is automatically translated to the equivalent, but usually much faster, data.table code.", + "License": "MIT + file LICENSE", + "URL": "https://dtplyr.tidyverse.org, https://github.com/tidyverse/dtplyr", + "BugReports": "https://github.com/tidyverse/dtplyr/issues", + "Depends": [ + "R (>= 4.0)" + ], + "Imports": [ + "cli (>= 3.4.0)", + "data.table (>= 1.13.0)", + "dplyr (>= 1.1.0)", "glue", "lifecycle", - "rlang", + "rlang (>= 1.0.4)", "tibble", - "tidyselect", - "vctrs" + "tidyselect (>= 1.2.0)", + "vctrs (>= 0.4.1)" + ], + "Suggests": [ + "bench", + "covr", + "knitr", + "rmarkdown", + "testthat (>= 3.1.2)", + "tidyr (>= 1.1.0)", + "waldo (>= 0.3.1)" + ], + "VignetteBuilder": "knitr", + "Config/Needs/website": "tidyverse/tidytemplate", + "Config/testthat/edition": "3", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.3", + "NeedsCompilation": "no", + "Author": "Hadley Wickham [cre, aut], Maximilian Girlich [aut], Mark Fairbanks [aut], Ryan Dickerson [aut], Posit Software, PBC [cph, fnd]", + "Maintainer": "Hadley Wickham ", + "Repository": "CRAN" + }, + "e1071": { + "Package": "e1071", + "Version": "1.7-17", + "Source": "Repository", + "Title": "Misc Functions of the Department of Statistics, Probability Theory Group (Formerly: E1071), TU Wien", + "Imports": [ + "graphics", + "grDevices", + "class", + "stats", + "methods", + "utils", + "proxy" + ], + "Suggests": [ + "cluster", + "mlbench", + "nnet", + "randomForest", + "rpart", + "SparseM", + "xtable", + "Matrix", + "MASS", + "slam" ], - "Hash": "54ed3ea01b11e81a86544faaecfef8e2" + "Authors@R": "c(person(given = \"David\", family = \"Meyer\", role = c(\"aut\", \"cre\"), email = \"David.Meyer@R-project.org\", comment = c(ORCID = \"0000-0002-5196-3048\")), person(given = \"Evgenia\", family = \"Dimitriadou\", role = c(\"aut\",\"cph\")), person(given = \"Kurt\", family = \"Hornik\", role = \"aut\", email = \"Kurt.Hornik@R-project.org\", comment = c(ORCID = \"0000-0003-4198-9911\")), person(given = \"Andreas\", family = \"Weingessel\", role = \"aut\"), person(given = \"Friedrich\", family = \"Leisch\", role = \"aut\"), person(given = \"Chih-Chung\", family = \"Chang\", role = c(\"ctb\",\"cph\"), comment = \"libsvm C++-code\"), person(given = \"Chih-Chen\", family = \"Lin\", role = c(\"ctb\",\"cph\"), comment = \"libsvm C++-code\"))", + "Description": "Functions for latent class analysis, short time Fourier transform, fuzzy clustering, support vector machines, shortest path computation, bagged clustering, naive Bayes classifier, generalized k-nearest neighbour ...", + "License": "GPL-2 | GPL-3", + "LazyLoad": "yes", + "NeedsCompilation": "yes", + "Author": "David Meyer [aut, cre] (ORCID: ), Evgenia Dimitriadou [aut, cph], Kurt Hornik [aut] (ORCID: ), Andreas Weingessel [aut], Friedrich Leisch [aut], Chih-Chung Chang [ctb, cph] (libsvm C++-code), Chih-Chen Lin [ctb, cph] (libsvm C++-code)", + "Maintainer": "David Meyer ", + "Repository": "CRAN", + "Encoding": "UTF-8" }, "edgeR": { "Package": "edgeR", - "Version": "4.2.0", + "Version": "4.8.2", "Source": "Bioconductor", - "Repository": "Bioconductor 3.19", - "Requirements": [ - "R", - "Rcpp", - "graphics", - "limma", - "locfit", + "Date": "2025-12-23", + "Title": "Empirical Analysis of Digital Gene Expression Data in R", + "Description": "Differential expression analysis of sequence count data. 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This is a low-level utility for reading in Alevin EDS format into R. This function is not designed for end-users but instead the package is predominantly for simplifying package dependency graph for other Bioconductor packages.", + "Depends": [ + "Matrix" + ], + "Imports": [ "Rcpp" ], - "Hash": "ab19d02b3418e44d6f32ffb8060427d2" - }, - "ellipsis": { - "Package": "ellipsis", - "Version": "0.3.2", - "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", - "rlang" + "Suggests": [ + "knitr", + "tximportData", + "testthat (>= 3.0.0)" + ], + "LinkingTo": [ + "Rcpp" ], - "Hash": "bb0eec2fe32e88d9e2836c2f73ea2077" + "SystemRequirements": "C++11", + "License": "GPL-2", + "Encoding": "UTF-8", + "URL": "https://github.com/mikelove/eds", + "biocViews": "Sequencing, RNASeq, GeneExpression, SingleCell", + "VignetteBuilder": "knitr", + "RoxygenNote": "7.1.2", + "Config/testthat/edition": "3", + "git_url": "https://git.bioconductor.org/packages/eds", + "git_branch": "RELEASE_3_22", + "git_last_commit": "d2b8448", + "git_last_commit_date": "2025-10-29", + "Repository": "Bioconductor 3.22", + "NeedsCompilation": "yes", + "Author": "Avi Srivastava [aut, cre], Michael Love [aut, ctb]", + "Maintainer": "Avi Srivastava " }, "emdbook": { "Package": "emdbook", - "Version": "1.3.13", - "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ + "Version": "1.3.14", + "Source": "Repository", + "Type": "Package", + "Title": "Support Functions and Data for \"Ecological Models and Data\"", + "LazyData": "yes", + "Authors@R": "c(person(\"Ben\",\"Bolker\",email=\"bolker@mcmaster.ca\", role=c(\"aut\",\"cre\")), person(\"Sang Woo\",\"Park\",role=\"ctb\"), person(\"James\",\"Vonesh\",role=\"dtc\"), person(\"Jacqueline\",\"Wilson\",role=\"dtc\"), person(\"Russ\",\"Schmitt\",role=\"dtc\"), person(\"Sally\",\"Holbrook\",role=\"dtc\"), person(\"James D.\",\"Thomson\",role=\"dtc\"), person(\"R. Scot\",\"Duncan\",role=\"dtc\") )", + "Description": "Auxiliary functions and data sets for \"Ecological Models and Data\", a book presenting maximum likelihood estimation and related topics for ecologists (ISBN 978-0-691-12522-0).", + "Suggests": [ + "R2jags", + "ellipse", + "SuppDists", + "numDeriv", + "testthat", + "rgl" + ], + "Imports": [ "MASS", - "bbmle", - "coda", "lattice", - "plyr" + "plyr", + "coda", + "bbmle" ], - "Hash": "ed650db9168aeca46d35aa373b12e056" + "License": "GPL", + "URL": "https://math.mcmaster.ca/bolker/emdbook", + "NeedsCompilation": "no", + "Author": "Ben Bolker [aut, cre], Sang Woo Park [ctb], James Vonesh [dtc], Jacqueline Wilson [dtc], Russ Schmitt [dtc], Sally Holbrook [dtc], James D. Thomson [dtc], R. 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It is mainly designed to work with the 'clusterProfiler' package suite. All the visualization methods are developed based on 'ggplot2' graphics.", + "Depends": [ + "R (>= 4.2.0)" + ], + "Imports": [ + "aplot (>= 0.2.1)", + "DOSE (>= 3.31.2)", + "ggfun (>= 0.1.7)", "ggnewscale", - "ggplot2", - "ggraph", - "ggtree", + "ggplot2 (>= 3.5.0)", + "ggrepel (>= 0.9.0)", + "ggtangle (>= 0.0.9)", "graphics", "grid", "igraph", - "magrittr", "methods", "plyr", "purrr", + "RColorBrewer", "reshape2", "rlang", - "scatterpie", - "shadowtext", "stats", + "tidydr", "utils", - "yulab.utils" + "scatterpie", + "GOSemSim (>= 2.31.2)", + "ggtree", + "yulab.utils (>= 0.1.6)" ], - "Hash": "19039e8a7075c61615d6f1459fb13ec6" + "Suggests": [ + "clusterProfiler", + "dplyr", + "europepmc", + "ggarchery", + "ggupset", + "glue", + "knitr", + "rmarkdown", + "org.Hs.eg.db", + "prettydoc", + "tibble", + "tidyr", + "ggforce", + "ggHoriPlot", + "AnnotationDbi", + "ggplotify", + "ggridges", + "grDevices", + "gridExtra", + "ggstar", + "scales", + "ggtreeExtra" + ], + "VignetteBuilder": "knitr", + "License": "Artistic-2.0", + "URL": "https://yulab-smu.top/contribution-knowledge-mining/", + "BugReports": "https://github.com/GuangchuangYu/enrichplot/issues", + "biocViews": "Annotation, GeneSetEnrichment, GO, KEGG, Pathways, Software, Visualization", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.3", + "git_url": "https://git.bioconductor.org/packages/enrichplot", + "git_branch": "RELEASE_3_22", + "git_last_commit": "920c1db", + "git_last_commit_date": "2025-11-30", + "Repository": "Bioconductor 3.22", + "NeedsCompilation": "no", + "Author": "Guangchuang Yu [aut, cre] (ORCID: ), Chun-Hui Gao [ctb] (ORCID: )", + "Maintainer": "Guangchuang Yu " }, "ensembldb": { "Package": "ensembldb", - "Version": "2.28.0", + "Version": "2.34.0", "Source": "Bioconductor", - "Repository": "Bioconductor 3.19", - "Requirements": [ - "AnnotationDbi", - "AnnotationFilter", - "Biobase", - "BiocGenerics", - "Biostrings", + "Type": "Package", + "Title": "Utilities to create and use Ensembl-based annotation databases", + "Authors@R": "c(person(given = \"Johannes\", family = \"Rainer\", email = \"johannes.rainer@eurac.edu\", role = c(\"aut\", \"cre\"), comment = c(ORCID = \"0000-0002-6977-7147\")), person(given = \"Tim\", family = \"Triche\", email = \"tim.triche@usc.edu\", role = \"ctb\"), person(given = \"Christian\", family = \"Weichenberger\", email = \"christian.weichenberger@eurac.edu\", role = \"ctb\", comment = c(ORCID = \"0000-0002-2176-0274\")), person(given = \"Sebastian\", family = \"Gibb\", email = \"mail@sebastiangibb.de\", role = \"ctb\", comment = c(ORCID = \"0000-0001-7406-4443\")), person(given = \"Laurent\", family = \"Gatto\", email = \"lg390@cam.ac.uk\", role = \"ctb\", comment = c(ORCID = \"0000-0002-1520-2268\")), person(given = \"Boyu\", family = \"Yu\", email = \"boyu.yu.tim@gmail.com\", role = \"ctb\"))", + "Author": "Johannes Rainer with contributions from Tim Triche, Sebastian Gibb, Laurent Gatto Christian Weichenberger and Boyu Yu.", + "Maintainer": "Johannes Rainer ", + "URL": "https://github.com/jorainer/ensembldb", + "BugReports": "https://github.com/jorainer/ensembldb/issues", + "Imports": [ + "methods", + "RSQLite (>= 1.1)", "DBI", - "GenomeInfoDb", - "GenomicFeatures", - "GenomicRanges", - "IRanges", - "ProtGenerics", - "R", - "RSQLite", + "Biobase", + "Seqinfo", + "GenomeInfoDb (>= 1.45.5)", + "AnnotationDbi (>= 1.31.19)", + "rtracklayer (>= 1.69.1)", + "S4Vectors (>= 0.23.10)", "Rsamtools", - "S4Vectors", - "curl", - "methods", - "rtracklayer" + "IRanges (>= 2.13.24)", + "ProtGenerics", + "Biostrings (>= 2.77.2)", + "curl" + ], + "Depends": [ + "R (>= 3.5.0)", + "BiocGenerics (>= 0.15.10)", + "GenomicRanges (>= 1.61.1)", + "GenomicFeatures (>= 1.61.4)", + "AnnotationFilter (>= 1.5.2)" + ], + "Suggests": [ + "BiocStyle", + "knitr", + "EnsDb.Hsapiens.v86 (>= 0.99.8)", + "testthat", + "BSgenome.Hsapiens.NCBI.GRCh38", + "ggbio (>= 1.24.0)", + "Gviz (>= 1.20.0)", + "rmarkdown", + "AnnotationHub" + ], + "Enhances": [ + "RMariaDB", + "shiny" + ], + "VignetteBuilder": "knitr", + "Description": "The package provides functions to create and use transcript centric annotation databases/packages. The annotation for the databases are directly fetched from Ensembl using their Perl API. The functionality and data is similar to that of the TxDb packages from the GenomicFeatures package, but, in addition to retrieve all gene/transcript models and annotations from the database, ensembldb provides a filter framework allowing to retrieve annotations for specific entries like genes encoded on a chromosome region or transcript models of lincRNA genes. EnsDb databases built with ensembldb contain also protein annotations and mappings between proteins and their encoding transcripts. Finally, ensembldb provides functions to map between genomic, transcript and protein coordinates.", + "Collate": "'Classes.R' 'Deprecated.R' 'Generics.R' 'Methods-Filter.R' 'Methods.R' 'dbhelpers.R' 'functions-EnsDb.R' 'functions-Filter.R' 'functions-create-EnsDb.R' 'functions-utils.R' 'proteinToX.R' 'transcriptToX.R' 'genomeToX.R' 'select-methods.R' 'seqname-utils.R' 'zzz.R'", + "biocViews": "Genetics, AnnotationData, Sequencing, Coverage", + "License": "LGPL", + "RoxygenNote": "7.3.2", + "git_url": "https://git.bioconductor.org/packages/ensembldb", + "git_branch": "RELEASE_3_22", + "git_last_commit": "a13967b", + "git_last_commit_date": "2025-10-29", + "Repository": "Bioconductor 3.22", + "NeedsCompilation": "no" + }, + "escheR": { + "Package": "escheR", + "Version": "1.10.0", + "Source": "Bioconductor", + "Title": "Unified multi-dimensional visualizations with Gestalt principles", + "Authors@R": "c( person(\"Boyi\", \"Guo\", email = \"boyi.guo.work@gmail.com\", role = c(\"aut\", \"cre\"), comment = c(ORCID = \"0000-0003-2950-2349\")), person(c(\"Stephanie\", \"C.\"), \"Hicks\", role = c(\"aut\"), email = \"shicks19@jhu.edu\", comment = c(ORCID = \"0000-0002-7858-0231\")), person(c(\"Erik\", \"D.\"), \"Nelson\", email = \"erik.nelson116@gmail.com\", role = c(\"ctb\"), comment = c(ORCID = \"0000-0001-8477-0982\")) )", + "Description": "The creation of effective visualizations is a fundamental component of data analysis. In biomedical research, new challenges are emerging to visualize multi-dimensional data in a 2D space, but current data visualization tools have limited capabilities. To address this problem, we leverage Gestalt principles to improve the design and interpretability of multi-dimensional data in 2D data visualizations, layering aesthetics to display multiple variables. The proposed visualization can be applied to spatially-resolved transcriptomics data, but also broadly to data visualized in 2D space, such as embedding visualizations. We provide this open source R package escheR, which is built off of the state-of-the-art ggplot2 visualization framework and can be seamlessly integrated into genomics toolboxes and workflows.", + "License": "MIT + file LICENSE", + "Encoding": "UTF-8", + "Roxygen": "list(markdown = TRUE)", + "RoxygenNote": "7.2.3", + "biocViews": "Spatial, SingleCell, Transcriptomics, Visualization, Software", + "Depends": [ + "ggplot2", + "R (>= 4.3)" + ], + "Imports": [ + "SpatialExperiment (>= 1.6.1)", + "SingleCellExperiment", + "rlang", + "SummarizedExperiment" ], - "Hash": "f9a5e52468ec832a839c012e15c41c15" + "BugReports": "https://github.com/boyiguo1/escheR/issues", + "URL": "https://github.com/boyiguo1/escheR", + "Suggests": [ + "STexampleData", + "BumpyMatrix", + "knitr", + "rmarkdown", + "BiocStyle", + "ggpubr", + "scran", + "scater", + "scuttle", + "Seurat", + "hexbin" + ], + "VignetteBuilder": "knitr", + "git_url": "https://git.bioconductor.org/packages/escheR", + "git_branch": "RELEASE_3_22", + "git_last_commit": "3f79e54", + "git_last_commit_date": "2025-10-29", + "Repository": "Bioconductor 3.22", + "NeedsCompilation": "no", + "Author": "Boyi Guo [aut, cre] (ORCID: ), Stephanie C. 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As different colour spaces have different uses, efficient conversion between these representations are important. The 'farver' package provides a set of functions that gives access to very fast colour space conversion and comparisons implemented in C++, and offers speed improvements over the 'convertColor' function in the 'grDevices' package.", + "License": "MIT + file LICENSE", + "URL": "https://farver.data-imaginist.com, https://github.com/thomasp85/farver", + "BugReports": "https://github.com/thomasp85/farver/issues", + "Suggests": [ + "covr", + "testthat (>= 3.0.0)" + ], + "Config/testthat/edition": "3", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.1", + "NeedsCompilation": "yes", + "Author": "Thomas Lin Pedersen [cre, aut] (), Berendea Nicolae [aut] (Author of the ColorSpace C++ library), Romain François [aut] (), Posit, PBC [cph, fnd]", + "Maintainer": "Thomas Lin Pedersen ", + "Repository": "CRAN" }, "fastmap": { "Package": "fastmap", - "Version": "1.1.1", + "Version": "1.2.0", "Source": "Repository", - "Repository": "CRAN", - "Hash": "f7736a18de97dea803bde0a2daaafb27" + "Title": "Fast Data Structures", + "Authors@R": "c( person(\"Winston\", \"Chang\", email = \"winston@posit.co\", role = c(\"aut\", \"cre\")), person(given = \"Posit Software, PBC\", role = c(\"cph\", \"fnd\")), person(given = \"Tessil\", role = \"cph\", comment = \"hopscotch_map library\") )", + "Description": "Fast implementation of data structures, including a key-value store, stack, and queue. 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Labels repel away from each other and away from the data points.", + "Depends": [ + "R (>= 3.0.0)", + "ggplot2 (>= 2.2.0)" + ], + "Imports": [ "grid", - "rlang", - "scales", - "withr" + "Rcpp", + "rlang (>= 0.3.0)", + "scales (>= 0.5.0)", + "withr (>= 2.5.0)" ], - "Hash": "cc3361e234c4a5050e29697d675764aa" - }, - "ggsignif": { - "Package": "ggsignif", - "Version": "0.6.4", - "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ + "Suggests": [ + "knitr", + "rmarkdown", + "testthat", + "svglite", + "vdiffr", + "gridExtra", + "ggpp", + "patchwork", + "devtools", + "prettydoc", + "ggbeeswarm", + "dplyr", + "magrittr", + "readr", + "stringr" + ], + "VignetteBuilder": "knitr", + "License": "GPL-3 | file LICENSE", + "URL": "https://ggrepel.slowkow.com/, https://github.com/slowkow/ggrepel", + "BugReports": "https://github.com/slowkow/ggrepel/issues", + "RoxygenNote": "7.3.1", + "LinkingTo": [ + "Rcpp" + ], + "Encoding": "UTF-8", + "NeedsCompilation": "yes", + "Author": "Kamil Slowikowski [aut, cre] (), Alicia Schep [ctb] (), Sean Hughes [ctb] (), Trung Kien Dang [ctb] (), Saulius Lukauskas [ctb], Jean-Olivier Irisson [ctb] (), Zhian N Kamvar [ctb] (), Thompson Ryan [ctb] (), Dervieux Christophe [ctb] (), Yutani Hiroaki [ctb], Pierre Gramme [ctb], Amir Masoud Abdol [ctb], Malcolm Barrett [ctb] (), Robrecht Cannoodt [ctb] (), Michał Krassowski [ctb] (), Michael Chirico [ctb] (), Pedro Aphalo [ctb] (), Francis Barton [ctb]", + "Maintainer": "Kamil Slowikowski ", + "Repository": "CRAN" + }, + "ggside": { + "Package": "ggside", + "Version": "0.4.1", + "Source": "Repository", + "Type": "Package", + "Title": "Side Grammar Graphics", + "Authors@R": "person(given = \"Justin\", family = \"Landis\", role = c(\"aut\", \"cre\"), email = \"jtlandis314@gmail.com\", comment = c(ORCID = \"0000-0001-5501-4934\"))", + "Maintainer": "Justin Landis ", + "Description": "The grammar of graphics as shown in 'ggplot2' has provided an expressive API for users to build plots. 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This package is particularly useful for visualizing metadata on a discrete axis, or summary graphics on a continuous axis such as a boxplot or a density distribution.", + "License": "MIT + file LICENSE", + "URL": "https://github.com/jtlandis/ggside", + "BugReports": "https://github.com/jtlandis/ggside/issues", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.3", + "VignetteBuilder": "knitr", + "Depends": [ + "R (>= 4.1)", + "ggplot2 (>= 4.0.0)" + ], + "Imports": [ + "grid", + "gtable", + "rlang", + "scales (>= 1.3.0)", + "cli", + "glue", + "stats", + "tibble", + "vctrs", + "S7", + "lifecycle" + ], + "Suggests": [ + "tidyr", + "dplyr", + "testthat (>= 3.0.3)", + "knitr", + "rmarkdown", + "vdiffr (>= 1.0.0)", + "ggdendro", + "viridis", + "waldo" + ], + "Config/testthat/edition": "3", + "Config/testthat/parallel": "true", + "Config/testthat/start-first": "all_ggside_layers, *themes", + "Collate": "'utils-ggproto.R' 'utils-calls.R' 'utils-ggplot2-reimpl-.R' 'utils-constructors.R' 'side-layer.R' 'constructor-.R' 'utils-.R' 'ggside.R' 'utils-side-facet.R' 'side-facet_.R' 'utils-side-coord.R' 'side-coord-cartesian.R' 'add_gg.R' 'ggplot_add.R' 'side-layout-.r' 'all_classes.r' 'geom-sideabline.r' 'geom-sidebar.r' 'geom-sideboxplot.r' 'geom-sidecol.r' 'geom-sidedensity.r' 'geom-sidefreqpoly.r' 'geom-sidefunction.r' 'geom-sidehistogram.r' 'geom-sidehline.r' 'geom-sidelabel.r' 'geom-sideline.r' 'geom-sidepath.r' 'geom-sidepoint.r' 'geom-sidesegment.r' 'geom-sidetext.r' 'geom-sidetile.r' 'geom-sideviolin.r' 'geom-sidevline.r' 'ggside-ggproto.r' 'ggside-package.r' 'ggside-themes.R' 'plot-construction.R' 'position_rescale.r' 'scales-sides-.R' 'scales-xycolour.R' 'scales-xyfill.R' 'utils-ggplot2-reimpl-facet.R' 'side-facet-wrap.R' 'side-facet-grid.R' 'side-facet-null.R' 'stats.r' 'update_ggplot.R' 'z-depricated.R' 'zzz.R'", + "NeedsCompilation": "no", + "Author": "Justin Landis [aut, cre] (ORCID: )", + "Repository": "CRAN" + }, + "ggsignif": { + "Package": "ggsignif", + "Version": "0.6.4", + "Source": "Repository", + "Type": "Package", + "Title": "Significance Brackets for 'ggplot2'", + "Authors@R": "c( person(given = \"Constantin\", family = \"Ahlmann-Eltze\", role = c(\"aut\", \"cre\", \"ctb\"), email = \"artjom31415@googlemail.com\", comment = c(ORCID = \"0000-0002-3762-068X\", Twitter = \"@const_ae\")), person(given = \"Indrajeet\", family = \"Patil\", role = c(\"aut\", \"ctb\"), email = \"patilindrajeet.science@gmail.com\", comment = c(ORCID = \"0000-0003-1995-6531\", Twitter = \"@patilindrajeets\")) )", + "Description": "Enrich your 'ggplots' with group-wise comparisons. This package provides an easy way to indicate if two groups are significantly different. Commonly this is shown by a bracket on top connecting the groups of interest which itself is annotated with the level of significance (NS, *, **, ***). The package provides a single layer (geom_signif()) that takes the groups for comparison and the test (t.test(), wilcox.text() etc.) as arguments and adds the annotation to the plot.", + "License": "GPL-3 | file LICENSE", + "URL": "https://const-ae.github.io/ggsignif/, https://github.com/const-ae/ggsignif", + "VignetteBuilder": "knitr", + "Encoding": "UTF-8", + "Language": "en-US", + "Imports": [ + "ggplot2 (>= 3.3.5)" + ], + "Suggests": [ + "knitr", + "rmarkdown", + "testthat", + "vdiffr (>= 1.0.2)" + ], + "RoxygenNote": "7.2.1", + "Config/testthat/edition": "3", + "Config/testthat/parallel": "true", + "NeedsCompilation": "no", + "Author": "Constantin Ahlmann-Eltze [aut, cre, ctb] (, @const_ae), Indrajeet Patil [aut, ctb] (, @patilindrajeets)", + "Maintainer": "Constantin Ahlmann-Eltze ", + "Repository": "CRAN" + }, + "ggspavis": { + "Package": "ggspavis", + "Version": "1.16.0", + "Source": "Bioconductor", + "Title": "Visualization functions for spatial transcriptomics data", + "Description": "Visualization functions for spatial transcriptomics data. Includes functions to generate several types of plots, including spot plots, feature (molecule) plots, reduced dimension plots, spot-level quality control (QC) plots, and feature-level QC plots, for datasets from the 10x Genomics Visium and other technological platforms. 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Drafts 04, 06 and 07 of 'JSON' schema are supported.", + "License": "MIT + file LICENSE", + "URL": "https://docs.ropensci.org/jsonvalidate/, https://github.com/ropensci/jsonvalidate", + "BugReports": "https://github.com/ropensci/jsonvalidate/issues", + "Imports": [ + "R6", "V8" ], - "Hash": "cdc2843ef7f44f157198bb99aea7552d" + "Suggests": [ + "knitr", + "jsonlite", + "rmarkdown", + "testthat", + "withr" + ], + "RoxygenNote": "7.3.2", + "VignetteBuilder": "knitr", + "Encoding": "UTF-8", + "Language": "en-GB", + "Config/testthat/edition": "3", + "NeedsCompilation": "no", + "Author": "Rich FitzJohn [aut, cre], Rob Ashton [aut], Alex Hill [ctb], Alicia Schep [ctb], Ian Lyttle [ctb], Kara Woo [ctb], Mathias Buus [aut, cph] (Author of bundled imjv library), Evgeny Poberezkin [aut, cph] (Author of bundled Ajv library)", + "Repository": "CRAN" }, "kernlab": { "Package": "kernlab", "Version": "0.9-33", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", - "grDevices", - "graphics", + "Title": "Kernel-Based Machine Learning Lab", + "Authors@R": "c(person(\"Alexandros\", \"Karatzoglou\", role = c(\"aut\", \"cre\"), email = \"alexandros.karatzoglou@gmail.com\"), person(\"Alex\", \"Smola\", role = \"aut\"), person(\"Kurt\", \"Hornik\", role = \"aut\", email = \"Kurt.Hornik@R-project.org\", comment = c(ORCID = \"0000-0003-4198-9911\")), person(\"National ICT Australia (NICTA)\", role = \"cph\"), person(c(\"Michael\", \"A.\"), \"Maniscalco\", role = c(\"ctb\", \"cph\")), person(c(\"Choon\", \"Hui\"), \"Teo\", role = \"ctb\"))", + "Description": "Kernel-based machine learning methods for classification, regression, clustering, novelty detection, quantile regression and dimensionality reduction. 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The function rst2pdf() requires rst2pdf (https://github.com/rst2pdf/rst2pdf).", + "Collate": "'block.R' 'cache.R' 'citation.R' 'hooks-html.R' 'plot.R' 'utils.R' 'defaults.R' 'concordance.R' 'engine.R' 'highlight.R' 'themes.R' 'header.R' 'hooks-asciidoc.R' 'hooks-chunk.R' 'hooks-extra.R' 'hooks-latex.R' 'hooks-md.R' 'hooks-rst.R' 'hooks-textile.R' 'hooks.R' 'otel.R' 'output.R' 'package.R' 'pandoc.R' 'params.R' 'parser.R' 'pattern.R' 'rocco.R' 'spin.R' 'table.R' 'template.R' 'utils-conversion.R' 'utils-rd2html.R' 'utils-string.R' 'utils-sweave.R' 'utils-upload.R' 'utils-vignettes.R' 'zzz.R'", + "RoxygenNote": "7.3.3", + "NeedsCompilation": "no", + "Author": "Yihui Xie [aut, cre] (ORCID: , URL: https://yihui.org), Abhraneel Sarma [ctb], Adam Vogt [ctb], Alastair Andrew [ctb], Alex Zvoleff [ctb], Amar Al-Zubaidi [ctb], Andre Simon [ctb] (the CSS files under inst/themes/ were derived from the Highlight package http://www.andre-simon.de), Aron Atkins [ctb], Aaron Wolen [ctb], Ashley Manton [ctb], Atsushi Yasumoto [ctb] (ORCID: ), Ben Baumer [ctb], Brian Diggs [ctb], Brian Zhang [ctb], Bulat Yapparov [ctb], Cassio Pereira [ctb], Christophe Dervieux [ctb], David Hall [ctb], David Hugh-Jones [ctb], David Robinson [ctb], Doug Hemken [ctb], Duncan Murdoch [ctb], Elio Campitelli [ctb], Ellis Hughes [ctb], Emily Riederer [ctb], Fabian Hirschmann [ctb], Fitch Simeon [ctb], Forest Fang [ctb], Frank E Harrell Jr [ctb] (the Sweavel package at inst/misc/Sweavel.sty), Garrick Aden-Buie [ctb], Gregoire Detrez [ctb], Hadley Wickham [ctb], Hao Zhu [ctb], Heewon Jeon [ctb], Henrik Bengtsson [ctb], Hiroaki Yutani [ctb], Ian Lyttle [ctb], Hodges Daniel [ctb], Jacob Bien [ctb], Jake Burkhead [ctb], James Manton [ctb], Jared Lander [ctb], Jason Punyon [ctb], Javier Luraschi [ctb], Jeff Arnold [ctb], Jenny Bryan [ctb], Jeremy Ashkenas [ctb, cph] (the CSS file at inst/misc/docco-classic.css), Jeremy Stephens [ctb], Jim Hester [ctb], Joe Cheng [ctb], Johannes Ranke [ctb], John Honaker [ctb], John Muschelli [ctb], Jonathan Keane [ctb], JJ Allaire [ctb], Johan Toloe [ctb], Jonathan Sidi [ctb], Joseph Larmarange [ctb], Julien Barnier [ctb], Kaiyin Zhong [ctb], Kamil Slowikowski [ctb], Karl Forner [ctb], Kevin K. 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Kamvar [ctb] (ORCID: ), Posit Software, PBC [cph, fnd]", + "Maintainer": "Yihui Xie ", + "Repository": "CRAN" }, "labeling": { "Package": "labeling", "Version": "0.4.3", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "graphics", - "stats" + "Type": "Package", + "Title": "Axis Labeling", + "Date": "2023-08-29", + "Author": "Justin Talbot,", + "Maintainer": "Nuno Sempere ", + "Description": "Functions which provide a range of axis labeling algorithms.", + "License": "MIT + file LICENSE | Unlimited", + "Collate": "'labeling.R'", + "NeedsCompilation": "no", + "Imports": [ + "stats", + "graphics" ], - "Hash": "b64ec208ac5bc1852b285f665d6368b3" - }, - "labelled": { - "Package": "labelled", - "Version": "2.13.0", - "Source": "Repository", "Repository": "CRAN", - "Requirements": [ - "R", - "dplyr", - "haven", - "lifecycle", - "rlang", - "stringr", - "tidyr", - "tidyselect", - "vctrs" - ], - "Hash": "ad4b6d757624221aec6220b8c78defeb" + "Encoding": "UTF-8" }, "lambda.r": { "Package": "lambda.r", "Version": "1.2.4", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", + "Type": "Package", + "Title": "Modeling Data with Functional Programming", + "Date": "2019-09-15", + "Depends": [ + "R (>= 3.0.0)" + ], + "Imports": [ "formatR" ], - "Hash": "b1e925c4b9ffeb901bacf812cbe9a6ad" + "Suggests": [ + "testit" + ], + "Author": "Brian Lee Yung Rowe", + "Maintainer": "Brian Lee Yung Rowe ", + "Description": "A language extension to efficiently write functional programs in R. 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Anitescu (2020) .", + "License": "MIT + file LICENSE", + "Imports": [ + "utils", "stats", - "utils" + "irlba", + "Rcpp (>= 0.12.15)" + ], + "Suggests": [ + "testthat", + "knitr", + "rmarkdown" + ], + "LinkingTo": [ + "Rcpp", + "RcppArmadillo" ], - "Hash": "62896dd832ccbf1091e7f13324c24160" + "LazyData": "true", + "NeedsCompilation": "yes", + "VignetteBuilder": "knitr", + "RoxygenNote": "7.1.2", + "Author": "Youngseok Kim [aut], Peter Carbonetto [aut, cre], Mihai Anitescu [aut], Matthew Stephens [aut], Jason Willwerscheid [ctb], Jean Morrison [ctb]", + "Maintainer": "Peter Carbonetto ", + "Repository": "CRAN" }, "mixtools": { "Package": "mixtools", - "Version": "2.0.0", + "Version": "2.0.0.1", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "MASS", - "R", + "Date": "2022-12-04", + "Title": "Tools for Analyzing Finite Mixture Models", + "Authors@R": "c(person(\"Derek\", \"Young\", role = c(\"aut\", \"cre\"), email = \"derek.young@uky.edu\", comment = c(ORCID = \"0000-0002-3048-3803\")), person(\"Tatiana\", \"Benaglia\", role = \"aut\"), person(\"Didier\", \"Chauveau\", role = \"aut\"), person(\"David\", \"Hunter\", role = \"aut\"), person(\"Kedai\", \"Cheng\", role = \"aut\"), person(\"Ryan\", \"Elmore\", role = \"ctb\"), person(\"Thomas\", \"Hettmansperger\", role = \"ctb\"), person(\"Hoben\", \"Thomas\", role = \"ctb\"), person(\"Fengjuan\", \"Xuan\", role = \"ctb\"))", + "Depends": [ + "R (>= 4.0.0)" + ], + "Imports": [ "kernlab", + "MASS", "plotly", "scales", "segmented", "stats", "survival" ], - "Hash": "2b9414057d7f3725130e2f743ea05a2f" + "URL": "https://github.com/dsy109/mixtools", + "Description": "Analyzes finite mixture models for various parametric and semiparametric settings. 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This package is based upon work supported by the National Science Foundation under Grant No. SES-0518772 and the Chan Zuckerberg Initiative: Essential Open Source Software for Science (Grant No. 2020-255193).", + "License": "GPL (>= 2)", + "NeedsCompilation": "yes", + "Author": "Derek Young [aut, cre] (), Tatiana Benaglia [aut], Didier Chauveau [aut], David Hunter [aut], Kedai Cheng [aut], Ryan Elmore [ctb], Thomas Hettmansperger [ctb], Hoben Thomas [ctb], Fengjuan Xuan [ctb]", + "Maintainer": "Derek Young ", + "Repository": "CRAN", + "Encoding": "UTF-8" }, "modelr": { "Package": "modelr", "Version": "0.1.11", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", + "Title": "Modelling Functions that Work with the Pipe", + "Authors@R": "c( person(\"Hadley\", \"Wickham\", , \"hadley@posit.co\", role = c(\"aut\", \"cre\")), person(\"Posit Software, PBC\", role = c(\"cph\", \"fnd\")) )", + "Description": "Functions for modelling that help you seamlessly integrate modelling into a pipeline of data manipulation and visualisation.", + "License": "GPL-3", + "URL": "https://modelr.tidyverse.org, https://github.com/tidyverse/modelr", + "BugReports": "https://github.com/tidyverse/modelr/issues", + "Depends": [ + "R (>= 3.2)" + ], + "Imports": [ "broom", "magrittr", - "purrr", - "rlang", + "purrr (>= 0.2.2)", + "rlang (>= 1.0.6)", "tibble", - "tidyr", + "tidyr (>= 0.8.0)", "tidyselect", "vctrs" ], - "Hash": "4f50122dc256b1b6996a4703fecea821" + "Suggests": [ + "compiler", + "covr", + "ggplot2", + "testthat (>= 3.0.0)" + ], + "Config/Needs/website": "tidyverse/tidytemplate", + "Encoding": "UTF-8", + "LazyData": "true", + "RoxygenNote": "7.2.3", + "Config/testthat/edition": "3", + "NeedsCompilation": "no", + "Author": "Hadley Wickham [aut, cre], Posit Software, PBC [cph, fnd]", + "Maintainer": "Hadley Wickham ", + "Repository": "CRAN" }, "modeltools": { "Package": "modeltools", - "Version": "0.2-23", + "Version": "0.2-24", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "methods", + "Title": "Tools and Classes for Statistical Models", + "Date": "2025-05-02", + "Authors@R": "c(person(given = \"Torsten\", family = \"Hothorn\", role = c(\"aut\", \"cre\"), email = \"Torsten.Hothorn@R-project.org\", comment = c(ORCID = \"0000-0001-8301-0471\")), person(given = \"Friedrich\", family = \"Leisch\", role = \"aut\", comment = c(ORCID = \"0000-0001-7278-1983\")), person(given = \"Achim\", family = \"Zeileis\", role = \"aut\", comment = c(ORCID = \"0000-0003-0918-3766\")))", + "Description": "A collection of tools to deal with statistical models. The functionality is experimental and the user interface is likely to change in the future. The documentation is rather terse, but packages `coin' and `party' have some working examples. However, if you find the implemented ideas interesting we would be very interested in a discussion of this proposal. 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Using 'renv', you can create and manage project-local R libraries, save the state of these libraries to a 'lockfile', and later restore your library as required. 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This package is its R interface. The package includes efficient linear model solver and tree learning algorithms. The package can automatically do parallel computation on a single machine which could be more than 10 times faster than existing gradient boosting packages. It supports various objective functions, including regression, classification and ranking. The package is made to be extensible, so that users are also allowed to define their own objectives easily.", + "License": "Apache License (== 2.0) | file LICENSE", + "URL": "https://github.com/dmlc/xgboost", + "BugReports": "https://github.com/dmlc/xgboost/issues", + "NeedsCompilation": "yes", + "VignetteBuilder": "knitr", + "Suggests": [ + "knitr", + "rmarkdown", + "ggplot2 (>= 1.0.1)", + "DiagrammeR (>= 0.9.0)", + "DiagrammeRsvg", + "rsvg", + "htmlwidgets", + "Ckmeans.1d.dp (>= 3.3.1)", + "vcd (>= 1.3)", + "testthat", + "igraph (>= 1.0.1)", + "float", + "titanic", + "RhpcBLASctl", + "survival" + ], + "Depends": [ + "R (>= 4.3.0)" + ], + "Imports": [ + "Matrix (>= 1.1-0)", + "methods", + "data.table (>= 1.9.6)", + "jsonlite (>= 1.0)" ], - "Hash": "6303e61eac62aef7bd2b396ef7e24386" + "RoxygenNote": "7.3.3", + "Encoding": "UTF-8", + "SystemRequirements": "GNU make, C++17", + "Author": "Tianqi Chen [aut], Tong He [aut], Michael Benesty [aut], Vadim Khotilovich [aut], Yuan Tang [aut] (ORCID: ), Hyunsu Cho [aut], Kailong Chen [aut], Rory Mitchell [aut], Ignacio Cano [aut], Tianyi Zhou [aut], Mu Li [aut], Junyuan Xie [aut], Min Lin [aut], Yifeng Geng [aut], Yutian Li [aut], Jiaming Yuan [aut, cre], David Cortes [aut], XGBoost contributors [cph] (base XGBoost implementation)", + "Repository": "CRAN" }, "xml2": { "Package": "xml2", - "Version": "1.3.6", + "Version": "1.5.2", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", + "Title": "Parse XML", + "Authors@R": "c( person(\"Hadley\", \"Wickham\", role = \"aut\"), person(\"Jim\", \"Hester\", role = \"aut\"), person(\"Jeroen\", \"Ooms\", email = \"jeroenooms@gmail.com\", role = c(\"aut\", \"cre\")), person(\"Posit Software, PBC\", role = c(\"cph\", \"fnd\")), person(\"R Foundation\", role = \"ctb\", comment = \"Copy of R-project homepage cached as example\") )", + "Description": "Bindings to 'libxml2' for working with XML data using a simple, consistent interface based on 'XPath' expressions. Also supports XML schema validation; for 'XSLT' transformations see the 'xslt' package.", + "License": "MIT + file LICENSE", + "URL": "https://xml2.r-lib.org, https://r-lib.r-universe.dev/xml2", + "BugReports": "https://github.com/r-lib/xml2/issues", + "Depends": [ + "R (>= 3.6.0)" + ], + "Imports": [ "cli", "methods", - "rlang" + "rlang (>= 1.1.0)" ], - "Hash": "1d0336142f4cd25d8d23cd3ba7a8fb61" + "Suggests": [ + "covr", + "curl", + "httr", + "knitr", + "mockery", + "rmarkdown", + "testthat (>= 3.2.0)", + "xslt" + ], + "VignetteBuilder": "knitr", + "Config/Needs/website": "tidyverse/tidytemplate", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.3", + "SystemRequirements": "libxml2: libxml2-dev (deb), libxml2-devel (rpm)", + "Collate": "'S4.R' 'as_list.R' 'xml_parse.R' 'as_xml_document.R' 'classes.R' 'format.R' 'import-standalone-obj-type.R' 'import-standalone-purrr.R' 'import-standalone-types-check.R' 'init.R' 'nodeset_apply.R' 'paths.R' 'utils.R' 'xml2-package.R' 'xml_attr.R' 'xml_children.R' 'xml_document.R' 'xml_find.R' 'xml_missing.R' 'xml_modify.R' 'xml_name.R' 'xml_namespaces.R' 'xml_node.R' 'xml_nodeset.R' 'xml_path.R' 'xml_schema.R' 'xml_serialize.R' 'xml_structure.R' 'xml_text.R' 'xml_type.R' 'xml_url.R' 'xml_write.R' 'zzz.R'", + "Config/testthat/edition": "3", + "NeedsCompilation": "yes", + "Author": "Hadley Wickham [aut], Jim Hester [aut], Jeroen Ooms [aut, cre], Posit Software, PBC [cph, fnd], R Foundation [ctb] (Copy of R-project homepage cached as example)", + "Maintainer": "Jeroen Ooms ", + "Repository": "CRAN" }, "xtable": { "Package": "xtable", "Version": "1.8-4", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ - "R", + "Date": "2019-04-08", + "Title": "Export Tables to LaTeX or HTML", + "Authors@R": "c(person(\"David B.\", \"Dahl\", role=\"aut\"), person(\"David\", \"Scott\", role=c(\"aut\",\"cre\"), email=\"d.scott@auckland.ac.nz\"), person(\"Charles\", \"Roosen\", role=\"aut\"), person(\"Arni\", \"Magnusson\", role=\"aut\"), person(\"Jonathan\", \"Swinton\", role=\"aut\"), person(\"Ajay\", \"Shah\", role=\"ctb\"), person(\"Arne\", \"Henningsen\", role=\"ctb\"), person(\"Benno\", \"Puetz\", role=\"ctb\"), person(\"Bernhard\", \"Pfaff\", role=\"ctb\"), person(\"Claudio\", \"Agostinelli\", role=\"ctb\"), person(\"Claudius\", \"Loehnert\", role=\"ctb\"), person(\"David\", \"Mitchell\", role=\"ctb\"), person(\"David\", \"Whiting\", role=\"ctb\"), person(\"Fernando da\", \"Rosa\", role=\"ctb\"), person(\"Guido\", \"Gay\", role=\"ctb\"), person(\"Guido\", \"Schulz\", role=\"ctb\"), person(\"Ian\", \"Fellows\", role=\"ctb\"), person(\"Jeff\", \"Laake\", role=\"ctb\"), person(\"John\", \"Walker\", role=\"ctb\"), person(\"Jun\", \"Yan\", role=\"ctb\"), person(\"Liviu\", \"Andronic\", role=\"ctb\"), person(\"Markus\", \"Loecher\", role=\"ctb\"), person(\"Martin\", \"Gubri\", role=\"ctb\"), person(\"Matthieu\", \"Stigler\", role=\"ctb\"), person(\"Robert\", \"Castelo\", role=\"ctb\"), person(\"Seth\", \"Falcon\", role=\"ctb\"), person(\"Stefan\", \"Edwards\", role=\"ctb\"), person(\"Sven\", \"Garbade\", role=\"ctb\"), person(\"Uwe\", \"Ligges\", role=\"ctb\"))", + "Maintainer": "David Scott ", + "Imports": [ "stats", "utils" ], - "Hash": "b8acdf8af494d9ec19ccb2481a9b11c2" + "Suggests": [ + "knitr", + "plm", + "zoo", + "survival" + ], + "VignetteBuilder": "knitr", + "Description": "Coerce data to LaTeX and HTML tables.", + "URL": "http://xtable.r-forge.r-project.org/", + "Depends": [ + "R (>= 2.10.0)" + ], + "License": "GPL (>= 2)", + "Repository": "CRAN", + "NeedsCompilation": "no", + "Author": "David B. Dahl [aut], David Scott [aut, cre], Charles Roosen [aut], Arni Magnusson [aut], Jonathan Swinton [aut], Ajay Shah [ctb], Arne Henningsen [ctb], Benno Puetz [ctb], Bernhard Pfaff [ctb], Claudio Agostinelli [ctb], Claudius Loehnert [ctb], David Mitchell [ctb], David Whiting [ctb], Fernando da Rosa [ctb], Guido Gay [ctb], Guido Schulz [ctb], Ian Fellows [ctb], Jeff Laake [ctb], John Walker [ctb], Jun Yan [ctb], Liviu Andronic [ctb], Markus Loecher [ctb], Martin Gubri [ctb], Matthieu Stigler [ctb], Robert Castelo [ctb], Seth Falcon [ctb], Stefan Edwards [ctb], Sven Garbade [ctb], Uwe Ligges [ctb]", + "Encoding": "UTF-8" }, "yaml": { "Package": "yaml", - "Version": "2.3.8", - "Source": "Repository", - "Repository": "CRAN", - "Hash": "29240487a071f535f5e5d5a323b7afbd" + "Version": "2.3.12", + "Source": "Repository", + "Type": "Package", + "Title": "Methods to Convert R Data to YAML and Back", + "Authors@R": "c( person(\"Hadley\", \"Wickham\", , \"hadley@posit.co\", role = \"cre\", comment = c(ORCID = \"0000-0003-4757-117X\")), person(\"Shawn\", \"Garbett\", , \"shawn.garbett@vumc.org\", role = \"ctb\", comment = c(ORCID = \"0000-0003-4079-5621\")), person(\"Jeremy\", \"Stephens\", role = c(\"aut\", \"ctb\")), person(\"Kirill\", \"Simonov\", role = \"aut\"), person(\"Yihui\", \"Xie\", role = \"ctb\", comment = c(ORCID = \"0000-0003-0645-5666\")), person(\"Zhuoer\", \"Dong\", role = \"ctb\"), person(\"Jeffrey\", \"Horner\", role = \"ctb\"), person(\"reikoch\", role = \"ctb\"), person(\"Will\", \"Beasley\", role = \"ctb\", comment = c(ORCID = \"0000-0002-5613-5006\")), person(\"Brendan\", \"O'Connor\", role = \"ctb\"), person(\"Michael\", \"Quinn\", role = \"ctb\"), person(\"Charlie\", \"Gao\", role = \"ctb\"), person(c(\"Gregory\", \"R.\"), \"Warnes\", role = \"ctb\"), person(c(\"Zhian\", \"N.\"), \"Kamvar\", role = \"ctb\") )", + "Description": "Implements the 'libyaml' 'YAML' 1.1 parser and emitter () for R.", + "License": "BSD_3_clause + file LICENSE", + "URL": "https://yaml.r-lib.org, https://github.com/r-lib/yaml/", + "BugReports": "https://github.com/r-lib/yaml/issues", + "Suggests": [ + "knitr", + "rmarkdown", + "testthat (>= 3.0.0)" + ], + "Config/testthat/edition": "3", + "Config/Needs/website": "tidyverse/tidytemplate", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.3", + "VignetteBuilder": "knitr", + "NeedsCompilation": "yes", + "Author": "Hadley Wickham [cre] (ORCID: ), Shawn Garbett [ctb] (ORCID: ), Jeremy Stephens [aut, ctb], Kirill Simonov [aut], Yihui Xie [ctb] (ORCID: ), Zhuoer Dong [ctb], Jeffrey Horner [ctb], reikoch [ctb], Will Beasley [ctb] (ORCID: ), Brendan O'Connor [ctb], Michael Quinn [ctb], Charlie Gao [ctb], Gregory R. Warnes [ctb], Zhian N. Kamvar [ctb]", + "Maintainer": "Hadley Wickham ", + "Repository": "CRAN" }, "yulab.utils": { "Package": "yulab.utils", - "Version": "0.1.4", + "Version": "0.2.4", "Source": "Repository", - "Repository": "CRAN", - "Requirements": [ + "Title": "Supporting Functions for Packages Maintained by 'YuLab-SMU'", + "Authors@R": "c(person(\"Guangchuang\", \"Yu\", email = \"guangchuangyu@gmail.com\", role = c(\"aut\", \"cre\"), comment = c(ORCID = \"0000-0002-6485-8781\")))", + "Description": "Miscellaneous functions commonly used by 'YuLab-SMU'.", + "Depends": [ + "R (>= 4.2.0)" + ], + "Imports": [ "cli", "digest", "fs", - "memoise", + "methods", + "rappdirs", "rlang", - "stats", "tools", "utils" ], - "Hash": "60ee2aaa179dc282e9fa7367bad76e89" - }, - "zlibbioc": { - "Package": "zlibbioc", - "Version": "1.50.0", - "Source": "Bioconductor", - "Repository": "Bioconductor 3.19", - "Hash": "3db02e3c460e1c852365df117a2b441b" + "Suggests": [ + "httr2", + "jsonlite", + "openssl", + "R.utils", + "testthat (>= 3.0.0)" + ], + "ByteCompile": "true", + "License": "Artistic-2.0", + "URL": "https://yulab-smu.top/", + "BugReports": "https://github.com/YuLab-SMU/yulab.utils/issues", + "Encoding": "UTF-8", + "RoxygenNote": "7.3.3", + "Config/testthat/edition": "3", + "NeedsCompilation": "no", + "Author": "Guangchuang Yu [aut, cre] (ORCID: )", + "Maintainer": "Guangchuang Yu ", + "Repository": "CRAN" } } } diff --git a/renv/.gitignore b/renv/.gitignore index 7c1c770d..0ec0cbba 100644 --- a/renv/.gitignore +++ b/renv/.gitignore @@ -1,8 +1,7 @@ -sandbox/ -cellar/ -local/ library/ +local/ +cellar/ lock/ python/ +sandbox/ staging/ - diff --git a/renv/activate.R b/renv/activate.R index 0eb51088..ef25ef83 100644 --- a/renv/activate.R +++ b/renv/activate.R @@ -2,7 +2,8 @@ local({ # the requested version of renv - version <- "1.0.11" + version <- "1.1.7" + attr(version, "md5") <- "dd5d60f155dadff4c88c2fc6680504b4" attr(version, "sha") <- NULL # the project directory @@ -42,7 +43,7 @@ local({ return(FALSE) # next, check environment variables - # TODO: prefer using the configuration one in the future + # prefer using the configuration one in the future envvars <- c( "RENV_CONFIG_AUTOLOADER_ENABLED", "RENV_AUTOLOADER_ENABLED", @@ -135,12 +136,12 @@ local({ # R help links pattern <- "`\\?(renv::(?:[^`])+)`" - replacement <- "`\033]8;;ide:help:\\1\a?\\1\033]8;;\a`" + replacement <- "`\033]8;;x-r-help:\\1\a?\\1\033]8;;\a`" text <- gsub(pattern, replacement, text, perl = TRUE) # runnable code pattern <- "`(renv::(?:[^`])+)`" - replacement <- "`\033]8;;ide:run:\\1\a\\1\033]8;;\a`" + replacement <- "`\033]8;;x-r-run:\\1\a\\1\033]8;;\a`" text <- gsub(pattern, replacement, text, perl = TRUE) # return ansified text @@ -168,6 +169,16 @@ local({ if (quiet) return(invisible()) + # also check for config environment variables that should suppress messages + # https://github.com/rstudio/renv/issues/2214 + enabled <- Sys.getenv("RENV_CONFIG_STARTUP_QUIET", unset = NA) + if (!is.na(enabled) && tolower(enabled) %in% c("true", "1")) + return(invisible()) + + enabled <- Sys.getenv("RENV_CONFIG_SYNCHRONIZED_CHECK", unset = NA) + if (!is.na(enabled) && tolower(enabled) %in% c("false", "0")) + return(invisible()) + msg <- sprintf(fmt, ...) cat(msg, file = stdout(), sep = if (appendLF) "\n" else "") @@ -209,16 +220,22 @@ local({ } - startswith <- function(string, prefix) { - substring(string, 1, nchar(prefix)) == prefix - } - bootstrap <- function(version, library) { friendly <- renv_bootstrap_version_friendly(version) section <- header(sprintf("Bootstrapping renv %s", friendly)) catf(section) + # try to install renv from cache + md5 <- attr(version, "md5", exact = TRUE) + if (length(md5)) { + pkgpath <- renv_bootstrap_find(version) + if (length(pkgpath) && file.exists(pkgpath)) { + file.copy(pkgpath, library, recursive = TRUE) + return(invisible()) + } + } + # attempt to download renv catf("- Downloading renv ... ", appendLF = FALSE) withCallingHandlers( @@ -244,7 +261,6 @@ local({ # add empty line to break up bootstrapping from normal output catf("") - return(invisible()) } @@ -261,12 +277,20 @@ local({ repos <- Sys.getenv("RENV_CONFIG_REPOS_OVERRIDE", unset = NA) if (!is.na(repos)) { - # check for RSPM; if set, use a fallback repository for renv - rspm <- Sys.getenv("RSPM", unset = NA) - if (identical(rspm, repos)) - repos <- c(RSPM = rspm, CRAN = cran) + # split on ';' if present + parts <- strsplit(repos, ";", fixed = TRUE)[[1L]] - return(repos) + # split into named repositories if present + idx <- regexpr("=", parts, fixed = TRUE) + keys <- substring(parts, 1L, idx - 1L) + vals <- substring(parts, idx + 1L) + names(vals) <- keys + + # if we have a single unnamed repository, call it CRAN + if (length(vals) == 1L && identical(keys, "")) + names(vals) <- "CRAN" + + return(vals) } @@ -515,6 +539,51 @@ local({ } + renv_bootstrap_find <- function(version) { + + path <- renv_bootstrap_find_cache(version) + if (length(path) && file.exists(path)) { + catf("- Using renv %s from global package cache", version) + return(path) + } + + } + + renv_bootstrap_find_cache <- function(version) { + + md5 <- attr(version, "md5", exact = TRUE) + if (is.null(md5)) + return() + + # infer path to renv cache + cache <- Sys.getenv("RENV_PATHS_CACHE", unset = "") + if (!nzchar(cache)) { + root <- Sys.getenv("RENV_PATHS_ROOT", unset = NA) + if (!is.na(root)) + cache <- file.path(root, "cache") + } + + if (!nzchar(cache)) { + tools <- asNamespace("tools") + if (is.function(tools$R_user_dir)) { + root <- tools$R_user_dir("renv", "cache") + cache <- file.path(root, "cache") + } + } + + # start completing path to cache + file.path( + cache, + renv_bootstrap_cache_version(), + renv_bootstrap_platform_prefix(), + "renv", + version, + md5, + "renv" + ) + + } + renv_bootstrap_download_tarball <- function(version) { # if the user has provided the path to a tarball via @@ -563,6 +632,9 @@ local({ # prepare download options token <- renv_bootstrap_github_token() + if (is.null(token)) + token <- "" + if (nzchar(Sys.which("curl")) && nzchar(token)) { fmt <- "--location --fail --header \"Authorization: token %s\"" extra <- sprintf(fmt, token) @@ -696,11 +768,19 @@ local({ } - renv_bootstrap_platform_prefix <- function() { + renv_bootstrap_platform_prefix_default <- function() { - # construct version prefix - version <- paste(R.version$major, R.version$minor, sep = ".") - prefix <- paste("R", numeric_version(version)[1, 1:2], sep = "-") + # read version component + version <- Sys.getenv("RENV_PATHS_VERSION", unset = "R-%v") + + # expand placeholders + placeholders <- list( + list("%v", format(getRversion()[1, 1:2])), + list("%V", format(getRversion()[1, 1:3])) + ) + + for (placeholder in placeholders) + version <- gsub(placeholder[[1L]], placeholder[[2L]], version, fixed = TRUE) # include SVN revision for development versions of R # (to avoid sharing platform-specific artefacts with released versions of R) @@ -709,10 +789,19 @@ local({ identical(R.version[["nickname"]], "Unsuffered Consequences") if (devel) - prefix <- paste(prefix, R.version[["svn rev"]], sep = "-r") + version <- paste(version, R.version[["svn rev"]], sep = "-r") + + version + + } + + renv_bootstrap_platform_prefix <- function() { + + # construct version prefix + version <- renv_bootstrap_platform_prefix_default() # build list of path components - components <- c(prefix, R.version$platform) + components <- c(version, R.version$platform) # include prefix if provided by user prefix <- renv_bootstrap_platform_prefix_impl() @@ -951,13 +1040,19 @@ local({ } renv_bootstrap_validate_version_dev <- function(version, description) { + expected <- description[["RemoteSha"]] - is.character(expected) && startswith(expected, version) + if (!is.character(expected)) + return(FALSE) + + pattern <- sprintf("^\\Q%s\\E", version) + grepl(pattern, expected, perl = TRUE) + } renv_bootstrap_validate_version_release <- function(version, description) { expected <- description[["Version"]] - is.character(expected) && identical(expected, version) + is.character(expected) && identical(c(expected), c(version)) } renv_bootstrap_hash_text <- function(text) { @@ -1132,10 +1227,10 @@ local({ renv_bootstrap_exec <- function(project, libpath, version) { if (!renv_bootstrap_load(project, libpath, version)) - renv_bootstrap_run(version, libpath) + renv_bootstrap_run(project, libpath, version) } - renv_bootstrap_run <- function(version, libpath) { + renv_bootstrap_run <- function(project, libpath, version) { # perform bootstrap bootstrap(version, libpath) @@ -1146,7 +1241,7 @@ local({ # try again to load if (requireNamespace("renv", lib.loc = libpath, quietly = TRUE)) { - return(renv::load(project = getwd())) + return(renv::load(project = project)) } # failed to download or load renv; warn the user @@ -1159,6 +1254,18 @@ local({ } + renv_bootstrap_cache_version <- function() { + # NOTE: users should normally not override the cache version; + # this is provided just to make testing easier + Sys.getenv("RENV_CACHE_VERSION", unset = "v5") + } + + renv_bootstrap_cache_version_previous <- function() { + version <- renv_bootstrap_cache_version() + number <- as.integer(substring(version, 2L)) + paste("v", number - 1L, sep = "") + } + renv_json_read <- function(file = NULL, text = NULL) { jlerr <- NULL @@ -1192,98 +1299,105 @@ local({ jsonlite::fromJSON(txt = text, simplifyVector = FALSE) } - renv_json_read_default <- function(file = NULL, text = NULL) { + renv_json_read_patterns <- function() { - # find strings in the JSON - text <- paste(text %||% readLines(file, warn = FALSE), collapse = "\n") - pattern <- '["](?:(?:\\\\.)|(?:[^"\\\\]))*?["]' - locs <- gregexpr(pattern, text, perl = TRUE)[[1]] + list( - # if any are found, replace them with placeholders - replaced <- text - strings <- character() - replacements <- character() + # objects + list("{", "\t\n\tobject(\t\n\t", TRUE), + list("}", "\t\n\t)\t\n\t", TRUE), - if (!identical(c(locs), -1L)) { + # arrays + list("[", "\t\n\tarray(\t\n\t", TRUE), + list("]", "\n\t\n)\n\t\n", TRUE), - # get the string values - starts <- locs - ends <- locs + attr(locs, "match.length") - 1L - strings <- substring(text, starts, ends) + # maps + list(":", "\t\n\t=\t\n\t", TRUE), - # only keep those requiring escaping - strings <- grep("[[\\]{}:]", strings, perl = TRUE, value = TRUE) + # newlines + list("\\u000a", "\n", FALSE) - # compute replacements - replacements <- sprintf('"\032%i\032"', seq_along(strings)) + ) - # replace the strings - mapply(function(string, replacement) { - replaced <<- sub(string, replacement, replaced, fixed = TRUE) - }, strings, replacements) + } - } + renv_json_read_envir <- function() { - # transform the JSON into something the R parser understands - transformed <- replaced - transformed <- gsub("{}", "`names<-`(list(), character())", transformed, fixed = TRUE) - transformed <- gsub("[[{]", "list(", transformed, perl = TRUE) - transformed <- gsub("[]}]", ")", transformed, perl = TRUE) - transformed <- gsub(":", "=", transformed, fixed = TRUE) - text <- paste(transformed, collapse = "\n") + envir <- new.env(parent = emptyenv()) - # parse it - json <- parse(text = text, keep.source = FALSE, srcfile = NULL)[[1L]] + envir[["+"]] <- `+` + envir[["-"]] <- `-` - # construct map between source strings, replaced strings - map <- as.character(parse(text = strings)) - names(map) <- as.character(parse(text = replacements)) + envir[["object"]] <- function(...) { + result <- list(...) + names(result) <- as.character(names(result)) + result + } - # convert to list - map <- as.list(map) + envir[["array"]] <- list - # remap strings in object - remapped <- renv_json_read_remap(json, map) + envir[["true"]] <- TRUE + envir[["false"]] <- FALSE + envir[["null"]] <- NULL - # evaluate - eval(remapped, envir = baseenv()) + envir } - renv_json_read_remap <- function(json, map) { + renv_json_read_remap <- function(object, patterns) { - # fix names - if (!is.null(names(json))) { - lhs <- match(names(json), names(map), nomatch = 0L) - rhs <- match(names(map), names(json), nomatch = 0L) - names(json)[rhs] <- map[lhs] - } + # repair names if necessary + if (!is.null(names(object))) { + + nms <- names(object) + for (pattern in patterns) + nms <- gsub(pattern[[2L]], pattern[[1L]], nms, fixed = TRUE) + names(object) <- nms - # fix values - if (is.character(json)) - return(map[[json]] %||% json) - - # handle true, false, null - if (is.name(json)) { - text <- as.character(json) - if (text == "true") - return(TRUE) - else if (text == "false") - return(FALSE) - else if (text == "null") - return(NULL) } - # recurse - if (is.recursive(json)) { - for (i in seq_along(json)) { - json[i] <- list(renv_json_read_remap(json[[i]], map)) - } + # repair strings if necessary + if (is.character(object)) { + for (pattern in patterns) + object <- gsub(pattern[[2L]], pattern[[1L]], object, fixed = TRUE) } - json + # recurse for other objects + if (is.recursive(object)) + for (i in seq_along(object)) + object[i] <- list(renv_json_read_remap(object[[i]], patterns)) + + # return remapped object + object + + } + + renv_json_read_default <- function(file = NULL, text = NULL) { + + # read json text + text <- paste(text %||% readLines(file, warn = FALSE), collapse = "\n") + + # convert into something the R parser will understand + patterns <- renv_json_read_patterns() + transformed <- text + for (pattern in patterns) + transformed <- gsub(pattern[[1L]], pattern[[2L]], transformed, fixed = TRUE) + + # parse it + rfile <- tempfile("renv-json-", fileext = ".R") + on.exit(unlink(rfile), add = TRUE) + writeLines(transformed, con = rfile) + json <- parse(rfile, keep.source = FALSE, srcfile = NULL)[[1L]] + + # evaluate in safe environment + result <- eval(json, envir = renv_json_read_envir()) + + # fix up strings if necessary -- do so only with reversible patterns + patterns <- Filter(function(pattern) pattern[[3L]], patterns) + renv_json_read_remap(result, patterns) } + # load the renv profile, if any renv_bootstrap_profile_load(project) diff --git a/renv/settings.json b/renv/settings.json index 0470d923..9566b6ff 100644 --- a/renv/settings.json +++ b/renv/settings.json @@ -1,5 +1,5 @@ { - "bioconductor.version": "3.19", + "bioconductor.version": "3.22", "external.libraries": [], "ignored.packages": [], "package.dependency.fields": [ @@ -9,7 +9,8 @@ ], "ppm.enabled": true, "ppm.ignored.urls": [], - "r.version": "4.4.0", + "r.version": "4.5.2", + "snapshot.dev": false, "snapshot.type": "implicit", "use.cache": true, "vcs.ignore.cellar": true, diff --git a/scRNA-seq-advanced/01-read_filter_normalize_scRNA-live.Rmd b/scRNA-seq-advanced/01-read_filter_normalize_scRNA-live.Rmd index 18140bb7..cf5c9cb3 100644 --- a/scRNA-seq-advanced/01-read_filter_normalize_scRNA-live.Rmd +++ b/scRNA-seq-advanced/01-read_filter_normalize_scRNA-live.Rmd @@ -93,8 +93,10 @@ normalized_dir <- file.path(data_dir, "normalized") fs::dir_create(normalized_dir) # output RDS file for normalized data -output_sce_file <- file.path(normalized_dir, - "glioblastoma_normalized_sce.rds") +output_sce_file <- file.path( + normalized_dir, + "glioblastoma_normalized_sce.rds" +) ``` @@ -106,7 +108,7 @@ Whether the 10x Cell Ranger data is in Matrix Exchange format or in an HDF5 file (Though again, we do not recommend using the `.h5` file if you can avoid it, _especially_ for raw (unfiltered) data.) If you used something other than Cell Ranger to process the raw data, you would need to use a different function to read it in and create the `SingleCellExperiment` object. -Some of these functions for other common data formats are discussed in [Chapter 3 of OSCA] (http://bioconductor.org/books/3.19/OSCA.intro/getting-scrna-seq-datasets.html#reading-counts-into-r). +Some of these functions for other common data formats are discussed in [Chapter 3 of OSCA](http://bioconductor.org/books/3.19/OSCA.intro/getting-scrna-seq-datasets.html#reading-counts-into-r). ```{r read SCE, live=TRUE} # read SCE from matrix directory @@ -327,8 +329,11 @@ We stick with that default, but for clarity, we will also include it in our code ```{r miQC plotFiltering} # look at miQC filtering -miQC::plotFiltering(filtered_sce, miqc_model, - posterior_cutoff = 0.75) + +miQC::plotFiltering( + filtered_sce, + miqc_model, + posterior_cutoff = 0.75 +) + theme_bw() ``` @@ -409,8 +414,10 @@ num_genes <- 2000 gene_variance <- scran::modelGeneVar(normalized_sce) # get the most variable genes -hv_genes <- scran::getTopHVGs(gene_variance, - n = num_genes) +hv_genes <- scran::getTopHVGs( + gene_variance, + n = num_genes +) ``` The result is a vector of gene ids (ordered from most to least variable): @@ -476,7 +483,7 @@ We can also visualize the UMAP results using the `plotReducedDim()` function. ```{r plotReducedDim, live=TRUE} # plot the UMAP - # color by the most variable gene + # color by the most variable gene ``` diff --git a/scRNA-seq-advanced/01-read_filter_normalize_scRNA.Rmd b/scRNA-seq-advanced/01-read_filter_normalize_scRNA.Rmd index bb7c0175..23991c70 100644 --- a/scRNA-seq-advanced/01-read_filter_normalize_scRNA.Rmd +++ b/scRNA-seq-advanced/01-read_filter_normalize_scRNA.Rmd @@ -96,8 +96,10 @@ normalized_dir <- file.path(data_dir, "normalized") fs::dir_create(normalized_dir) # output RDS file for normalized data -output_sce_file <- file.path(normalized_dir, - "glioblastoma_normalized_sce.rds") +output_sce_file <- file.path( + normalized_dir, + "glioblastoma_normalized_sce.rds" +) ``` @@ -109,7 +111,7 @@ Whether the 10x Cell Ranger data is in Matrix Exchange format or in an HDF5 file (Though again, we do not recommend using the `.h5` file if you can avoid it, _especially_ for raw (unfiltered) data.) If you used something other than Cell Ranger to process the raw data, you would need to use a different function to read it in and create the `SingleCellExperiment` object. -Some of these functions for other common data formats are discussed in [Chapter 3 of OSCA] (http://bioconductor.org/books/3.19/OSCA.intro/getting-scrna-seq-datasets.html#reading-counts-into-r). +Some of these functions for other common data formats are discussed in [Chapter 3 of OSCA](http://bioconductor.org/books/3.19/OSCA.intro/getting-scrna-seq-datasets.html#reading-counts-into-r). ```{r read SCE, live=TRUE} # read SCE from matrix directory @@ -273,8 +275,10 @@ The `mito` name is important in that it is the name that will be expected by a l (We could define more subsets, but for now this one will do.) ```{r per cell QC, live=TRUE} -filtered_sce <- scuttle::addPerCellQC(filtered_sce, - subsets = list(mito = mito_genes)) +filtered_sce <- scuttle::addPerCellQC( + filtered_sce, + subsets = list(mito = mito_genes) +) ``` Now we can look at the colData to see what was added: @@ -337,8 +341,11 @@ We stick with that default, but for clarity, we will also include it in our code ```{r miQC plotFiltering} # look at miQC filtering -miQC::plotFiltering(filtered_sce, miqc_model, - posterior_cutoff = 0.75) + +miQC::plotFiltering( + filtered_sce, + miqc_model, + posterior_cutoff = 0.75 +) + theme_bw() ``` @@ -349,8 +356,10 @@ At this point, we can perform the actual filtering using the `filterCells()` fun ```{r miQC filtercells, live=TRUE} # perform miQC filtering -qcfiltered_sce <- miQC::filterCells(filtered_sce, - model = miqc_model) +qcfiltered_sce <- miQC::filterCells( + filtered_sce, + model = miqc_model +) ``` #### One more filter: unique gene count @@ -383,8 +392,10 @@ Finally, we apply the scaling factor to the expression values for each cell and qclust <- scran::quickCluster(qcfiltered_sce) # use clusters to compute scaling factors and add to SCE object -qcfiltered_sce <- scran::computeSumFactors(qcfiltered_sce, - clusters = qclust) +qcfiltered_sce <- scran::computeSumFactors( + qcfiltered_sce, + clusters = qclust +) # perform normalization using scaling factors # and save as a new SCE object @@ -424,8 +435,10 @@ num_genes <- 2000 gene_variance <- scran::modelGeneVar(normalized_sce) # get the most variable genes -hv_genes <- scran::getTopHVGs(gene_variance, - n = num_genes) +hv_genes <- scran::getTopHVGs( + gene_variance, + n = num_genes +) ``` The result is a vector of gene ids (ordered from most to least variable): @@ -486,8 +499,10 @@ Since the UMAP procedure would be slow to calculate with the full data, so the ` Since we already have a PCA matrix, we will tell the function use that instead of recalculating it. ```{r runUMAP, live=TRUE} -normalized_sce <- scater::runUMAP(normalized_sce, - dimred = "PCA") +normalized_sce <- scater::runUMAP( + normalized_sce, + dimred = "PCA" +) ``` As before, we could extract the UMAP matrix from our SCE object with the `reducedDim()` function. @@ -495,10 +510,12 @@ We can also visualize the UMAP results using the `plotReducedDim()` function. ```{r plotReducedDim, live=TRUE} # plot the UMAP -scater::plotReducedDim(normalized_sce, - "UMAP", - # color by the most variable gene - color_by = hv_genes[1]) +scater::plotReducedDim( + normalized_sce, + "UMAP", + # color by the most variable gene + color_by = hv_genes[1] +) ``` @@ -552,8 +569,10 @@ Here rather than the general `plotReducedDim()` function, we will use `plotUMAP( ```{r plot clusters, live=TRUE} # plot UMAP with assigned clusters -scater::plotUMAP(normalized_sce, - color_by = "nn_cluster") +scater::plotUMAP( + normalized_sce, + color_by = "nn_cluster" +) ``` What do you see in these results? diff --git a/scRNA-seq-advanced/01-read_filter_normalize_scRNA.nb.html b/scRNA-seq-advanced/01-read_filter_normalize_scRNA.nb.html index dd2bf48f..6f2dc05a 100644 --- a/scRNA-seq-advanced/01-read_filter_normalize_scRNA.nb.html +++ b/scRNA-seq-advanced/01-read_filter_normalize_scRNA.nb.html @@ -3107,7 +3107,7 @@

Directories and files

- +
# Outputs ------------------------------------
 
 # Directory and file to save output
@@ -3117,8 +3117,10 @@ 

Directories and files

fs::dir_create(normalized_dir) # output RDS file for normalized data -output_sce_file <- file.path(normalized_dir, - "glioblastoma_normalized_sce.rds")
+output_sce_file <- file.path( + normalized_dir, + "glioblastoma_normalized_sce.rds" +) @@ -3139,7 +3141,8 @@

Reading Cell Ranger data

If you used something other than Cell Ranger to process the raw data, you would need to use a different function to read it in and create the SingleCellExperiment object. Some of these functions for -other common data formats are discussed in [Chapter 3 of OSCA] (http://bioconductor.org/books/3.19/OSCA.intro/getting-scrna-seq-datasets.html#reading-counts-into-r).

+other common data formats are discussed in Chapter +3 of OSCA.

@@ -3490,9 +3493,11 @@

Calculating summary QC statistics

more subsets, but for now this one will do.)

- -
filtered_sce <- scuttle::addPerCellQC(filtered_sce,
-                                      subsets = list(mito = mito_genes))
+ +
filtered_sce <- scuttle::addPerCellQC(
+  filtered_sce,
+  subsets = list(mito = mito_genes)
+)
@@ -3616,10 +3621,13 @@

Filtering compromised cells

also include it in our code!

- +
# look at miQC filtering
-miQC::plotFiltering(filtered_sce, miqc_model,
-                    posterior_cutoff = 0.75) +
+miQC::plotFiltering(
+  filtered_sce,
+  miqc_model,
+  posterior_cutoff = 0.75
+) +
   theme_bw()
@@ -3637,10 +3645,12 @@

Filtering compromised cells

object.

- +
# perform miQC filtering
-qcfiltered_sce <- miQC::filterCells(filtered_sce,
-                                    model = miqc_model)
+qcfiltered_sce <- miQC::filterCells( + filtered_sce, + model = miqc_model +)
Removing 387 out of 1626 cells.
@@ -3705,13 +3715,15 @@

Normalization

scuttle::logNormCounts() function.

- +
# Perform rough clustering
 qclust <- scran::quickCluster(qcfiltered_sce)
 
 # use clusters to compute scaling factors and add to SCE object
-qcfiltered_sce <- scran::computeSumFactors(qcfiltered_sce,
-                                           clusters = qclust)
+qcfiltered_sce <- scran::computeSumFactors(
+  qcfiltered_sce,
+  clusters = qclust
+)
 
 # perform normalization using scaling factors
 # and save as a new SCE object
@@ -3777,7 +3789,7 @@ 

Selecting highly variable genes

variation, which is what we are most interested in.

- +
# identify 2000 genes
 num_genes <- 2000
 
@@ -3785,8 +3797,10 @@ 

Selecting highly variable genes

gene_variance <- scran::modelGeneVar(normalized_sce) # get the most variable genes -hv_genes <- scran::getTopHVGs(gene_variance, - n = num_genes)
+hv_genes <- scran::getTopHVGs( + gene_variance, + n = num_genes +)
@@ -3907,9 +3921,11 @@

UMAP

recalculating it.

- -
normalized_sce <- scater::runUMAP(normalized_sce,
-                                  dimred = "PCA")
+ +
normalized_sce <- scater::runUMAP(
+  normalized_sce,
+  dimred = "PCA"
+)
@@ -3918,12 +3934,14 @@

UMAP

results using the plotReducedDim() function.

- +
# plot the UMAP
-scater::plotReducedDim(normalized_sce,
-                       "UMAP",
-                       # color by the most variable gene
-                       color_by = hv_genes[1])
+scater::plotReducedDim( + normalized_sce, + "UMAP", + # color by the most variable gene + color_by = hv_genes[1] +)

@@ -4003,10 +4021,12 @@

Unsupervised clustering

we can skip that argument.

- +
# plot UMAP with assigned clusters
-scater::plotUMAP(normalized_sce,
-                 color_by = "nn_cluster")
+scater::plotUMAP( + normalized_sce, + color_by = "nn_cluster" +)

@@ -4141,7 +4161,7 @@

Print session info

-
---
title: "Reading, filtering, and normalizing scRNA-seq data"
author: Data Lab for ALSF
date: 2023
output:
  html_notebook:
    toc: true
    toc_float: true
---

## Objectives

This notebook will demonstrate how to:

- Read Cell Ranger data into R
- Filter to cells using `emptyDropsCellRanger()`
- Calculate quality control measures on scRNA-seq data
- Remove likely compromised cells with `miQC()`
- Normalize expression data across cells
- Calculate and plot reduced dimension representations of expression data (PCA, UMAP)

---

In this notebook, we will review basic processing for single-cell RNA-seq data, starting with the output from Cell Ranger, and proceeding through filtering, quality control, normalization, and dimension reduction. We will perform these tasks using tools from the [Bioconductor project](https://bioconductor.org), in particular [`SingleCellExperiment` objects](https://bioconductor.org/packages/release/bioc/html/SingleCellExperiment.html) and functions that work with those objects.
Much of the material in this notebook is directly inspired by, and draws heavily on, material presented in the book [_Orchestrating Single Cell Analysis with Bioconductor_ (OSCA)](http://bioconductor.org/books/release/OSCA/).

![Single-cell roadmap: Overview](diagrams/roadmap_single_overview.png)

The data we will use for this notebook is derived from a human glioblastoma specimen.
The sample was processed by 10x Genomics using a 3' RNA kit (v3.1), sequenced, and quantified with Cell Ranger 6.0.
Further details about the sample and processing can be found on the [10x website](https://www.10xgenomics.com/resources/datasets/2-k-sorted-cells-from-human-glioblastoma-multiforme-3-v-3-1-3-1-standard-6-0-0).


## Set Up

To start, we will load some of the libraries we will need later, and set a random number seed for reproducibility.

```{r setup}
# Load libraries

# Plotting functions
library(ggplot2)

# The main class we will use for Single Cell data
library(SingleCellExperiment)

# Setting the seed for reproducibility
set.seed(12345)
```

### Directories and files

Before we get too far, we like to define the input and output files that the notebook will use near the top of the document.
While you might not know the names of all of the files you will need or create output files when you start an analysis, we have found it helpful to keep all file and directory names in a single place near the top of the document.
This makes it easier for somebody coming to the code later to quickly see what files are needed as input and what will be produced as output.
More often than not, that somebody is you!

The gene expression data were processed to create a gene-by-cell expression matrix of counts for using Cell Ranger 6.0.
We have provided the raw data directory, `raw_feature_bc_matrix`, which is usually produced by Cell Ranger and placed in its `outs` directory.
This directory usually contains three files:
- `barcodes.tsv.gz`, a table of the cell barcodes that 10x uses, corresponding to the columns of the count matrix.
- `features.tsv.gz`, a table of the features (genes in this case) for which expression was quantified.
This will usually also include a bit of metadata about the features, including gene symbols (if the features are genes) and the type of data they represent (e.g., gene expression or antibody capture).
- `matrix.mtx.gz`, The counts themselves, stored in a sparse ["Matrix Exchange" format](https://math.nist.gov/MatrixMarket/formats.html).

Cell Ranger will also export these data in a single `HDF5` format file with a `.h5` extension, which can also be imported with the same commands we will use below.
However, we have found that processing large `.h5` files is often _much_ less efficient in R, so we prefer to start with the matrix files when possible.
In particular, we would not recommend working with `.h5` files for raw data; the filtering steps we will use below can sometimes take hours when using those files as input.

We will also need a table of mitochondrial genes, which we have stored in the `data/reference/` directory.

Finally, we will set up the our output directory, creating it if it does not yet exist, and define the name for the files we will save after all of our initial processing is complete.

```{r inputs, live=TRUE}
# Inputs --------------------------------------
# main data directory
data_dir <- file.path("data", "glioblastoma-10x")

# Path to the Cell Ranger matrix directory
raw_matrix_dir <- file.path(data_dir, "raw_feature_bc_matrix")

# reference data directory
ref_dir <- file.path("data", "reference")

# Path to mitochondrial genes table
mito_file <- file.path(ref_dir, "hs_mitochondrial_genes.tsv")
```

```{r outputs}
# Outputs ------------------------------------

# Directory and file to save output
normalized_dir <- file.path(data_dir, "normalized")

# create the directory if it does not exist
fs::dir_create(normalized_dir)

# output RDS file for normalized data
output_sce_file <- file.path(normalized_dir,
                             "glioblastoma_normalized_sce.rds")
```


## Reading Cell Ranger data

![Single-cell roadmap: Preprocess and Import](diagrams/roadmap_single_preprocess.png)

Whether the 10x Cell Ranger data is in Matrix Exchange format or in an HDF5 file, we can use the `read10xCounts()` function from the `DropletUtils` package to read the data into R and create a `SingleCellExperiment` object.
(Though again, we do not recommend using the `.h5` file if you can avoid it, _especially_ for raw (unfiltered) data.)

If you used something other than Cell Ranger to process the raw data, you would need to use a different function to read it in and create the `SingleCellExperiment` object.
Some of these functions for other common data formats are discussed in [Chapter 3 of OSCA] (http://bioconductor.org/books/3.19/OSCA.intro/getting-scrna-seq-datasets.html#reading-counts-into-r).

```{r read SCE, live=TRUE}
# read SCE from matrix directory
raw_sce <- DropletUtils::read10xCounts(
  raw_matrix_dir,
  col.names = TRUE # ensure barcodes are set as column names in the SCE object
)
```

Let's look at the contents of the object after reading it in:

```{r view SCE, live=TRUE}
# view SCE object
raw_sce
```

We can see from this summary that this `SingleCellExperiment` (SCE) object contains 36,601 rows, which correspond to the features (genes) that were analyzed, and 734,492 columns, which correspond to the possible barcode tags that were used in the experiment.
Note that not all of these barcode tags will have been used, and many of the features may never have been seen either.
One of our first steps will be to filter out barcodes that were never seen, or that may have only been seen in a droplet that did not contain a cell (an "empty droplet").

### Structure of the `SingleCellExperiment` object

In addition to the main `counts` matrix, listed as an `assay` in the SCE summary above, the SCE object can contain a number of other tables and matrices, each stored in a "slot" with a particular format.
The overall structure of the object can be seen in the figure below, which comes from an [OSCA Introduction chapter](http://bioconductor.org/books/3.19/OSCA.intro/the-singlecellexperiment-class.html).

![Structure of a SingleCellExperiment object](diagrams/SingleCellExperiment.png)

We have just mentioned the main `assay` slot, which contains full matrices of data (such as transcript counts) with each row a feature and each column a cell.
There are also a couple of tables for metadata, and a slot to store reduced-dimension representations (e.g., PCA and/or UMAP) of the expression data.

We'll start with the `rowData` slot, which is a table of metadata for each feature in our object.
For now that contains the contents of the `features.tsv.gz` file that we discussed earlier.
If we had read the data from something other than Cell Ranger output, we might have different contents, but each row would still correspond to a single feature of the SCE object.

Let's look at this table, extracting it from the SCE object with the `rowData()` function and using `head()` to view only the first 6 rows.

```{r rowdata}
# view rowData (features)
head(rowData(raw_sce))
```

You can see that this table includes an `ID` for each feature, which is usually the Ensembl gene ID, as well as the corresponding gene symbol in the `Symbol` column.
Finally there is a column for `Type`, which in this case is always "Gene Expression", as all of the features in this data set are genes.
If there were another modality of data that had been assayed in this experiment, there might be other values in this column, such as "Antibody Capture" for CITE-seq experiments.

The second slot is the `colData` table, which now corresponds to the `barcodes.tsv.gz` file, containing one row per cell barcode, or, more generally, one row per column of the `counts` assay.
We can look at this table using the `colData()` function (and `head()` again to prevent printing the whole table):

```{r coldata}
# view colData (cell barcodes)
head(colData(raw_sce))
```

Here we see that there are currently two columns:

- the `Sample` column has the path of the file that we read in (you may not see the whole path in this display); this should be identical in all rows from a single sample.
- the `Barcode` column contains the sequence that was used to identify each potential droplet for sequencing (and a numeric tag, in this case).
These will be unique within a sample.

As we proceed to calculate per-cell statistics, we will be adding new data to this table.

## Quality control and filtering

![Single-cell roadmap: QC, Filter, and Normalize](diagrams/roadmap_single_qc_norm.png)

### Filtering empty droplets

Most of the barcodes in any given 10x experiment will not be seen at all, so our first step can be to filter this raw data to only the cells where there is at least one transcript that was counted with that barcode.

To do this, we will use the `colSums()` function to quickly add up all the counts that correspond to each possible cell barcode, then filter our `raw_sce` down to just those columns where there are non-zero total counts.
We will need to extract the `counts` matrix from our SCE object, which we can do using the `counts()` function, conveniently enough.

```{r remove zeros, live=TRUE}
# sum columns from counts matrix
barcode_counts <- colSums(counts(raw_sce))

# filter SCE object to only rows with counts > 0
raw_sce <- raw_sce[, which(barcode_counts > 0)]
```

Now we can look at how our SCE object has changed:

```{r zero-filtered SCE}
raw_sce
```

But barcodes with zero counts are not the only ones that correspond to droplets without cells in them!
Even if a droplet does not have a cell in it, there will often be spurious reads from RNA sequences that were present in the extracellular solution, whether from the original sample or from cells that were damaged during single-cell library preparation.

We could identify these barcodes simply as those with low transcript counts.
Or, we can be a bit more clever!
We can look at the transcript counts _from_ the lowest-count droplets to create an expected distribution of transcripts in droplets that don't contain cells.
Then we can test each droplet to determine whether or not its transcript distribution deviates from that expectation.
If it does, then we have pretty good evidence that there _is_ a cell in there.

This test was first proposed by [Lun _et al._ (2019)](https://doi.org/10.1186/s13059-019-1662-y) and implemented as `emptyDrops()` in the `DropletUtils` package.
This method was then adopted, with some modifications, as the default cell filtering method used by Cell Ranger.
Here we will use the [`emptyDropsCellRanger()` function](https://rdrr.io/github/MarioniLab/DropletUtils/man/emptyDropsCellRanger.html) to perform filtering that more closely matches the Cell Ranger implementation.


```{r calculate droplet stats, live=TRUE}
# create a table of statistics using emptyDropsCellRanger
droplet_df <- DropletUtils::emptyDropsCellRanger(raw_sce)
```

Most values in this table are `NA`, because individual statistics were not calculated for the low-count droplets that were used to generate the background distribution.
(Most droplets don't have cells, so this makes some sense!)

We can look at just the rows without `NA` values by selected the ones where the FDR (which we will use again soon), is not `NA`.

```{r droplet stats}
# view rows where FDR is not `NA`
droplet_df[!is.na(droplet_df$FDR), ]
```
You will notice that some cells with high counts also have `NA` values for many statistics.
In those cases, `NA` values are actually present _because_ of the high counts - `emptyDropsCellRanger()` automatically assumed cells were present, so they were also not tested.

Now we can filter our `raw_sce` object _by column_ to only keep the cells with a small FDR: those that are quite unlikely to be empty droplets.

```{r filter emptydrops, live=TRUE}
# filter droplets using `which` to prevent NA trouble
cells_to_retain <- which(droplet_df$FDR < 0.01)
filtered_sce <- raw_sce[, cells_to_retain]
```

How many cells do we have now?

```{r filtered summary}
filtered_sce
```

### Additional quality control

In addition to filtering out empty droplets, we also will want to filter out cells that may have been damaged during library preparation.
These will often be characterized by a high proportion of mitochondrial transcripts and a smaller overall number of unique transcripts.
When a cell ruptures, cytoplasmic transcripts will leak out, but mitochondrial transcripts, still protected by the mitochondrial membrane, may remain.
As a consequence, there will be an over-abundance of mitochondrial reads, and fewer unique transcripts expressed.

Our first step then, is create a vector of the mitochondrial genes that are present in our dataset.
The mitochondrial file we defined during setup (`mito_file`) is a TSV file containing all of the human mitochondrial genes with additional annotation information for each gene, such as the gene location and alternative names.
(For more detail on the steps we took to create this file, you can look at [one of our setup notebooks](https://github.com/AlexsLemonade/training-modules/blob/master/scRNA-seq-advanced/setup/mito_gene_lists.Rmd))

All we need now is the `gene_id`, and only for the genes that are present in our SCE, so we will do some filtering with `dplyr` to pull out a vector with just those ids.

```{r get mitochondrial genes}
# read in a table of mitochondrial genes and extract ids
mito_genes <- readr::read_tsv(mito_file) |>
  # filter to only the genes that are found in our dataset
  dplyr::filter(gene_id %in% rownames(filtered_sce)) |>
  # create a vector from the gene_id column
  dplyr::pull(gene_id)
```

### Calculating summary QC statistics

We can now use the `scuttle` function `addPerCellQC()` to calculate some statistics based on the counts matrix, which will be added to the `colData` table.

In addition to calculating statistics like the total read count for each cell and the number of transcripts that are detected, we can also calculate those statistics for defined subsets of genes.
In this case, we will use our `mito_genes` vector to define a subset called `mito`.
The `mito` name is important in that it is the name that will be expected by a later function.
(We could define more subsets, but for now this one will do.)

```{r per cell QC, live=TRUE}
filtered_sce <- scuttle::addPerCellQC(filtered_sce,
                                      subsets = list(mito = mito_genes))
```

Now we can look at the colData to see what was added:

```{r view colData stats}
head(colData(filtered_sce))
```

We can also plot some of these statistics, here using the `plotMetrics()` function from the `miQC` package to plot the percent of reads that are mitochondrial (the `subsets_mito_percent` column) against the number of unique genes detected (the `detected` column) for each cell.

```{r miQC plotMetrics}
# use miQC::plotMetrics()
miQC::plotMetrics(filtered_sce) + theme_bw()
```

We can see that there is a range of mitochondrial percentages, and it does also seem that cells with high percentages of mitochondrial genes don't seem to contain very many unique genes.

How do we filter with this information?
One option is to define a cutoff for the mitochondrial percentage above which we call a cell compromised and exclude it from further analysis.
However, choosing that cutoff can be a bit fraught, as the expected percentage of mitochondrial reads can vary depending on the cell type and library preparation methods.
So it might be nice to have a method to determine that cutoff from the data itself.

### Filtering compromised cells

Determining mitochondrial cutoffs is exactly what the `miQC` package does ([Hippen _et al._ 2021](https://doi.org/10.1371/journal.pcbi.1009290))!
In truth, it does something possibly even a bit better: it fits a mixture model to the data that consists of distributions of healthy cells and compromised cells.
Then we can calculate whether each cell is more likely to belong to the healthy or compromised distribution.
We can then exclude the cells that are more likely to be compromised.

To use `miQC`, we first fit a model to the data in our SCE object:

```{r miQC model, live=TRUE}
# fit the miQC model
miqc_model <- miQC::mixtureModel(filtered_sce)
```

Now we can plot the model results using the `plotModel()` function to see how it corresponds to our data.
We should expect to see two fit lines:

- One line will correspond the the "healthy" cells and should show little to no relationship between the number of unique genes detected and the mitochondrial percentage.
- By contrast, the line that corresponds to "compromised" cells will show a negative relationship between the number of unique genes detected and the mitochondrial percentage.

This plot will also show, for each cell, the posterior probability that the cell is derived from the compromised distribution; a higher score indicates that a cell is more likely to be compromised.

It is also critical to note that this model can _and does_ fail at times.
Plotting the results as we have done here is not a step to skip.
**Always look at your data!**

```{r miQC plotModel, live=TRUE}
# plot the miQC model
miQC::plotModel(filtered_sce, miqc_model) +
  theme_bw()
```

We can now filter our data based on the probability compromised as calculated from the model.
But before we do that, we might want to quickly plot to see what would be filtered out with a given cutoff, using the `plotFiltering()` function.
The default is to exclude cells that have a posterior probability of 0.75 or greater of being compromised.
We stick with that default, but for clarity, we will also include it in our code!


```{r miQC plotFiltering}
# look at miQC filtering
miQC::plotFiltering(filtered_sce, miqc_model,
                    posterior_cutoff = 0.75) +
  theme_bw()
```

In this case, the line between the cells to be kept and those that will be removed seems to correspond to a mitochondrial percentage of about 12.5%, but note that this will not always be constant.
The cutoff point can vary for different numbers of unique genes within a sample, and it will certainly vary among samples!

At this point, we can perform the actual filtering using the `filterCells()` function, giving us a further filtered SCE object.

```{r miQC filtercells, live=TRUE}
# perform miQC filtering
qcfiltered_sce <- miQC::filterCells(filtered_sce,
                                    model = miqc_model)
```

#### One more filter: unique gene count

While the miQC filtering is pretty good, you may have noticed that it still left some cells that had very low numbers of unique genes.
While these cells may not be compromised, the information from them is also not likely to be useful, so we will filter those as well.
We will only keep cells that have at least 200 unique genes.

```{r unique cutoff, live=TRUE}
# filter cells by unique gene count (`detected`)
qcfiltered_sce <- qcfiltered_sce[, which(qcfiltered_sce$detected >= 200)]
qcfiltered_sce
```


## Normalization

Now that we have done our filtering, we can start analyzing the expression counts for the remaining cells.

The next step at this point is to convert the raw counts into a measure that accounts for differences in sequencing depth between cells, and to convert the distribution of expression values from the skewed distribution we expect to see in raw counts to one that is more normally distributed.

We will do this using functions from the `scran` and `scuttle` packages.
The procedure we will use here is derived from the [OSCA chapter on normalization](http://bioconductor.org/books/3.19/OSCA.basic/normalization.html#normalization-by-deconvolution).
The idea is that the varying expression patterns that different cell types exhibit will affect the scaling factors that we would apply.
To account for that variation, we first do a rough clustering of cells by their expression with `scran::quickCluster()`, then use that clustering to calculate the scaling factor for each cell within the clusters using `scran::computeSumFactors()`.
Finally, we apply the scaling factor to the expression values for each cell and calculate the log-scaled expression values using the `scuttle::logNormCounts()` function.

```{r normalization, live=TRUE}
# Perform rough clustering
qclust <- scran::quickCluster(qcfiltered_sce)

# use clusters to compute scaling factors and add to SCE object
qcfiltered_sce <- scran::computeSumFactors(qcfiltered_sce,
                                           clusters = qclust)

# perform normalization using scaling factors
# and save as a new SCE object
normalized_sce <- scuttle::logNormCounts(qcfiltered_sce)
```

This creates a new "assay" in the `normalized_sce` object, `logcounts`, which contains the normalized count values for each cell and gene.
(The data here are not _actually_ the log of the counts, since we also applied the scaling factors, but that name is used for historical reasons.)

Let's take a look:

```{r normalized sce}
normalized_sce
```

## Dimension reduction

![Single-cell roadmap: Dimension reduction](diagrams/roadmap_single_dimension_reduction.png)

Now that we have normalized expression values, we would like to produce some reduced-dimension representations of the data.
These will allow us to perform some downstream calculations more quickly, reduce some of the noise in the data, and allow us to visualize overall relationships among cells more easily (though with many caveats!).

### Selecting highly variable genes

While we could calculate the reduced dimensions using all of the genes that we have assayed, in practice most of the genes will have very little variation in expression, so doing so will not provide much additional signal.
Reducing the number of genes we include will also speed up some of the calculations.

To identify the most variable genes, we will use functions from the `scran` package.
The first function, `modelGeneVar()`, attempts to divide the variation observed for each gene into a biological and technical component, with the intuition that genes with lower mean expression tend to have lower variance for purely technical reasons.
We then provide the `modelGeneVar()` output to the `getTopHVGs()` function to identify the genes with the highest _biological_ variation, which is what we are most interested in.

```{r select HVGs}
# identify 2000 genes
num_genes <- 2000

# model variance, partitioning into biological and technical variation
gene_variance <- scran::modelGeneVar(normalized_sce)

# get the most variable genes
hv_genes <- scran::getTopHVGs(gene_variance,
                              n = num_genes)
```

The result is a vector of gene ids (ordered from most to least variable):

```{r view HVGs}
head(hv_genes)
```

### Principal components analysis

Now that we have selected the genes we would like to use for the reduced-dimension representations of the expression data, we can start to calculate them.
First we will use the `scater::runPCA()` function to calculate the principal components from the expression matrix.
This representation is fast and fairly robust, but the result is still quite multidimensional.
We want keep a fair number of components (dimensions) in order to accurately represent the variation in the data, but doing so means that plotting only a few of these dimensions (in 2D) is not likely to provide a full view of the data.

The default number of components is 50, which we will stick with, but let's enter it manually just for the record.

```{r runPCA, live=TRUE}
# calculate and save PCA results
normalized_sce <- scater::runPCA(
  normalized_sce,
  ncomponents = 50, # how many components to keep
  subset_row = hv_genes # use only the variable genes we chose
)
```

These reduced-dimension results will be stored in a `reducedDim` slot in the SCE object.
We can see the names of the `reducedDim`s that we have by looking at the object summary:

```{r view reduced dimensions}
normalized_sce
```

If we want to extract the PCA results, we can do that with the `reducedDim()` function:
Note that for these reduced-dimensionality matrices, the rows are the cells and the columns are the PC dimensions.

```{r getReducedDim}
# extract the PCA matrix
pca_matrix <- reducedDim(normalized_sce, "PCA")

# look at the shape of the matrix
dim(pca_matrix)
```

### UMAP

Finally, we will calculate a UMAP (Uniform Manifold Approximation and Projection) representation of our data.
This is a machine-learning-based method that is useful for performing dimensionality reduction suitable for visualization.
It's goal is to provide a representation of the data in two dimensions (typically, more are possible) that preserves as much of the distance relationships among cells as possible.
While this does make for visually appealing and useful plots, it is important not to overinterpret the results!
In particular, while you will often see some apparent clustering of cells in the resulting output, those clusters may not be particularly valid, and the spacing within or between clusters may not reflect true distances.

In many ways this is analogous to the problem of projecting a map of the earth onto a flat surface; any choice will result in some distortions.
However, with UMAP, we rarely know exactly what choices were made and what distortions might have resulted.
The UMAP coordinates themselves should never be used for downstream analysis.

Since the UMAP procedure would be slow to calculate with the full data, so the `runUMAP()` function first calculates a PCA matrix and then uses _that_ to calculate the UMAP.
Since we already have a PCA matrix, we will tell the function use that instead of recalculating it.

```{r runUMAP, live=TRUE}
normalized_sce <- scater::runUMAP(normalized_sce,
                                  dimred = "PCA")
```

As before, we could extract the UMAP matrix from our SCE object with the `reducedDim()` function.
We can also visualize the UMAP results using the `plotReducedDim()` function.

```{r plotReducedDim, live=TRUE}
# plot the UMAP
scater::plotReducedDim(normalized_sce,
                       "UMAP",
                       # color by the most variable gene
                       color_by = hv_genes[1])
```


## Unsupervised clustering

As a final analysis step at this stage, we will return to the PCA results to perform unsupervised clustering.
Here we will use a graph-based clustering method, which starts by identifying cells that are close together in the multidimensional space.
It then identifies "communities" of highly connected cells, and breaks them apart by regions of lower connection.

There are a number of algorithms that can perform this clustering, each with parameters that can affect how many clusters are identified and which cells belong to each cluster.
It is also worth noting that these clusters may or may not correspond to "cell types" by whatever definition you might prefer to use.
Interpretation of these clusters, or other measures of cell type, are something that will require more careful and likely more customized analysis.

We will perform our clustering using the function `scran::clusterCells`, which can perform many different types of clustering using the Bioconductor `bluster` package under the hood.
As mentioned earlier, we are using "graph" clustering, which we define using the `bluster::NNGraphParam()` function.
Within that are a number of further options, such as the weighting used for building the network graph and the algorithm used for dividing the graph into clusters.

Modifying these parameters can result in quite different cluster assignments!
For the clustering below we will use Jaccard weighting and Louvain clustering, which correspond more closely to the default methods used by `Seurat` than the default parameters.
It is also worth noting that the the cluster assignments are somewhat stochastic.
In particular, the names/numbers of the clusters can be quite inconsistent between runs!


```{r clustering, live=TRUE}
# perform graph-based clustering
nn_clusters <- scran::clusterCells(
  normalized_sce, # SCE to perform clustering on
  use.dimred = "PCA", # perform clustering on the PCA matrix
  BLUSPARAM = bluster::NNGraphParam( # clustering parameters to pass to bluster
    # number of neighbors to use in network graph
    k = 20,
    # weighting scheme for building the network graph
    # default is "rank"
    type = "jaccard",
    # cluster detection algorithm
    # default is "walktrap"
    cluster.fun = "louvain"
  )
)
```

We can save the cluster assignments back into the `colData` of the SCE object with a little shortcut: the `$` followed by the name of the new column we want to add.

```{r add clusters to SCE, live=TRUE}
# save clusters to SCE colData
normalized_sce$nn_cluster <- nn_clusters
```

Now we can plot the UMAP again, this time colored by the cluster assignments that we just created.
Here rather than the general `plotReducedDim()` function, we will use `plotUMAP()`, which is exactly the same, except it always plots from the `reducedDim` slot named `UMAP`, so we can skip that argument.

```{r plot clusters, live=TRUE}
# plot UMAP with assigned clusters
scater::plotUMAP(normalized_sce,
                 color_by = "nn_cluster")
```

What do you see in these results?

What would you want to do next?

## Save SCE object for later

We will now save our filtered and normalized object, including the dimension reduction and clustering results to an `RDS` file, using the file path that we defined at the start of the notebook.
If we were to want to return to this data, we could load this file directly into a new R session and not have to repeat the processing that we have done up to this point.

The data in these objects tends to be quite large, but very compressible.
To save space on disk (at the expense of time), we will make sure that the data is compressed internally before writing it out to a file.
Note that the file we write is still going to be an `.rds` file with no additional extension.
(Further note: The base R function `saveRDS()` uses compression by default, but the `tidyverse` function `readr::write_rds()` does not.)

```{r save SCE, live=TRUE}
# write RDS file with compression
readr::write_rds(normalized_sce, file = output_sce_file, compress = "gz")
```


## Print session info

As is our habit at the Data Lab, we will save information about the computing environment, the packages we have used in this notebook, and their versions using the `sessionInfo()` command.

```{r session info}
sessionInfo()
```


+
---
title: "Reading, filtering, and normalizing scRNA-seq data"
author: Data Lab for ALSF
date: 2023
output:
  html_notebook:
    toc: true
    toc_float: true
---

## Objectives

This notebook will demonstrate how to:

- Read Cell Ranger data into R
- Filter to cells using `emptyDropsCellRanger()`
- Calculate quality control measures on scRNA-seq data
- Remove likely compromised cells with `miQC()`
- Normalize expression data across cells
- Calculate and plot reduced dimension representations of expression data (PCA, UMAP)

---

In this notebook, we will review basic processing for single-cell RNA-seq data, starting with the output from Cell Ranger, and proceeding through filtering, quality control, normalization, and dimension reduction. We will perform these tasks using tools from the [Bioconductor project](https://bioconductor.org), in particular [`SingleCellExperiment` objects](https://bioconductor.org/packages/release/bioc/html/SingleCellExperiment.html) and functions that work with those objects.
Much of the material in this notebook is directly inspired by, and draws heavily on, material presented in the book [_Orchestrating Single Cell Analysis with Bioconductor_ (OSCA)](http://bioconductor.org/books/release/OSCA/).

![Single-cell roadmap: Overview](diagrams/roadmap_single_overview.png)

The data we will use for this notebook is derived from a human glioblastoma specimen.
The sample was processed by 10x Genomics using a 3' RNA kit (v3.1), sequenced, and quantified with Cell Ranger 6.0.
Further details about the sample and processing can be found on the [10x website](https://www.10xgenomics.com/resources/datasets/2-k-sorted-cells-from-human-glioblastoma-multiforme-3-v-3-1-3-1-standard-6-0-0).


## Set Up

To start, we will load some of the libraries we will need later, and set a random number seed for reproducibility.

```{r setup}
# Load libraries

# Plotting functions
library(ggplot2)

# The main class we will use for Single Cell data
library(SingleCellExperiment)

# Setting the seed for reproducibility
set.seed(12345)
```

### Directories and files

Before we get too far, we like to define the input and output files that the notebook will use near the top of the document.
While you might not know the names of all of the files you will need or create output files when you start an analysis, we have found it helpful to keep all file and directory names in a single place near the top of the document.
This makes it easier for somebody coming to the code later to quickly see what files are needed as input and what will be produced as output.
More often than not, that somebody is you!

The gene expression data were processed to create a gene-by-cell expression matrix of counts for using Cell Ranger 6.0.
We have provided the raw data directory, `raw_feature_bc_matrix`, which is usually produced by Cell Ranger and placed in its `outs` directory.
This directory usually contains three files:
- `barcodes.tsv.gz`, a table of the cell barcodes that 10x uses, corresponding to the columns of the count matrix.
- `features.tsv.gz`, a table of the features (genes in this case) for which expression was quantified.
This will usually also include a bit of metadata about the features, including gene symbols (if the features are genes) and the type of data they represent (e.g., gene expression or antibody capture).
- `matrix.mtx.gz`, The counts themselves, stored in a sparse ["Matrix Exchange" format](https://math.nist.gov/MatrixMarket/formats.html).

Cell Ranger will also export these data in a single `HDF5` format file with a `.h5` extension, which can also be imported with the same commands we will use below.
However, we have found that processing large `.h5` files is often _much_ less efficient in R, so we prefer to start with the matrix files when possible.
In particular, we would not recommend working with `.h5` files for raw data; the filtering steps we will use below can sometimes take hours when using those files as input.

We will also need a table of mitochondrial genes, which we have stored in the `data/reference/` directory.

Finally, we will set up the our output directory, creating it if it does not yet exist, and define the name for the files we will save after all of our initial processing is complete.

```{r inputs, live=TRUE}
# Inputs --------------------------------------
# main data directory
data_dir <- file.path("data", "glioblastoma-10x")

# Path to the Cell Ranger matrix directory
raw_matrix_dir <- file.path(data_dir, "raw_feature_bc_matrix")

# reference data directory
ref_dir <- file.path("data", "reference")

# Path to mitochondrial genes table
mito_file <- file.path(ref_dir, "hs_mitochondrial_genes.tsv")
```

```{r outputs}
# Outputs ------------------------------------

# Directory and file to save output
normalized_dir <- file.path(data_dir, "normalized")

# create the directory if it does not exist
fs::dir_create(normalized_dir)

# output RDS file for normalized data
output_sce_file <- file.path(
  normalized_dir,
  "glioblastoma_normalized_sce.rds"
)
```


## Reading Cell Ranger data

![Single-cell roadmap: Preprocess and Import](diagrams/roadmap_single_preprocess.png)

Whether the 10x Cell Ranger data is in Matrix Exchange format or in an HDF5 file, we can use the `read10xCounts()` function from the `DropletUtils` package to read the data into R and create a `SingleCellExperiment` object.
(Though again, we do not recommend using the `.h5` file if you can avoid it, _especially_ for raw (unfiltered) data.)

If you used something other than Cell Ranger to process the raw data, you would need to use a different function to read it in and create the `SingleCellExperiment` object.
Some of these functions for other common data formats are discussed in [Chapter 3 of OSCA](http://bioconductor.org/books/3.19/OSCA.intro/getting-scrna-seq-datasets.html#reading-counts-into-r).

```{r read SCE, live=TRUE}
# read SCE from matrix directory
raw_sce <- DropletUtils::read10xCounts(
  raw_matrix_dir,
  col.names = TRUE # ensure barcodes are set as column names in the SCE object
)
```

Let's look at the contents of the object after reading it in:

```{r view SCE, live=TRUE}
# view SCE object
raw_sce
```

We can see from this summary that this `SingleCellExperiment` (SCE) object contains 36,601 rows, which correspond to the features (genes) that were analyzed, and 734,492 columns, which correspond to the possible barcode tags that were used in the experiment.
Note that not all of these barcode tags will have been used, and many of the features may never have been seen either.
One of our first steps will be to filter out barcodes that were never seen, or that may have only been seen in a droplet that did not contain a cell (an "empty droplet").

### Structure of the `SingleCellExperiment` object

In addition to the main `counts` matrix, listed as an `assay` in the SCE summary above, the SCE object can contain a number of other tables and matrices, each stored in a "slot" with a particular format.
The overall structure of the object can be seen in the figure below, which comes from an [OSCA Introduction chapter](http://bioconductor.org/books/3.19/OSCA.intro/the-singlecellexperiment-class.html).

![Structure of a SingleCellExperiment object](diagrams/SingleCellExperiment.png)

We have just mentioned the main `assay` slot, which contains full matrices of data (such as transcript counts) with each row a feature and each column a cell.
There are also a couple of tables for metadata, and a slot to store reduced-dimension representations (e.g., PCA and/or UMAP) of the expression data.

We'll start with the `rowData` slot, which is a table of metadata for each feature in our object.
For now that contains the contents of the `features.tsv.gz` file that we discussed earlier.
If we had read the data from something other than Cell Ranger output, we might have different contents, but each row would still correspond to a single feature of the SCE object.

Let's look at this table, extracting it from the SCE object with the `rowData()` function and using `head()` to view only the first 6 rows.

```{r rowdata}
# view rowData (features)
head(rowData(raw_sce))
```

You can see that this table includes an `ID` for each feature, which is usually the Ensembl gene ID, as well as the corresponding gene symbol in the `Symbol` column.
Finally there is a column for `Type`, which in this case is always "Gene Expression", as all of the features in this data set are genes.
If there were another modality of data that had been assayed in this experiment, there might be other values in this column, such as "Antibody Capture" for CITE-seq experiments.

The second slot is the `colData` table, which now corresponds to the `barcodes.tsv.gz` file, containing one row per cell barcode, or, more generally, one row per column of the `counts` assay.
We can look at this table using the `colData()` function (and `head()` again to prevent printing the whole table):

```{r coldata}
# view colData (cell barcodes)
head(colData(raw_sce))
```

Here we see that there are currently two columns:

- the `Sample` column has the path of the file that we read in (you may not see the whole path in this display); this should be identical in all rows from a single sample.
- the `Barcode` column contains the sequence that was used to identify each potential droplet for sequencing (and a numeric tag, in this case).
These will be unique within a sample.

As we proceed to calculate per-cell statistics, we will be adding new data to this table.

## Quality control and filtering

![Single-cell roadmap: QC, Filter, and Normalize](diagrams/roadmap_single_qc_norm.png)

### Filtering empty droplets

Most of the barcodes in any given 10x experiment will not be seen at all, so our first step can be to filter this raw data to only the cells where there is at least one transcript that was counted with that barcode.

To do this, we will use the `colSums()` function to quickly add up all the counts that correspond to each possible cell barcode, then filter our `raw_sce` down to just those columns where there are non-zero total counts.
We will need to extract the `counts` matrix from our SCE object, which we can do using the `counts()` function, conveniently enough.

```{r remove zeros, live=TRUE}
# sum columns from counts matrix
barcode_counts <- colSums(counts(raw_sce))

# filter SCE object to only rows with counts > 0
raw_sce <- raw_sce[, which(barcode_counts > 0)]
```

Now we can look at how our SCE object has changed:

```{r zero-filtered SCE}
raw_sce
```

But barcodes with zero counts are not the only ones that correspond to droplets without cells in them!
Even if a droplet does not have a cell in it, there will often be spurious reads from RNA sequences that were present in the extracellular solution, whether from the original sample or from cells that were damaged during single-cell library preparation.

We could identify these barcodes simply as those with low transcript counts.
Or, we can be a bit more clever!
We can look at the transcript counts _from_ the lowest-count droplets to create an expected distribution of transcripts in droplets that don't contain cells.
Then we can test each droplet to determine whether or not its transcript distribution deviates from that expectation.
If it does, then we have pretty good evidence that there _is_ a cell in there.

This test was first proposed by [Lun _et al._ (2019)](https://doi.org/10.1186/s13059-019-1662-y) and implemented as `emptyDrops()` in the `DropletUtils` package.
This method was then adopted, with some modifications, as the default cell filtering method used by Cell Ranger.
Here we will use the [`emptyDropsCellRanger()` function](https://rdrr.io/github/MarioniLab/DropletUtils/man/emptyDropsCellRanger.html) to perform filtering that more closely matches the Cell Ranger implementation.


```{r calculate droplet stats, live=TRUE}
# create a table of statistics using emptyDropsCellRanger
droplet_df <- DropletUtils::emptyDropsCellRanger(raw_sce)
```

Most values in this table are `NA`, because individual statistics were not calculated for the low-count droplets that were used to generate the background distribution.
(Most droplets don't have cells, so this makes some sense!)

We can look at just the rows without `NA` values by selected the ones where the FDR (which we will use again soon), is not `NA`.

```{r droplet stats}
# view rows where FDR is not `NA`
droplet_df[!is.na(droplet_df$FDR), ]
```
You will notice that some cells with high counts also have `NA` values for many statistics.
In those cases, `NA` values are actually present _because_ of the high counts - `emptyDropsCellRanger()` automatically assumed cells were present, so they were also not tested.

Now we can filter our `raw_sce` object _by column_ to only keep the cells with a small FDR: those that are quite unlikely to be empty droplets.

```{r filter emptydrops, live=TRUE}
# filter droplets using `which` to prevent NA trouble
cells_to_retain <- which(droplet_df$FDR < 0.01)
filtered_sce <- raw_sce[, cells_to_retain]
```

How many cells do we have now?

```{r filtered summary}
filtered_sce
```

### Additional quality control

In addition to filtering out empty droplets, we also will want to filter out cells that may have been damaged during library preparation.
These will often be characterized by a high proportion of mitochondrial transcripts and a smaller overall number of unique transcripts.
When a cell ruptures, cytoplasmic transcripts will leak out, but mitochondrial transcripts, still protected by the mitochondrial membrane, may remain.
As a consequence, there will be an over-abundance of mitochondrial reads, and fewer unique transcripts expressed.

Our first step then, is create a vector of the mitochondrial genes that are present in our dataset.
The mitochondrial file we defined during setup (`mito_file`) is a TSV file containing all of the human mitochondrial genes with additional annotation information for each gene, such as the gene location and alternative names.
(For more detail on the steps we took to create this file, you can look at [one of our setup notebooks](https://github.com/AlexsLemonade/training-modules/blob/master/scRNA-seq-advanced/setup/mito_gene_lists.Rmd))

All we need now is the `gene_id`, and only for the genes that are present in our SCE, so we will do some filtering with `dplyr` to pull out a vector with just those ids.

```{r get mitochondrial genes}
# read in a table of mitochondrial genes and extract ids
mito_genes <- readr::read_tsv(mito_file) |>
  # filter to only the genes that are found in our dataset
  dplyr::filter(gene_id %in% rownames(filtered_sce)) |>
  # create a vector from the gene_id column
  dplyr::pull(gene_id)
```

### Calculating summary QC statistics

We can now use the `scuttle` function `addPerCellQC()` to calculate some statistics based on the counts matrix, which will be added to the `colData` table.

In addition to calculating statistics like the total read count for each cell and the number of transcripts that are detected, we can also calculate those statistics for defined subsets of genes.
In this case, we will use our `mito_genes` vector to define a subset called `mito`.
The `mito` name is important in that it is the name that will be expected by a later function.
(We could define more subsets, but for now this one will do.)

```{r per cell QC, live=TRUE}
filtered_sce <- scuttle::addPerCellQC(
  filtered_sce,
  subsets = list(mito = mito_genes)
)
```

Now we can look at the colData to see what was added:

```{r view colData stats}
head(colData(filtered_sce))
```

We can also plot some of these statistics, here using the `plotMetrics()` function from the `miQC` package to plot the percent of reads that are mitochondrial (the `subsets_mito_percent` column) against the number of unique genes detected (the `detected` column) for each cell.

```{r miQC plotMetrics}
# use miQC::plotMetrics()
miQC::plotMetrics(filtered_sce) + theme_bw()
```

We can see that there is a range of mitochondrial percentages, and it does also seem that cells with high percentages of mitochondrial genes don't seem to contain very many unique genes.

How do we filter with this information?
One option is to define a cutoff for the mitochondrial percentage above which we call a cell compromised and exclude it from further analysis.
However, choosing that cutoff can be a bit fraught, as the expected percentage of mitochondrial reads can vary depending on the cell type and library preparation methods.
So it might be nice to have a method to determine that cutoff from the data itself.

### Filtering compromised cells

Determining mitochondrial cutoffs is exactly what the `miQC` package does ([Hippen _et al._ 2021](https://doi.org/10.1371/journal.pcbi.1009290))!
In truth, it does something possibly even a bit better: it fits a mixture model to the data that consists of distributions of healthy cells and compromised cells.
Then we can calculate whether each cell is more likely to belong to the healthy or compromised distribution.
We can then exclude the cells that are more likely to be compromised.

To use `miQC`, we first fit a model to the data in our SCE object:

```{r miQC model, live=TRUE}
# fit the miQC model
miqc_model <- miQC::mixtureModel(filtered_sce)
```

Now we can plot the model results using the `plotModel()` function to see how it corresponds to our data.
We should expect to see two fit lines:

- One line will correspond the the "healthy" cells and should show little to no relationship between the number of unique genes detected and the mitochondrial percentage.
- By contrast, the line that corresponds to "compromised" cells will show a negative relationship between the number of unique genes detected and the mitochondrial percentage.

This plot will also show, for each cell, the posterior probability that the cell is derived from the compromised distribution; a higher score indicates that a cell is more likely to be compromised.

It is also critical to note that this model can _and does_ fail at times.
Plotting the results as we have done here is not a step to skip.
**Always look at your data!**

```{r miQC plotModel, live=TRUE}
# plot the miQC model
miQC::plotModel(filtered_sce, miqc_model) +
  theme_bw()
```

We can now filter our data based on the probability compromised as calculated from the model.
But before we do that, we might want to quickly plot to see what would be filtered out with a given cutoff, using the `plotFiltering()` function.
The default is to exclude cells that have a posterior probability of 0.75 or greater of being compromised.
We stick with that default, but for clarity, we will also include it in our code!


```{r miQC plotFiltering}
# look at miQC filtering
miQC::plotFiltering(
  filtered_sce,
  miqc_model,
  posterior_cutoff = 0.75
) +
  theme_bw()
```

In this case, the line between the cells to be kept and those that will be removed seems to correspond to a mitochondrial percentage of about 12.5%, but note that this will not always be constant.
The cutoff point can vary for different numbers of unique genes within a sample, and it will certainly vary among samples!

At this point, we can perform the actual filtering using the `filterCells()` function, giving us a further filtered SCE object.

```{r miQC filtercells, live=TRUE}
# perform miQC filtering
qcfiltered_sce <- miQC::filterCells(
  filtered_sce,
  model = miqc_model
)
```

#### One more filter: unique gene count

While the miQC filtering is pretty good, you may have noticed that it still left some cells that had very low numbers of unique genes.
While these cells may not be compromised, the information from them is also not likely to be useful, so we will filter those as well.
We will only keep cells that have at least 200 unique genes.

```{r unique cutoff, live=TRUE}
# filter cells by unique gene count (`detected`)
qcfiltered_sce <- qcfiltered_sce[, which(qcfiltered_sce$detected >= 200)]
qcfiltered_sce
```


## Normalization

Now that we have done our filtering, we can start analyzing the expression counts for the remaining cells.

The next step at this point is to convert the raw counts into a measure that accounts for differences in sequencing depth between cells, and to convert the distribution of expression values from the skewed distribution we expect to see in raw counts to one that is more normally distributed.

We will do this using functions from the `scran` and `scuttle` packages.
The procedure we will use here is derived from the [OSCA chapter on normalization](http://bioconductor.org/books/3.19/OSCA.basic/normalization.html#normalization-by-deconvolution).
The idea is that the varying expression patterns that different cell types exhibit will affect the scaling factors that we would apply.
To account for that variation, we first do a rough clustering of cells by their expression with `scran::quickCluster()`, then use that clustering to calculate the scaling factor for each cell within the clusters using `scran::computeSumFactors()`.
Finally, we apply the scaling factor to the expression values for each cell and calculate the log-scaled expression values using the `scuttle::logNormCounts()` function.

```{r normalization, live=TRUE}
# Perform rough clustering
qclust <- scran::quickCluster(qcfiltered_sce)

# use clusters to compute scaling factors and add to SCE object
qcfiltered_sce <- scran::computeSumFactors(
  qcfiltered_sce,
  clusters = qclust
)

# perform normalization using scaling factors
# and save as a new SCE object
normalized_sce <- scuttle::logNormCounts(qcfiltered_sce)
```

This creates a new "assay" in the `normalized_sce` object, `logcounts`, which contains the normalized count values for each cell and gene.
(The data here are not _actually_ the log of the counts, since we also applied the scaling factors, but that name is used for historical reasons.)

Let's take a look:

```{r normalized sce}
normalized_sce
```

## Dimension reduction

![Single-cell roadmap: Dimension reduction](diagrams/roadmap_single_dimension_reduction.png)

Now that we have normalized expression values, we would like to produce some reduced-dimension representations of the data.
These will allow us to perform some downstream calculations more quickly, reduce some of the noise in the data, and allow us to visualize overall relationships among cells more easily (though with many caveats!).

### Selecting highly variable genes

While we could calculate the reduced dimensions using all of the genes that we have assayed, in practice most of the genes will have very little variation in expression, so doing so will not provide much additional signal.
Reducing the number of genes we include will also speed up some of the calculations.

To identify the most variable genes, we will use functions from the `scran` package.
The first function, `modelGeneVar()`, attempts to divide the variation observed for each gene into a biological and technical component, with the intuition that genes with lower mean expression tend to have lower variance for purely technical reasons.
We then provide the `modelGeneVar()` output to the `getTopHVGs()` function to identify the genes with the highest _biological_ variation, which is what we are most interested in.

```{r select HVGs}
# identify 2000 genes
num_genes <- 2000

# model variance, partitioning into biological and technical variation
gene_variance <- scran::modelGeneVar(normalized_sce)

# get the most variable genes
hv_genes <- scran::getTopHVGs(
  gene_variance,
  n = num_genes
)
```

The result is a vector of gene ids (ordered from most to least variable):

```{r view HVGs}
head(hv_genes)
```

### Principal components analysis

Now that we have selected the genes we would like to use for the reduced-dimension representations of the expression data, we can start to calculate them.
First we will use the `scater::runPCA()` function to calculate the principal components from the expression matrix.
This representation is fast and fairly robust, but the result is still quite multidimensional.
We want keep a fair number of components (dimensions) in order to accurately represent the variation in the data, but doing so means that plotting only a few of these dimensions (in 2D) is not likely to provide a full view of the data.

The default number of components is 50, which we will stick with, but let's enter it manually just for the record.

```{r runPCA, live=TRUE}
# calculate and save PCA results
normalized_sce <- scater::runPCA(
  normalized_sce,
  ncomponents = 50, # how many components to keep
  subset_row = hv_genes # use only the variable genes we chose
)
```

These reduced-dimension results will be stored in a `reducedDim` slot in the SCE object.
We can see the names of the `reducedDim`s that we have by looking at the object summary:

```{r view reduced dimensions}
normalized_sce
```

If we want to extract the PCA results, we can do that with the `reducedDim()` function:
Note that for these reduced-dimensionality matrices, the rows are the cells and the columns are the PC dimensions.

```{r getReducedDim}
# extract the PCA matrix
pca_matrix <- reducedDim(normalized_sce, "PCA")

# look at the shape of the matrix
dim(pca_matrix)
```

### UMAP

Finally, we will calculate a UMAP (Uniform Manifold Approximation and Projection) representation of our data.
This is a machine-learning-based method that is useful for performing dimensionality reduction suitable for visualization.
It's goal is to provide a representation of the data in two dimensions (typically, more are possible) that preserves as much of the distance relationships among cells as possible.
While this does make for visually appealing and useful plots, it is important not to overinterpret the results!
In particular, while you will often see some apparent clustering of cells in the resulting output, those clusters may not be particularly valid, and the spacing within or between clusters may not reflect true distances.

In many ways this is analogous to the problem of projecting a map of the earth onto a flat surface; any choice will result in some distortions.
However, with UMAP, we rarely know exactly what choices were made and what distortions might have resulted.
The UMAP coordinates themselves should never be used for downstream analysis.

Since the UMAP procedure would be slow to calculate with the full data, so the `runUMAP()` function first calculates a PCA matrix and then uses _that_ to calculate the UMAP.
Since we already have a PCA matrix, we will tell the function use that instead of recalculating it.

```{r runUMAP, live=TRUE}
normalized_sce <- scater::runUMAP(
  normalized_sce,
  dimred = "PCA"
)
```

As before, we could extract the UMAP matrix from our SCE object with the `reducedDim()` function.
We can also visualize the UMAP results using the `plotReducedDim()` function.

```{r plotReducedDim, live=TRUE}
# plot the UMAP
scater::plotReducedDim(
  normalized_sce,
  "UMAP",
  # color by the most variable gene
  color_by = hv_genes[1]
)
```


## Unsupervised clustering

As a final analysis step at this stage, we will return to the PCA results to perform unsupervised clustering.
Here we will use a graph-based clustering method, which starts by identifying cells that are close together in the multidimensional space.
It then identifies "communities" of highly connected cells, and breaks them apart by regions of lower connection.

There are a number of algorithms that can perform this clustering, each with parameters that can affect how many clusters are identified and which cells belong to each cluster.
It is also worth noting that these clusters may or may not correspond to "cell types" by whatever definition you might prefer to use.
Interpretation of these clusters, or other measures of cell type, are something that will require more careful and likely more customized analysis.

We will perform our clustering using the function `scran::clusterCells`, which can perform many different types of clustering using the Bioconductor `bluster` package under the hood.
As mentioned earlier, we are using "graph" clustering, which we define using the `bluster::NNGraphParam()` function.
Within that are a number of further options, such as the weighting used for building the network graph and the algorithm used for dividing the graph into clusters.

Modifying these parameters can result in quite different cluster assignments!
For the clustering below we will use Jaccard weighting and Louvain clustering, which correspond more closely to the default methods used by `Seurat` than the default parameters.
It is also worth noting that the the cluster assignments are somewhat stochastic.
In particular, the names/numbers of the clusters can be quite inconsistent between runs!


```{r clustering, live=TRUE}
# perform graph-based clustering
nn_clusters <- scran::clusterCells(
  normalized_sce, # SCE to perform clustering on
  use.dimred = "PCA", # perform clustering on the PCA matrix
  BLUSPARAM = bluster::NNGraphParam( # clustering parameters to pass to bluster
    # number of neighbors to use in network graph
    k = 20,
    # weighting scheme for building the network graph
    # default is "rank"
    type = "jaccard",
    # cluster detection algorithm
    # default is "walktrap"
    cluster.fun = "louvain"
  )
)
```

We can save the cluster assignments back into the `colData` of the SCE object with a little shortcut: the `$` followed by the name of the new column we want to add.

```{r add clusters to SCE, live=TRUE}
# save clusters to SCE colData
normalized_sce$nn_cluster <- nn_clusters
```

Now we can plot the UMAP again, this time colored by the cluster assignments that we just created.
Here rather than the general `plotReducedDim()` function, we will use `plotUMAP()`, which is exactly the same, except it always plots from the `reducedDim` slot named `UMAP`, so we can skip that argument.

```{r plot clusters, live=TRUE}
# plot UMAP with assigned clusters
scater::plotUMAP(
  normalized_sce,
  color_by = "nn_cluster"
)
```

What do you see in these results?

What would you want to do next?

## Save SCE object for later

We will now save our filtered and normalized object, including the dimension reduction and clustering results to an `RDS` file, using the file path that we defined at the start of the notebook.
If we were to want to return to this data, we could load this file directly into a new R session and not have to repeat the processing that we have done up to this point.

The data in these objects tends to be quite large, but very compressible.
To save space on disk (at the expense of time), we will make sure that the data is compressed internally before writing it out to a file.
Note that the file we write is still going to be an `.rds` file with no additional extension.
(Further note: The base R function `saveRDS()` uses compression by default, but the `tidyverse` function `readr::write_rds()` does not.)

```{r save SCE, live=TRUE}
# write RDS file with compression
readr::write_rds(normalized_sce, file = output_sce_file, compress = "gz")
```


## Print session info

As is our habit at the Data Lab, we will save information about the computing environment, the packages we have used in this notebook, and their versions using the `sessionInfo()` command.

```{r session info}
sessionInfo()
```


diff --git a/scRNA-seq-advanced/02-dataset_integration-live.Rmd b/scRNA-seq-advanced/02-dataset_integration-live.Rmd index ce4cbb89..64a4cd7e 100644 --- a/scRNA-seq-advanced/02-dataset_integration-live.Rmd +++ b/scRNA-seq-advanced/02-dataset_integration-live.Rmd @@ -55,7 +55,7 @@ set.seed(12345) ``` -### Directories and files +### Define directories and files We have already prepared count data for the four samples we'll be integrating (i.e., filtered cells, normalized counts, and calculated PCA & UMAP). @@ -66,17 +66,22 @@ These SCE objects, stored as RDS files, are available in the `data/rms/processed - `SCPCL000481.rds` (Patient B) - `SCPCL000482.rds` (Patient B) +Both Patient A (18 year old male) and Patient B (4 year old female) had recurrent embryonal rhabdomyosarcoma when samples were taken. + To begin, let's set up our directories and files: -```{r directories, live = TRUE} +```{r directories} # Define directory where processed SCE objects to be integrated are stored +input_dir <- file.path("data", "rms", "processed") # Define directory to save integrated SCE object to +output_dir <- file.path("data", "rms", "integrated") # Create output directory if it doesn't exist +fs::dir_create(output_dir) # Define output file name for the integrated object - +integrated_sce_file <- file.path(output_dir, "rms_integrated_subset.rds") ``` @@ -126,29 +131,6 @@ length(histologies) purrr::map(histologies, length) ``` -One other new coding strategy we'll learn in this notebook is using the [`glue`](https://glue.tidyverse.org/) package to combine strings. -This package offers a convenient function `glue::glue()` that can be used instead of the base R `paste()` function. - -```{r paste} -# Define a variable for example: -org_name <- "Data Lab" - -# We can use paste to combine strings and variables: -paste("Welcome to the", org_name, "workshop on Advanced scRNA-seq!") -``` - -We can use `glue::glue()` to accomplish the same goal with some different syntax: - -```{r glue} -# glue::glue takes a single string argument (only one set of quotes!), and -# variables can easily be included inside {curly braces} -glue::glue("Welcome to the {org_name} workshop on Advanced scRNA-seq!") -``` - -(Note that even though the `glue::glue()` output isn't in quotes, it still behaves like a string!) - - -Alright, time for the good stuff! Let's use `purrr::map()` to read in our SCE objects so that they are immediately stored together in a list. @@ -210,7 +192,7 @@ That said, the integration methods we will be applying _do not actually use_ any If we have annotations, they are a helpful "bonus" for assessing the integration's performance, but they are not part of the integration itself. -## Prepare the SCE list for integration +## Merge the SCE list into one object ![Single-cell roadmap: Merge](diagrams/roadmap_multi_merge.png) @@ -220,115 +202,136 @@ A word of caution before we begin: **This merged SCE object is NOT an integrated Merging SCEs does not perform any batch correction, but just reorganizes the data to allow us to proceed to integration next. To merge SCE objects, we do need to do some wrangling and bookkeeping to ensure compatibility and that we don't lose important information. -Overall we'll want to take care of these items: +Overall, we'll want to make sure that: -1. We should be able to trace sample-specific information back to the originating sample, including... - - Cell-level information: Which sample is each cell from? - - Library-specific feature statistics, e.g., gene-level statistics for a given library found in `rowData`. - Which sample is a given feature statistic from? -2. SCE objects should contain the same genes: Each SCE object should have the same row names. -3. SCE cell metadata columns should match: The `colData` for each SCE object should have the same column names. +1. All objects have compatible dimensions. +This means that all objects should... + + Have the same genes (aka row names), in the same order + + Have the same `colData` slot columns, in the same order + + Have the same assays +2. After merging, we'll still be able to identify which sample different pieces of information came from +As we saw in the slides, this means we'll have to... + + Attach sample names to the barcodes (aka column names) + This also ensures that column names are unique; while a single sample (library) is guaranteed to have unique barcodes, technically they can be repeated across samples! + + Attach sample names to `rowData` slot column names and `metadata` field names (if you care to keep this information around - today, we will!) + + Add a new column indicating the sample to the `colData` slot +We'll approach this merge in two parts: -We'll begin by taking some time to thoroughly explore our SCE objects and figure out what wrangling steps we need to take for these specific data. -Don't skip this exploration! -Bear in mind that the exact wrangling shown here will not be the same for other SCE objects you work with, but the same general principles apply. ++ First, we'll take some time to thoroughly explore the our SCE objects to determine what wrangling we need to do to make all the objects _compatible_ for merging ++ Then, we'll write (ok, we've written it for you) a _custom function_ to format each SCE object for merging, including: + + Making any changes to ensure objects are compatible + + Adding in identifying information so we know which sample the cells and other metadata came from + + Removing the existing reduced dimension matrices (PCA and UMAP). + This is because we'll want to recalculate these matrices on the merged objects, taking batch into account +When merging objects on your own, don't skip these data exploration steps! +The steps we take to prepare our SCEs will probably be different from the steps you need to take with other SCEs, and only by carefully exploring the objects can you figure out what steps you'll need to take to meet all of our conditions. -#### Preserving sample information at the cell level -How will we be able to tell which sample a given cell came from? +### Prepare to merge SCEs -The best way to do this is simply to add a `colData` column with the sample information, so that we can know which sample each row came from. +#### Create unique cell identifiers -In addition, we want to pay some attention to the SCE object's column names (the cell ids), which must remain unique after merging since duplicate ids will cause an R error. -In this case, the SCE column names are barcodes (which is usually but not always the case in SCE objects), which are only guaranteed to be unique _within_ a sample but may be repeated across samples. -So, after merging, it's technically possible that multiple cells will have the same barcode. -This would be a problem for two reasons: -First, the cell id would not be able to point us back to cell's originating sample. -Second, it would literally cause an error in R, which does not allow duplicate column names. +As part of the custom function we'll write, we'll include a step to create unique cell identifiers by attaching sample names to the SCE column names (cell barcodes). +For example, we would update the column name for a cell from `Sample1` with the barcode `ACGT` to `Sample1-ACGT`. +When merging, there can't be any duplicate column names (barcodes) across _all_ the objects or R will throw an error. +While you're guaranteed to have unique barcodes in a given SCE object, there is _no guarantee_ that they are unique across multiple samples - it is absolutely possible to have cells from two different samples share the same barcode (and we've seen it happen!). -One way to ensure that cell ids remain unique even after merging is to actually modify them by _prepending_ the relevant sample name. -For example, consider these barcodes for the `SCPCL000479` sample: +Adding the sample id to the column names (barcodes) is therefore a crucial step in our merging bookkeeping. -```{r barcodes} -# Look at the column names for the `SCPCL000479` sample, for example -colnames(sce_list$SCPCL000479) |> - # Only print out the first 6 for convenience - head() -``` -These ids will be updated to `SCPCL000479-GGGACCTCAAGCGGAT`, `SCPCL000479-CACAGATAGTGAGTGC`, and so on, thereby ensuring fully unique ids for all cells across all samples. +#### Explore the SCE objects -#### Preserving sample information at the gene level +##### Check the genes -The `rowData` table in SCE objects will often contain both "general" and "library-specific" information, for example: +First, we'll compare the object's genes (aka, their row names). +We can use some `purrr` magic to help us find the set of shared genes among all objects: -```{r rowdata} -rowData(sce_list$SCPCL000479) |> - head() +```{r shared genes} +# Define vector of shared genes +shared_genes <- sce_list |> + # get rownames (genes) for each SCE in sce_list + purrr::map(rownames) |> + # reduce to the _intersection_ among lists + purrr::reduce(intersect) + +# How many shared genes are there? +length(shared_genes) ``` -Here, the rownames are Ensembl gene ids, and columns are `gene_symbol`, `mean`, and `detected`. -The `gene_symbol` column is general information about all genes, not specific to any library or experiment, but `mean` and `detected` are library-specific gene statistics. -So, `gene_symbol` does not need to be traced back to its originating sample, but `mean` and `detected` do. -To this end, we can take a similar approach to what we'll do for cell ids: -We can change the sample-specific `rowData` column names by prepending the sample name. -For example, rather than being called `mean`, this column will be named `SCPCL000479-mean` for the `SCPCL000479` sample. +That's quite a lot! +In fact, because these objects were all uniformly processed by the same workflow (which did not filter out any genes!), we expect them to all have the same genes. +We can map over the list to confirm that indeed, they have the same number of rows (genes): -All our SCE objects have the same `rowData` columns (as we can see in the next chunk), so we'll perform this renaming across all SCEs. -```{r compare rowdata, live = TRUE} -# Use `purrr::map()` to quickly extract rowData column names for all SCEs +```{r check shared genes, live = TRUE} +# The number of genes in an SCE corresponds to its number of rows: ``` +Even though we know the genes already match, we need to also be sure they are in the same _order_ among all objects. +So, we'll hold onto that `shared_genes` variable we defined and use it soon in our custom formatting function to make sure all objects fully match. -#### Ensuring that only shared genes are used +It's worth noting that the intersection isn't the only option here, though! +Using the intersection means a lot of genes will get discarded if the objects have different genes. +We could instead take the _union_ of genes so nothing gets thrown out. +In this case, you'd need to create "dummy" assay rows for genes that a given SCE doesn't have and fill it with `NA` expression values. +You'll still have to make sure the SCEs have the same rows in the same order before merging, so you may need to do a decent bit of matrix wrangling. -The next step in ensuring SCE compatibility is to make sure they all contain the same genes, which are stored as the SCE object's row names (these names are also found the `rowData` slot's row names). -Here, those gene ids are unique Ensembl gene ids. +##### Check the `colData` column names -We can use some `purrr` magic to quickly find the set of shared genes among our samples, and then ask how many there are. +Next up, we'll check the `colData` columns: we need these to be the same, and in the same order. +Let's print out each object's `colData` column name to see where we stand: -```{r shared genes} -# Define vector of shared genes -shared_genes <- sce_list |> - # get rownames (genes) for each SCE in sce_list - purrr::map(rownames) |> - # reduce to the _intersection_ among lists - purrr::reduce(intersect) +```{r coldata colnames} +sce_list |> + purrr::map( + \(sce) colnames(colData(sce)) + ) ``` +We see the same columns all around in the same order, which is great! -```{r print shared genes, live = TRUE} -# How many shared genes are there? +But what if there were different columns across objects, or they were differently ordered? +In that case, we could find the intersection of column names like we did above for genes, and use that to re-order and subset all `colData` slots in our custom formatting function. -``` -In this case, we happen to know that all SCE objects we're working with already contained the same genes. -We do a quick-and-dirty check for this by looking at the number of rows across SCE objects, and we'll see that they are all the same: +##### Check the assays -```{r check shared genes, live = TRUE} -# The number of genes in an SCE corresponds to its number of rows: +Next, we'll make sure that all objects share the same assays: + +```{r assay names, live = TRUE} +# print all the assay names ``` +Again, all objects are compatible already with both having a `counts` and `logcounts` assay. -So, for our data, we will not have to subset to shared genes since they are already shared! +In your own data exploration, if you encounter SCEs to merge that have extraneous assays that you don't need, you can remove them by setting them to `NULL` in your custom formatting function, e.g. `assay(sce, "assay_to_remove") <- NULL`. -#### Ensuring matching columns in `colData` +##### Check the `rowData` contents -Finally, we'll need to have the same column names across all SCE `colData` tables, so let's look at all those column names. -We can use similar syntax here to what we used to look at all the `rowData` column names. +One of the other items we said we'd need to think about is the `rowData`, which contains gene metadata. +This slot is interesting because some of its columns are specific to the given sample, while others are general: -```{r compare coldata} -purrr::map(sce_list, - \(sce) colnames(colData(sce)) ) +```{r little head rowdata} +sce_list |> + purrr::map( + \(sce) head(rowData(sce), 3) # only print 3 rows for space! + ) ``` -It looks like the column names are all already matching among SCEs, so there's no specific preparation we'll need to do there. +The column `gene_symbol` is not sample-specific - it just provides the corresponding gene symbol to the Ensembl ids seen here as row names. +The columns `mean` and `detected`, however, are sample-specific - they contain sample-specific statistics about gene expression. -### Perform SCE merging +This means we definitely need to update the column names `mean` and `detected` to include the sample id. +But, we don't need a separate `gene_symbol` column for each sample, so we can leave that one alone as just `gene_symbol`. +Once we eventually merge, only one `gene_symbol` column will be left in the final object since it is the same across all the SCEs. + +We'll show one way to do this in our custom function, but it's worth noting there's nothing _wrong_ with also adding the sample id to the `gene_symbol` column; you'll just end up with a bunch of redundant gene symbol columns. + + +#### Reformat the SCE objects As you can see, there's a lot of moving parts to consider! Again, these moving parts may (will!) differ for SCEs that you are working with, so you have to explore your own SCEs in depth to prepare for merging. @@ -343,33 +346,61 @@ We'll then use our new `purrr::map()` programming skills to run this function ov This will give us a new list of formatted SCEs that we can proceed to merge. It's important to remember that the `format_sce()` function written below is not a function for general use – it's been precisely written to match the processing we need to do for _these_ SCEs, and different SCEs you work with will require different types of processing. +We also include roxygen-style comments for this function, which can be a helpful consistent way to document your code if you like it - we've even written a blog post about it :) (). + ```{r format_sce function} -format_sce <- function(sce, sample_name) { - # Input arguments: - ## sce: An SCE object to format - ## sample_name: The SCE object's name - # This function returns a formatted SCE object. - - ###### Ensure that we can identify the originating sample information ###### - # Add a column called `sample` that stores this information - # This will be stored in `colData` +#' Custom function to format an SCE before merging +#' +#' @param sce SCE object to format +#' @param sample_name Name of the sample +#' @param shared_genes Vector of shared genes across all SCE objects +#' +#' @returns An updated SCE object ready for merging +format_sce <- function( + sce, + sample_name, + shared_genes +) { + + ### Remove the single-sample reduced dimensions + # We do this first since it makes the object a lot smaller for the rest of this code! + reducedDims(sce) <- NULL + + ### Add dedicated sample indicator column to the colData slot + # Recall, the `sce$` shortcut points to the colData sce$sample <- sample_name - - ###### Ensure cell ids will be unique ###### - # Update the SCE object column names (cell ids) by prepending `sample_name` + ### Ensure objects have the same genes in the same order + # Use the shared_genes vector to index genes to the intersection + # Doing this both subsets to just those genes, and reorders! + sce <- sce[shared_genes, ] + + ### There is no additional wrangling to do for the colData column names or assays. + ### But if there were, you could add your custom code to do so here. + ### Your custom function may need additional arguments for this, too. + + ### Ensure cell ids are identifiable and fully unique + # Update the SCE object column names (cell ids) by prepending the `sample_name` colnames(sce) <- glue::glue("{sample_name}-{colnames(sce)}") - - ###### Ensure gene-level statistics can be identified in `rowData` ###### - # We want to rename the columns `mean` and `detected` to contain the `sample_name` - # Recall the names are: "gene_symbol", "mean", "detected" - colnames(rowData(sce)) <- c("gene_symbol", - glue::glue("{sample_name}-mean"), - glue::glue("{sample_name}-detected")) - - # Return the formatted SCE object - return(sce) + ### Ensure the rowData columns can be identified + # Recall, we want to leave `gene_symbol` alone, but add the `sample_name` to the rest + rowdata_names <- colnames(rowData(sce)) + # prefix rowData names with the sample name, except for gene symbols + new_rowdata_names <- ifelse( + rowdata_names == "gene_symbol", + "gene_symbol", + glue::glue("{sample_name}-{rowdata_names}") + ) + colnames(rowData(sce)) <- new_rowdata_names + + ### Ensure metadata slot fields can be identified + # We'll simply prepend the `sample_name` to all fields for this slot + names(metadata(sce)) <- glue::glue("{sample_name}-{names(metadata(sce))}") + + + ### Finally, we can return the formatted SCE object + return(sce) } ``` @@ -382,15 +413,16 @@ In our case, we want to run `format_sce()` over paired `sce_list` items and `sce # Each "iteration" will march down the first two # arguments `sce_list` and `names(sce_list)` in order - # Name of the function to run - -# Print resulting list +# Print formatted SCE list ``` (Psst, like `purrr` and want to dive deeper? Check out [the `purrr::imap()` function](https://purrr.tidyverse.org/reference/imap.html)!) +### Perform the merging + + At long last, we are ready to merge the SCEs, which we'll do using the R function `cbind()`. The `cbind()` function is often used to combine data frames or matrices by column, i.e. "stack" them next to each other. The same principle applies here, but when run on SCE objects, `cbind()` will create a new SCE object by combining `counts` and `logcounts` matrices by column. @@ -405,7 +437,7 @@ Since we need to apply `cbind()` to a _list_ of objects, we need to use some sli ``` -We now have a single SCE object that contains all cells from all samples we'd like to integrate. +We now have a single merged SCE object that contains all cells from all samples we'd like to integrate. Let's take a peek at some of the innards of this new SCE object: @@ -419,54 +451,28 @@ Let's take a peek at some of the innards of this new SCE object: ``` -## Integration +## Integrate samples ![Single-cell roadmap: Integrate](diagrams/roadmap_multi_integrate.png) - So far, we've created a `merged_sce` object which is (almost!) ready for integration. The integration methods we'll be using here actually perform batch correction on a reduced dimension representation of the normalized gene expression values, which is more efficient. `fastMNN` and `harmony` specifically use PCA for this, but be aware that different integration methods may use other kinds of reduced dimensions. -You'll notice that the merged SCE object object already contains PCA and UMAP reduced dimensions, which were calculated during our pre-processing: - -```{r merged_sce reddim, live = TRUE} -# Print the reducedDimNames of the merged_sce - -``` - -These represent the original dimension reductions that were performed on _each individual SCE_ before merging, but we actually need to calculate PCA (and UMAP for visualization) from the merged object directly. - -Why can't we use the sample-specific PCA and UMAP matrices? -Part of these calculations themselves involves scaling the raw data to center the mean. -When samples are separately centered but plotting together, you will see samples "overlapping" in space, but this placement is actually just an artifact of the individual centering. -In addition, the mathematical relationship between the original expression data and reduced dimension version of that data will differ across samples, meaning we can't interpret them all together. -To see how this looks, let's look at the UMAP when calculated from individual samples: - -```{r plot individual UMAPs} -# Plot UMAP calculated from individual samples with separate scaling -scater::plotReducedDim(merged_sce, - dimred = "UMAP", - color_by = "sample", - point_size = 0.5, - point_alpha = 0.2) + - # Use a CVD-friendly color scheme and specify legend name - scale_color_brewer(palette = "Dark2", name = "sample") + - # Modify the legend key with larger, easier to see points - guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) + - ggtitle("UMAP calculated on each sample separately") -``` - -As we see in this UMAP, all samples are centered at zero and all overlapping. -This visual artifact can give the _incorrect impression_ that data is integrated - to be clear, this data is NOT integrated! +Before merging, our objects had reduced dimension representations calculated on each individual SCE, and we removed them when preparing for merge. +We removed them because we don't actually want to use them anymore! +This is because part of their calculation involves scaling the raw data to center the mean. +When samples are separately centered, _all_ of them will be centered at zero, making it look like the datasets are already pretty overlapping when you plot their UMAPs together. +But, this is just a mathematical artifact of how dimension reduction is performed. +So, we'll begin by re-calculating PCA and UMAP on the merged object in a way that takes batches into consideration. For input to integration, we'll want the reduced dimension calculations to consider normalized gene expression values from all samples simultaneously. So we'll need to recalculate PCA (and UMAP for visualization) on the merged object. -We'll also save these new reduced dimensions with different names, `merged_PCA` and `merged_UMAP`, to distinguish them from already-present `PCA` and `UMAP`. First, as usual, we'll determine the high-variance genes to use for PCA from the `merged_sce` object. For this, we'll need to provide the argument `block = merged_sce$sample` when modeling gene variance, which tells `scran::modelGeneVar()` to first model variance separately for each batch and then combine those modeling statistics. +(Psst: isn't it handy we created that `sample` column when merging?!) ```{r calc merged hv genes} # Specify the number of genes to identify @@ -515,41 +521,32 @@ We can now include this PCA matrix in our `merged_sce` object: Now that we have the PCA matrix, we can proceed to calculate UMAP to visualize the uncorrected merged data. -We'll calculate UMAP as "usual", but in this case we'll specify two additional arguments: - -- `dimred = "merged_PCA"`, which specifies which existing reduced dimension should be used for the calculation. -We want to use the batch-weighted PCA, which we named above as `"merged_PCA"`. -- `name = "merged_UMAP"`, which names the final UMAP that this function calculates. -This argument will prevent us from overwriting the existing UMAP which is already named "UMAP" and instead create a separate `"merged_UMAP"`. - ```{r calculate merged umap, live = TRUE} -# add merged_UMAP from merged_PCA ``` -Now, let's see how this new `merged_UMAP` looks compared to the `UMAP` calculated from individual samples: - ```{r plot uncorrected merged UMAP} # UMAPs scaled together when calculated from the merged SCE -scater::plotReducedDim(merged_sce, - dimred = "merged_UMAP", - color_by = "sample", - # Some styling to help us see the points: - point_size = 0.5, - point_alpha = 0.2) + +scater::plotUMAP( + merged_sce, + color_by = "sample", + # Some styling to help us see the points: + point_size = 0.5, + point_alpha = 0.2 +) + scale_color_brewer(palette = "Dark2", name = "sample") + guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) + ggtitle("UMAP calculated on merged_sce") ``` -Samples are now separated, which more reasonably reflects that this data is _not yet batch-corrected_. +We see (mostly) four separate clumps representing the four different _merged but not yet integrated_ samples. We can think of this UMAP as our "before" UMAP, and we can compare this to the "after" UMAP we see post-integration. Let's discuss a little first: What visual differences do you think the UMAP on the integrated version of data will have? What similarities do you think the integrated UMAP will have to this plot? -### Integration with `fastMNN` +### Integrate with `fastMNN` Finally, we're ready to integrate! To start, we'll use the `fastMNN` approach from the Bioconductor [`batchelor` package](http://www.bioconductor.org/packages/release/bioc/html/batchelor.html). @@ -588,6 +585,10 @@ We're mostly interested in the PCA that `fastMNN` calculated, so let's save that ``` Finally, we'll calculate UMAP from these corrected PCA matrix for visualization. +In this case we need to specify two additional arguments since we're working with non-standard reduced dimension names: + ++ `dimred = "fastmnn_PCA"`, which specifies the existing reduced dimension to use for the calculation ++ `name = "fastmnn_UMAP"`, which names the final UMAP that this function calculates ```{r calculate fastmnn umap, live = TRUE} # Calculate UMAP @@ -612,7 +613,7 @@ scater::plotReducedDim(merged_sce, ggtitle("UMAP after integration with fastMNN") ``` -This `fastmnn_UMAP` certainly looks different from the one we made from `merged_UMAP`! +This `fastmnn_UMAP` certainly looks different from the one we made before integrating! What different trends do you see? Do all samples look "equally well" integrated, from a first look? @@ -684,12 +685,11 @@ scater::plotReducedDim(merged_sce, What trends do you observe between tumor and healthy tissues among these integrated samples? - -### Integration with `harmony` +### Integrate with `harmony` `fastMNN` is only one of many approaches to perform integration, and different methods have different capabilities and may give different results. For example, some methods can accommodate additional covariates (e.g., technology, patient, diagnosis, etc.) that can influence integration. -In fact the data we are using has a known _patient_ covariate; `SCPCL000479` and `SCPCL000480` are from the first patient, and `SCPCL000481` and `SCPCL000482` are from the second patient. +In fact the data we are using has a known _patient_ covariate; `SCPCL000479` and `SCPCL000480` are from Patient A, and `SCPCL000481` and `SCPCL000482` are from Patient B. So, let's perform integration with a method that can use this information - [`harmony`](https://portals.broadinstitute.org/harmony/)! @@ -713,7 +713,7 @@ However, unlike `fastMNN`, `harmony` does not "back-calculate" corrected express For input, `harmony` needs a couple pieces of information: - First, `harmony` takes a batch-weighted PCA matrix to perform integration. -We already calculated a batch-weighted PCA matrix (our `merged_PCA` reduced dimension), we'll provide this as the the input. +We already calculated a batch-weighted PCA matrix so we'll provide this as the the input. - Second, we need to tell `harmony` about the covariates to use - `sample` and `patient`. To do this, we provide two arguments: - `meta_data`, a data frame that contains covariates across samples. @@ -773,7 +773,7 @@ scater::plotReducedDim(merged_sce, # Specify variable for faceting other_fields = "sample") + scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") + - guides(color = guide_legend(override.aes = list(size = 3))) + + guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) + ggtitle("UMAP after integration with harmony") + facet_wrap(vars(sample)) ``` @@ -782,7 +782,7 @@ What do you now notice in this faceted view that wasn't clear previously? Are there other patterns you see that are similar or different from the `fastMNN` UMAP? How do you think `fastMNN` vs. `harmony` performed in integrating these samples? -### Export +## Export Finally, we'll export the final SCE object with both `fastMNN` and `harmony` integration to a file. Since this object is very large (over 1 GB!), we'll export it to a file with some compression, which, in this case, will reduce the final size to a smaller ~360 MB. diff --git a/scRNA-seq-advanced/02-dataset_integration.Rmd b/scRNA-seq-advanced/02-dataset_integration.Rmd index 07c9078b..849f4761 100644 --- a/scRNA-seq-advanced/02-dataset_integration.Rmd +++ b/scRNA-seq-advanced/02-dataset_integration.Rmd @@ -55,7 +55,7 @@ set.seed(12345) ``` -### Directories and files +### Define directories and files We have already prepared count data for the four samples we'll be integrating (i.e., filtered cells, normalized counts, and calculated PCA & UMAP). @@ -66,9 +66,11 @@ These SCE objects, stored as RDS files, are available in the `data/rms/processed - `SCPCL000481.rds` (Patient B) - `SCPCL000482.rds` (Patient B) +Both Patient A (18 year old male) and Patient B (4 year old female) had recurrent embryonal rhabdomyosarcoma when samples were taken. + To begin, let's set up our directories and files: -```{r directories, live = TRUE} +```{r directories} # Define directory where processed SCE objects to be integrated are stored input_dir <- file.path("data", "rms", "processed") @@ -129,29 +131,6 @@ length(histologies) purrr::map(histologies, length) ``` -One other new coding strategy we'll learn in this notebook is using the [`glue`](https://glue.tidyverse.org/) package to combine strings. -This package offers a convenient function `glue::glue()` that can be used instead of the base R `paste()` function. - -```{r paste} -# Define a variable for example: -org_name <- "Data Lab" - -# We can use paste to combine strings and variables: -paste("Welcome to the", org_name, "workshop on Advanced scRNA-seq!") -``` - -We can use `glue::glue()` to accomplish the same goal with some different syntax: - -```{r glue} -# glue::glue takes a single string argument (only one set of quotes!), and -# variables can easily be included inside {curly braces} -glue::glue("Welcome to the {org_name} workshop on Advanced scRNA-seq!") -``` - -(Note that even though the `glue::glue()` output isn't in quotes, it still behaves like a string!) - - -Alright, time for the good stuff! Let's use `purrr::map()` to read in our SCE objects so that they are immediately stored together in a list. @@ -218,7 +197,7 @@ That said, the integration methods we will be applying _do not actually use_ any If we have annotations, they are a helpful "bonus" for assessing the integration's performance, but they are not part of the integration itself. -## Prepare the SCE list for integration +## Merge the SCE list into one object ![Single-cell roadmap: Merge](diagrams/roadmap_multi_merge.png) @@ -228,78 +207,52 @@ A word of caution before we begin: **This merged SCE object is NOT an integrated Merging SCEs does not perform any batch correction, but just reorganizes the data to allow us to proceed to integration next. To merge SCE objects, we do need to do some wrangling and bookkeeping to ensure compatibility and that we don't lose important information. -Overall we'll want to take care of these items: +Overall, we'll want to make sure that: -1. We should be able to trace sample-specific information back to the originating sample, including... - - Cell-level information: Which sample is each cell from? - - Library-specific feature statistics, e.g., gene-level statistics for a given library found in `rowData`. - Which sample is a given feature statistic from? -2. SCE objects should contain the same genes: Each SCE object should have the same row names. -3. SCE cell metadata columns should match: The `colData` for each SCE object should have the same column names. +1. All objects have compatible dimensions. +This means that all objects should... + + Have the same genes (aka row names), in the same order + + Have the same `colData` slot columns, in the same order + + Have the same assays +2. After merging, we'll still be able to identify which sample different pieces of information came from +As we saw in the slides, this means we'll have to... + + Attach sample names to the barcodes (aka column names) + This also ensures that column names are unique; while a single sample (library) is guaranteed to have unique barcodes, technically they can be repeated across samples! + + Attach sample names to `rowData` slot column names and `metadata` field names (if you care to keep this information around - today, we will!) + + Add a new column indicating the sample to the `colData` slot +We'll approach this merge in two parts: -We'll begin by taking some time to thoroughly explore our SCE objects and figure out what wrangling steps we need to take for these specific data. -Don't skip this exploration! -Bear in mind that the exact wrangling shown here will not be the same for other SCE objects you work with, but the same general principles apply. - - -#### Preserving sample information at the cell level - -How will we be able to tell which sample a given cell came from? - -The best way to do this is simply to add a `colData` column with the sample information, so that we can know which sample each row came from. - -In addition, we want to pay some attention to the SCE object's column names (the cell ids), which must remain unique after merging since duplicate ids will cause an R error. -In this case, the SCE column names are barcodes (which is usually but not always the case in SCE objects), which are only guaranteed to be unique _within_ a sample but may be repeated across samples. -So, after merging, it's technically possible that multiple cells will have the same barcode. -This would be a problem for two reasons: -First, the cell id would not be able to point us back to cell's originating sample. -Second, it would literally cause an error in R, which does not allow duplicate column names. - - -One way to ensure that cell ids remain unique even after merging is to actually modify them by _prepending_ the relevant sample name. -For example, consider these barcodes for the `SCPCL000479` sample: - -```{r barcodes} -# Look at the column names for the `SCPCL000479` sample, for example -colnames(sce_list$SCPCL000479) |> - # Only print out the first 6 for convenience - head() -``` ++ First, we'll take some time to thoroughly explore the our SCE objects to determine what wrangling we need to do to make all the objects _compatible_ for merging ++ Then, we'll write (ok, we've written it for you) a _custom function_ to format each SCE object for merging, including: + + Making any changes to ensure objects are compatible + + Adding in identifying information so we know which sample the cells and other metadata came from + + Removing the existing reduced dimension matrices (PCA and UMAP). + This is because we'll want to recalculate these matrices on the merged objects, taking batch into account -These ids will be updated to `SCPCL000479-GGGACCTCAAGCGGAT`, `SCPCL000479-CACAGATAGTGAGTGC`, and so on, thereby ensuring fully unique ids for all cells across all samples. +When merging objects on your own, don't skip these data exploration steps! +The steps we take to prepare our SCEs will probably be different from the steps you need to take with other SCEs, and only by carefully exploring the objects can you figure out what steps you'll need to take to meet all of our conditions. -#### Preserving sample information at the gene level -The `rowData` table in SCE objects will often contain both "general" and "library-specific" information, for example: +### Prepare to merge SCEs -```{r rowdata} -rowData(sce_list$SCPCL000479) |> - head() -``` +#### Create unique cell identifiers -Here, the rownames are Ensembl gene ids, and columns are `gene_symbol`, `mean`, and `detected`. -The `gene_symbol` column is general information about all genes, not specific to any library or experiment, but `mean` and `detected` are library-specific gene statistics. -So, `gene_symbol` does not need to be traced back to its originating sample, but `mean` and `detected` do. -To this end, we can take a similar approach to what we'll do for cell ids: -We can change the sample-specific `rowData` column names by prepending the sample name. -For example, rather than being called `mean`, this column will be named `SCPCL000479-mean` for the `SCPCL000479` sample. +As part of the custom function we'll write, we'll include a step to create unique cell identifiers by attaching sample names to the SCE column names (cell barcodes). +For example, we would update the column name for a cell from `Sample1` with the barcode `ACGT` to `Sample1-ACGT`. -All our SCE objects have the same `rowData` columns (as we can see in the next chunk), so we'll perform this renaming across all SCEs. +When merging, there can't be any duplicate column names (barcodes) across _all_ the objects or R will throw an error. +While you're guaranteed to have unique barcodes in a given SCE object, there is _no guarantee_ that they are unique across multiple samples - it is absolutely possible to have cells from two different samples share the same barcode (and we've seen it happen!). -```{r compare rowdata, live = TRUE} -# Use `purrr::map()` to quickly extract rowData column names for all SCEs -purrr::map(sce_list, - \(sce) colnames(rowData(sce))) -``` +Adding the sample id to the column names (barcodes) is therefore a crucial step in our merging bookkeeping. -#### Ensuring that only shared genes are used +#### Explore the SCE objects -The next step in ensuring SCE compatibility is to make sure they all contain the same genes, which are stored as the SCE object's row names (these names are also found the `rowData` slot's row names). -Here, those gene ids are unique Ensembl gene ids. +##### Check the genes -We can use some `purrr` magic to quickly find the set of shared genes among our samples, and then ask how many there are. +First, we'll compare the object's genes (aka, their row names). +We can use some `purrr` magic to help us find the set of shared genes among all objects: ```{r shared genes} # Define vector of shared genes @@ -308,15 +261,15 @@ shared_genes <- sce_list |> purrr::map(rownames) |> # reduce to the _intersection_ among lists purrr::reduce(intersect) -``` -```{r print shared genes, live = TRUE} # How many shared genes are there? length(shared_genes) ``` -In this case, we happen to know that all SCE objects we're working with already contained the same genes. -We do a quick-and-dirty check for this by looking at the number of rows across SCE objects, and we'll see that they are all the same: +That's quite a lot! +In fact, because these objects were all uniformly processed by the same workflow (which did not filter out any genes!), we expect them to all have the same genes. +We can map over the list to confirm that indeed, they have the same number of rows (genes): + ```{r check shared genes, live = TRUE} # The number of genes in an SCE corresponds to its number of rows: @@ -324,21 +277,68 @@ sce_list |> purrr::map(nrow) ``` -So, for our data, we will not have to subset to shared genes since they are already shared! +Even though we know the genes already match, we need to also be sure they are in the same _order_ among all objects. +So, we'll hold onto that `shared_genes` variable we defined and use it soon in our custom formatting function to make sure all objects fully match. + +It's worth noting that the intersection isn't the only option here, though! +Using the intersection means a lot of genes will get discarded if the objects have different genes. +We could instead take the _union_ of genes so nothing gets thrown out. +In this case, you'd need to create "dummy" assay rows for genes that a given SCE doesn't have and fill it with `NA` expression values. +You'll still have to make sure the SCEs have the same rows in the same order before merging, so you may need to do a decent bit of matrix wrangling. + +##### Check the `colData` column names + +Next up, we'll check the `colData` columns: we need these to be the same, and in the same order. +Let's print out each object's `colData` column name to see where we stand: + +```{r coldata colnames} +sce_list |> + purrr::map( + \(sce) colnames(colData(sce)) + ) +``` +We see the same columns all around in the same order, which is great! -#### Ensuring matching columns in `colData` +But what if there were different columns across objects, or they were differently ordered? +In that case, we could find the intersection of column names like we did above for genes, and use that to re-order and subset all `colData` slots in our custom formatting function. -Finally, we'll need to have the same column names across all SCE `colData` tables, so let's look at all those column names. -We can use similar syntax here to what we used to look at all the `rowData` column names. -```{r compare coldata} -purrr::map(sce_list, - \(sce) colnames(colData(sce)) ) +##### Check the assays + +Next, we'll make sure that all objects share the same assays: + +```{r assay names, live = TRUE} +# print all the assay names +sce_list |> + purrr::map(assayNames) +``` +Again, all objects are compatible already with both having a `counts` and `logcounts` assay. + +In your own data exploration, if you encounter SCEs to merge that have extraneous assays that you don't need, you can remove them by setting them to `NULL` in your custom formatting function, e.g. `assay(sce, "assay_to_remove") <- NULL`. + +##### Check the `rowData` contents + +One of the other items we said we'd need to think about is the `rowData`, which contains gene metadata. +This slot is interesting because some of its columns are specific to the given sample, while others are general: + +```{r little head rowdata} +sce_list |> + purrr::map( + \(sce) head(rowData(sce), 3) # only print 3 rows for space! + ) ``` -It looks like the column names are all already matching among SCEs, so there's no specific preparation we'll need to do there. +The column `gene_symbol` is not sample-specific - it just provides the corresponding gene symbol to the Ensembl ids seen here as row names. +The columns `mean` and `detected`, however, are sample-specific - they contain sample-specific statistics about gene expression. -### Perform SCE merging +This means we definitely need to update the column names `mean` and `detected` to include the sample id. +But, we don't need a separate `gene_symbol` column for each sample, so we can leave that one alone as just `gene_symbol`. +Once we eventually merge, only one `gene_symbol` column will be left in the final object since it is the same across all the SCEs. + +We'll show one way to do this in our custom function, but it's worth noting there's nothing _wrong_ with also adding the sample id to the `gene_symbol` column; you'll just end up with a bunch of redundant gene symbol columns. + + +#### Reformat the SCE objects As you can see, there's a lot of moving parts to consider! Again, these moving parts may (will!) differ for SCEs that you are working with, so you have to explore your own SCEs in depth to prepare for merging. @@ -353,33 +353,61 @@ We'll then use our new `purrr::map()` programming skills to run this function ov This will give us a new list of formatted SCEs that we can proceed to merge. It's important to remember that the `format_sce()` function written below is not a function for general use – it's been precisely written to match the processing we need to do for _these_ SCEs, and different SCEs you work with will require different types of processing. +We also include roxygen-style comments for this function, which can be a helpful consistent way to document your code if you like it - we've even written a blog post about it :) (). + ```{r format_sce function} -format_sce <- function(sce, sample_name) { - # Input arguments: - ## sce: An SCE object to format - ## sample_name: The SCE object's name - # This function returns a formatted SCE object. - - ###### Ensure that we can identify the originating sample information ###### - # Add a column called `sample` that stores this information - # This will be stored in `colData` +#' Custom function to format an SCE before merging +#' +#' @param sce SCE object to format +#' @param sample_name Name of the sample +#' @param shared_genes Vector of shared genes across all SCE objects +#' +#' @returns An updated SCE object ready for merging +format_sce <- function( + sce, + sample_name, + shared_genes +) { + + ### Remove the single-sample reduced dimensions + # We do this first since it makes the object a lot smaller for the rest of this code! + reducedDims(sce) <- NULL + + ### Add dedicated sample indicator column to the colData slot + # Recall, the `sce$` shortcut points to the colData sce$sample <- sample_name - - ###### Ensure cell ids will be unique ###### - # Update the SCE object column names (cell ids) by prepending `sample_name` + ### Ensure objects have the same genes in the same order + # Use the shared_genes vector to index genes to the intersection + # Doing this both subsets to just those genes, and reorders! + sce <- sce[shared_genes, ] + + ### There is no additional wrangling to do for the colData column names or assays. + ### But if there were, you could add your custom code to do so here. + ### Your custom function may need additional arguments for this, too. + + ### Ensure cell ids are identifiable and fully unique + # Update the SCE object column names (cell ids) by prepending the `sample_name` colnames(sce) <- glue::glue("{sample_name}-{colnames(sce)}") - - ###### Ensure gene-level statistics can be identified in `rowData` ###### - # We want to rename the columns `mean` and `detected` to contain the `sample_name` - # Recall the names are: "gene_symbol", "mean", "detected" - colnames(rowData(sce)) <- c("gene_symbol", - glue::glue("{sample_name}-mean"), - glue::glue("{sample_name}-detected")) - - # Return the formatted SCE object - return(sce) + ### Ensure the rowData columns can be identified + # Recall, we want to leave `gene_symbol` alone, but add the `sample_name` to the rest + rowdata_names <- colnames(rowData(sce)) + # prefix rowData names with the sample name, except for gene symbols + new_rowdata_names <- ifelse( + rowdata_names == "gene_symbol", + "gene_symbol", + glue::glue("{sample_name}-{rowdata_names}") + ) + colnames(rowData(sce)) <- new_rowdata_names + + ### Ensure metadata slot fields can be identified + # We'll simply prepend the `sample_name` to all fields for this slot + names(metadata(sce)) <- glue::glue("{sample_name}-{names(metadata(sce))}") + + + ### Finally, we can return the formatted SCE object + return(sce) } ``` @@ -393,17 +421,19 @@ sce_list_formatted <- purrr::map2( # arguments `sce_list` and `names(sce_list)` in order sce_list, names(sce_list), - # Name of the function to run - format_sce + \(sce, sample_name) format_sce(sce, sample_name, shared_genes) ) -# Print resulting list +# Print formatted SCE list sce_list_formatted ``` (Psst, like `purrr` and want to dive deeper? Check out [the `purrr::imap()` function](https://purrr.tidyverse.org/reference/imap.html)!) +### Perform the merging + + At long last, we are ready to merge the SCEs, which we'll do using the R function `cbind()`. The `cbind()` function is often used to combine data frames or matrices by column, i.e. "stack" them next to each other. The same principle applies here, but when run on SCE objects, `cbind()` will create a new SCE object by combining `counts` and `logcounts` matrices by column. @@ -419,70 +449,45 @@ merged_sce <- do.call(cbind, sce_list_formatted) merged_sce ``` -We now have a single SCE object that contains all cells from all samples we'd like to integrate. +We now have a single merged SCE object that contains all cells from all samples we'd like to integrate. Let's take a peek at some of the innards of this new SCE object: ```{r explore merged_sce, live = TRUE} # How many samples, and cells per sample? -table( colData(merged_sce)$sample ) +table(colData(merged_sce)$sample) # What are the new cell ids (column names)? -head( colnames(merged_sce) ) +head(colnames(merged_sce)) +tail(colnames(merged_sce)) # What does rowData look like? -head( rowData(merged_sce) ) +head(rowData(merged_sce)) ``` -## Integration +## Integrate samples ![Single-cell roadmap: Integrate](diagrams/roadmap_multi_integrate.png) - So far, we've created a `merged_sce` object which is (almost!) ready for integration. The integration methods we'll be using here actually perform batch correction on a reduced dimension representation of the normalized gene expression values, which is more efficient. `fastMNN` and `harmony` specifically use PCA for this, but be aware that different integration methods may use other kinds of reduced dimensions. -You'll notice that the merged SCE object object already contains PCA and UMAP reduced dimensions, which were calculated during our pre-processing: - -```{r merged_sce reddim, live = TRUE} -# Print the reducedDimNames of the merged_sce -reducedDimNames(merged_sce) -``` - -These represent the original dimension reductions that were performed on _each individual SCE_ before merging, but we actually need to calculate PCA (and UMAP for visualization) from the merged object directly. - -Why can't we use the sample-specific PCA and UMAP matrices? -Part of these calculations themselves involves scaling the raw data to center the mean. -When samples are separately centered but plotting together, you will see samples "overlapping" in space, but this placement is actually just an artifact of the individual centering. -In addition, the mathematical relationship between the original expression data and reduced dimension version of that data will differ across samples, meaning we can't interpret them all together. -To see how this looks, let's look at the UMAP when calculated from individual samples: - -```{r plot individual UMAPs} -# Plot UMAP calculated from individual samples with separate scaling -scater::plotReducedDim(merged_sce, - dimred = "UMAP", - color_by = "sample", - point_size = 0.5, - point_alpha = 0.2) + - # Use a CVD-friendly color scheme and specify legend name - scale_color_brewer(palette = "Dark2", name = "sample") + - # Modify the legend key with larger, easier to see points - guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) + - ggtitle("UMAP calculated on each sample separately") -``` - -As we see in this UMAP, all samples are centered at zero and all overlapping. -This visual artifact can give the _incorrect impression_ that data is integrated - to be clear, this data is NOT integrated! +Before merging, our objects had reduced dimension representations calculated on each individual SCE, and we removed them when preparing for merge. +We removed them because we don't actually want to use them anymore! +This is because part of their calculation involves scaling the raw data to center the mean. +When samples are separately centered, _all_ of them will be centered at zero, making it look like the datasets are already pretty overlapping when you plot their UMAPs together. +But, this is just a mathematical artifact of how dimension reduction is performed. +So, we'll begin by re-calculating PCA and UMAP on the merged object in a way that takes batches into consideration. For input to integration, we'll want the reduced dimension calculations to consider normalized gene expression values from all samples simultaneously. So we'll need to recalculate PCA (and UMAP for visualization) on the merged object. -We'll also save these new reduced dimensions with different names, `merged_PCA` and `merged_UMAP`, to distinguish them from already-present `PCA` and `UMAP`. First, as usual, we'll determine the high-variance genes to use for PCA from the `merged_sce` object. For this, we'll need to provide the argument `block = merged_sce$sample` when modeling gene variance, which tells `scran::modelGeneVar()` to first model variance separately for each batch and then combine those modeling statistics. +(Psst: isn't it handy we created that `sample` column when merging?!) ```{r calc merged hv genes} # Specify the number of genes to identify @@ -529,48 +534,37 @@ We can now include this PCA matrix in our `merged_sce` object: ```{r add merged_pca, live = TRUE} # add PCA results to merged SCE object -reducedDim(merged_sce, "merged_PCA") <- merged_pca[[1]] +reducedDim(merged_sce, "PCA") <- merged_pca[[1]] ``` Now that we have the PCA matrix, we can proceed to calculate UMAP to visualize the uncorrected merged data. -We'll calculate UMAP as "usual", but in this case we'll specify two additional arguments: - -- `dimred = "merged_PCA"`, which specifies which existing reduced dimension should be used for the calculation. -We want to use the batch-weighted PCA, which we named above as `"merged_PCA"`. -- `name = "merged_UMAP"`, which names the final UMAP that this function calculates. -This argument will prevent us from overwriting the existing UMAP which is already named "UMAP" and instead create a separate `"merged_UMAP"`. - ```{r calculate merged umap, live = TRUE} -# add merged_UMAP from merged_PCA -merged_sce <- scater::runUMAP(merged_sce, - dimred = "merged_PCA", - name = "merged_UMAP") +merged_sce <- scater::runUMAP(merged_sce) ``` -Now, let's see how this new `merged_UMAP` looks compared to the `UMAP` calculated from individual samples: - ```{r plot uncorrected merged UMAP} # UMAPs scaled together when calculated from the merged SCE -scater::plotReducedDim(merged_sce, - dimred = "merged_UMAP", - color_by = "sample", - # Some styling to help us see the points: - point_size = 0.5, - point_alpha = 0.2) + +scater::plotUMAP( + merged_sce, + color_by = "sample", + # Some styling to help us see the points: + point_size = 0.5, + point_alpha = 0.2 +) + scale_color_brewer(palette = "Dark2", name = "sample") + guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) + ggtitle("UMAP calculated on merged_sce") ``` -Samples are now separated, which more reasonably reflects that this data is _not yet batch-corrected_. +We see (mostly) four separate clumps representing the four different _merged but not yet integrated_ samples. We can think of this UMAP as our "before" UMAP, and we can compare this to the "after" UMAP we see post-integration. Let's discuss a little first: What visual differences do you think the UMAP on the integrated version of data will have? What similarities do you think the integrated UMAP will have to this plot? -### Integration with `fastMNN` +### Integrate with `fastMNN` Finally, we're ready to integrate! To start, we'll use the `fastMNN` approach from the Bioconductor [`batchelor` package](http://www.bioconductor.org/packages/release/bioc/html/batchelor.html). @@ -613,6 +607,10 @@ reducedDim(merged_sce, "fastmnn_PCA") <- reducedDim(integrated_sce, "corrected") ``` Finally, we'll calculate UMAP from these corrected PCA matrix for visualization. +In this case we need to specify two additional arguments since we're working with non-standard reduced dimension names: + ++ `dimred = "fastmnn_PCA"`, which specifies the existing reduced dimension to use for the calculation ++ `name = "fastmnn_UMAP"`, which names the final UMAP that this function calculates ```{r calculate fastmnn umap, live = TRUE} # Calculate UMAP @@ -641,7 +639,7 @@ scater::plotReducedDim(merged_sce, ggtitle("UMAP after integration with fastMNN") ``` -This `fastmnn_UMAP` certainly looks different from the one we made from `merged_UMAP`! +This `fastmnn_UMAP` certainly looks different from the one we made before integrating! What different trends do you see? Do all samples look "equally well" integrated, from a first look? @@ -713,12 +711,11 @@ scater::plotReducedDim(merged_sce, What trends do you observe between tumor and healthy tissues among these integrated samples? - -### Integration with `harmony` +### Integrate with `harmony` `fastMNN` is only one of many approaches to perform integration, and different methods have different capabilities and may give different results. For example, some methods can accommodate additional covariates (e.g., technology, patient, diagnosis, etc.) that can influence integration. -In fact the data we are using has a known _patient_ covariate; `SCPCL000479` and `SCPCL000480` are from the first patient, and `SCPCL000481` and `SCPCL000482` are from the second patient. +In fact the data we are using has a known _patient_ covariate; `SCPCL000479` and `SCPCL000480` are from Patient A, and `SCPCL000481` and `SCPCL000482` are from Patient B. So, let's perform integration with a method that can use this information - [`harmony`](https://portals.broadinstitute.org/harmony/)! @@ -742,7 +739,7 @@ However, unlike `fastMNN`, `harmony` does not "back-calculate" corrected express For input, `harmony` needs a couple pieces of information: - First, `harmony` takes a batch-weighted PCA matrix to perform integration. -We already calculated a batch-weighted PCA matrix (our `merged_PCA` reduced dimension), we'll provide this as the the input. +We already calculated a batch-weighted PCA matrix so we'll provide this as the the input. - Second, we need to tell `harmony` about the covariates to use - `sample` and `patient`. To do this, we provide two arguments: - `meta_data`, a data frame that contains covariates across samples. @@ -755,7 +752,7 @@ Let's go! ```{r run harmony, live = TRUE} # Run harmony integration harmony_pca <- harmony::RunHarmony( - data_mat = reducedDim(merged_sce, "merged_PCA"), + data_mat = reducedDim(merged_sce, "PCA"), meta_data = colData(merged_sce), vars_use = c("sample", "patient") ) @@ -809,7 +806,7 @@ scater::plotReducedDim(merged_sce, # Specify variable for faceting other_fields = "sample") + scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") + - guides(color = guide_legend(override.aes = list(size = 3))) + + guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) + ggtitle("UMAP after integration with harmony") + facet_wrap(vars(sample)) ``` @@ -818,7 +815,7 @@ What do you now notice in this faceted view that wasn't clear previously? Are there other patterns you see that are similar or different from the `fastMNN` UMAP? How do you think `fastMNN` vs. `harmony` performed in integrating these samples? -### Export +## Export Finally, we'll export the final SCE object with both `fastMNN` and `harmony` integration to a file. Since this object is very large (over 1 GB!), we'll export it to a file with some compression, which, in this case, will reduce the final size to a smaller ~360 MB. @@ -826,9 +823,11 @@ This will take a couple minutes to save while compression is performed. ```{r save integration, live = TRUE} # Export to RDS file with "gz" compression -readr::write_rds(merged_sce, - integrated_sce_file, - compress = "gz") +readr::write_rds( + merged_sce, + integrated_sce_file, + compress = "gz" +) ``` diff --git a/scRNA-seq-advanced/02-dataset_integration.nb.html b/scRNA-seq-advanced/02-dataset_integration.nb.html index 1b5b26bf..28d07264 100644 --- a/scRNA-seq-advanced/02-dataset_integration.nb.html +++ b/scRNA-seq-advanced/02-dataset_integration.nb.html @@ -3075,8 +3075,8 @@

Set up

-
-

Directories and files

+
+

Define directories and files

We have already prepared count data for the four samples we’ll be integrating (i.e., filtered cells, normalized counts, and calculated PCA & UMAP). These SCE objects, stored as RDS files, are available in @@ -3088,6 +3088,8 @@

Directories and files

  • SCPCL000481.rds (Patient B)
  • SCPCL000482.rds (Patient B)
  • +

    Both Patient A (18 year old male) and Patient B (4 year old female) +had recurrent embryonal rhabdomyosarcoma when samples were taken.

    To begin, let’s set up our directories and files:

    @@ -3205,50 +3207,8 @@

    Directories and files

    -

    One other new coding strategy we’ll learn in this notebook is using -the glue package -to combine strings. This package offers a convenient function -glue::glue() that can be used instead of the base R -paste() function.

    - - - -
    # Define a variable for example:
    -org_name <- "Data Lab"
    -
    -# We can use paste to combine strings and variables:
    -paste("Welcome to the", org_name, "workshop on Advanced scRNA-seq!")
    - - -
    [1] "Welcome to the Data Lab workshop on Advanced scRNA-seq!"
    - - -
    Welcome to the Data Lab workshop on Advanced scRNA-seq!
    - - - -

    We can use glue::glue() to accomplish the same goal with -some different syntax:

    - - - -
    # glue::glue takes a single string argument (only one set of quotes!), and
    -#  variables can easily be included inside {curly braces}
    -glue::glue("Welcome to the {org_name} workshop on Advanced scRNA-seq!")
    - - -
    Welcome to the Data Lab workshop on Advanced scRNA-seq!
    - - -
    Welcome to the Data Lab workshop on Advanced scRNA-seq!
    - - - -

    (Note that even though the glue::glue() output isn’t in -quotes, it still behaves like a string!)

    -

    Alright, time for the good stuff! Let’s use purrr::map() -to read in our SCE objects so that they are immediately stored together -in a list.

    +

    Let’s use purrr::map() to read in our SCE objects so +that they are immediately stored together in a list.

    We’ll first need to define a vector of the file paths to read in. We’ll start by creating a vector of sample names themselves and then formatting them into the correct paths. This way (foreshadowing!) we @@ -3416,8 +3376,8 @@

    Directories and files

    performance, but they are not part of the integration itself.

    -
    -

    Prepare the SCE list for integration

    +
    +

    Merge the SCE list into one object

    Single-cell roadmap: Merge
    Single-cell roadmap: Merge
    @@ -3430,179 +3390,88 @@

    Prepare the SCE list for integration

    integration next.

    To merge SCE objects, we do need to do some wrangling and bookkeeping to ensure compatibility and that we don’t lose important information. -Overall we’ll want to take care of these items:

    +Overall, we’ll want to make sure that:

      -
    1. We should be able to trace sample-specific information back to the -originating sample, including… +
    2. All objects have compatible dimensions. This means that all objects +should…
        -
      • Cell-level information: Which sample is each cell from?
      • -
      • Library-specific feature statistics, e.g., gene-level statistics for -a given library found in rowData. Which sample is a given -feature statistic from?
      • +
      • Have the same genes (aka row names), in the same order
      • +
      • Have the same colData slot columns, in the same +order
      • +
      • Have the same assays
      • +
    3. +
    4. After merging, we’ll still be able to identify which sample +different pieces of information came from As we saw in the slides, this +means we’ll have to… +
        +
      • Attach sample names to the barcodes (aka column names) This also +ensures that column names are unique; while a single sample (library) is +guaranteed to have unique barcodes, technically they can be repeated +across samples!
      • +
      • Attach sample names to rowData slot column names and +metadata field names (if you care to keep this information +around - today, we will!)
      • +
      • Add a new column indicating the sample to the colData +slot
    5. -
    6. SCE objects should contain the same genes: Each SCE object should -have the same row names.
    7. -
    8. SCE cell metadata columns should match: The colData for -each SCE object should have the same column names.
    -

    We’ll begin by taking some time to thoroughly explore our SCE objects -and figure out what wrangling steps we need to take for these specific -data. Don’t skip this exploration! Bear in mind that the exact wrangling -shown here will not be the same for other SCE objects you work with, but -the same general principles apply.

    -
    -

    Preserving sample information at the cell level

    -

    How will we be able to tell which sample a given cell came from?

    -

    The best way to do this is simply to add a colData -column with the sample information, so that we can know which sample -each row came from.

    -

    In addition, we want to pay some attention to the SCE object’s column -names (the cell ids), which must remain unique after merging since -duplicate ids will cause an R error. In this case, the SCE column names -are barcodes (which is usually but not always the case in SCE objects), -which are only guaranteed to be unique within a sample but may -be repeated across samples. So, after merging, it’s technically possible -that multiple cells will have the same barcode. This would be a problem -for two reasons: First, the cell id would not be able to point us back -to cell’s originating sample. Second, it would literally cause an error -in R, which does not allow duplicate column names.

    -

    One way to ensure that cell ids remain unique even after merging is -to actually modify them by prepending the relevant sample name. -For example, consider these barcodes for the SCPCL000479 -sample:

    - - - -
    # Look at the column names for the `SCPCL000479` sample, for example
    -colnames(sce_list$SCPCL000479) |>
    -  # Only print out the first 6 for convenience
    -  head()
    - - -
    [1] "GGGACCTCAAGCGGAT" "CACAGATAGTGAGTGC" "TGTGGCGGTGAATTGA" "GCCGATGGTACATACC"
    -[5] "ATTATCCCAGTTGGTT" "TCCGAAATCACACCGG"
    - - -
    GGGACCTCAAGCGGAT
    -CACAGATAGTGAGTGC
    -TGTGGCGGTGAATTGA
    -GCCGATGGTACATACC
    -ATTATCCCAGTTGGTT
    -TCCGAAATCACACCGG
    - - - -

    These ids will be updated to -SCPCL000479-GGGACCTCAAGCGGAT, -SCPCL000479-CACAGATAGTGAGTGC, and so on, thereby ensuring -fully unique ids for all cells across all samples.

    -
    -
    -

    Preserving sample information at the gene level

    -

    The rowData table in SCE objects will often contain both -“general” and “library-specific” information, for example:

    - - - -
    rowData(sce_list$SCPCL000479) |>
    -  head()
    - - -
    DataFrame with 6 rows and 3 columns
    -                gene_symbol       mean  detected
    -                <character>  <numeric> <numeric>
    -ENSG00000000003      TSPAN6 0.01772018  1.639778
    -ENSG00000000005        TNMD 0.00264480  0.158688
    -ENSG00000000419        DPM1 0.07299656  6.109495
    -ENSG00000000457       SCYL3 0.02300979  1.983602
    -ENSG00000000460    C1orf112 0.08119545  6.426871
    -ENSG00000000938         FGR 0.00317376  0.238032
    - - - -

    Here, the rownames are Ensembl gene ids, and columns are -gene_symbol, mean, and detected. -The gene_symbol column is general information about all -genes, not specific to any library or experiment, but mean -and detected are library-specific gene statistics. So, -gene_symbol does not need to be traced back to its -originating sample, but mean and detected do. -To this end, we can take a similar approach to what we’ll do for cell -ids: We can change the sample-specific rowData column names -by prepending the sample name. For example, rather than being called -mean, this column will be named -SCPCL000479-mean for the SCPCL000479 -sample.

    -

    All our SCE objects have the same rowData columns (as we -can see in the next chunk), so we’ll perform this renaming across all -SCEs.

    - - - -
    # Use `purrr::map()` to quickly extract rowData column names for all SCEs
    -purrr::map(sce_list,
    -           \(sce) colnames(rowData(sce)))
    - - -
    $SCPCL000479
    -[1] "gene_symbol" "mean"        "detected"   
    -
    -$SCPCL000480
    -[1] "gene_symbol" "mean"        "detected"   
    -
    -$SCPCL000481
    -[1] "gene_symbol" "mean"        "detected"   
    -
    -$SCPCL000482
    -[1] "gene_symbol" "mean"        "detected"   
    - - -
    gene_symbol
    -mean
    -detected
    - - -
    gene_symbol
    -mean
    -detected
    - - -
    gene_symbol
    -mean
    -detected
    - - -
    gene_symbol
    -mean
    -detected
    - - - +

    We’ll approach this merge in two parts:

    +
      +
    • First, we’ll take some time to thoroughly explore the our SCE +objects to determine what wrangling we need to do to make all the +objects compatible for merging
    • +
    • Then, we’ll write (ok, we’ve written it for you) a custom +function to format each SCE object for merging, including: +
        +
      • Making any changes to ensure objects are compatible
      • +
      • Adding in identifying information so we know which sample the cells +and other metadata came from
      • +
      • Removing the existing reduced dimension matrices (PCA and UMAP). +This is because we’ll want to recalculate these matrices on the merged +objects, taking batch into account
      • +
    • +
    +

    When merging objects on your own, don’t skip these data exploration +steps! The steps we take to prepare our SCEs will probably be different +from the steps you need to take with other SCEs, and only by carefully +exploring the objects can you figure out what steps you’ll need to take +to meet all of our conditions.

    +
    +

    Prepare to merge SCEs

    +
    +

    Create unique cell identifiers

    +

    As part of the custom function we’ll write, we’ll include a step to +create unique cell identifiers by attaching sample names to the SCE +column names (cell barcodes). For example, we would update the column +name for a cell from Sample1 with the barcode +ACGT to Sample1-ACGT.

    +

    When merging, there can’t be any duplicate column names (barcodes) +across all the objects or R will throw an error. While you’re +guaranteed to have unique barcodes in a given SCE object, there is +no guarantee that they are unique across multiple samples - it +is absolutely possible to have cells from two different samples share +the same barcode (and we’ve seen it happen!).

    +

    Adding the sample id to the column names (barcodes) is therefore a +crucial step in our merging bookkeeping.

    -
    -

    Ensuring that only shared genes are used

    -

    The next step in ensuring SCE compatibility is to make sure they all -contain the same genes, which are stored as the SCE object’s row names -(these names are also found the rowData slot’s row names). -Here, those gene ids are unique Ensembl gene ids.

    -

    We can use some purrr magic to quickly find the set of -shared genes among our samples, and then ask how many there are.

    +
    +

    Explore the SCE objects

    +
    +
    Check the genes
    +

    First, we’ll compare the object’s genes (aka, their row names). We +can use some purrr magic to help us find the set of shared +genes among all objects:

    - +
    # Define vector of shared genes
     shared_genes <- sce_list |>
       # get rownames (genes) for each SCE in sce_list
       purrr::map(rownames) |>
       # reduce to the _intersection_ among lists
    -  purrr::reduce(intersect)
    - - - - - - -
    # How many shared genes are there?
    +  purrr::reduce(intersect)
    +
    +# How many shared genes are there?
     length(shared_genes)
    @@ -3610,10 +3479,10 @@

    Ensuring that only shared genes are used

    -

    In this case, we happen to know that all SCE objects we’re working -with already contained the same genes. We do a quick-and-dirty check for -this by looking at the number of rows across SCE objects, and we’ll see -that they are all the same:

    +

    That’s quite a lot! In fact, because these objects were all uniformly +processed by the same workflow (which did not filter out any genes!), we +expect them to all have the same genes. We can map over the list to +confirm that indeed, they have the same number of rows (genes):

    @@ -3636,20 +3505,32 @@

    Ensuring that only shared genes are used

    -

    So, for our data, we will not have to subset to shared genes since -they are already shared!

    +

    Even though we know the genes already match, we need to also be sure +they are in the same order among all objects. So, we’ll hold +onto that shared_genes variable we defined and use it soon +in our custom formatting function to make sure all objects fully +match.

    +

    It’s worth noting that the intersection isn’t the only option here, +though! Using the intersection means a lot of genes will get discarded +if the objects have different genes. We could instead take the +union of genes so nothing gets thrown out. In this case, you’d +need to create “dummy” assay rows for genes that a given SCE doesn’t +have and fill it with NA expression values. You’ll still +have to make sure the SCEs have the same rows in the same order before +merging, so you may need to do a decent bit of matrix wrangling.

    -
    -

    Ensuring matching columns in colData

    -

    Finally, we’ll need to have the same column names across all SCE -colData tables, so let’s look at all those column names. We -can use similar syntax here to what we used to look at all the -rowData column names.

    +
    +
    Check the colData column names
    +

    Next up, we’ll check the colData columns: we need these +to be the same, and in the same order. Let’s print out each object’s +colData column name to see where we stand:

    - -
    purrr::map(sce_list,
    -           \(sce) colnames(colData(sce)) )
    + +
    sce_list |>
    +  purrr::map(
    +    \(sce) colnames(colData(sce)) 
    +  ) 
    $SCPCL000479
    @@ -3734,18 +3615,138 @@ 

    Ensuring matching columns in colData

    -

    It looks like the column names are all already matching among SCEs, -so there’s no specific preparation we’ll need to do there.

    +

    We see the same columns all around in the same order, which is +great!

    +

    But what if there were different columns across objects, or they were +differently ordered? In that case, we could find the intersection of +column names like we did above for genes, and use that to re-order and +subset all colData slots in our custom formatting +function.

    -
    -

    Perform SCE merging

    +
    +
    Check the assays
    +

    Next, we’ll make sure that all objects share the same assays:

    + + + +
    # print all the assay names
    +sce_list |>
    +  purrr::map(assayNames)
    + + +
    $SCPCL000479
    +[1] "counts"    "logcounts"
    +
    +$SCPCL000480
    +[1] "counts"    "logcounts"
    +
    +$SCPCL000481
    +[1] "counts"    "logcounts"
    +
    +$SCPCL000482
    +[1] "counts"    "logcounts"
    + + +
    counts
    +logcounts
    + + +
    counts
    +logcounts
    + + +
    counts
    +logcounts
    + + +
    counts
    +logcounts
    + + + +

    Again, all objects are compatible already with both having a +counts and logcounts assay.

    +

    In your own data exploration, if you encounter SCEs to merge that +have extraneous assays that you don’t need, you can remove them by +setting them to NULL in your custom formatting function, +e.g. assay(sce, "assay_to_remove") <- NULL.

    +
    +
    +
    Check the rowData contents
    +

    One of the other items we said we’d need to think about is the +rowData, which contains gene metadata. This slot is +interesting because some of its columns are specific to the given +sample, while others are general:

    + + + +
    sce_list |>
    +  purrr::map(
    +    \(sce) head(rowData(sce), 3) # only print 3 rows for space!
    +  )
    + + +
    $SCPCL000479
    +DataFrame with 3 rows and 3 columns
    +                gene_symbol      mean  detected
    +                <character> <numeric> <numeric>
    +ENSG00000000003      TSPAN6 0.0177202  1.639778
    +ENSG00000000005        TNMD 0.0026448  0.158688
    +ENSG00000000419        DPM1 0.0729966  6.109495
    +
    +$SCPCL000480
    +DataFrame with 3 rows and 3 columns
    +                gene_symbol       mean  detected
    +                <character>  <numeric> <numeric>
    +ENSG00000000003      TSPAN6 0.05209841  4.828312
    +ENSG00000000005        TNMD 0.00855151  0.828838
    +ENSG00000000419        DPM1 0.08919879  8.301539
    +
    +$SCPCL000481
    +DataFrame with 3 rows and 3 columns
    +                gene_symbol      mean  detected
    +                <character> <numeric> <numeric>
    +ENSG00000000003      TSPAN6 0.0637984  5.989666
    +ENSG00000000005        TNMD 0.0054835  0.495624
    +ENSG00000000419        DPM1 0.2401139 20.056944
    +
    +$SCPCL000482
    +DataFrame with 3 rows and 3 columns
    +                gene_symbol      mean  detected
    +                <character> <numeric> <numeric>
    +ENSG00000000003      TSPAN6 0.0289113  2.659838
    +ENSG00000000005        TNMD 0.0100776  0.759954
    +ENSG00000000419        DPM1 0.1391046 11.217578
    + + + +

    The column gene_symbol is not sample-specific - it just +provides the corresponding gene symbol to the Ensembl ids seen here as +row names. The columns mean and detected, +however, are sample-specific - they contain sample-specific statistics +about gene expression.

    +

    This means we definitely need to update the column names +mean and detected to include the sample id. +But, we don’t need a separate gene_symbol column for each +sample, so we can leave that one alone as just gene_symbol. +Once we eventually merge, only one gene_symbol column will +be left in the final object since it is the same across all the +SCEs.

    +

    We’ll show one way to do this in our custom function, but it’s worth +noting there’s nothing wrong with also adding the sample id to +the gene_symbol column; you’ll just end up with a bunch of +redundant gene symbol columns.

    +
    +
    +
    +

    Reformat the SCE objects

    As you can see, there’s a lot of moving parts to consider! Again, these moving parts may (will!) differ for SCEs that you are working with, so you have to explore your own SCEs in depth to prepare for merging.

    Based on our exploration, here is a schematic of how one of the SCE objects will ultimately be modified into the final merged SCE:

    -

    +

    We’ll write a custom function (seen in the chunk below) tailored to our wrangling steps that prepares a single SCE object for merging. We’ll then use our new purrr::map() programming @@ -3756,35 +3757,64 @@

    Perform SCE merging

    written to match the processing we need to do for these SCEs, and different SCEs you work with will require different types of processing.

    +

    We also include roxygen-style comments for this function, which can +be a helpful consistent way to document your code if you like it - we’ve +even written a blog post about it :) (https://www.ccdatalab.org/blog/dont-make-me-write-tips-for-avoiding-typing-in-rstudio).

    - -
    format_sce <- function(sce, sample_name) {
    -  # Input arguments:
    -  ## sce: An SCE object to format
    -  ## sample_name: The SCE object's name
    -  # This function returns a formatted SCE object.
    -
    -  ###### Ensure that we can identify the originating sample information ######
    -  # Add a column called `sample` that stores this information
    -  # This will be stored in `colData`
    +
    +
    #' Custom function to format an SCE before merging
    +#'
    +#' @param sce SCE object to format
    +#' @param sample_name Name of the sample
    +#' @param shared_genes Vector of shared genes across all SCE objects
    +#'
    +#' @returns An updated SCE object ready for merging
    +format_sce <- function(
    +  sce, 
    +  sample_name, 
    +  shared_genes
    +) {
    +  
    +  ### Remove the single-sample reduced dimensions 
    +  # We do this first since it makes the object a lot smaller for the rest of this code!
    +  reducedDims(sce) <- NULL
    +  
    +  ### Add dedicated sample indicator column to the colData slot
    +  # Recall, the `sce$` shortcut points to the colData
       sce$sample <- sample_name
     
    -
    -  ###### Ensure cell ids will be unique ######
    -  # Update the SCE object column names (cell ids) by prepending `sample_name`
    +  ### Ensure objects have the same genes in the same order
    +  # Use the shared_genes vector to index genes to the intersection
    +  # Doing this both subsets to just those genes, and reorders!
    +  sce <- sce[shared_genes, ]
    +  
    +  ### There is no additional wrangling to do for the colData column names or assays.
    +  ### But if there were, you could add your custom code to do so here.
    +  ### Your custom function may need additional arguments for this, too.
    +
    +  ### Ensure cell ids are identifiable and fully unique 
    +  # Update the SCE object column names (cell ids) by prepending the `sample_name`
       colnames(sce) <- glue::glue("{sample_name}-{colnames(sce)}")
     
    -
    -  ###### Ensure gene-level statistics can be identified in `rowData` ######
    -  # We want to rename the columns `mean` and `detected` to contain the `sample_name`
    -  # Recall the names are: "gene_symbol", "mean", "detected"
    -  colnames(rowData(sce)) <- c("gene_symbol",
    -                              glue::glue("{sample_name}-mean"),
    -                              glue::glue("{sample_name}-detected"))
    -
    -  # Return the formatted SCE object
    -  return(sce)
    +  ### Ensure the rowData columns can be identified
    +  # Recall, we want to leave `gene_symbol` alone, but add the `sample_name` to the rest
    +  rowdata_names <- colnames(rowData(sce))
    +  # prefix rowData names with the sample name, except for gene symbols
    +  new_rowdata_names <- ifelse(
    +    rowdata_names == "gene_symbol",
    +    "gene_symbol",
    +    glue::glue("{sample_name}-{rowdata_names}")
    +  )
    +  colnames(rowData(sce)) <- new_rowdata_names
    +  
    +  ### Ensure metadata slot fields can be identified
    +  # We'll simply prepend the `sample_name` to all fields for this slot
    +  names(metadata(sce)) <- glue::glue("{sample_name}-{names(metadata(sce))}")
    +  
    +  
    +  ### Finally, we can return the formatted SCE object
    +  return(sce)  
     }
    @@ -3796,26 +3826,25 @@

    Perform SCE merging

    sce_list names.

    - +
    # We can use `purrr::map2()` to loop over two list/vector arguments
     sce_list_formatted <- purrr::map2(
       # Each "iteration" will march down the first two
       #  arguments `sce_list` and `names(sce_list)` in order
       sce_list,
       names(sce_list),
    -  # Name of the function to run
    -  format_sce
    +  \(sce, sample_name) format_sce(sce, sample_name, shared_genes) 
     )
     
    -# Print resulting list
    +# Print formatted SCE list
     sce_list_formatted
    - +
    $SCPCL000479
     class: SingleCellExperiment 
     dim: 60319 1918 
    -metadata(14): salmon_version reference_index ... filtering_method
    -  miQC_model
    +metadata(14): SCPCL000479-salmon_version SCPCL000479-reference_index
    +  ... SCPCL000479-filtering_method SCPCL000479-miQC_model
     assays(2): counts logcounts
     rownames(60319): ENSG00000000003 ENSG00000000005 ... ENSG00000288724
       ENSG00000288725
    @@ -3824,15 +3853,15 @@ 

    Perform SCE merging

    SCPCL000479-CACAGATAGTGAGTGC ... SCPCL000479-GTTGTCCCACGTACAT SCPCL000479-TCCGATCGTCGTGCCA colData names(13): sum detected ... celltype_broad sample -reducedDimNames(2): PCA UMAP +reducedDimNames(0): mainExpName: NULL altExpNames(0): $SCPCL000480 class: SingleCellExperiment dim: 60319 4428 -metadata(14): salmon_version reference_index ... filtering_method - miQC_model +metadata(14): SCPCL000480-salmon_version SCPCL000480-reference_index + ... SCPCL000480-filtering_method SCPCL000480-miQC_model assays(2): counts logcounts rownames(60319): ENSG00000000003 ENSG00000000005 ... ENSG00000288724 ENSG00000288725 @@ -3841,15 +3870,15 @@

    Perform SCE merging

    SCPCL000480-AACTTCTTCCCTCAAC ... SCPCL000480-AGGGAGTAGCCTCATA SCPCL000480-TCGGATACATTGCAAC colData names(13): sum detected ... celltype_broad sample -reducedDimNames(2): PCA UMAP +reducedDimNames(0): mainExpName: NULL altExpNames(0): $SCPCL000481 class: SingleCellExperiment dim: 60319 5236 -metadata(14): salmon_version reference_index ... filtering_method - miQC_model +metadata(14): SCPCL000481-salmon_version SCPCL000481-reference_index + ... SCPCL000481-filtering_method SCPCL000481-miQC_model assays(2): counts logcounts rownames(60319): ENSG00000000003 ENSG00000000005 ... ENSG00000288724 ENSG00000288725 @@ -3858,15 +3887,15 @@

    Perform SCE merging

    SCPCL000481-GGGTATTTCGTTGTGA ... SCPCL000481-AAAGAACCACTTCAAG SCPCL000481-CAGCAGCTCGTGCATA colData names(13): sum detected ... celltype_broad sample -reducedDimNames(2): PCA UMAP +reducedDimNames(0): mainExpName: NULL altExpNames(0): $SCPCL000482 class: SingleCellExperiment dim: 60319 4372 -metadata(14): salmon_version reference_index ... filtering_method - miQC_model +metadata(14): SCPCL000482-salmon_version SCPCL000482-reference_index + ... SCPCL000482-filtering_method SCPCL000482-miQC_model assays(2): counts logcounts rownames(60319): ENSG00000000003 ENSG00000000005 ... ENSG00000288724 ENSG00000288725 @@ -3875,7 +3904,7 @@

    Perform SCE merging

    SCPCL000482-CAACCTCTCCGATCGG ... SCPCL000482-TGATTTCCACAAGTTC SCPCL000482-ACCAAACGTTCTCAGA colData names(13): sum detected ... celltype_broad sample -reducedDimNames(2): PCA UMAP +reducedDimNames(0): mainExpName: NULL altExpNames(0):
    @@ -3883,6 +3912,10 @@

    Perform SCE merging

    (Psst, like purrr and want to dive deeper? Check out the purrr::imap() function!)

    +
    +
    +
    +

    Perform the merging

    At long last, we are ready to merge the SCEs, which we’ll do using the R function cbind(). The cbind() function is often used to combine data frames or matrices by column, i.e. “stack” @@ -3905,11 +3938,11 @@

    Perform SCE merging

    # Print the merged_sce object merged_sce
    - +
    class: SingleCellExperiment 
     dim: 60319 15954 
    -metadata(56): salmon_version reference_index ... filtering_method
    -  miQC_model
    +metadata(56): SCPCL000479-salmon_version SCPCL000479-reference_index
    +  ... SCPCL000482-filtering_method SCPCL000482-miQC_model
     assays(2): counts logcounts
     rownames(60319): ENSG00000000003 ENSG00000000005 ... ENSG00000288724
       ENSG00000288725
    @@ -3919,29 +3952,29 @@ 

    Perform SCE merging

    SCPCL000479-CACAGATAGTGAGTGC ... SCPCL000482-TGATTTCCACAAGTTC SCPCL000482-ACCAAACGTTCTCAGA colData names(13): sum detected ... celltype_broad sample -reducedDimNames(2): PCA UMAP +reducedDimNames(0): mainExpName: NULL altExpNames(0):
    -

    We now have a single SCE object that contains all cells from all -samples we’d like to integrate.

    +

    We now have a single merged SCE object that contains all cells from +all samples we’d like to integrate.

    Let’s take a peek at some of the innards of this new SCE object:

    - +
    # How many samples, and cells per sample?
    -table( colData(merged_sce)$sample )
    +table(colData(merged_sce)$sample)
    
     SCPCL000479 SCPCL000480 SCPCL000481 SCPCL000482 
            1918        4428        5236        4372 
    - +
    # What are the new cell ids (column names)?
    -head( colnames(merged_sce) )
    +head(colnames(merged_sce))
    [1] "SCPCL000479-GGGACCTCAAGCGGAT" "SCPCL000479-CACAGATAGTGAGTGC"
    @@ -3956,9 +3989,25 @@ 

    Perform SCE merging

    SCPCL000479-ATTATCCCAGTTGGTT SCPCL000479-TCCGAAATCACACCGG
    - + +
    tail(colnames(merged_sce))
    + + +
    [1] "SCPCL000482-GATCACACAGCTAACT" "SCPCL000482-GACGCTGAGACTCTAC"
    +[3] "SCPCL000482-GTGAGGAGTCAACCTA" "SCPCL000482-ATTCCTAGTGTACATC"
    +[5] "SCPCL000482-TGATTTCCACAAGTTC" "SCPCL000482-ACCAAACGTTCTCAGA"
    + + +
    SCPCL000482-GATCACACAGCTAACT
    +SCPCL000482-GACGCTGAGACTCTAC
    +SCPCL000482-GTGAGGAGTCAACCTA
    +SCPCL000482-ATTCCTAGTGTACATC
    +SCPCL000482-TGATTTCCACAAGTTC
    +SCPCL000482-ACCAAACGTTCTCAGA
    + +
    # What does rowData look like?
    -head( rowData(merged_sce) )
    +head(rowData(merged_sce))
    DataFrame with 6 rows and 9 columns
    @@ -3991,8 +4040,8 @@ 

    Perform SCE merging

    -
    -

    Integration

    +
    +

    Integrate samples

    Single-cell roadmap: Integrate
    Single-cell roadmap: Integrate
    @@ -4005,78 +4054,28 @@

    Integration

    harmony specifically use PCA for this, but be aware that different integration methods may use other kinds of reduced dimensions.

    -

    You’ll notice that the merged SCE object object already contains PCA -and UMAP reduced dimensions, which were calculated during our -pre-processing:

    - - - -
    # Print the reducedDimNames of the merged_sce
    -reducedDimNames(merged_sce)
    - - -
    [1] "PCA"  "UMAP"
    - - -
    PCA
    -UMAP
    - - - -

    These represent the original dimension reductions that were performed -on each individual SCE before merging, but we actually need to -calculate PCA (and UMAP for visualization) from the merged object -directly.

    -

    Why can’t we use the sample-specific PCA and UMAP matrices? Part of -these calculations themselves involves scaling the raw data to center -the mean. When samples are separately centered but plotting together, -you will see samples “overlapping” in space, but this placement is -actually just an artifact of the individual centering. In addition, the -mathematical relationship between the original expression data and -reduced dimension version of that data will differ across samples, -meaning we can’t interpret them all together. To see how this looks, -let’s look at the UMAP when calculated from individual samples:

    - - - -
    # Plot UMAP calculated from individual samples with separate scaling
    -scater::plotReducedDim(merged_sce,
    -                       dimred = "UMAP",
    -                       color_by = "sample",
    -                       point_size = 0.5,
    -                       point_alpha = 0.2) +
    -  # Use a CVD-friendly color scheme and specify legend name
    -  scale_color_brewer(palette = "Dark2", name = "sample") + 
    -  # Modify the legend key with larger, easier to see points
    -  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
    -  ggtitle("UMAP calculated on each sample separately")
    - - -
    Scale for colour is already present.
    -Adding another scale for colour, which will replace the existing scale.
    - - -

    - - - -

    As we see in this UMAP, all samples are centered at zero and all -overlapping. This visual artifact can give the incorrect -impression that data is integrated - to be clear, this data is NOT -integrated!

    -

    For input to integration, we’ll want the reduced dimension -calculations to consider normalized gene expression values from all -samples simultaneously. So we’ll need to recalculate PCA (and UMAP for -visualization) on the merged object. We’ll also save these new reduced -dimensions with different names, merged_PCA and -merged_UMAP, to distinguish them from already-present -PCA and UMAP.

    +

    Before merging, our objects had reduced dimension representations +calculated on each individual SCE, and we removed them when preparing +for merge. We removed them because we don’t actually want to use them +anymore! This is because part of their calculation involves scaling the +raw data to center the mean. When samples are separately centered, +all of them will be centered at zero, making it look like the +datasets are already pretty overlapping when you plot their UMAPs +together. But, this is just a mathematical artifact of how dimension +reduction is performed.

    +

    So, we’ll begin by re-calculating PCA and UMAP on the merged object +in a way that takes batches into consideration. For input to +integration, we’ll want the reduced dimension calculations to consider +normalized gene expression values from all samples simultaneously. So +we’ll need to recalculate PCA (and UMAP for visualization) on the merged +object.

    First, as usual, we’ll determine the high-variance genes to use for PCA from the merged_sce object. For this, we’ll need to provide the argument block = merged_sce$sample when modeling gene variance, which tells scran::modelGeneVar() to first model variance separately for each batch and then combine those -modeling statistics.

    +modeling statistics. (Psst: isn’t it handy we created that +sample column when merging?!)

    @@ -4147,48 +4146,32 @@

    Integration

    object:

    - +
    # add PCA results to merged SCE object
    -reducedDim(merged_sce, "merged_PCA") <- merged_pca[[1]]
    +reducedDim(merged_sce, "PCA") <- merged_pca[[1]]

    Now that we have the PCA matrix, we can proceed to calculate UMAP to visualize the uncorrected merged data.

    -

    We’ll calculate UMAP as “usual”, but in this case we’ll specify two -additional arguments:

    -
      -
    • dimred = "merged_PCA", which specifies which existing -reduced dimension should be used for the calculation. We want to use the -batch-weighted PCA, which we named above as -"merged_PCA".
    • -
    • name = "merged_UMAP", which names the final UMAP that -this function calculates. This argument will prevent us from overwriting -the existing UMAP which is already named “UMAP” and instead create a -separate "merged_UMAP".
    • -
    - -
    # add merged_UMAP from merged_PCA
    -merged_sce <- scater::runUMAP(merged_sce,
    -                              dimred = "merged_PCA",
    -                              name = "merged_UMAP")
    + +
    merged_sce <- scater::runUMAP(merged_sce)
    -

    Now, let’s see how this new merged_UMAP looks compared -to the UMAP calculated from individual samples:

    - +
    # UMAPs scaled together when calculated from the merged SCE
    -scater::plotReducedDim(merged_sce,
    -                       dimred = "merged_UMAP",
    -                       color_by = "sample",
    -                       # Some styling to help us see the points:
    -                       point_size = 0.5,
    -                       point_alpha = 0.2) +
    +scater::plotUMAP(
    +  merged_sce,
    +  color_by = "sample",
    +  # Some styling to help us see the points:
    +  point_size = 0.5,
    +  point_alpha = 0.2
    +) +
       scale_color_brewer(palette = "Dark2", name = "sample") +
       guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
       ggtitle("UMAP calculated on merged_sce")
    @@ -4198,19 +4181,19 @@

    Integration

    Adding another scale for colour, which will replace the existing scale. -

    +

    -

    Samples are now separated, which more reasonably reflects that this -data is not yet batch-corrected. We can think of this UMAP as -our “before” UMAP, and we can compare this to the “after” UMAP we see -post-integration.

    +

    We see (mostly) four separate clumps representing the four different +merged but not yet integrated samples. We can think of this +UMAP as our “before” UMAP, and we can compare this to the “after” UMAP +we see post-integration.

    Let’s discuss a little first: What visual differences do you think the UMAP on the integrated version of data will have? What similarities do you think the integrated UMAP will have to this plot?

    -
    -

    Integration with fastMNN

    +
    +

    Integrate with fastMNN

    Finally, we’re ready to integrate! To start, we’ll use the fastMNN approach from the Bioconductor batchelor package.

    @@ -4292,7 +4275,14 @@

    Integration with fastMNN

    Finally, we’ll calculate UMAP from these corrected PCA matrix for -visualization.

    +visualization. In this case we need to specify two additional arguments +since we’re working with non-standard reduced dimension names:

    +
      +
    • dimred = "fastmnn_PCA", which specifies the existing +reduced dimension to use for the calculation
    • +
    • name = "fastmnn_UMAP", which names the final UMAP that +this function calculates
    • +
    @@ -4332,13 +4322,13 @@

    Integration with fastMNN

    Adding another scale for colour, which will replace the existing scale. -

    +

    This fastmnn_UMAP certainly looks different from the one -we made from merged_UMAP! What different trends do you see? -Do all samples look “equally well” integrated, from a first look?

    +we made before integrating! What different trends do you see? Do all +samples look “equally well” integrated, from a first look?

    Importantly, one reason that batches may still appear separated in the corrected UMAP is if they should be separated - for example, maybe two batches contain very different cell types, have very @@ -4398,7 +4388,7 @@

    Integration with fastMNN

    Adding another scale for colour, which will replace the existing scale. -

    +

    @@ -4437,24 +4427,24 @@

    Integration with fastMNN

    Adding another scale for colour, which will replace the existing scale. -

    +

    What trends do you observe between tumor and healthy tissues among these integrated samples?

    -
    -

    Integration with harmony

    +
    +

    Integrate with harmony

    fastMNN is only one of many approaches to perform integration, and different methods have different capabilities and may give different results. For example, some methods can accommodate additional covariates (e.g., technology, patient, diagnosis, etc.) that can influence integration. In fact the data we are using has a known patient covariate; SCPCL000479 and -SCPCL000480 are from the first patient, and -SCPCL000481 and SCPCL000482 are from the -second patient.

    +SCPCL000480 are from Patient A, and +SCPCL000481 and SCPCL000482 are from Patient +B.

    So, let’s perform integration with a method that can use this information - harmony!

    To begin setting up for harmony integration, we need to @@ -4487,8 +4477,7 @@

    Integration with harmony

    • First, harmony takes a batch-weighted PCA matrix to perform integration. We already calculated a batch-weighted PCA matrix -(our merged_PCA reduced dimension), we’ll provide this as -the the input.
    • +so we’ll provide this as the the input.
    • Second, we need to tell harmony about the covariates to use - sample and patient. To do this, we provide two arguments: @@ -4505,10 +4494,10 @@

      Integration with harmony

      Let’s go!

      - +
      # Run harmony integration
       harmony_pca <- harmony::RunHarmony(
      -  data_mat = reducedDim(merged_sce, "merged_PCA"),
      +  data_mat = reducedDim(merged_sce, "PCA"),
         meta_data = colData(merged_sce),
         vars_use = c("sample", "patient")
       )
      @@ -4538,13 +4527,13 @@

      Integration with harmony

      # Print the harmony result
       harmony_pca[1:5, 1:5]
      - +
                                        [,1]      [,2]      [,3]     [,4]       [,5]
      -SCPCL000479-GGGACCTCAAGCGGAT -7.032044 -7.271764 -3.713022 5.477643  0.6781986
      -SCPCL000479-CACAGATAGTGAGTGC -8.237485 -7.436515 -5.579578 6.101144 -0.1577978
      -SCPCL000479-TGTGGCGGTGAATTGA -7.893770 -6.595934 -2.042903 4.706149  0.9176450
      -SCPCL000479-GCCGATGGTACATACC -6.741593 -7.537079 -5.997807 2.609384 -4.1100267
      -SCPCL000479-ATTATCCCAGTTGGTT -5.534487 -4.887054 -1.897409 2.588823  1.5458433
      +SCPCL000479-GGGACCTCAAGCGGAT -7.036850 -7.280253 -3.825712 5.564675 0.5984375 +SCPCL000479-CACAGATAGTGAGTGC -8.232417 -7.422768 -5.641379 6.128005 -0.2208036 +SCPCL000479-TGTGGCGGTGAATTGA -7.928602 -6.591079 -2.140173 4.775223 0.8639596 +SCPCL000479-GCCGATGGTACATACC -6.821359 -7.465958 -5.872961 2.313765 -4.1492218 +SCPCL000479-ATTATCCCAGTTGGTT -5.611779 -4.916862 -1.894365 2.626925 1.5535017 @@ -4584,7 +4573,7 @@

      Integration with harmony

      Adding another scale for colour, which will replace the existing scale. -

      +

      @@ -4594,7 +4583,7 @@

      Integration with harmony

      visibility:

      - +
      scater::plotReducedDim(merged_sce,
                              dimred = "harmony_UMAP",
                              color_by = "celltype_broad",
      @@ -4603,7 +4592,7 @@ 

      Integration with harmony

      # Specify variable for faceting other_fields = "sample") + scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") + - guides(color = guide_legend(override.aes = list(size = 3))) + + guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) + ggtitle("UMAP after integration with harmony") + facet_wrap(vars(sample))
      @@ -4612,7 +4601,7 @@

      Integration with harmony

      Adding another scale for colour, which will replace the existing scale. -

      +

      @@ -4622,8 +4611,9 @@

      Integration with harmony

      fastMNN vs. harmony performed in integrating these samples?

    -
    -

    Export

    +
    +
    +

    Export

    Finally, we’ll export the final SCE object with both fastMNN and harmony integration to a file. Since this object is very large (over 1 GB!), we’ll export it to a file @@ -4632,16 +4622,17 @@

    Export

    compression is performed.

    - +
    # Export to RDS file with "gz" compression
    -readr::write_rds(merged_sce,
    -                 integrated_sce_file,
    -                 compress = "gz")
    +readr::write_rds( + merged_sce, + integrated_sce_file, + compress = "gz" +)
    -
    -
    ---
title: "Integrating scRNA-seq datasets"
author: Data Lab for ALSF
date: 2023
output:
  html_notebook:
    toc: true
    toc_depth: 3
    toc_float: true
---

## Objectives

This notebook will demonstrate how to:

- Prepare SCE objects for integration
- Apply integration methods including `fastMNN` and `harmony`
- Visually explore the results of integration
- Use `purrr::map()` functions for iterating over lists

---

In this notebook, we'll perform integration on scRNA-seq datasets from the [Single-cell Pediatric Cancer Atlas (`ScPCA`)](https://scpca.alexslemonade.org/), a database of uniformly-processed pediatric scRNA-seq data built and maintained by the Data Lab.
The `ScPCA` database currently hosts single-cell pediatric cancer transcriptomic data generated by ALSF-funded labs, with the goal of making this data easily accessible to investigators (like you!).
The expression data in `ScPCA` were mapping and quantified with [`alevin-fry`](https://doi.org/10.1038/s41592-022-01408-3), followed by processing with Bioconductor tools using the same general procedures that we have covered in this workshop.
The processing pipeline used `emptyDropsCellRanger()` and `miQC` to filter the raw counts matrix, `scuttle` to log-normalize the counts, and `scater` for dimension reduction.
The processed data are stored as `.rds` files containing `SingleCellExperiment` objects.
You can read more about how data in the `ScPCA` is processed in [the associated documentation](https://scpca.readthedocs.io/en/latest/).


![Single-cell roadmap: Integration Overview](diagrams/roadmap_multi_merge-integrate.png)

To learn about integration, we'll have a look at four samples from the [`SCPCP000005` project](https://scpca.alexslemonade.org/projects/SCPCP000005) ([Patel _et al._ 2022](https://doi.org/10.1016/j.devcel.2022.04.003)), an investigation of pediatric solid tumors led by the [Dyer](https://www.stjude.org/research/labs/dyer-lab.html) and [Chen](https://www.stjude.org/research/labs/chen-lab-taosheng.html) labs at St. Jude Children's Research Hospital.
The particular libraries we'll integrate come from two rhabdomyosarcoma (RMS) patients, with two samples from each of two patients, all sequenced with 10x Chromium v3 technology.
Each library is from a separate biological sample.

We'll be integrating these samples with two different tools, [`fastMNN`](http://www.bioconductor.org/packages/release/bioc/html/batchelor.html) ([Haghverdi _et al._ 2018](https://doi.org/10.1038/nbt.4091)) and [`harmony`](https://portals.broadinstitute.org/harmony/) ([Korsunsky _et al._ 2019](https://doi.org/10.1038/s41592-019-0619-0)).
Integration corrects for batch effects that arise from different library preparations, genetic backgrounds, and other sample-specific factors, so that datasets can be jointly analyzed at the cell level.
`fastMNN` corrects for batch effects using a faster variant of the mutual-nearest neighbors algorithm, the technical details of which you can learn more about from this [vignette by Lun (2019)](https://marionilab.github.io/FurtherMNN2018/theory/description.html).
`harmony`, on the other hand, corrects for batch effects using an iterative clustering approach, and unlike `fastMNN`, it is also able to consider additional covariates beyond just the batch groupings.

Regardless of which integration tool is used, the `SingleCellExperiment` (SCE) objects first need to be reformatted and merged into a single (uncorrected!) SCE object that contains all cells from all samples.
This merged SCE can then be used for integration to obtain a formally batch-corrected SCE object.


## Set up

```{r setup}
# Load libraries
library(ggplot2)  # plotting tools
library(SingleCellExperiment) # work with SCE objects

# Set the seed for reproducibility
set.seed(12345)
```


### Directories and files


We have already prepared count data for the four samples we'll be integrating (i.e., filtered cells, normalized counts, and calculated PCA & UMAP).
These SCE objects, stored as RDS files, are available in the `data/rms/processed/` directory and are named according to their `ScPCA` library ids :

- `SCPCL000479.rds` (Patient A)
- `SCPCL000480.rds` (Patient A)
- `SCPCL000481.rds` (Patient B)
- `SCPCL000482.rds` (Patient B)

To begin, let's set up our directories and files:

```{r directories, live = TRUE}
# Define directory where processed SCE objects to be integrated are stored
input_dir <- file.path("data", "rms", "processed")

# Define directory to save integrated SCE object to
output_dir <- file.path("data", "rms", "integrated")

# Create output directory if it doesn't exist
fs::dir_create(output_dir)

# Define output file name for the integrated object
integrated_sce_file <- file.path(output_dir, "rms_integrated_subset.rds")
```


We can use the `dir()` function to list all contents of a given directory, for example to see all the files in our `input_dir`:

```{r input dir, live = TRUE}
dir(input_dir)
```

We want to read in just four of these files, as listed previously.
To read in these files, we could use the `readr::read_rds()` function (or the base R `readRDS()`) four times, once for each of the files.
We could also use a `for` loop, which is the approach that many programming languages would lean toward.
A different and more modular coding approach to reading in these files (and more!) is to leverage the [`purrr`](https://purrr.tidyverse.org/) `tidyverse` package, which provides a convenient set of functions for operating on lists.
You can read more about the `purrr` functions and their power and utility in R in [the "Functionals" chapter of the _Advanced R_ e-book](https://adv-r.hadley.nz/functionals.html).

Of particular interest is the [`purrr::map()`](https://purrr.tidyverse.org/reference/map.html) family of functions, which can be used to run a given function on each element of a list (or vector) in one call.
The general syntax for `purrr::map()` and friends is:

```
# Syntax for using the map function:
purrr::map(<input list or vector>,
           <function to apply to each item in the input>,
           <any additional arguments to the function can go here>,
           <and also here if there are even more arguments, and so on>)
```


The output from running `purrr::map()` is always a list (but note that there are other `purrr::map()` relatives which return other object types, as you can read about in [the `purrr::map()` documentation](https://purrr.tidyverse.org/reference/index.html)).
If this concept sounds a little familiar to you, that's because it probably is!
Base R's `lapply()` function can provide similar utility, and the `purrr::map()` family of functions can (in part) be thought of as an alternative to some of the base R `apply` functions, with more consistent behavior.

Let's see a very simple example of `purrr::map()` in action, inspired by cancer groups the Data Lab has analyzed through the [OpenPBTA](https://github.com/AlexsLemonade/OpenPBTA-analysis/) project:

```{r map example}
# Define a list of cancer histologies
histologies <- list(
  "low-grade gliomas"  = c("SEGA", "PA", "GNG", "PXA"),
  "high-grade gliomas" = c("DMG", "DIPG"),
  "embryonal tumors"   = c("MB", "ATRT", "ETMR")
 )

# The overall length of the list is 3
length(histologies)

# How can we run `length()` on each item of the list?
# We can use our new friend purrr::map():
purrr::map(histologies, length)
```

One other new coding strategy we'll learn in this notebook is using the [`glue`](https://glue.tidyverse.org/) package to combine strings.
This package offers a convenient function `glue::glue()` that can be used instead of the base R `paste()` function.

```{r paste}
# Define a variable for example:
org_name <- "Data Lab"

# We can use paste to combine strings and variables:
paste("Welcome to the", org_name, "workshop on Advanced scRNA-seq!")
```

We can use `glue::glue()` to accomplish the same goal with some different syntax:

```{r glue}
# glue::glue takes a single string argument (only one set of quotes!), and
#  variables can easily be included inside {curly braces}
glue::glue("Welcome to the {org_name} workshop on Advanced scRNA-seq!")
```

(Note that even though the `glue::glue()` output isn't in quotes, it still behaves like a string!)


Alright, time for the good stuff!
Let's use `purrr::map()` to read in our SCE objects so that they are immediately stored together in a list.


We'll first need to define a vector of the file paths to read in.
We'll start by creating a vector of sample names themselves and then formatting them into the correct paths.
This way (foreshadowing!) we also have a stand-alone vector of just sample names, which will come in handy!

```{r sample names}
# Vector of all the samples to read in:
sample_names <- c("SCPCL000479",
                  "SCPCL000480",
                  "SCPCL000481",
                  "SCPCL000482")
```


```{r define sce_paths, live = TRUE}
# Now, convert these to file paths: <input_dir>/<sample_name>.rds
sce_paths <- file.path(input_dir,
                       glue::glue("{sample_names}.rds")
)
# Print the sce_paths vector
sce_paths
```

Let's make this a named vector using the sample names.
This will help us keep track of which objects are which after we read the SCE objects in:

```{r add list names, live = TRUE}
# Assign the sample names as the names for sce_paths
names(sce_paths) <- sample_names
```

We can now read these files in and create a list of four SCE objects. 
Since `readr::read_rds()` can only operate on one input at a time, we'll need to use `purrr::map()` to run it on all input file paths in one command.
Although `sce_paths` is a vector (not a list), it will still work as input to `purrr:map()`.
The output from this code will still be a list, since that's what `purrr::map()` always returns, and it will retain the sample names as the list names for convenient bookkeeping:

```{r read sce paths, live = TRUE}
# Use purrr::map() to read all files into a list at once
sce_list <- purrr::map(
  sce_paths,
  readr::read_rds
)
```

Let's have a look at our named list of SCE objects:

```{r print sce list, live=TRUE}
# Print sce_list
sce_list
```

If you look closely at the printed SCE objects, you may notice that they all contain `colData` table columns `celltype_fine` and `celltype_broad`.
These columns (which we added to SCE objects during [pre-processing](https://github.com/AlexsLemonade/training-modules/tree/master/scRNA-seq-advanced/setup/rms)) contain putative cell type annotations as assigned by [Patel _et al._ (2022)](https://doi.org/10.1016/j.devcel.2022.04.003):


> For each cell subset identified by clustering, we used a combination of `SingleR` version 1.0.1 ([Aran et al., 2019](https://www.nature.com/articles/s41590-018-0276-y)) and manual inspection of differentially expressed genes to annotate whether a cluster belongs to stromal, immune or malignant subpopulations. 
Malignant cells were confirmed in patient tumor data by inference of copy-number variation using `inferCNV` version 1.1.3 of the TrinityCTAT Project (https://github.com/broadinstitute/infercnv). 

We will end up leveraging these cell type annotations to explore the integration results; after integration, we expect cell types from different samples to group together, rather than being separated by batches. 

That said, the integration methods we will be applying _do not actually use_ any existing cell type annotations.
If we have annotations, they are a helpful "bonus" for assessing the integration's performance, but they are not part of the integration itself.


## Prepare the SCE list for integration

![Single-cell roadmap: Merge](diagrams/roadmap_multi_merge.png)


Now that we have a list of processed SCE objects, we need to merge the objects into one overall SCE object for input to integration.
A word of caution before we begin: **This merged SCE object is NOT an integrated SCE!**
Merging SCEs does not perform any batch correction, but just reorganizes the data to allow us to proceed to integration next.

To merge SCE objects, we do need to do some wrangling and bookkeeping to ensure compatibility and that we don't lose important information.
Overall we'll want to take care of these items:

1. We should be able to trace sample-specific information back to the originating sample, including...
    - Cell-level information: Which sample is each cell from?
    - Library-specific feature statistics, e.g., gene-level statistics for a given library found in `rowData`.
    Which sample is a given feature statistic from?
2. SCE objects should contain the same genes: Each SCE object should have the same row names.
3. SCE cell metadata columns should match: The `colData` for each SCE object should have the same column names.


We'll begin by taking some time to thoroughly explore our SCE objects and figure out what wrangling steps we need to take for these specific data.
Don't skip this exploration!
Bear in mind that the exact wrangling shown here will not be the same for other SCE objects you work with, but the same general principles apply.


#### Preserving sample information at the cell level

How will we be able to tell which sample a given cell came from?

The best way to do this is simply to add a `colData` column with the sample information, so that we can know which sample each row came from.

In addition, we want to pay some attention to the SCE object's column names (the cell ids), which must remain unique after merging since duplicate ids will cause an R error.
In this case, the SCE column names are barcodes (which is usually but not always the case in SCE objects), which are only guaranteed to be unique _within_ a sample but may be repeated across samples.
So, after merging, it's technically possible that multiple cells will have the same barcode.
This would be a problem for two reasons:
First, the cell id would not be able to point us back to cell's originating sample.
Second, it would literally cause an error in R, which does not allow duplicate column names.


One way to ensure that cell ids remain unique even after merging is to actually modify them by _prepending_ the relevant sample name.
For example, consider these barcodes for the `SCPCL000479` sample:

```{r barcodes}
# Look at the column names for the `SCPCL000479` sample, for example
colnames(sce_list$SCPCL000479) |>
  # Only print out the first 6 for convenience
  head()
```

These ids will be updated to `SCPCL000479-GGGACCTCAAGCGGAT`, `SCPCL000479-CACAGATAGTGAGTGC`, and so on, thereby ensuring fully unique ids for all cells across all samples.

#### Preserving sample information at the gene level

The `rowData` table in SCE objects will often contain both "general" and "library-specific" information, for example:

```{r rowdata}
rowData(sce_list$SCPCL000479) |>
  head()
```

Here, the rownames are Ensembl gene ids, and columns are `gene_symbol`, `mean`, and `detected`.
The `gene_symbol` column is general information about all genes, not specific to any library or experiment, but `mean` and `detected` are library-specific gene statistics.
So, `gene_symbol` does not need to be traced back to its originating sample, but `mean` and `detected` do.
To this end, we can take a similar approach to what we'll do for cell ids:
We can change the sample-specific `rowData` column names by prepending the sample name.
For example, rather than being called `mean`, this column will be named `SCPCL000479-mean` for the `SCPCL000479` sample.

All our SCE objects have the same `rowData` columns (as we can see in the next chunk), so we'll perform this renaming across all SCEs.

```{r compare rowdata, live = TRUE}
# Use `purrr::map()` to quickly extract rowData column names for all SCEs
purrr::map(sce_list,
           \(sce) colnames(rowData(sce)))
```


#### Ensuring that only shared genes are used

The next step in ensuring SCE compatibility is to make sure they all contain the same genes, which are stored as the SCE object's row names (these names are also found the `rowData` slot's row names).
Here, those gene ids are unique Ensembl gene ids.

We can use some `purrr` magic to quickly find the set of shared genes among our samples, and then ask how many there are.

```{r shared genes}
# Define vector of shared genes
shared_genes <- sce_list |>
  # get rownames (genes) for each SCE in sce_list
  purrr::map(rownames) |>
  # reduce to the _intersection_ among lists
  purrr::reduce(intersect)
```

```{r print shared genes, live = TRUE}
# How many shared genes are there?
length(shared_genes)
```

In this case, we happen to know that all SCE objects we're working with already contained the same genes.
We do a quick-and-dirty check for this by looking at the number of rows across SCE objects, and we'll see that they are all the same:

```{r check shared genes, live = TRUE}
# The number of genes in an SCE corresponds to its number of rows:
sce_list |>
  purrr::map(nrow)
```

So, for our data, we will not have to subset to shared genes since they are already shared!

#### Ensuring matching columns in `colData`

Finally, we'll need to have the same column names across all SCE `colData` tables, so let's look at all those column names.
We can use similar syntax here to what we used to look at all the `rowData` column names.

```{r compare coldata}
purrr::map(sce_list,
           \(sce) colnames(colData(sce)) )
```

It looks like the column names are all already matching among SCEs, so there's no specific preparation we'll need to do there.

### Perform SCE merging

As you can see, there's a lot of moving parts to consider!
Again, these moving parts may (will!) differ for SCEs that you are working with, so you have to explore your own SCEs in depth to prepare for merging.

Based on our exploration, here is a schematic of how one of the SCE objects will ultimately be modified into the final merged SCE:

![](diagrams/technical_merge_sce.png)


We'll write a _custom function_ (seen in the chunk below) tailored to our wrangling steps that prepares a single SCE object for merging.
We'll then use our new `purrr::map()` programming skills to run this function over the `sce_list`.
This will give us a new list of formatted SCEs that we can proceed to merge.
It's important to remember that the `format_sce()` function written below is not a function for general use – it's been precisely written to match the processing we need to do for _these_ SCEs, and different SCEs you work with will require different types of processing.

```{r format_sce function}
format_sce <- function(sce, sample_name) {
  # Input arguments:
  ## sce: An SCE object to format
  ## sample_name: The SCE object's name
  # This function returns a formatted SCE object.

  ###### Ensure that we can identify the originating sample information ######
  # Add a column called `sample` that stores this information
  # This will be stored in `colData`
  sce$sample <- sample_name


  ###### Ensure cell ids will be unique ######
  # Update the SCE object column names (cell ids) by prepending `sample_name`
  colnames(sce) <- glue::glue("{sample_name}-{colnames(sce)}")


  ###### Ensure gene-level statistics can be identified in `rowData` ######
  # We want to rename the columns `mean` and `detected` to contain the `sample_name`
  # Recall the names are: "gene_symbol", "mean", "detected"
  colnames(rowData(sce)) <- c("gene_symbol",
                              glue::glue("{sample_name}-mean"),
                              glue::glue("{sample_name}-detected"))

  # Return the formatted SCE object
  return(sce)
}
```

To run this function, we'll use the `purrr::map2()` function, a relative of `purrr::map()` that allows you to loop over _two_ input lists/vectors.
In our case, we want to run `format_sce()` over paired `sce_list` items and `sce_list` names.

```{r format sces for merge, live = TRUE}
# We can use `purrr::map2()` to loop over two list/vector arguments
sce_list_formatted <- purrr::map2(
  # Each "iteration" will march down the first two
  #  arguments `sce_list` and `names(sce_list)` in order
  sce_list,
  names(sce_list),
  # Name of the function to run
  format_sce
)

# Print resulting list
sce_list_formatted
```

(Psst, like `purrr` and want to dive deeper? Check out [the `purrr::imap()` function](https://purrr.tidyverse.org/reference/imap.html)!)


At long last, we are ready to merge the SCEs, which we'll do using the R function `cbind()`.
The `cbind()` function is often used to combine data frames or matrices by column, i.e. "stack" them next to each other.
The same principle applies here, but when run on SCE objects, `cbind()` will create a new SCE object by combining `counts` and `logcounts` matrices by column.
Following that structure, other SCE slots (`colData`, `rowData`, reduced dimensions, and other metadata) are combined appropriately.

Since we need to apply `cbind()` to a _list_ of objects, we need to use some slightly-gnarly syntax: We'll use the function `do.call()`, which allows the `cbind()` input to be a list of objects to combine.

```{r merges sces, live = TRUE}
# Merge SCE objects
merged_sce <- do.call(cbind, sce_list_formatted)

# Print the merged_sce object
merged_sce
```

We now have a single SCE object that contains all cells from all samples we'd like to integrate.

Let's take a peek at some of the innards of this new SCE object:

```{r explore merged_sce, live = TRUE}
# How many samples, and cells per sample?
table( colData(merged_sce)$sample )

# What are the new cell ids (column names)?
head( colnames(merged_sce) )

# What does rowData look like?
head( rowData(merged_sce) )
```


## Integration

![Single-cell roadmap: Integrate](diagrams/roadmap_multi_integrate.png)


So far, we've created a `merged_sce` object which is (almost!) ready for integration.

The integration methods we'll be using here actually perform batch correction on a reduced dimension representation of the normalized gene expression values, which is more efficient.
`fastMNN` and `harmony` specifically use PCA for this, but be aware that different integration methods may use other kinds of reduced dimensions.

You'll notice that the merged SCE object object already contains PCA and UMAP reduced dimensions, which were calculated during our pre-processing:

```{r merged_sce reddim, live = TRUE}
# Print the reducedDimNames of the merged_sce
reducedDimNames(merged_sce)
```

These represent the original dimension reductions that were performed on _each individual SCE_ before merging, but we actually need to calculate PCA (and UMAP for visualization) from the merged object directly.

Why can't we use the sample-specific PCA and UMAP matrices?
Part of these calculations themselves involves scaling the raw data to center the mean.
When samples are separately centered but plotting together, you will see samples "overlapping" in space, but this placement is actually just an artifact of the individual centering.
In addition, the mathematical relationship between the original expression data and reduced dimension version of that data will differ across samples, meaning we can't interpret them all together.
To see how this looks, let's look at the UMAP when calculated from individual samples:

```{r plot individual UMAPs}
# Plot UMAP calculated from individual samples with separate scaling
scater::plotReducedDim(merged_sce,
                       dimred = "UMAP",
                       color_by = "sample",
                       point_size = 0.5,
                       point_alpha = 0.2) +
  # Use a CVD-friendly color scheme and specify legend name
  scale_color_brewer(palette = "Dark2", name = "sample") + 
  # Modify the legend key with larger, easier to see points
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP calculated on each sample separately")
```

As we see in this UMAP, all samples are centered at zero and all overlapping.
This visual artifact can give the _incorrect impression_ that data is integrated - to be clear, this data is NOT integrated!

For input to integration, we'll want the reduced dimension calculations to consider normalized gene expression values from all samples simultaneously.
So we'll need to recalculate PCA (and UMAP for visualization) on the merged object.
We'll also save these new reduced dimensions with different names, `merged_PCA` and `merged_UMAP`, to distinguish them from already-present `PCA` and `UMAP`.

First, as usual, we'll determine the high-variance genes to use for PCA from the `merged_sce` object.
For this, we'll need to provide the argument `block = merged_sce$sample` when modeling gene variance, which tells `scran::modelGeneVar()` to first model variance separately for each batch and then combine those modeling statistics.

```{r calc merged hv genes}
# Specify the number of genes to identify
num_genes <- 2000

# Calculate variation for each gene
gene_variance <- scran::modelGeneVar(merged_sce,
                                     # specify the grouping column:
                                     block = merged_sce$sample)

# Get the top `num_genes` high-variance genes to use for dimension reduction
hv_genes <- scran::getTopHVGs(gene_variance,
                              n = num_genes)
```

To calculate the PCA matrix itself, we'll use an approach from the `batchelor` package, which is the R package that contains the `fastMNN` method.
The [`batchelor::multiBatchPCA()`](https://rdrr.io/bioc/batchelor/man/multiBatchPCA.html) function calculates a batch-weighted PCA matrix.
This weighting ensures that all batches, which may have very different numbers of cells, contribute equally to the overall scaling.

```{r make merged_pca, live = TRUE}
# Use batchelor to calculate PCA for merged_sce, considering only
#  the high-variance genes
# We'll need to include the argument `preserve.single = TRUE` to get
#  a single matrix with all samples and not separate matrices for each sample
merged_pca <- batchelor::multiBatchPCA(merged_sce,
                                       subset.row = hv_genes,
                                       batch = merged_sce$sample,
                                       preserve.single = TRUE)
```

Let's have a look at the output:
```{r print merged_pca, live = TRUE}
# This output is not very interesting!
merged_pca
```

We can use indexing `[[1]]` to see the PCA matrix calculated, looking at a small subset for convenience:

```{r print merged_pca indexed, live = TRUE}
merged_pca[[1]][1:5,1:5]
```

We can now include this PCA matrix in our `merged_sce` object:

```{r add merged_pca, live = TRUE}
# add PCA results to merged SCE object
reducedDim(merged_sce, "merged_PCA") <- merged_pca[[1]]
```

Now that we have the PCA matrix, we can proceed to calculate UMAP to visualize the uncorrected merged data.

We'll calculate UMAP as "usual", but in this case we'll specify two additional arguments:

- `dimred = "merged_PCA"`, which specifies which existing reduced dimension should be used for the calculation.
We want to use the batch-weighted PCA, which we named above as `"merged_PCA"`.
- `name = "merged_UMAP"`, which names the final UMAP that this function calculates.
This argument will prevent us from overwriting the existing UMAP which is already named "UMAP" and instead create a separate `"merged_UMAP"`.

```{r calculate merged umap, live = TRUE}
# add merged_UMAP from merged_PCA
merged_sce <- scater::runUMAP(merged_sce,
                              dimred = "merged_PCA",
                              name = "merged_UMAP")
```

Now, let's see how this new `merged_UMAP` looks compared to the `UMAP` calculated from individual samples:

```{r plot uncorrected merged UMAP}
# UMAPs scaled together when calculated from the merged SCE
scater::plotReducedDim(merged_sce,
                       dimred = "merged_UMAP",
                       color_by = "sample",
                       # Some styling to help us see the points:
                       point_size = 0.5,
                       point_alpha = 0.2) +
  scale_color_brewer(palette = "Dark2", name = "sample") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP calculated on merged_sce")
```

Samples are now separated, which more reasonably reflects that this data is _not yet batch-corrected_.
We can think of this UMAP as our "before" UMAP, and we can compare this to the "after" UMAP we see post-integration.

Let's discuss a little first: What visual differences do you think the UMAP on the integrated version of data will have?
What similarities do you think the integrated UMAP will have to this plot?


### Integration with `fastMNN`

Finally, we're ready to integrate!
To start, we'll use the `fastMNN` approach from the Bioconductor [`batchelor` package](http://www.bioconductor.org/packages/release/bioc/html/batchelor.html).

`fastMNN` takes as input the `merged_sce` object to integrate, and the first step it performs is actually to run `batchelor::multiBatchPCA()` on that SCE.
It then uses that batch-weighted PCA matrix to perform the actual batch correction.
The `batch` argument is used to specify the different groupings within the `merged_sce` (i.e. the original sample that each cell belongs to), and the `subset.row` argument can optionally be used to provide a vector of high-variance genes that should be considered for this PCA calculation.
`fastMNN` will return an SCE object that contains a batch-corrected PCA.
Let's run it and save the result to a variable called `integrated_sce`.


```{r run fastmnn, live = TRUE}
# integrate with fastMNN, again specifying only our high-variance genes
integrated_sce <- batchelor::fastMNN(
  merged_sce,
  batch = merged_sce$sample,
  subset.row = hv_genes
)
```

Let's have a look at the result:

```{r fastmnn result, live = TRUE}
# Print the integrated_sce object
integrated_sce
```

There are couple pieces of information here of interest:

- The `corrected` reduced dimension represents the batch-corrected PCA that `fastMNN` calculated.
- The `reconstructed` assay represents the batch-corrected normalized expression values, which `fastMNN` "back-calculated" from the batch-corrected PCA (`corrected`).
Generally speaking, these expression values are not stand-alone values that you should use for other applications like differential gene expression, as described in [_Orchestrating Single Cell Analyses_](http://bioconductor.org/books/3.19/OSCA.multisample/using-corrected-values.html).
If the `subset.row` argument is provided (as it was here), only genes present in `subset.row` will be included in these reconstructed expression values, but this setting can be overridden so that all genes have reconstructed expression with the argument `correct.all = TRUE`.

We're mostly interested in the PCA that `fastMNN` calculated, so let's save that information (with an informative and unique name) into our `merged_sce` object:

```{r fastmnn pcs, live = TRUE}
# Make a new reducedDim named fastmnn_PCA from the corrected reducedDim in integrated_sce
reducedDim(merged_sce, "fastmnn_PCA") <- reducedDim(integrated_sce, "corrected")
```

Finally, we'll calculate UMAP from these corrected PCA matrix for visualization.

```{r calculate fastmnn umap, live = TRUE}
# Calculate UMAP
merged_sce <- scater::runUMAP(
  merged_sce,
  dimred = "fastmnn_PCA",
  name = "fastmnn_UMAP"
)
```

First, let's plot the integrated UMAP highlighting the different batches.
A well-integrated dataset will show batch mixing, but a poorly-integrated dataset will show more separation among batches, similar to the uncorrected UMAP.
Note that this is a more qualitative way to assess the success of integration, but there are formal metrics one can use to assess batch mixing, which you can read more about in [this chapter of OSCA](http://bioconductor.org/books/3.19/OSCA.multisample/correction-diagnostics.html).

```{r plot fastmnn umap batches}
scater::plotReducedDim(merged_sce,
                       # plot the fastMNN coordinates
                       dimred = "fastmnn_UMAP",
                       # color by sample
                       color_by = "sample",
                       # Some styling to help us see the points:
                       point_size = 0.5,
                       point_alpha = 0.2) +
  scale_color_brewer(palette = "Dark2", name = "sample") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with fastMNN")
```

This `fastmnn_UMAP` certainly looks different from the one we made from `merged_UMAP`!
What different trends do you see?
Do all samples look "equally well" integrated, from a first look?

Importantly, one reason that batches may still appear separated in the corrected UMAP is if they _should_ be separated - for example, maybe two batches contain very different cell types, have very different diagnoses, or may be from different patients.

Recall from earlier that we conveniently have cell type annotations in our SCEs, so we can explore those here!
Let's take a quick detour to see what kinds of cell types are in this data by making a barplot of the cell types across samples:

```{r explore celltypes}
# Cell types are in the `celltype_broad` and `celltype_fine` columns
merged_sce_df <- as.data.frame(colData(merged_sce))

# Use ggplot2 to make a barplot the cell types across samples
ggplot(merged_sce_df,
       aes(x = sample,
           fill = celltype_broad)) +
  # Barplot of celltype proportions
  geom_bar(position = "fill") +
  # Use a CVD-friendly color scheme
  scale_fill_brewer(palette = "Dark2", na.value = "grey80") +
  # customize y-axis label
  labs(y = "Proportion") +
  # nicer theme
  theme_bw()
```

We see that Tumor cell types are by far the most prevalent across all samples, and normal tissue cell types are not very common.
We see also that `SCPCL000481` has a larger `Tumor_Myocyte` population, while all other samples have larger `Tumor_Mesoderm` populations.
This difference _may_ explain why we observe that `SCPCL000481` is somewhat more separated from the other samples in the `fastMNN` UMAP.

Let's re-plot this UMAP to highlight cell types:


```{r plot fastmnn umap celltypes}
scater::plotReducedDim(merged_sce,
                       dimred = "fastmnn_UMAP",
                       # color by broad celltypes
                       color_by = "celltype_broad",
                       point_size = 0.5,
                       point_alpha = 0.2) +
  # include argument to specify color of NA values
  scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with fastMNN")
```

This UMAP shows that the normal tissue cell types (mostly vascular endothelium, muscle cells, and monocytes) tend to cluster together and are generally separated from the tumor cell types, which is an encouraging pattern!
Tumor cell types from different samples are all also clustering together, which is even more encouraging that we had successful integration.

However, it's a bit challenging to see all the points given the amount of overlap in the plot.
One way we can see all the points a bit better is to facet the plot by sample, using `facet_wrap()` from the `ggplot2` package (which we can do because `scater::plotReducedDim()` returns a `ggplot2` object):

```{r plot fastmnn umap celltypes faceted}
scater::plotReducedDim(merged_sce,
                       dimred = "fastmnn_UMAP",
                       color_by = "celltype_broad",
                       point_size = 0.5,
                       point_alpha = 0.2,
                       # Allow for faceting by a variable using `other_fields`:
                       other_fields = "sample") +
  scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with fastMNN") +
  # Facet by sample
  facet_wrap(vars(sample)) +
  # Use a theme with background grid to more easily compare panel coordinates
  theme_bw()
```

What trends do you observe between tumor and healthy tissues among these integrated samples?


### Integration with `harmony`

`fastMNN` is only one of many approaches to perform integration, and different methods have different capabilities and may give different results.
For example, some methods can accommodate additional covariates (e.g., technology, patient, diagnosis, etc.) that can influence integration.
In fact the data we are using has a known _patient_ covariate; `SCPCL000479` and `SCPCL000480` are from the first patient, and `SCPCL000481` and `SCPCL000482` are from the second patient.

So, let's perform integration with a method that can use this information - [`harmony`](https://portals.broadinstitute.org/harmony/)!

To begin setting up for `harmony` integration, we need to add explicit patient information into our merged SCE.
We'll create a new column `patient` whose value is either "A" or "B" depending on the given sample name, using the [`dplyr::case_when()`](https://dplyr.tidyverse.org/reference/case_when.html) function.
We provide this function with a set of logical expressions and each assigned value is designated by `~`.
The expressions are evaluated in order, stopping at the _first_ one that evaluates as `TRUE` and returning the associated value.

```{r add patient info}
# Create patient column with values "A" or "B" for the two patients
merged_sce$patient <- dplyr::case_when(
  merged_sce$sample %in% c("SCPCL000479", "SCPCL000480") ~ "A",
  merged_sce$sample %in% c("SCPCL000481", "SCPCL000482") ~ "B",
)
```


Unlike `fastMNN`, `harmony` does not calculate corrected expression values nor does it return an SCE object.
Like `fastMNN`, `harmony` performs integration on a merged PCA matrix.
However, unlike `fastMNN`, `harmony` does not "back-calculate" corrected expression from the corrected PCA matrix and it only returns the corrected PCA matrix itself.
For input, `harmony` needs a couple pieces of information:

- First, `harmony` takes a batch-weighted PCA matrix to perform integration.
We already calculated a batch-weighted PCA matrix (our `merged_PCA` reduced dimension), we'll provide this as the the input.
- Second, we need to tell `harmony` about the covariates to use - `sample` and `patient`.
To do this, we provide two arguments:
  - `meta_data`, a data frame that contains covariates across samples.
  We can simply specify the SCE `colData` here since it contains `sample` and `patient` columns.
  - `vars_use`, a vector of which column names in `meta_data` should actually be used as covariates.
  Other columns in `meta_data` which are not in `vars_use` are ignored.

Let's go!

```{r run harmony, live = TRUE}
# Run harmony integration
harmony_pca <- harmony::RunHarmony(
  data_mat = reducedDim(merged_sce, "merged_PCA"),
  meta_data = colData(merged_sce),
  vars_use = c("sample", "patient")
)
```

The result is a PCA matrix.
Let's print a subset of this matrix to see it:

```{r print harmony result, live = TRUE}
# Print the harmony result
harmony_pca[1:5, 1:5]
```

As we did with `fastMNN` results, let's store this PCA matrix directly in our `merged_sce` object with an informative name that won't overwrite any of the existing PCA matrices.
We'll also calculate UMAP from it.

```{r save harmony, live = TRUE}
# Store PCA as `harmony_PCA`
reducedDim(merged_sce, "harmony_PCA") <- harmony_pca

# As before, calculate UMAP on this PCA matrix with appropriate names
merged_sce <- scater::runUMAP(merged_sce,
                              dimred = "harmony_PCA",
                              name   = "harmony_UMAP")
```


Let's see how the `harmony` UMAP, colored by sample, looks compared to the `fastMNN` UMAP:

```{r plot harmony umap batches}
scater::plotReducedDim(merged_sce,
                       dimred = "harmony_UMAP",
                       color_by = "sample",
                       point_size = 0.5,
                       point_alpha = 0.2) +
  scale_color_brewer(palette = "Dark2", name = "sample") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with harmony")
```

How do you think this `harmony` UMAP compares to that from `fastMNN` integration?

Let's see how this UMAP looks colored by cell type, and faceted for visibility:

```{r plot harmony umap celltypes}
scater::plotReducedDim(merged_sce,
                       dimred = "harmony_UMAP",
                       color_by = "celltype_broad",
                       point_size = 0.5,
                       point_alpha = 0.2,
                       # Specify variable for faceting
                       other_fields = "sample") +
  scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") +
  guides(color = guide_legend(override.aes = list(size = 3))) +
  ggtitle("UMAP after integration with harmony") +
  facet_wrap(vars(sample))
```

What do you now notice in this faceted view that wasn't clear previously?
Are there other patterns you see that are similar or different from the `fastMNN` UMAP?
How do you think `fastMNN` vs. `harmony` performed in integrating these samples?

### Export

Finally, we'll export the final SCE object with both `fastMNN` and `harmony` integration to a file.
Since this object is very large (over 1 GB!), we'll export it to a file with some compression, which, in this case, will reduce the final size to a smaller ~360 MB.
This will take a couple minutes to save while compression is performed.

```{r save integration, live = TRUE}
# Export to RDS file with "gz" compression
readr::write_rds(merged_sce,
                 integrated_sce_file,
                 compress = "gz")
```


## Print session info

As always, we'll print the session info to be transparent about what packages, and which versions, were used during this R session.

```{r sessioninfo}
sessionInfo()
```

    +
    ---
title: "Integrating scRNA-seq datasets"
author: Data Lab for ALSF
date: 2023
output:
  html_notebook:
    toc: true
    toc_depth: 3
    toc_float: true
---

## Objectives

This notebook will demonstrate how to:

- Prepare SCE objects for integration
- Apply integration methods including `fastMNN` and `harmony`
- Visually explore the results of integration
- Use `purrr::map()` functions for iterating over lists

---

In this notebook, we'll perform integration on scRNA-seq datasets from the [Single-cell Pediatric Cancer Atlas (`ScPCA`)](https://scpca.alexslemonade.org/), a database of uniformly-processed pediatric scRNA-seq data built and maintained by the Data Lab.
The `ScPCA` database currently hosts single-cell pediatric cancer transcriptomic data generated by ALSF-funded labs, with the goal of making this data easily accessible to investigators (like you!).
The expression data in `ScPCA` were mapping and quantified with [`alevin-fry`](https://doi.org/10.1038/s41592-022-01408-3), followed by processing with Bioconductor tools using the same general procedures that we have covered in this workshop.
The processing pipeline used `emptyDropsCellRanger()` and `miQC` to filter the raw counts matrix, `scuttle` to log-normalize the counts, and `scater` for dimension reduction.
The processed data are stored as `.rds` files containing `SingleCellExperiment` objects.
You can read more about how data in the `ScPCA` is processed in [the associated documentation](https://scpca.readthedocs.io/en/latest/).


![Single-cell roadmap: Integration Overview](diagrams/roadmap_multi_merge-integrate.png)

To learn about integration, we'll have a look at four samples from the [`SCPCP000005` project](https://scpca.alexslemonade.org/projects/SCPCP000005) ([Patel _et al._ 2022](https://doi.org/10.1016/j.devcel.2022.04.003)), an investigation of pediatric solid tumors led by the [Dyer](https://www.stjude.org/research/labs/dyer-lab.html) and [Chen](https://www.stjude.org/research/labs/chen-lab-taosheng.html) labs at St. Jude Children's Research Hospital.
The particular libraries we'll integrate come from two rhabdomyosarcoma (RMS) patients, with two samples from each of two patients, all sequenced with 10x Chromium v3 technology.
Each library is from a separate biological sample.

We'll be integrating these samples with two different tools, [`fastMNN`](http://www.bioconductor.org/packages/release/bioc/html/batchelor.html) ([Haghverdi _et al._ 2018](https://doi.org/10.1038/nbt.4091)) and [`harmony`](https://portals.broadinstitute.org/harmony/) ([Korsunsky _et al._ 2019](https://doi.org/10.1038/s41592-019-0619-0)).
Integration corrects for batch effects that arise from different library preparations, genetic backgrounds, and other sample-specific factors, so that datasets can be jointly analyzed at the cell level.
`fastMNN` corrects for batch effects using a faster variant of the mutual-nearest neighbors algorithm, the technical details of which you can learn more about from this [vignette by Lun (2019)](https://marionilab.github.io/FurtherMNN2018/theory/description.html).
`harmony`, on the other hand, corrects for batch effects using an iterative clustering approach, and unlike `fastMNN`, it is also able to consider additional covariates beyond just the batch groupings.

Regardless of which integration tool is used, the `SingleCellExperiment` (SCE) objects first need to be reformatted and merged into a single (uncorrected!) SCE object that contains all cells from all samples.
This merged SCE can then be used for integration to obtain a formally batch-corrected SCE object.


## Set up

```{r setup}
# Load libraries
library(ggplot2)  # plotting tools
library(SingleCellExperiment) # work with SCE objects

# Set the seed for reproducibility
set.seed(12345)
```


### Define directories and files


We have already prepared count data for the four samples we'll be integrating (i.e., filtered cells, normalized counts, and calculated PCA & UMAP).
These SCE objects, stored as RDS files, are available in the `data/rms/processed/` directory and are named according to their `ScPCA` library ids :

- `SCPCL000479.rds` (Patient A)
- `SCPCL000480.rds` (Patient A)
- `SCPCL000481.rds` (Patient B)
- `SCPCL000482.rds` (Patient B)

Both Patient A (18 year old male) and Patient B (4 year old female) had recurrent embryonal rhabdomyosarcoma when samples were taken.

To begin, let's set up our directories and files:

```{r directories}
# Define directory where processed SCE objects to be integrated are stored
input_dir <- file.path("data", "rms", "processed")

# Define directory to save integrated SCE object to
output_dir <- file.path("data", "rms", "integrated")

# Create output directory if it doesn't exist
fs::dir_create(output_dir)

# Define output file name for the integrated object
integrated_sce_file <- file.path(output_dir, "rms_integrated_subset.rds")
```


We can use the `dir()` function to list all contents of a given directory, for example to see all the files in our `input_dir`:

```{r input dir, live = TRUE}
dir(input_dir)
```

We want to read in just four of these files, as listed previously.
To read in these files, we could use the `readr::read_rds()` function (or the base R `readRDS()`) four times, once for each of the files.
We could also use a `for` loop, which is the approach that many programming languages would lean toward.
A different and more modular coding approach to reading in these files (and more!) is to leverage the [`purrr`](https://purrr.tidyverse.org/) `tidyverse` package, which provides a convenient set of functions for operating on lists.
You can read more about the `purrr` functions and their power and utility in R in [the "Functionals" chapter of the _Advanced R_ e-book](https://adv-r.hadley.nz/functionals.html).

Of particular interest is the [`purrr::map()`](https://purrr.tidyverse.org/reference/map.html) family of functions, which can be used to run a given function on each element of a list (or vector) in one call.
The general syntax for `purrr::map()` and friends is:

```
# Syntax for using the map function:
purrr::map(<input list or vector>,
           <function to apply to each item in the input>,
           <any additional arguments to the function can go here>,
           <and also here if there are even more arguments, and so on>)
```


The output from running `purrr::map()` is always a list (but note that there are other `purrr::map()` relatives which return other object types, as you can read about in [the `purrr::map()` documentation](https://purrr.tidyverse.org/reference/index.html)).
If this concept sounds a little familiar to you, that's because it probably is!
Base R's `lapply()` function can provide similar utility, and the `purrr::map()` family of functions can (in part) be thought of as an alternative to some of the base R `apply` functions, with more consistent behavior.

Let's see a very simple example of `purrr::map()` in action, inspired by cancer groups the Data Lab has analyzed through the [OpenPBTA](https://github.com/AlexsLemonade/OpenPBTA-analysis/) project:

```{r map example}
# Define a list of cancer histologies
histologies <- list(
  "low-grade gliomas"  = c("SEGA", "PA", "GNG", "PXA"),
  "high-grade gliomas" = c("DMG", "DIPG"),
  "embryonal tumors"   = c("MB", "ATRT", "ETMR")
 )

# The overall length of the list is 3
length(histologies)

# How can we run `length()` on each item of the list?
# We can use our new friend purrr::map():
purrr::map(histologies, length)
```

Let's use `purrr::map()` to read in our SCE objects so that they are immediately stored together in a list.


We'll first need to define a vector of the file paths to read in.
We'll start by creating a vector of sample names themselves and then formatting them into the correct paths.
This way (foreshadowing!) we also have a stand-alone vector of just sample names, which will come in handy!

```{r sample names}
# Vector of all the samples to read in:
sample_names <- c("SCPCL000479",
                  "SCPCL000480",
                  "SCPCL000481",
                  "SCPCL000482")
```


```{r define sce_paths, live = TRUE}
# Now, convert these to file paths: <input_dir>/<sample_name>.rds
sce_paths <- file.path(input_dir,
                       glue::glue("{sample_names}.rds")
)
# Print the sce_paths vector
sce_paths
```

Let's make this a named vector using the sample names.
This will help us keep track of which objects are which after we read the SCE objects in:

```{r add list names, live = TRUE}
# Assign the sample names as the names for sce_paths
names(sce_paths) <- sample_names
```

We can now read these files in and create a list of four SCE objects. 
Since `readr::read_rds()` can only operate on one input at a time, we'll need to use `purrr::map()` to run it on all input file paths in one command.
Although `sce_paths` is a vector (not a list), it will still work as input to `purrr:map()`.
The output from this code will still be a list, since that's what `purrr::map()` always returns, and it will retain the sample names as the list names for convenient bookkeeping:

```{r read sce paths, live = TRUE}
# Use purrr::map() to read all files into a list at once
sce_list <- purrr::map(
  sce_paths,
  readr::read_rds
)
```

Let's have a look at our named list of SCE objects:

```{r print sce list, live=TRUE}
# Print sce_list
sce_list
```

If you look closely at the printed SCE objects, you may notice that they all contain `colData` table columns `celltype_fine` and `celltype_broad`.
These columns (which we added to SCE objects during [pre-processing](https://github.com/AlexsLemonade/training-modules/tree/master/scRNA-seq-advanced/setup/rms)) contain putative cell type annotations as assigned by [Patel _et al._ (2022)](https://doi.org/10.1016/j.devcel.2022.04.003):


> For each cell subset identified by clustering, we used a combination of `SingleR` version 1.0.1 ([Aran et al., 2019](https://www.nature.com/articles/s41590-018-0276-y)) and manual inspection of differentially expressed genes to annotate whether a cluster belongs to stromal, immune or malignant subpopulations. 
Malignant cells were confirmed in patient tumor data by inference of copy-number variation using `inferCNV` version 1.1.3 of the TrinityCTAT Project (https://github.com/broadinstitute/infercnv). 

We will end up leveraging these cell type annotations to explore the integration results; after integration, we expect cell types from different samples to group together, rather than being separated by batches. 

That said, the integration methods we will be applying _do not actually use_ any existing cell type annotations.
If we have annotations, they are a helpful "bonus" for assessing the integration's performance, but they are not part of the integration itself.


## Merge the SCE list into one object

![Single-cell roadmap: Merge](diagrams/roadmap_multi_merge.png)


Now that we have a list of processed SCE objects, we need to merge the objects into one overall SCE object for input to integration.
A word of caution before we begin: **This merged SCE object is NOT an integrated SCE!**
Merging SCEs does not perform any batch correction, but just reorganizes the data to allow us to proceed to integration next.

To merge SCE objects, we do need to do some wrangling and bookkeeping to ensure compatibility and that we don't lose important information.
Overall, we'll want to make sure that:

1. All objects have compatible dimensions.
This means that all objects should...
    + Have the same genes (aka row names), in the same order
    + Have the same `colData` slot columns, in the same order
    + Have the same assays
2. After merging, we'll still be able to identify which sample different pieces of information came from
As we saw in the slides, this means we'll have to...
    + Attach sample names to the barcodes (aka column names)
    This also ensures that column names are unique; while a single sample (library) is guaranteed to have unique barcodes, technically they can be repeated across samples!
    + Attach sample names to `rowData` slot column names and `metadata` field names (if you care to keep this information around - today, we will!)
    + Add a new column indicating the sample to the `colData` slot

We'll approach this merge in two parts:

+ First, we'll take some time to thoroughly explore the our SCE objects to determine what wrangling we need to do to make all the objects _compatible_ for merging
+ Then, we'll write (ok, we've written it for you) a _custom function_ to format each SCE object for merging, including:
    + Making any changes to ensure objects are compatible
    + Adding in identifying information so we know which sample the cells and other metadata came from 
    + Removing the existing reduced dimension matrices (PCA and UMAP).
    This is because we'll want to recalculate these matrices on the merged objects, taking batch into account

When merging objects on your own, don't skip these data exploration steps!
The steps we take to prepare our SCEs will probably be different from the steps you need to take with other SCEs, and only by carefully exploring the objects can you figure out what steps you'll need to take to meet all of our conditions.


### Prepare to merge SCEs

#### Create unique cell identifiers

As part of the custom function we'll write, we'll include a step to create unique cell identifiers by attaching sample names to the SCE column names (cell barcodes).
For example, we would update the column name for a cell from `Sample1` with the barcode `ACGT` to `Sample1-ACGT`.

When merging, there can't be any duplicate column names (barcodes) across _all_ the objects or R will throw an error.
While you're guaranteed to have unique barcodes in a given SCE object, there is _no guarantee_ that they are unique across multiple samples - it is absolutely possible to have cells from two different samples share the same barcode (and we've seen it happen!).

Adding the sample id to the column names (barcodes) is therefore a crucial step in our merging bookkeeping.


#### Explore the SCE objects

##### Check the genes

First, we'll compare the object's genes (aka, their row names).
We can use some `purrr` magic to help us find the set of shared genes among all objects:

```{r shared genes}
# Define vector of shared genes
shared_genes <- sce_list |>
  # get rownames (genes) for each SCE in sce_list
  purrr::map(rownames) |>
  # reduce to the _intersection_ among lists
  purrr::reduce(intersect)

# How many shared genes are there?
length(shared_genes)
```

That's quite a lot!
In fact, because these objects were all uniformly processed by the same workflow (which did not filter out any genes!), we expect them to all have the same genes.
We can map over the list to confirm that indeed, they have the same number of rows (genes):


```{r check shared genes, live = TRUE}
# The number of genes in an SCE corresponds to its number of rows:
sce_list |>
  purrr::map(nrow)
```

Even though we know the genes already match, we need to also be sure they are in the same _order_ among all objects.
So, we'll hold onto that `shared_genes` variable we defined and use it soon in our custom formatting function to make sure all objects fully match.

It's worth noting that the intersection isn't the only option here, though!
Using the intersection means a lot of genes will get discarded if the objects have different genes.
We could instead take the _union_ of genes so nothing gets thrown out.
In this case, you'd need to create "dummy" assay rows for genes that a given SCE doesn't have and fill it with `NA` expression values.
You'll still have to make sure the SCEs have the same rows in the same order before merging, so you may need to do a decent bit of matrix wrangling.

##### Check the `colData` column names

Next up, we'll check the `colData` columns: we need these to be the same, and in the same order.
Let's print out each object's `colData` column name to see where we stand:

```{r coldata colnames}
sce_list |>
  purrr::map(
    \(sce) colnames(colData(sce)) 
  ) 
```
We see the same columns all around in the same order, which is great!

But what if there were different columns across objects, or they were differently ordered?
In that case, we could find the intersection of column names like we did above for genes, and use that to re-order and subset all `colData` slots in our custom formatting function.


##### Check the assays

Next, we'll make sure that all objects share the same assays:

```{r assay names, live = TRUE}
# print all the assay names
sce_list |>
  purrr::map(assayNames)
```
Again, all objects are compatible already with both having a `counts` and `logcounts` assay.

In your own data exploration, if you encounter SCEs to merge that have extraneous assays that you don't need, you can remove them by setting them to `NULL` in your custom formatting function, e.g. `assay(sce, "assay_to_remove") <- NULL`.

##### Check the `rowData` contents

One of the other items we said we'd need to think about is the `rowData`, which contains gene metadata.
This slot is interesting because some of its columns are specific to the given sample, while others are general:

```{r little head rowdata}
sce_list |>
  purrr::map(
    \(sce) head(rowData(sce), 3) # only print 3 rows for space!
  )
```

The column `gene_symbol` is not sample-specific - it just provides the corresponding gene symbol to the Ensembl ids seen here as row names.
The columns `mean` and `detected`, however, are sample-specific - they contain sample-specific statistics about gene expression.

This means we definitely need to update the column names `mean` and `detected` to include the sample id.
But, we don't need a separate `gene_symbol` column for each sample, so we can leave that one alone as just `gene_symbol`. 
Once we eventually merge, only one `gene_symbol` column will be left in the final object since it is the same across all the SCEs.

We'll show one way to do this in our custom function, but it's worth noting there's nothing _wrong_ with also adding the sample id to the `gene_symbol` column; you'll just end up with a bunch of redundant gene symbol columns.


#### Reformat the SCE objects

As you can see, there's a lot of moving parts to consider!
Again, these moving parts may (will!) differ for SCEs that you are working with, so you have to explore your own SCEs in depth to prepare for merging.

Based on our exploration, here is a schematic of how one of the SCE objects will ultimately be modified into the final merged SCE:

![](diagrams/technical_merge_sce.png)


We'll write a _custom function_ (seen in the chunk below) tailored to our wrangling steps that prepares a single SCE object for merging.
We'll then use our new `purrr::map()` programming skills to run this function over the `sce_list`.
This will give us a new list of formatted SCEs that we can proceed to merge.
It's important to remember that the `format_sce()` function written below is not a function for general use – it's been precisely written to match the processing we need to do for _these_ SCEs, and different SCEs you work with will require different types of processing.

We also include roxygen-style comments for this function, which can be a helpful consistent way to document your code if you like it - we've even written a blog post about it :) (<https://www.ccdatalab.org/blog/dont-make-me-write-tips-for-avoiding-typing-in-rstudio>).

```{r format_sce function}
#' Custom function to format an SCE before merging
#'
#' @param sce SCE object to format
#' @param sample_name Name of the sample
#' @param shared_genes Vector of shared genes across all SCE objects
#'
#' @returns An updated SCE object ready for merging
format_sce <- function(
  sce, 
  sample_name, 
  shared_genes
) {
  
  ### Remove the single-sample reduced dimensions 
  # We do this first since it makes the object a lot smaller for the rest of this code!
  reducedDims(sce) <- NULL
  
  ### Add dedicated sample indicator column to the colData slot
  # Recall, the `sce$` shortcut points to the colData
  sce$sample <- sample_name

  ### Ensure objects have the same genes in the same order
  # Use the shared_genes vector to index genes to the intersection
  # Doing this both subsets to just those genes, and reorders!
  sce <- sce[shared_genes, ]
  
  ### There is no additional wrangling to do for the colData column names or assays.
  ### But if there were, you could add your custom code to do so here.
  ### Your custom function may need additional arguments for this, too.

  ### Ensure cell ids are identifiable and fully unique 
  # Update the SCE object column names (cell ids) by prepending the `sample_name`
  colnames(sce) <- glue::glue("{sample_name}-{colnames(sce)}")

  ### Ensure the rowData columns can be identified
  # Recall, we want to leave `gene_symbol` alone, but add the `sample_name` to the rest
  rowdata_names <- colnames(rowData(sce))
  # prefix rowData names with the sample name, except for gene symbols
  new_rowdata_names <- ifelse(
    rowdata_names == "gene_symbol",
    "gene_symbol",
    glue::glue("{sample_name}-{rowdata_names}")
  )
  colnames(rowData(sce)) <- new_rowdata_names
  
  ### Ensure metadata slot fields can be identified
  # We'll simply prepend the `sample_name` to all fields for this slot
  names(metadata(sce)) <- glue::glue("{sample_name}-{names(metadata(sce))}")
  
  
  ### Finally, we can return the formatted SCE object
  return(sce)  
}
```

To run this function, we'll use the `purrr::map2()` function, a relative of `purrr::map()` that allows you to loop over _two_ input lists/vectors.
In our case, we want to run `format_sce()` over paired `sce_list` items and `sce_list` names.

```{r format sces for merge, live = TRUE}
# We can use `purrr::map2()` to loop over two list/vector arguments
sce_list_formatted <- purrr::map2(
  # Each "iteration" will march down the first two
  #  arguments `sce_list` and `names(sce_list)` in order
  sce_list,
  names(sce_list),
  \(sce, sample_name) format_sce(sce, sample_name, shared_genes) 
)

# Print formatted SCE list
sce_list_formatted
```

(Psst, like `purrr` and want to dive deeper? Check out [the `purrr::imap()` function](https://purrr.tidyverse.org/reference/imap.html)!)


### Perform the merging


At long last, we are ready to merge the SCEs, which we'll do using the R function `cbind()`.
The `cbind()` function is often used to combine data frames or matrices by column, i.e. "stack" them next to each other.
The same principle applies here, but when run on SCE objects, `cbind()` will create a new SCE object by combining `counts` and `logcounts` matrices by column.
Following that structure, other SCE slots (`colData`, `rowData`, reduced dimensions, and other metadata) are combined appropriately.

Since we need to apply `cbind()` to a _list_ of objects, we need to use some slightly-gnarly syntax: We'll use the function `do.call()`, which allows the `cbind()` input to be a list of objects to combine.

```{r merges sces, live = TRUE}
# Merge SCE objects
merged_sce <- do.call(cbind, sce_list_formatted)

# Print the merged_sce object
merged_sce
```

We now have a single merged SCE object that contains all cells from all samples we'd like to integrate.

Let's take a peek at some of the innards of this new SCE object:

```{r explore merged_sce, live = TRUE}
# How many samples, and cells per sample?
table(colData(merged_sce)$sample)

# What are the new cell ids (column names)?
head(colnames(merged_sce))
tail(colnames(merged_sce))

# What does rowData look like?
head(rowData(merged_sce))
```


## Integrate samples

![Single-cell roadmap: Integrate](diagrams/roadmap_multi_integrate.png)

So far, we've created a `merged_sce` object which is (almost!) ready for integration.

The integration methods we'll be using here actually perform batch correction on a reduced dimension representation of the normalized gene expression values, which is more efficient.
`fastMNN` and `harmony` specifically use PCA for this, but be aware that different integration methods may use other kinds of reduced dimensions.

Before merging, our objects had reduced dimension representations calculated on each individual SCE, and we removed them when preparing for merge.
We removed them because we don't actually want to use them anymore!
This is because part of their calculation involves scaling the raw data to center the mean.
When samples are separately centered, _all_ of them will be centered at zero, making it look like the datasets are already pretty overlapping when you plot their UMAPs together.
But, this is just a mathematical artifact of how dimension reduction is performed.

So, we'll begin by re-calculating PCA and UMAP on the merged object in a way that takes batches into consideration. 
For input to integration, we'll want the reduced dimension calculations to consider normalized gene expression values from all samples simultaneously.
So we'll need to recalculate PCA (and UMAP for visualization) on the merged object.

First, as usual, we'll determine the high-variance genes to use for PCA from the `merged_sce` object.
For this, we'll need to provide the argument `block = merged_sce$sample` when modeling gene variance, which tells `scran::modelGeneVar()` to first model variance separately for each batch and then combine those modeling statistics.
(Psst: isn't it handy we created that `sample` column when merging?!)

```{r calc merged hv genes}
# Specify the number of genes to identify
num_genes <- 2000

# Calculate variation for each gene
gene_variance <- scran::modelGeneVar(merged_sce,
                                     # specify the grouping column:
                                     block = merged_sce$sample)

# Get the top `num_genes` high-variance genes to use for dimension reduction
hv_genes <- scran::getTopHVGs(gene_variance,
                              n = num_genes)
```

To calculate the PCA matrix itself, we'll use an approach from the `batchelor` package, which is the R package that contains the `fastMNN` method.
The [`batchelor::multiBatchPCA()`](https://rdrr.io/bioc/batchelor/man/multiBatchPCA.html) function calculates a batch-weighted PCA matrix.
This weighting ensures that all batches, which may have very different numbers of cells, contribute equally to the overall scaling.

```{r make merged_pca, live = TRUE}
# Use batchelor to calculate PCA for merged_sce, considering only
#  the high-variance genes
# We'll need to include the argument `preserve.single = TRUE` to get
#  a single matrix with all samples and not separate matrices for each sample
merged_pca <- batchelor::multiBatchPCA(merged_sce,
                                       subset.row = hv_genes,
                                       batch = merged_sce$sample,
                                       preserve.single = TRUE)
```

Let's have a look at the output:
```{r print merged_pca, live = TRUE}
# This output is not very interesting!
merged_pca
```

We can use indexing `[[1]]` to see the PCA matrix calculated, looking at a small subset for convenience:

```{r print merged_pca indexed, live = TRUE}
merged_pca[[1]][1:5,1:5]
```

We can now include this PCA matrix in our `merged_sce` object:

```{r add merged_pca, live = TRUE}
# add PCA results to merged SCE object
reducedDim(merged_sce, "PCA") <- merged_pca[[1]]
```

Now that we have the PCA matrix, we can proceed to calculate UMAP to visualize the uncorrected merged data.

```{r calculate merged umap, live = TRUE}
merged_sce <- scater::runUMAP(merged_sce)
```

```{r plot uncorrected merged UMAP}
# UMAPs scaled together when calculated from the merged SCE
scater::plotUMAP(
  merged_sce,
  color_by = "sample",
  # Some styling to help us see the points:
  point_size = 0.5,
  point_alpha = 0.2
) +
  scale_color_brewer(palette = "Dark2", name = "sample") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP calculated on merged_sce")
```

We see (mostly) four separate clumps representing the four different _merged but not yet integrated_ samples.
We can think of this UMAP as our "before" UMAP, and we can compare this to the "after" UMAP we see post-integration.

Let's discuss a little first: What visual differences do you think the UMAP on the integrated version of data will have?
What similarities do you think the integrated UMAP will have to this plot?


### Integrate with `fastMNN`

Finally, we're ready to integrate!
To start, we'll use the `fastMNN` approach from the Bioconductor [`batchelor` package](http://www.bioconductor.org/packages/release/bioc/html/batchelor.html).

`fastMNN` takes as input the `merged_sce` object to integrate, and the first step it performs is actually to run `batchelor::multiBatchPCA()` on that SCE.
It then uses that batch-weighted PCA matrix to perform the actual batch correction.
The `batch` argument is used to specify the different groupings within the `merged_sce` (i.e. the original sample that each cell belongs to), and the `subset.row` argument can optionally be used to provide a vector of high-variance genes that should be considered for this PCA calculation.
`fastMNN` will return an SCE object that contains a batch-corrected PCA.
Let's run it and save the result to a variable called `integrated_sce`.


```{r run fastmnn, live = TRUE}
# integrate with fastMNN, again specifying only our high-variance genes
integrated_sce <- batchelor::fastMNN(
  merged_sce,
  batch = merged_sce$sample,
  subset.row = hv_genes
)
```

Let's have a look at the result:

```{r fastmnn result, live = TRUE}
# Print the integrated_sce object
integrated_sce
```

There are couple pieces of information here of interest:

- The `corrected` reduced dimension represents the batch-corrected PCA that `fastMNN` calculated.
- The `reconstructed` assay represents the batch-corrected normalized expression values, which `fastMNN` "back-calculated" from the batch-corrected PCA (`corrected`).
Generally speaking, these expression values are not stand-alone values that you should use for other applications like differential gene expression, as described in [_Orchestrating Single Cell Analyses_](http://bioconductor.org/books/3.19/OSCA.multisample/using-corrected-values.html).
If the `subset.row` argument is provided (as it was here), only genes present in `subset.row` will be included in these reconstructed expression values, but this setting can be overridden so that all genes have reconstructed expression with the argument `correct.all = TRUE`.

We're mostly interested in the PCA that `fastMNN` calculated, so let's save that information (with an informative and unique name) into our `merged_sce` object:

```{r fastmnn pcs, live = TRUE}
# Make a new reducedDim named fastmnn_PCA from the corrected reducedDim in integrated_sce
reducedDim(merged_sce, "fastmnn_PCA") <- reducedDim(integrated_sce, "corrected")
```

Finally, we'll calculate UMAP from these corrected PCA matrix for visualization.
In this case we need to specify two additional arguments since we're working with non-standard reduced dimension names:

+ `dimred = "fastmnn_PCA"`, which specifies the existing reduced dimension to use for the calculation
+ `name = "fastmnn_UMAP"`, which names the final UMAP that this function calculates

```{r calculate fastmnn umap, live = TRUE}
# Calculate UMAP
merged_sce <- scater::runUMAP(
  merged_sce,
  dimred = "fastmnn_PCA",
  name = "fastmnn_UMAP"
)
```

First, let's plot the integrated UMAP highlighting the different batches.
A well-integrated dataset will show batch mixing, but a poorly-integrated dataset will show more separation among batches, similar to the uncorrected UMAP.
Note that this is a more qualitative way to assess the success of integration, but there are formal metrics one can use to assess batch mixing, which you can read more about in [this chapter of OSCA](http://bioconductor.org/books/3.19/OSCA.multisample/correction-diagnostics.html).

```{r plot fastmnn umap batches}
scater::plotReducedDim(merged_sce,
                       # plot the fastMNN coordinates
                       dimred = "fastmnn_UMAP",
                       # color by sample
                       color_by = "sample",
                       # Some styling to help us see the points:
                       point_size = 0.5,
                       point_alpha = 0.2) +
  scale_color_brewer(palette = "Dark2", name = "sample") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with fastMNN")
```

This `fastmnn_UMAP` certainly looks different from the one we made before integrating!
What different trends do you see?
Do all samples look "equally well" integrated, from a first look?

Importantly, one reason that batches may still appear separated in the corrected UMAP is if they _should_ be separated - for example, maybe two batches contain very different cell types, have very different diagnoses, or may be from different patients.

Recall from earlier that we conveniently have cell type annotations in our SCEs, so we can explore those here!
Let's take a quick detour to see what kinds of cell types are in this data by making a barplot of the cell types across samples:

```{r explore celltypes}
# Cell types are in the `celltype_broad` and `celltype_fine` columns
merged_sce_df <- as.data.frame(colData(merged_sce))

# Use ggplot2 to make a barplot the cell types across samples
ggplot(merged_sce_df,
       aes(x = sample,
           fill = celltype_broad)) +
  # Barplot of celltype proportions
  geom_bar(position = "fill") +
  # Use a CVD-friendly color scheme
  scale_fill_brewer(palette = "Dark2", na.value = "grey80") +
  # customize y-axis label
  labs(y = "Proportion") +
  # nicer theme
  theme_bw()
```

We see that Tumor cell types are by far the most prevalent across all samples, and normal tissue cell types are not very common.
We see also that `SCPCL000481` has a larger `Tumor_Myocyte` population, while all other samples have larger `Tumor_Mesoderm` populations.
This difference _may_ explain why we observe that `SCPCL000481` is somewhat more separated from the other samples in the `fastMNN` UMAP.

Let's re-plot this UMAP to highlight cell types:


```{r plot fastmnn umap celltypes}
scater::plotReducedDim(merged_sce,
                       dimred = "fastmnn_UMAP",
                       # color by broad celltypes
                       color_by = "celltype_broad",
                       point_size = 0.5,
                       point_alpha = 0.2) +
  # include argument to specify color of NA values
  scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with fastMNN")
```

This UMAP shows that the normal tissue cell types (mostly vascular endothelium, muscle cells, and monocytes) tend to cluster together and are generally separated from the tumor cell types, which is an encouraging pattern!
Tumor cell types from different samples are all also clustering together, which is even more encouraging that we had successful integration.

However, it's a bit challenging to see all the points given the amount of overlap in the plot.
One way we can see all the points a bit better is to facet the plot by sample, using `facet_wrap()` from the `ggplot2` package (which we can do because `scater::plotReducedDim()` returns a `ggplot2` object):

```{r plot fastmnn umap celltypes faceted}
scater::plotReducedDim(merged_sce,
                       dimred = "fastmnn_UMAP",
                       color_by = "celltype_broad",
                       point_size = 0.5,
                       point_alpha = 0.2,
                       # Allow for faceting by a variable using `other_fields`:
                       other_fields = "sample") +
  scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with fastMNN") +
  # Facet by sample
  facet_wrap(vars(sample)) +
  # Use a theme with background grid to more easily compare panel coordinates
  theme_bw()
```

What trends do you observe between tumor and healthy tissues among these integrated samples?

### Integrate with `harmony`

`fastMNN` is only one of many approaches to perform integration, and different methods have different capabilities and may give different results.
For example, some methods can accommodate additional covariates (e.g., technology, patient, diagnosis, etc.) that can influence integration.
In fact the data we are using has a known _patient_ covariate; `SCPCL000479` and `SCPCL000480` are from Patient A, and `SCPCL000481` and `SCPCL000482` are from Patient B.

So, let's perform integration with a method that can use this information - [`harmony`](https://portals.broadinstitute.org/harmony/)!

To begin setting up for `harmony` integration, we need to add explicit patient information into our merged SCE.
We'll create a new column `patient` whose value is either "A" or "B" depending on the given sample name, using the [`dplyr::case_when()`](https://dplyr.tidyverse.org/reference/case_when.html) function.
We provide this function with a set of logical expressions and each assigned value is designated by `~`.
The expressions are evaluated in order, stopping at the _first_ one that evaluates as `TRUE` and returning the associated value.

```{r add patient info}
# Create patient column with values "A" or "B" for the two patients
merged_sce$patient <- dplyr::case_when(
  merged_sce$sample %in% c("SCPCL000479", "SCPCL000480") ~ "A",
  merged_sce$sample %in% c("SCPCL000481", "SCPCL000482") ~ "B",
)
```


Unlike `fastMNN`, `harmony` does not calculate corrected expression values nor does it return an SCE object.
Like `fastMNN`, `harmony` performs integration on a merged PCA matrix.
However, unlike `fastMNN`, `harmony` does not "back-calculate" corrected expression from the corrected PCA matrix and it only returns the corrected PCA matrix itself.
For input, `harmony` needs a couple pieces of information:

- First, `harmony` takes a batch-weighted PCA matrix to perform integration.
We already calculated a batch-weighted PCA matrix so we'll provide this as the the input.
- Second, we need to tell `harmony` about the covariates to use - `sample` and `patient`.
To do this, we provide two arguments:
  - `meta_data`, a data frame that contains covariates across samples.
  We can simply specify the SCE `colData` here since it contains `sample` and `patient` columns.
  - `vars_use`, a vector of which column names in `meta_data` should actually be used as covariates.
  Other columns in `meta_data` which are not in `vars_use` are ignored.

Let's go!

```{r run harmony, live = TRUE}
# Run harmony integration
harmony_pca <- harmony::RunHarmony(
  data_mat = reducedDim(merged_sce, "PCA"),
  meta_data = colData(merged_sce),
  vars_use = c("sample", "patient")
)
```

The result is a PCA matrix.
Let's print a subset of this matrix to see it:

```{r print harmony result, live = TRUE}
# Print the harmony result
harmony_pca[1:5, 1:5]
```

As we did with `fastMNN` results, let's store this PCA matrix directly in our `merged_sce` object with an informative name that won't overwrite any of the existing PCA matrices.
We'll also calculate UMAP from it.

```{r save harmony, live = TRUE}
# Store PCA as `harmony_PCA`
reducedDim(merged_sce, "harmony_PCA") <- harmony_pca

# As before, calculate UMAP on this PCA matrix with appropriate names
merged_sce <- scater::runUMAP(merged_sce,
                              dimred = "harmony_PCA",
                              name   = "harmony_UMAP")
```


Let's see how the `harmony` UMAP, colored by sample, looks compared to the `fastMNN` UMAP:

```{r plot harmony umap batches}
scater::plotReducedDim(merged_sce,
                       dimred = "harmony_UMAP",
                       color_by = "sample",
                       point_size = 0.5,
                       point_alpha = 0.2) +
  scale_color_brewer(palette = "Dark2", name = "sample") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with harmony")
```

How do you think this `harmony` UMAP compares to that from `fastMNN` integration?

Let's see how this UMAP looks colored by cell type, and faceted for visibility:

```{r plot harmony umap celltypes}
scater::plotReducedDim(merged_sce,
                       dimred = "harmony_UMAP",
                       color_by = "celltype_broad",
                       point_size = 0.5,
                       point_alpha = 0.2,
                       # Specify variable for faceting
                       other_fields = "sample") +
  scale_color_brewer(palette = "Dark2", name = "Broad celltype", na.value = "grey80") +
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) +
  ggtitle("UMAP after integration with harmony") +
  facet_wrap(vars(sample))
```

What do you now notice in this faceted view that wasn't clear previously?
Are there other patterns you see that are similar or different from the `fastMNN` UMAP?
How do you think `fastMNN` vs. `harmony` performed in integrating these samples?

## Export

Finally, we'll export the final SCE object with both `fastMNN` and `harmony` integration to a file.
Since this object is very large (over 1 GB!), we'll export it to a file with some compression, which, in this case, will reduce the final size to a smaller ~360 MB.
This will take a couple minutes to save while compression is performed.

```{r save integration, live = TRUE}
# Export to RDS file with "gz" compression
readr::write_rds(
  merged_sce,
  integrated_sce_file,
  compress = "gz"
)
```


## Print session info

As always, we'll print the session info to be transparent about what packages, and which versions, were used during this R session.

```{r sessioninfo}
sessionInfo()
```

    diff --git a/scRNA-seq-advanced/03-differential_expression-live.Rmd b/scRNA-seq-advanced/03-differential_expression-live.Rmd index 82022af2..fd878bf0 100644 --- a/scRNA-seq-advanced/03-differential_expression-live.Rmd +++ b/scRNA-seq-advanced/03-differential_expression-live.Rmd @@ -68,27 +68,35 @@ To begin, let's set up our directories and files: data_dir <- file.path("data", "rms") # integrated file containing samples to use for DE analysis -integrated_sce_file <- file.path(data_dir, - "integrated", - "rms_all_sce.rds") +integrated_sce_file <- file.path( + data_dir, + "integrated", + "rms_all_sce.rds" +) # sample metadata to set up DE analysis -sample_metadata_file <- file.path(data_dir, - "annotations", - "rms_sample_metadata.tsv") +sample_metadata_file <- file.path( + data_dir, + "annotations", + "rms_sample_metadata.tsv" +) # directory to store output deseq_dir <- file.path("analysis", "rms", "deseq") fs::dir_create(deseq_dir) # results file to output from DE analysis -deseq_output_file <- file.path(deseq_dir, - "rms_myoblast_deseq_results.tsv") +deseq_output_file <- file.path( + deseq_dir, + "rms_myoblast_deseq_results.tsv" +) # output integrated sce object -output_sce_file <- file.path(data_dir, - "integrated", - "rms_subset_sce.rds") +output_sce_file <- file.path( + data_dir, + "integrated", + "rms_subset_sce.rds" +) ``` We can go ahead and read in the SCE object and the metadata file. @@ -172,17 +180,21 @@ coldata_df <- colData(integrated_sce) |> dplyr::left_join(sample_metadata, by = c("sample" = "library_id")) |> # create new columns # cell_id is a combination of barcode and sample - dplyr::mutate(cell_id = glue::glue("{sample}-{barcode}"), - # simplify subdiagnosis - diagnosis_group = forcats::fct_recode( - subdiagnosis, - "ARMS" = "Alveolar rhabdomyosarcoma", - "ERMS" = "Embryonal rhabdomyosarcoma" - )) + dplyr::mutate( + cell_id = glue::glue("{sample}-{barcode}"), + # simplify subdiagnosis + diagnosis_group = forcats::fct_recode( + subdiagnosis, + "ARMS" = "Alveolar rhabdomyosarcoma", + "ERMS" = "Embryonal rhabdomyosarcoma" + ) + ) # add modified data frame back to SCE as DataFrame -colData(integrated_sce) <- DataFrame(coldata_df, - row.names = coldata_df$cell_id) +colData(integrated_sce) <- DataFrame( + coldata_df, + row.names = coldata_df$cell_id +) ``` Now when we look at the `colData` of the SCE object we should see new columns, including the `diagnosis_group` column which indicates if each cell comes from an ERMS or ARMS sample. @@ -218,12 +230,14 @@ The samples which could be further classified have a mix of `Tumor_Mesoderm`, `T ```{r celltype UMAP} # UMAP of all samples labeled by cell type -scater::plotReducedDim(integrated_sce, - dimred = "fastmnn_UMAP", - # color each point by cell type - color_by = "celltype_broad", - point_size = 0.5, - point_alpha = 0.4) + +scater::plotReducedDim( + integrated_sce, + dimred = "fastmnn_UMAP", + # color each point by cell type + color_by = "celltype_broad", + point_size = 0.5, + point_alpha = 0.4 +) + # Modify the legend key with larger, easier to see points guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) ``` @@ -241,9 +255,9 @@ In the below plot we will color our cells by cell type while also using `facet_g # UMAP of all samples # separating by diagnosis group and labeling cell type - # color each point by cell type + # color each point by cell type - # tell scater to use diagnosis_group for plotting + # tell scater to use diagnosis_group for plotting # include each diagnosis group as its own column @@ -261,20 +275,22 @@ tumor_cells_df <- coldata_df |> # create a stacked barplot ggplot(tumor_cells_df, aes(x = sample, fill = celltype_broad)) + - geom_bar(position = "fill", color = "black", size = 0.2) + - labs( - x = "Sample", - y = "Proportion of cells", - fill = "Cell type" - ) + + geom_bar(position = "fill", color = "black", size = 0.2) + + labs( + x = "Sample", + y = "Proportion of cells", + fill = "Cell type" + ) + scale_fill_brewer(palette = "Dark2") + theme_bw() + - theme(axis.text.x = element_text(angle = 90, vjust = 0.5))+ + theme(axis.text.x = element_text(angle = 90, vjust = 0.5)) + # facet by diagnosis group - facet_grid(cols = vars(diagnosis_group), - # only show non-NA values on x-axis - scales = "free_x", - space = "free_x") + facet_grid( + cols = vars(diagnosis_group), + # only show non-NA values on x-axis + scales = "free_x", + space = "free_x" + ) ``` Similar to the UMAP, this plot shows that ARMS and ERMS share a lot of the same cell types. @@ -366,8 +382,10 @@ We'll start by creating a fake matrix of counts. counts_mtx <- matrix( 1:12, ncol = 4, - dimnames = list(c("geneA", "geneB", "geneC"), - c("A-cell1", "A-cell2", "B-cell1", "B-cell2")) + dimnames = list( + c("geneA", "geneB", "geneC"), + c("A-cell1", "A-cell2", "B-cell1", "B-cell2") + ) ) counts_mtx ``` @@ -401,8 +419,10 @@ pb_groups <- colData(rms_sce)[, c("celltype_broad", "sample")] # create a new SCE object that contains # the pseudo-bulked counts across the provided groups -pb_sce <- scuttle::aggregateAcrossCells(rms_sce, - id = pb_groups) +pb_sce <- scuttle::aggregateAcrossCells( + rms_sce, + id = pb_groups +) # column names aren't automatically added to the pseudo-bulked sce, # so let's add them in @@ -519,8 +539,10 @@ As a reminder, this is NOT required for running `DESeq2` analysis; we are just u deseq_object <- DESeq2::estimateSizeFactors(deseq_object) # normalize and log transform to use for visualization -normalized_object <- DESeq2::rlog(deseq_object, - blind = TRUE) +normalized_object <- DESeq2::rlog( + deseq_object, + blind = TRUE +) normalized_object ``` @@ -632,18 +654,19 @@ This package automatically colors the points by cutoffs for both significance an Even better, it outputs a `ggplot2` object, so if we want to customize the plot further, we can use the same `ggplot2` commands we have used before. ```{r volcano} -EnhancedVolcano::EnhancedVolcano(deseq_results, - x = 'log2FoldChange', # fold change statistic to plot - y = 'pvalue', # significance values - lab = deseq_results$gene_symbol, # labels for points - pCutoff = 1e-05, # p value cutoff (default) - FCcutoff = 1, # fold change cutoff (default) - title = NULL, # no title - subtitle = NULL, # or subtitle - caption = NULL, # or caption - drawConnectors = TRUE, # add some fun arrows - labSize = 3 # smaller labels - ) + +EnhancedVolcano::EnhancedVolcano( + deseq_results, + x = "log2FoldChange", # fold change statistic to plot + y = "pvalue", # significance values + lab = deseq_results$gene_symbol, # labels for points + pCutoff = 1e-05, # p value cutoff (default) + FCcutoff = 1, # fold change cutoff (default) + title = NULL, # no title + subtitle = NULL, # or subtitle + caption = NULL, # or caption + drawConnectors = TRUE, # add some fun arrows + labSize = 3 # smaller labels +) + # change the overall theme theme_bw() + # move the legend to the bottom @@ -669,10 +692,12 @@ However, we might be interested to see the expression of genes that are differen ```{r celltype comparison} # let's compare gene expression across some other cell types # look at all tumor cells and pick one normal cell type -celltypes <- c("Tumor_Myoblast", - "Tumor_Mesoderm", - "Tumor_Myocyte", - "Vascular Endothelium") +celltypes <- c( + "Tumor_Myoblast", + "Tumor_Mesoderm", + "Tumor_Myocyte", + "Vascular Endothelium" +) # subset to just celltypes that we are interested in tumor_sce <- rms_sce[, which(rms_sce$celltype_broad %in% celltypes)] @@ -687,28 +712,37 @@ We can then directly reference that `Feature` column when plotting, instead of u ```{r multi-gene plot} # pick a couple genes to look at -genes_to_plot <- c("ENSG00000196090", #PTPRT - "ENSG00000148935") #GAS2 +genes_to_plot <- c( + "ENSG00000196090", # PTPRT + "ENSG00000148935" +) # GAS2 # create a violin plot -scater::plotExpression(tumor_sce, - # a vector of genes to plot - features = genes_to_plot, - x = "diagnosis_group", - color_by = "diagnosis_group", - other_fields = "celltype_broad", - point_size = 0.1) + +scater::plotExpression( + tumor_sce, + # a vector of genes to plot + features = genes_to_plot, + x = "diagnosis_group", + color_by = "diagnosis_group", + other_fields = "celltype_broad", + point_size = 0.1 +) + # each celltype is its own column - facet_grid(cols = vars(celltype_broad), - # each feature (gene) is its own row - rows = vars(Feature)) + + facet_grid( + cols = vars(celltype_broad), + # each feature (gene) is its own row + rows = vars(Feature) + ) + # change the font size of the facet labels theme(strip.text = element_text(size = 7)) + - guides(color = guide_legend( - title = "Subtype", # update the legend title - # change the size of the legend colors - override.aes = list(size = 3, alpha = 1)) + guides( + color = guide_legend( + # update the legend title + title = "Subtype", + # change the size of the legend colors + override.aes = list(size = 3, alpha = 1) ) + ) ``` How do the expression of these genes change across cell types and RMS subtypes? diff --git a/scRNA-seq-advanced/03-differential_expression.Rmd b/scRNA-seq-advanced/03-differential_expression.Rmd index 43d0cb9e..55f1ea68 100644 --- a/scRNA-seq-advanced/03-differential_expression.Rmd +++ b/scRNA-seq-advanced/03-differential_expression.Rmd @@ -68,27 +68,35 @@ To begin, let's set up our directories and files: data_dir <- file.path("data", "rms") # integrated file containing samples to use for DE analysis -integrated_sce_file <- file.path(data_dir, - "integrated", - "rms_all_sce.rds") +integrated_sce_file <- file.path( + data_dir, + "integrated", + "rms_all_sce.rds" +) # sample metadata to set up DE analysis -sample_metadata_file <- file.path(data_dir, - "annotations", - "rms_sample_metadata.tsv") +sample_metadata_file <- file.path( + data_dir, + "annotations", + "rms_sample_metadata.tsv" +) # directory to store output deseq_dir <- file.path("analysis", "rms", "deseq") fs::dir_create(deseq_dir) # results file to output from DE analysis -deseq_output_file <- file.path(deseq_dir, - "rms_myoblast_deseq_results.tsv") +deseq_output_file <- file.path( + deseq_dir, + "rms_myoblast_deseq_results.tsv" +) # output integrated sce object -output_sce_file <- file.path(data_dir, - "integrated", - "rms_subset_sce.rds") +output_sce_file <- file.path( + data_dir, + "integrated", + "rms_subset_sce.rds" +) ``` We can go ahead and read in the SCE object and the metadata file. @@ -174,17 +182,21 @@ coldata_df <- colData(integrated_sce) |> dplyr::left_join(sample_metadata, by = c("sample" = "library_id")) |> # create new columns # cell_id is a combination of barcode and sample - dplyr::mutate(cell_id = glue::glue("{sample}-{barcode}"), - # simplify subdiagnosis - diagnosis_group = forcats::fct_recode( - subdiagnosis, - "ARMS" = "Alveolar rhabdomyosarcoma", - "ERMS" = "Embryonal rhabdomyosarcoma" - )) + dplyr::mutate( + cell_id = glue::glue("{sample}-{barcode}"), + # simplify subdiagnosis + diagnosis_group = forcats::fct_recode( + subdiagnosis, + "ARMS" = "Alveolar rhabdomyosarcoma", + "ERMS" = "Embryonal rhabdomyosarcoma" + ) + ) # add modified data frame back to SCE as DataFrame -colData(integrated_sce) <- DataFrame(coldata_df, - row.names = coldata_df$cell_id) +colData(integrated_sce) <- DataFrame( + coldata_df, + row.names = coldata_df$cell_id +) ``` Now when we look at the `colData` of the SCE object we should see new columns, including the `diagnosis_group` column which indicates if each cell comes from an ERMS or ARMS sample. @@ -204,11 +216,13 @@ In the chunk below we will start by taking a look at our integration results and ```{r diagnosis group UMAP, live=TRUE} # UMAP of all samples, separating by diagnosis group -scater::plotReducedDim(integrated_sce, - dimred = "fastmnn_UMAP", - color_by = "diagnosis_group", - point_size= 0.5, - point_alpha = 0.2) +scater::plotReducedDim( + integrated_sce, + dimred = "fastmnn_UMAP", + color_by = "diagnosis_group", + point_size = 0.5, + point_alpha = 0.2 +) ``` Interestingly, it looks like samples from the ARMS and ERMS subtypes tend to group with samples of the same subtype rather than all together. @@ -225,12 +239,14 @@ The samples which could be further classified have a mix of `Tumor_Mesoderm`, `T ```{r celltype UMAP} # UMAP of all samples labeled by cell type -scater::plotReducedDim(integrated_sce, - dimred = "fastmnn_UMAP", - # color each point by cell type - color_by = "celltype_broad", - point_size = 0.5, - point_alpha = 0.4) + +scater::plotReducedDim( + integrated_sce, + dimred = "fastmnn_UMAP", + # color each point by cell type + color_by = "celltype_broad", + point_size = 0.5, + point_alpha = 0.4 +) + # Modify the legend key with larger, easier to see points guides(color = guide_legend(override.aes = list(size = 3, alpha = 1))) ``` @@ -247,14 +263,16 @@ In the below plot we will color our cells by cell type while also using `facet_g ```{r celltype subdiagnosis UMAP, live=TRUE} # UMAP of all samples # separating by diagnosis group and labeling cell type -scater::plotReducedDim(integrated_sce, - dimred = "fastmnn_UMAP", - # color each point by cell type - color_by = "celltype_broad", - point_size= 0.5, - point_alpha = 0.4, - # tell scater to use diagnosis_group for plotting - other_fields = "diagnosis_group") + +scater::plotReducedDim( + integrated_sce, + dimred = "fastmnn_UMAP", + # color each point by cell type + color_by = "celltype_broad", + point_size = 0.5, + point_alpha = 0.4, + # tell scater to use diagnosis_group for plotting + other_fields = "diagnosis_group" +) + # include each diagnosis group as its own column facet_grid(cols = vars(diagnosis_group)) ``` @@ -271,20 +289,22 @@ tumor_cells_df <- coldata_df |> # create a stacked barplot ggplot(tumor_cells_df, aes(x = sample, fill = celltype_broad)) + - geom_bar(position = "fill", color = "black", size = 0.2) + - labs( - x = "Sample", - y = "Proportion of cells", - fill = "Cell type" - ) + + geom_bar(position = "fill", color = "black", size = 0.2) + + labs( + x = "Sample", + y = "Proportion of cells", + fill = "Cell type" + ) + scale_fill_brewer(palette = "Dark2") + theme_bw() + - theme(axis.text.x = element_text(angle = 90, vjust = 0.5))+ + theme(axis.text.x = element_text(angle = 90, vjust = 0.5)) + # facet by diagnosis group - facet_grid(cols = vars(diagnosis_group), - # only show non-NA values on x-axis - scales = "free_x", - space = "free_x") + facet_grid( + cols = vars(diagnosis_group), + # only show non-NA values on x-axis + scales = "free_x", + space = "free_x" + ) ``` Similar to the UMAP, this plot shows that ARMS and ERMS share a lot of the same cell types. @@ -376,8 +396,10 @@ We'll start by creating a fake matrix of counts. counts_mtx <- matrix( 1:12, ncol = 4, - dimnames = list(c("geneA", "geneB", "geneC"), - c("A-cell1", "A-cell2", "B-cell1", "B-cell2")) + dimnames = list( + c("geneA", "geneB", "geneC"), + c("A-cell1", "A-cell2", "B-cell1", "B-cell2") + ) ) counts_mtx ``` @@ -390,8 +412,7 @@ To do this we will use the `DelayedArray::colsum()` function, which allows us to groups <- c("A", "A", "B", "B") # sum counts across cells (columns) by group label -pb_counts <- DelayedArray::colsum(counts_mtx, - groups) +pb_counts <- DelayedArray::colsum(counts_mtx, groups) pb_counts ``` @@ -414,8 +435,10 @@ pb_groups <- colData(rms_sce)[, c("celltype_broad", "sample")] # create a new SCE object that contains # the pseudo-bulked counts across the provided groups -pb_sce <- scuttle::aggregateAcrossCells(rms_sce, - id = pb_groups) +pb_sce <- scuttle::aggregateAcrossCells( + rms_sce, + id = pb_groups +) # column names aren't automatically added to the pseudo-bulked sce, # so let's add them in @@ -514,8 +537,10 @@ The subtype information is stored in the `diagnosis_group` column of the `colDat ```{r deseq object, live=TRUE} # set up the deseq object, group by diagnosis -deseq_object <- DESeq2::DESeqDataSet(tumor_myoblast_sce, - design = ~ diagnosis_group) +deseq_object <- DESeq2::DESeqDataSet( + tumor_myoblast_sce, + design = ~diagnosis_group +) ``` The pseudo-bulked SCE object contains only one assay: the `counts` assay. @@ -536,8 +561,10 @@ As a reminder, this is NOT required for running `DESeq2` analysis; we are just u deseq_object <- DESeq2::estimateSizeFactors(deseq_object) # normalize and log transform to use for visualization -normalized_object <- DESeq2::rlog(deseq_object, - blind = TRUE) +normalized_object <- DESeq2::rlog( + deseq_object, + blind = TRUE +) normalized_object ``` @@ -651,18 +678,19 @@ This package automatically colors the points by cutoffs for both significance an Even better, it outputs a `ggplot2` object, so if we want to customize the plot further, we can use the same `ggplot2` commands we have used before. ```{r volcano} -EnhancedVolcano::EnhancedVolcano(deseq_results, - x = 'log2FoldChange', # fold change statistic to plot - y = 'pvalue', # significance values - lab = deseq_results$gene_symbol, # labels for points - pCutoff = 1e-05, # p value cutoff (default) - FCcutoff = 1, # fold change cutoff (default) - title = NULL, # no title - subtitle = NULL, # or subtitle - caption = NULL, # or caption - drawConnectors = TRUE, # add some fun arrows - labSize = 3 # smaller labels - ) + +EnhancedVolcano::EnhancedVolcano( + deseq_results, + x = "log2FoldChange", # fold change statistic to plot + y = "pvalue", # significance values + lab = deseq_results$gene_symbol, # labels for points + pCutoff = 1e-05, # p value cutoff (default) + FCcutoff = 1, # fold change cutoff (default) + title = NULL, # no title + subtitle = NULL, # or subtitle + caption = NULL, # or caption + drawConnectors = TRUE, # add some fun arrows + labSize = 3 # smaller labels +) + # change the overall theme theme_bw() + # move the legend to the bottom @@ -680,12 +708,14 @@ This can help us validate the `DESeq2` results so that we can visualize gene exp myoblast_combined_sce <- rms_sce[, which(rms_sce$celltype_broad == "Tumor_Myoblast")] # plot PTPRT (ENSG00000196090) expression in ARMS vs. ERMS -scater::plotReducedDim(myoblast_combined_sce, - dimred = "fastmnn_UMAP", - color_by = "ENSG00000196090", #PTPRT - point_size= 0.5, - point_alpha = 0.4, - other_fields = "diagnosis_group") + +scater::plotReducedDim( + myoblast_combined_sce, + dimred = "fastmnn_UMAP", + color_by = "ENSG00000196090", # PTPRT + point_size = 0.5, + point_alpha = 0.4, + other_fields = "diagnosis_group" +) + facet_grid(cols = vars(diagnosis_group)) ``` @@ -695,10 +725,12 @@ However, we might be interested to see the expression of genes that are differen ```{r celltype comparison} # let's compare gene expression across some other cell types # look at all tumor cells and pick one normal cell type -celltypes <- c("Tumor_Myoblast", - "Tumor_Mesoderm", - "Tumor_Myocyte", - "Vascular Endothelium") +celltypes <- c( + "Tumor_Myoblast", + "Tumor_Mesoderm", + "Tumor_Myocyte", + "Vascular Endothelium" +) # subset to just celltypes that we are interested in tumor_sce <- rms_sce[, which(rms_sce$celltype_broad %in% celltypes)] @@ -713,28 +745,37 @@ We can then directly reference that `Feature` column when plotting, instead of u ```{r multi-gene plot} # pick a couple genes to look at -genes_to_plot <- c("ENSG00000196090", #PTPRT - "ENSG00000148935") #GAS2 +genes_to_plot <- c( + "ENSG00000196090", # PTPRT + "ENSG00000148935" +) # GAS2 # create a violin plot -scater::plotExpression(tumor_sce, - # a vector of genes to plot - features = genes_to_plot, - x = "diagnosis_group", - color_by = "diagnosis_group", - other_fields = "celltype_broad", - point_size = 0.1) + +scater::plotExpression( + tumor_sce, + # a vector of genes to plot + features = genes_to_plot, + x = "diagnosis_group", + color_by = "diagnosis_group", + other_fields = "celltype_broad", + point_size = 0.1 +) + # each celltype is its own column - facet_grid(cols = vars(celltype_broad), - # each feature (gene) is its own row - rows = vars(Feature)) + + facet_grid( + cols = vars(celltype_broad), + # each feature (gene) is its own row + rows = vars(Feature) + ) + # change the font size of the facet labels theme(strip.text = element_text(size = 7)) + - guides(color = guide_legend( - title = "Subtype", # update the legend title - # change the size of the legend colors - override.aes = list(size = 3, alpha = 1)) + guides( + color = guide_legend( + # update the legend title + title = "Subtype", + # change the size of the legend colors + override.aes = list(size = 3, alpha = 1) ) + ) ``` How do the expression of these genes change across cell types and RMS subtypes? diff --git a/scRNA-seq-advanced/03-differential_expression.nb.html b/scRNA-seq-advanced/03-differential_expression.nb.html index 4a6d15be..33bb78ec 100644 --- a/scRNA-seq-advanced/03-differential_expression.nb.html +++ b/scRNA-seq-advanced/03-differential_expression.nb.html @@ -2979,33 +2979,41 @@

    Directories and files

    To begin, let’s set up our directories and files:

    - +
    # set up file paths
     # data directory for RMS data
     data_dir <- file.path("data", "rms")
     
     # integrated file containing samples to use for DE analysis
    -integrated_sce_file <- file.path(data_dir,
    -                                 "integrated",
    -                                 "rms_all_sce.rds")
    +integrated_sce_file <- file.path(
    +  data_dir,
    +  "integrated",
    +  "rms_all_sce.rds"
    +)
     
     # sample metadata to set up DE analysis
    -sample_metadata_file <- file.path(data_dir,
    -                                  "annotations",
    -                                  "rms_sample_metadata.tsv")
    +sample_metadata_file <- file.path(
    +  data_dir,
    +  "annotations",
    +  "rms_sample_metadata.tsv"
    +)
     
     # directory to store output
     deseq_dir <- file.path("analysis", "rms", "deseq")
     fs::dir_create(deseq_dir)
     
     # results file to output from DE analysis
    -deseq_output_file <- file.path(deseq_dir,
    -                               "rms_myoblast_deseq_results.tsv")
    +deseq_output_file <- file.path(
    +  deseq_dir,
    +  "rms_myoblast_deseq_results.tsv"
    +)
     
     # output integrated sce object
    -output_sce_file <- file.path(data_dir,
    -                             "integrated",
    -                             "rms_subset_sce.rds")
    +output_sce_file <- file.path( + data_dir, + "integrated", + "rms_subset_sce.rds" +)
    @@ -3178,7 +3186,7 @@

    Cell type annotations

    plotting.

    - +
    # add sample metadata to colData from the integrated SCE object
     coldata_df <- colData(integrated_sce) |>
       # convert from DataFrame to data.frame
    @@ -3187,17 +3195,21 @@ 

    Cell type annotations

    dplyr::left_join(sample_metadata, by = c("sample" = "library_id")) |> # create new columns # cell_id is a combination of barcode and sample - dplyr::mutate(cell_id = glue::glue("{sample}-{barcode}"), - # simplify subdiagnosis - diagnosis_group = forcats::fct_recode( - subdiagnosis, - "ARMS" = "Alveolar rhabdomyosarcoma", - "ERMS" = "Embryonal rhabdomyosarcoma" - )) + dplyr::mutate( + cell_id = glue::glue("{sample}-{barcode}"), + # simplify subdiagnosis + diagnosis_group = forcats::fct_recode( + subdiagnosis, + "ARMS" = "Alveolar rhabdomyosarcoma", + "ERMS" = "Embryonal rhabdomyosarcoma" + ) + ) # add modified data frame back to SCE as DataFrame -colData(integrated_sce) <- DataFrame(coldata_df, - row.names = coldata_df$cell_id)
    +colData(integrated_sce) <- DataFrame( + coldata_df, + row.names = coldata_df$cell_id +) @@ -3230,13 +3242,15 @@

    Plotting with annotations

    multiple libraries or samples.

    - +
    # UMAP of all samples, separating by diagnosis group
    -scater::plotReducedDim(integrated_sce,
    -                       dimred = "fastmnn_UMAP",
    -                       color_by = "diagnosis_group",
    -                       point_size= 0.5,
    -                       point_alpha = 0.2)
    +scater::plotReducedDim( + integrated_sce, + dimred = "fastmnn_UMAP", + color_by = "diagnosis_group", + point_size = 0.5, + point_alpha = 0.2 +)

    @@ -3262,14 +3276,16 @@

    Plotting with annotations

    Tumor_Myocyte.

    - +
    # UMAP of all samples labeled by cell type
    -scater::plotReducedDim(integrated_sce,
    -                       dimred = "fastmnn_UMAP",
    -                       # color each point by cell type
    -                       color_by = "celltype_broad",
    -                       point_size = 0.5,
    -                       point_alpha = 0.4) +
    +scater::plotReducedDim(
    +  integrated_sce,
    +  dimred = "fastmnn_UMAP",
    +  # color each point by cell type
    +  color_by = "celltype_broad",
    +  point_size = 0.5,
    +  point_alpha = 0.4
    +) +
       # Modify the legend key with larger, easier to see points
       guides(color = guide_legend(override.aes = list(size = 3, alpha = 1)))
    @@ -3298,17 +3314,19 @@

    Plotting with annotations

    be in their own plot panel.

    - +
    # UMAP of all samples
     # separating by diagnosis group and labeling cell type
    -scater::plotReducedDim(integrated_sce,
    -                       dimred = "fastmnn_UMAP",
    -                       # color each point by cell type
    -                       color_by = "celltype_broad",
    -                       point_size= 0.5,
    -                       point_alpha = 0.4,
    -                       # tell scater to use diagnosis_group for plotting
    -                       other_fields = "diagnosis_group") +
    +scater::plotReducedDim(
    +  integrated_sce,
    +  dimred = "fastmnn_UMAP",
    +  # color each point by cell type
    +  color_by = "celltype_broad",
    +  point_size = 0.5,
    +  point_alpha = 0.4,
    +  # tell scater to use diagnosis_group for plotting
    +  other_fields = "diagnosis_group"
    +) +
       # include each diagnosis group as its own column
       facet_grid(cols = vars(diagnosis_group))
    @@ -3324,7 +3342,7 @@

    Plotting with annotations

    first.

    - +
    # filter coldata to only include tumor cells
     tumor_cells_df <- coldata_df |>
       # find rows where the cell type name contains the string "Tumor"
    @@ -3332,20 +3350,22 @@ 

    Plotting with annotations

    # create a stacked barplot ggplot(tumor_cells_df, aes(x = sample, fill = celltype_broad)) + - geom_bar(position = "fill", color = "black", size = 0.2) + - labs( - x = "Sample", - y = "Proportion of cells", - fill = "Cell type" - ) + + geom_bar(position = "fill", color = "black", size = 0.2) + + labs( + x = "Sample", + y = "Proportion of cells", + fill = "Cell type" + ) + scale_fill_brewer(palette = "Dark2") + theme_bw() + - theme(axis.text.x = element_text(angle = 90, vjust = 0.5))+ + theme(axis.text.x = element_text(angle = 90, vjust = 0.5)) + # facet by diagnosis group - facet_grid(cols = vars(diagnosis_group), - # only show non-NA values on x-axis - scales = "free_x", - space = "free_x")
    + facet_grid( + cols = vars(diagnosis_group), + # only show non-NA values on x-axis + scales = "free_x", + space = "free_x" + )
    Warning: Using `size` aesthetic for lines was deprecated in ggplot2 3.4.0.
    @@ -3512,13 +3532,15 @@ 

    Pseudo-bulking

    matrix of counts.

    - +
    # create an example counts matrix
     counts_mtx <- matrix(
       1:12,
       ncol = 4,
    -  dimnames = list(c("geneA", "geneB", "geneC"),
    -                  c("A-cell1", "A-cell2", "B-cell1", "B-cell2"))
    +  dimnames = list(
    +    c("geneA", "geneB", "geneC"),
    +    c("A-cell1", "A-cell2", "B-cell1", "B-cell2")
    +  )
     )
     counts_mtx
    @@ -3537,13 +3559,12 @@

    Pseudo-bulking

    columns.

    - +
    # define the group that each column belongs to
     groups <- c("A", "A", "B", "B")
     
     # sum counts across cells (columns) by group label
    -pb_counts <- DelayedArray::colsum(counts_mtx,
    -                                  groups)
    +pb_counts <- DelayedArray::colsum(counts_mtx, groups)
     pb_counts
    @@ -3572,15 +3593,17 @@

    Pseudo-bulking

    type and original sample.

    - +
    # first subset the coldata
     # to only have the columns we care about for pseudo-bulking
     pb_groups <- colData(rms_sce)[, c("celltype_broad", "sample")]
     
     # create a new SCE object that contains
     # the pseudo-bulked counts across the provided groups
    -pb_sce <- scuttle::aggregateAcrossCells(rms_sce,
    -                                        id = pb_groups)
    +pb_sce <- scuttle::aggregateAcrossCells(
    +  rms_sce,
    +  id = pb_groups
    +)
     
     # column names aren't automatically added to the pseudo-bulked sce,
     # so let's add them in
    @@ -3752,10 +3775,12 @@ 

    Create the DESeqDataSet object

    colData in the pseudo-bulked SCE.

    - +
    # set up the deseq object, group by diagnosis
    -deseq_object <- DESeq2::DESeqDataSet(tumor_myoblast_sce,
    -                                     design = ~ diagnosis_group)
    +deseq_object <- DESeq2::DESeqDataSet( + tumor_myoblast_sce, + design = ~diagnosis_group +)
    converting counts to integer mode
    @@ -3782,13 +3807,15 @@

    Create the DESeqDataSet object

    we are just using it to visualize our data prior to DE analysis.

    - +
    # estimate size factors first
     deseq_object <- DESeq2::estimateSizeFactors(deseq_object)
     
     # normalize and log transform to use for visualization
    -normalized_object <- DESeq2::rlog(deseq_object,
    -                                  blind = TRUE)
    +normalized_object <- DESeq2::rlog(
    +  deseq_object,
    +  blind = TRUE
    +)
     normalized_object
    @@ -4028,19 +4055,20 @@

    Exploring the identified differentially expressed genes

    ggplot2 commands we have used before.

    - -
    EnhancedVolcano::EnhancedVolcano(deseq_results,
    -                x = 'log2FoldChange', # fold change statistic to plot
    -                y = 'pvalue', # significance values
    -                lab = deseq_results$gene_symbol, # labels for points
    -                pCutoff = 1e-05, # p value cutoff (default)
    -                FCcutoff = 1, # fold change cutoff (default)
    -                title = NULL, # no title
    -                subtitle = NULL, # or subtitle
    -                caption = NULL, # or caption
    -                drawConnectors = TRUE, # add some fun arrows
    -                labSize = 3  # smaller labels
    -                ) +
    +
    +
    EnhancedVolcano::EnhancedVolcano(
    +  deseq_results,
    +  x = "log2FoldChange", # fold change statistic to plot
    +  y = "pvalue", # significance values
    +  lab = deseq_results$gene_symbol, # labels for points
    +  pCutoff = 1e-05, # p value cutoff (default)
    +  FCcutoff = 1, # fold change cutoff (default)
    +  title = NULL, # no title
    +  subtitle = NULL, # or subtitle
    +  caption = NULL, # or caption
    +  drawConnectors = TRUE, # add some fun arrows
    +  labSize = 3 # smaller labels
    +) +
       # change the overall theme
       theme_bw() +
       # move the legend to the bottom
    @@ -4067,17 +4095,19 @@ 

    Exploring the identified differentially expressed genes

    interest on a single-cell level.

    - +
    # filter to just myoblast cells and remove any NA's before plotting
     myoblast_combined_sce <- rms_sce[, which(rms_sce$celltype_broad == "Tumor_Myoblast")]
     
     # plot PTPRT (ENSG00000196090) expression in ARMS vs. ERMS
    -scater::plotReducedDim(myoblast_combined_sce,
    -                       dimred = "fastmnn_UMAP",
    -                       color_by = "ENSG00000196090", #PTPRT
    -                       point_size= 0.5,
    -                       point_alpha = 0.4,
    -                       other_fields = "diagnosis_group") +
    +scater::plotReducedDim(
    +  myoblast_combined_sce,
    +  dimred = "fastmnn_UMAP",
    +  color_by = "ENSG00000196090", # PTPRT
    +  point_size = 0.5,
    +  point_alpha = 0.4,
    +  other_fields = "diagnosis_group"
    +) +
       facet_grid(cols = vars(diagnosis_group))
    @@ -4091,13 +4121,15 @@

    Exploring the identified differentially expressed genes

    types present in our samples.

    - +
    # let's compare gene expression across some other cell types
     # look at all tumor cells and pick one normal cell type
    -celltypes <- c("Tumor_Myoblast",
    -               "Tumor_Mesoderm",
    -               "Tumor_Myocyte",
    -               "Vascular Endothelium")
    +celltypes <- c(
    +  "Tumor_Myoblast",
    +  "Tumor_Mesoderm",
    +  "Tumor_Myocyte",
    +  "Vascular Endothelium"
    +)
     
     # subset to just celltypes that we are interested in
     tumor_sce <- rms_sce[, which(rms_sce$celltype_broad %in% celltypes)]
    @@ -4121,30 +4153,39 @@

    Exploring the identified differentially expressed genes

    previously.

    - +
    # pick a couple genes to look at
    -genes_to_plot <- c("ENSG00000196090", #PTPRT
    -                   "ENSG00000148935") #GAS2
    +genes_to_plot <- c(
    +  "ENSG00000196090", # PTPRT
    +  "ENSG00000148935"
    +) # GAS2
     
     # create a violin plot
    -scater::plotExpression(tumor_sce,
    -                       # a vector of genes to plot
    -                       features = genes_to_plot,
    -                       x = "diagnosis_group",
    -                       color_by = "diagnosis_group",
    -                       other_fields = "celltype_broad",
    -                       point_size = 0.1) +
    +scater::plotExpression(
    +  tumor_sce,
    +  # a vector of genes to plot
    +  features = genes_to_plot,
    +  x = "diagnosis_group",
    +  color_by = "diagnosis_group",
    +  other_fields = "celltype_broad",
    +  point_size = 0.1
    +) +
       # each celltype is its own column
    -  facet_grid(cols = vars(celltype_broad),
    -             # each feature (gene) is its own row
    -             rows = vars(Feature)) +
    +  facet_grid(
    +    cols = vars(celltype_broad),
    +    # each feature (gene) is its own row
    +    rows = vars(Feature)
    +  ) +
       # change the font size of the facet labels
       theme(strip.text = element_text(size = 7)) +
    -  guides(color = guide_legend(
    -    title = "Subtype", # update the legend title
    -    # change the size of the legend colors
    -    override.aes = list(size = 3, alpha = 1))
    -    )
    + guides( + color = guide_legend( + # update the legend title + title = "Subtype", + # change the size of the legend colors + override.aes = list(size = 3, alpha = 1) + ) + )

    @@ -4263,7 +4304,7 @@

    Print session info

    -
    ---
title: "Differential expression analysis for scRNA-seq data"
author: "Data Lab for ALSF"
date: 2023
output:
  html_notebook:
    toc: yes
    toc_float: yes
---

## Objectives

This notebook will demonstrate how to:

- Use pseudo-bulking to prepare scRNA-seq libraries for differential expression
- Perform differential expression with the `DESeq2` package
- Use `ggplot2` and `EnhancedVolcano` to visualize gene expression changes across cell types and samples

---

Just like bulk RNA-seq, it is likely that one of the goals when performing scRNA-seq will be to compare the gene expression of multiple samples to each other.
Unlike bulk RNA-seq analysis, scRNA-seq analysis allows us to identify and annotate cell types or subpopulations of cells present in each of our samples.
This means that we can account for differences in cell type composition across samples and specifically focus on cell types or populations of interest when performing differential expression (DE) analysis.
In this notebook, we will work with multiple samples to identify differentially expressed genes across cell types of interest using the [`DESeq2`](https://bioconductor.org/packages/release/bioc/html/DESeq2.html) package.

![Single-cell roadmap: Differential expression](diagrams/roadmap_differential_expression.png)

We will continue working with samples from the [`SCPCP000005` project](https://scpca.alexslemonade.org/projects/SCPCP000005), an investigation of pediatric solid tumors led by the Dyer and Chen labs at St. Jude Children's Research Hospital.
This particular dataset contains 10 different samples that have been integrated using `fastMNN`, following the same procedure we outlined in `02-dataset_integration.Rmd`.
These 10 samples represent two different types of rhabdomyosarcoma (RMS): embryonal rhabdomyosarcoma (ERMS) and alveolar rhabdomyosarcoma (ARMS).
These two subtypes are distinguished by the presence of the `PAX3/PAX7-FOXO1` fusion gene, which is present only in ARMS patients.
Additionally, cells found in ARMS tumors tend to have an increased mutational burden with cells in a more differentiated state compared to ERMS tumor cells ([Shern _et al._ 2014](https://doi.org/10.1158/2159-8290.CD-13-0639); [Stewart _et al._ 2018](https://doi.org/10.1016/j.ccell.2018.07.012)).
RMS tumors, regardless of subtype, are made up of cells typically associated with development of skeletal muscle: mesoderm, myoblasts, and myocytes ([Sebire and Malone 2003](https://doi.org/10.1136/jcp.56.6.412)).
[Patel _et al._ (2022)](https://doi.org/10.1016/j.devcel.2022.04.003) tested the hypothesis that cell types have distinct gene expression patterns in ARMS vs. ERMS samples.
Here we will look at a subset of the samples they sequenced and identify differentially expressed genes in tumor cells between ARMS and ERMS samples.

## Set up

```{r setup, message=FALSE}
# set seed for reproducibility
set.seed(2022)

# load libraries
library(ggplot2) # plotting functions
library(SingleCellExperiment)

# package used for differential expression analysis
library(DESeq2)
```

### Directories and files

We will start by reading in a `SingleCellExperiment` (SCE) object that contains both the uncorrected (merged but not integrated) and corrected (integrated) gene expression data for all 10 samples.

Prior to integration, all 10 samples went through the same filtering, normalization, and dimensionality reduction.
These 10 samples were then merged into one `SingleCellExperiment` object following the same steps outlined in `03-dataset_integration.Rmd`.
The merged object was then integrated with `fastMNN` to obtain a corrected gene expression assay and corrected reduced dimensionality results.
The final SCE object was stored in `data/rms/integrated/rms_all_sce.rds`.

We also have provided a metadata file, `data/rms/annotations/rms_sample_metadata.tsv`, that contains information from each sample, such as diagnosis, sex, age, etc.
Each row in this file corresponds to a sample found in the integrated SCE object.

To begin, let's set up our directories and files:

```{r filepaths}
# set up file paths
# data directory for RMS data
data_dir <- file.path("data", "rms")

# integrated file containing samples to use for DE analysis
integrated_sce_file <- file.path(data_dir,
                                 "integrated",
                                 "rms_all_sce.rds")

# sample metadata to set up DE analysis
sample_metadata_file <- file.path(data_dir,
                                  "annotations",
                                  "rms_sample_metadata.tsv")

# directory to store output
deseq_dir <- file.path("analysis", "rms", "deseq")
fs::dir_create(deseq_dir)

# results file to output from DE analysis
deseq_output_file <- file.path(deseq_dir,
                               "rms_myoblast_deseq_results.tsv")

# output integrated sce object
output_sce_file <- file.path(data_dir,
                             "integrated",
                             "rms_subset_sce.rds")
```

We can go ahead and read in the SCE object and the metadata file.

```{r read files, live=TRUE}
# read in the SCE object that has already been integrated
integrated_sce <- readr::read_rds(integrated_sce_file)

# read in sample metadata file
sample_metadata <- readr::read_tsv(sample_metadata_file)
```

## Dataset exploration

Before we dive into differential expression, let's explore our integrated SCE object and the dataset a little more.

We'll start by looking at what's inside the object.
Here we should have both the original (uncorrected) data and the integrated (corrected) data for both the gene expression and the reduced dimensionality results.
How are those stored in our object?

```{r print sce, live=TRUE}
# print out entire object
integrated_sce
```


```{r print assay names, live=TRUE}
# look at the assay names in our object
assayNames(integrated_sce)
```

When we look at the assay names we should see that there are 3 matrices, `counts`, `logcounts`, and `fastmnn_corrected`.
The `counts` and `logcounts` assays correspond to the uncorrected gene expression data that has been merged but NOT integrated.
The `fastmnn_corrected` data contains the corrected gene expression data obtained from integration.
For this exercise we will not be using the `fastmnn_corrected` data (more on why not once we get to setting up the differential expression), but we need to be aware that it is present and be able to distinguish it from our uncorrected data.


```{r print reducedDim names, live=TRUE}
# look at the names of the dimension reductions
reducedDimNames(integrated_sce)
```

In the `reducedDim` slots you should see `PCA` and `UMAP`, which were both calculated from the combined data _before_ integration.
You should also see `fastmnn_PCA` and `fastmnn_UMAP` reduced dimensions, which correspond to the integrated results.

### Cell type annotations

Just like in the integration notebook, this dataset also contains the cell type annotations found in the `celltype_fine` and `celltype_broad` columns of the `colData`.
These cell types were originally assigned in [Patel _et al._ (2022)](https://doi.org/10.1016/j.devcel.2022.04.003).
We will use these cell type assignments to set up the DE analysis below, but they are not required for DE analysis itself.
It's important to note that DE analysis can be applied to any subpopulation of interest that is shared across samples besides just cell types.

Because we are going to be doing DE analysis between ARMS and ERMS samples, let's start by labeling cells in the integrated dataset based on their RMS subtype.
To do this we will need to be sure that the subtype is present in the `colData` of the integrated SCE object.
If it's not there, we need to add it in.

```{r coldata head, live=TRUE}
# look at the head of the coldata
head(colData(integrated_sce)) |>
  as.data.frame()
```

Uh oh, it looks like the RMS subtype is not found in the SCE object.
Fortunately we also have the sample metadata table that we read in earlier, which contains information about each of the samples present in the dataset.

```{r sample metadata, live=TRUE}
# print out sample metadata
head(sample_metadata)
```

Looking at this sample table, we see a column named `subdiagnosis` which accounts for the RMS subtype, ARMS or ERMS.
We also see other columns that contain information about each specific sample.

We can incorporate the information in this sample metadata table into the `colData` of the integrated SCE object.
This will allow us to match each of the samples in the SCE object with the RMS subtype and also allow us to use any of the columns in the sample metadata for plotting.

```{r modify coldata}
# add sample metadata to colData from the integrated SCE object
coldata_df <- colData(integrated_sce) |>
  # convert from DataFrame to data.frame
  as.data.frame() |>
  # merge with sample metadata
  dplyr::left_join(sample_metadata, by = c("sample" = "library_id")) |>
  # create new columns
  # cell_id is a combination of barcode and sample
  dplyr::mutate(cell_id = glue::glue("{sample}-{barcode}"),
                # simplify subdiagnosis
                diagnosis_group = forcats::fct_recode(
                  subdiagnosis,
                  "ARMS" = "Alveolar rhabdomyosarcoma",
                  "ERMS" = "Embryonal rhabdomyosarcoma"
                ))

# add modified data frame back to SCE as DataFrame
colData(integrated_sce) <- DataFrame(coldata_df,
                                     row.names = coldata_df$cell_id)
```

Now when we look at the `colData` of the SCE object we should see new columns, including the `diagnosis_group` column which indicates if each cell comes from an ERMS or ARMS sample.

```{r print new coldata, live=TRUE}
# take a look at the new modified colData
head(colData(integrated_sce)) |>
  as.data.frame()
```

### Plotting with annotations

We can now use that column to label any UMAP plots (or other plot types) that we make.
In the chunk below we will start by taking a look at our integration results and color our cells by RMS subtype.

**Reminder: You should always use the batch-corrected dimensionality reduction results for visualizing datasets containing multiple libraries or samples.**

```{r diagnosis group UMAP, live=TRUE}
# UMAP of all samples, separating by diagnosis group
scater::plotReducedDim(integrated_sce,
                       dimred = "fastmnn_UMAP",
                       color_by = "diagnosis_group",
                       point_size= 0.5,
                       point_alpha = 0.2)
```

Interestingly, it looks like samples from the ARMS and ERMS subtypes tend to group with samples of the same subtype rather than all together.

In the integration notebook we also looked at the distribution of cell types after integration.
In that notebook, we discussed that cells of the same cell type are expected to integrate with other cells of the same type.
Is that the case with this dataset?

A word of caution when evaluating the cell type results for this dataset: The cell types for this dataset were assigned in a two stage process as described in [Patel _et al._ (2022)](https://doi.org/10.1016/j.devcel.2022.04.003).
The first stage assigned cells as tumor or non-tumor.
The next stage further classified tumor cells into one of three types of tumor cells: myoblast, myocyte, or mesoderm.
Some samples could not be further classified, so all of their tumor cells are denoted `Tumor`.
The samples which could be further classified have a mix of `Tumor_Mesoderm`, `Tumor_Myoblast`, and `Tumor_Myocyte`.

```{r celltype UMAP}
# UMAP of all samples labeled by cell type
scater::plotReducedDim(integrated_sce,
                       dimred = "fastmnn_UMAP",
                       # color each point by cell type
                       color_by = "celltype_broad",
                       point_size = 0.5,
                       point_alpha = 0.4) +
  # Modify the legend key with larger, easier to see points
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1)))
```

Unlike with the previous datasets we have seen where all cells of the same cell type always grouped together, this dataset shows some slightly different patterns and not all cells of the same cell type cluster together.
One reason is that tumor data can be heterogeneous and every tumor is unique.
Depending on the tumor type we may not expect every sample to integrate perfectly and more heterogeneous tumor types will be more difficult to integrate together.
In this particular case we are looking at two subtypes of RMS that have distinct mutation burdens and differentiation states, so it's likely that those differences contribute to how well they integrate.

To explore whether cells are grouping together both by cell type and by RMS subtype, we can create a plot that incorporates both pieces of metadata.
We will take advantage of the `facet_grid()` function from `ggplot2` to look at two variables in the `colData` at once - the cell type and the subdiagnosis.
In the below plot we will color our cells by cell type while also using `facet_grid()` so that cells from different subdiagnoses will be in their own plot panel.

```{r celltype subdiagnosis UMAP, live=TRUE}
# UMAP of all samples
# separating by diagnosis group and labeling cell type
scater::plotReducedDim(integrated_sce,
                       dimred = "fastmnn_UMAP",
                       # color each point by cell type
                       color_by = "celltype_broad",
                       point_size= 0.5,
                       point_alpha = 0.4,
                       # tell scater to use diagnosis_group for plotting
                       other_fields = "diagnosis_group") +
  # include each diagnosis group as its own column
  facet_grid(cols = vars(diagnosis_group))
```

As expected, we see that cell types are separated, most likely due to different RMS subtypes.

We can also use a stacked barplot to look at the distribution of cell types across each sample, which will require a bit of wrangling first.

```{r celltype barplot}
# filter coldata to only include tumor cells
tumor_cells_df <- coldata_df |>
  # find rows where the cell type name contains the string "Tumor"
  dplyr::filter(stringr::str_detect(celltype_broad, "Tumor"))

# create a stacked barplot
ggplot(tumor_cells_df, aes(x = sample, fill = celltype_broad)) +
    geom_bar(position = "fill", color = "black", size = 0.2) +
    labs(
      x = "Sample",
      y = "Proportion of cells",
      fill = "Cell type"
    ) +
  scale_fill_brewer(palette = "Dark2") +
  theme_bw() +
  theme(axis.text.x = element_text(angle = 90, vjust = 0.5))+
  # facet by diagnosis group
  facet_grid(cols = vars(diagnosis_group),
             # only show non-NA values on x-axis
             scales = "free_x",
             space = "free_x")
```

Similar to the UMAP, this plot shows that ARMS and ERMS share a lot of the same cell types.

We also see that only 6 of these libraries have tumor cells that have been further classified into mesoderm, myoblast, and myocyte.
3 libraries contain cells that are only classified as tumor or non-tumor, and tumor cells are not further classified, and the remaining library is not even present in our plot because it was not assigned any cell types (all are `NA`).
We will continue our analysis only using the 6 libraries with fully classified cell types, removing the other 4 before we proceed with differential expression.

### Filtering samples

The reason we want to pare down our list of samples to consider is that we want to ensure that the cell types (or subpopulations) that we are interested in are present in all samples included in our DE analysis.
We want to remove any samples that do not contain our cell population(s) of interest as they have no counts to contribute to the DE analysis.

```{r subset sce}
# define samples to keep
library_ids <- c(
  "SCPCL000479",
  "SCPCL000480",
  "SCPCL000481",
  "SCPCL000484",
  "SCPCL000488",
  "SCPCL000491"
)

# subset sce to only contain samples of interest
samples_to_keep <- integrated_sce$sample %in% library_ids
rms_sce <- integrated_sce[, samples_to_keep]

# print out our new SCE
rms_sce
```

Before we move on, we'll remove the original integrated object from our environment to save some memory.

```{r remove sce}
rm(integrated_sce)
```

We will also save our new object in case we want to use it for other analysis later on.

```{r save sce}
# write RDS file with compression
readr::write_rds(rms_sce, file = output_sce_file, compress = "gz")
```

We now have an updated SCE object that contains 6 samples that were obtained from a mix of ARMS and ERMS patients.
We can then ask the question, do specific tumor cell types contain sets of differentially expressed genes between ARMS and ERMS samples?

We should make sure that we have enough biological replicates from each group to set up our experiment.
It is imperative to consider good experimental design and ensure that we have enough biological replicates (at least 3 for each group) when performing differential gene expression analysis.

If we look back at our stacked barplot we see that we picked 3 ARMS and 3 ERMS samples.
We can also see that the majority of cells are tumor cells, in particular the largest population of cells appears to be the `Tumor_Myoblast`.
For this example we will focus on identifying DE genes in these `Tumor_Myoblast` cells, but the principles applied below can be applied to any cell types or subpopulations of interest.

## Differential expression analysis

Now we are ready to start preparing for our DE analysis, where we will compare the gene expression of tumor myoblast cells between ARMS and ERMS samples.

Throughout the notebook we have been working with an integrated dataset that contains corrected gene expression data (`fastmnn_corrected` assay) and a corrected UMAP.
As a reminder, the uncorrected gene expression data, found in the `counts` and `logcounts` assays, correspond to data that has been merged (the first step we walked through prior to integration) into the same SCE but not yet integrated.
We do not want to use corrected gene expression values for differential expression; `DESeq2` expects the original raw counts as input so we will be using data found in the `counts` assay of the `SingleCellExperiment` object.

It is advised to only use the corrected values for any analyses being performed at the cell level, e.g., dimensionality reduction.
In contrast, it is not advised to use corrected values for any analyses that are gene-based, such as differential expression or marker gene detection, because within-batch and between-batch gene expression differences are no longer preserved.
The reason for this is two-fold – many of the DE models will expect uncorrected counts because they will account for between-sample variation within the model, and we want to ensure we are preserving variation that is present so as not to artificially inflate differences between populations.
See the [OSCA chapter on Using the corrected values](https://bioconductor.org/books/3.19/OSCA.multisample/using-corrected-values.html#using-corrected-values) for more insight.

### Pseudo-bulking

Before we can compare the gene expression profiles of myoblasts in ARMS vs. ERMS samples, we will need to "pseudo-bulk" the gene counts.
Pseudo-bulking creates a new counts matrix that contains the sum of the counts from all cells with a given label (e.g., cell type) for each sample ([Tung _et al._ 2017](https://doi.org/10.1038/srep39921)).
If we were to keep each cell's counts separate, they would be treated as replicates, leading to inflated statistics.
By pseudo-bulking first, we will now have one count for each gene for each sample and we can take advantage of well-established methods for differential expression with bulk RNA-seq.

Pseudo-bulking is implemented prior to differential expression analysis on single-cell data because it:

- Produces larger and less sparse counts, which allows us to use standard normalization and differential expression methods used by bulk RNA-seq.
- Collapses gene expression counts by sample, so that samples, rather than cells, represent replicates.
- Masks variance within a sample to emphasize variance across samples.
This can be both good and bad!
Masking intra-sample variation means you might not identify genes where average expression doesn't change between samples but the degree of cell-to-cell variation does.

Before we apply pseudo-bulking to our dataset, let's look at a simple example of how pseudo-bulking works.
We'll start by creating a fake matrix of counts.

```{r create matrix}
# create an example counts matrix
counts_mtx <- matrix(
  1:12,
  ncol = 4,
  dimnames = list(c("geneA", "geneB", "geneC"),
                  c("A-cell1", "A-cell2", "B-cell1", "B-cell2"))
)
counts_mtx
```

Next we will create a pseudo-bulked version of this matrix with only 2 columns: 1 for group `A` and 1 for group `B`.
To do this we will use the `DelayedArray::colsum()` function, which allows us to sum the counts for each row across groups of columns.

```{r pseudobulk matrix, live=TRUE}
# define the group that each column belongs to
groups <- c("A", "A", "B", "B")

# sum counts across cells (columns) by group label
pb_counts <- DelayedArray::colsum(counts_mtx,
                                  groups)
pb_counts
```

Looking at this output, you should see that the original 4 columns have been condensed to only 2 columns: 1 column to represent all cells from group `A`, and 1 column to represent all cells from group `B`.

Now the actual pseudo-bulking for our dataset!

We will use the [`scuttle::aggregateAcrossCells()` function](https://rdrr.io/github/LTLA/scuttle/man/aggregateAcrossCells.html) to pseudo-bulk our dataset.
This function takes as input an SCE object and the grouping assignments for each cell.
The output will be an SCE object that contains only the pseudo-bulked counts for all genes across all specified groups, rather than across all cells.
We can then subset this SCE to just include our cell type of interest (tumor myoblasts) for input to the DE analysis.

We can pseudo-bulk using any grouping that we are interested in.
For right now, we are interested in looking at gene expression across cell types, so we want to group the pseudo-bulked counts matrix by both cell type and original sample.

```{r pseudobulk sce}
# first subset the coldata
# to only have the columns we care about for pseudo-bulking
pb_groups <- colData(rms_sce)[, c("celltype_broad", "sample")]

# create a new SCE object that contains
# the pseudo-bulked counts across the provided groups
pb_sce <- scuttle::aggregateAcrossCells(rms_sce,
                                        id = pb_groups)

# column names aren't automatically added to the pseudo-bulked sce,
# so let's add them in
colnames(pb_sce) <- glue::glue(
  "{pb_sce$celltype_broad}_{pb_sce$sample}"
)

pb_sce
```

How does the new pseudo-bulked `SingleCellExperiment` look different?
How many columns does it have?

Let's take a look at what the `colData` looks like in the pseudo-bulked SCE object.

```{r pseudobulk colData, live=TRUE}
# note the new column with number of cells per group
head(colData(pb_sce)) |>
  as.data.frame()
```

You should see that columns such as `sum`, `detected`, `subsets_mito_sum`, and other columns that typically contain per cell QC statistics now contain `NA` rather than numeric values.
This is because these values were initially calculated on a per cell level (we did this using `scuttle::addPerCellQCMetrics()`), but we no longer have a single column per cell.
Instead, each column now represents a _group_ of cells, in this case comprised of cells of a given cell type and sample combination.
Therefore, the values that we calculated on a per-cell level are no longer applicable to this pseudo-bulked SCE object.

You should also see a new column that wasn't present previously, the `ncells` column.
This column was added during pseudo-bulking and indicates the total number of cells that were summed together to form each column of the SCE object.

Before we proceed we will want to filter out any columns that have a low number of cells.
A low number of cells will usually result in small counts that can cause issues with the statistical approximations made during differential expression analysis.
This is equivalent to filtering out any libraries in bulk RNA-seq analysis that have low library sizes.

We can set a threshold for the number of cells required to continue with our analysis and remove any groups that do not meet the minimum threshold.
Here we will use 10, but the threshold you use for your dataset can vary depending on the composition of cell types.

```{r filter pseudobulk, live=TRUE}
# remove any groups with fewer than 10 cells
filter_pb_sce <- pb_sce[, pb_sce$ncells >= 10]
```

We can then take a look and see how many cell type-sample columns we removed, if any.

```{r print dim, live=TRUE}
# print out dimensions of unfiltered pseudobulk sce
dim(pb_sce)

# dimensions of filtered pseudobulk sce
dim(filter_pb_sce)
```

It looks like we only got rid of one group.
We can do a quick check to see which group was removed by finding which column is no longer present in the filtered object.

```{r removed columns, live=TRUE}
# find removed columns
removed_cols <- !(colnames(pb_sce) %in% colnames(filter_pb_sce))

# print out missing columns
colnames(pb_sce)[removed_cols]
```

The last step we want to do to prepare our dataset for DE is to subset the pseudo-bulked SCE object to contain only the cell type that we are interested in comparing across the two RMS subtypes.
As mentioned previously, we are specifically interested in the `Tumor_Myoblast` cell type.

```{r filter celltype}
# logical vector indicating if cells are tumor myoblast or not
myoblast_cells <- filter_pb_sce$celltype_broad == "Tumor_Myoblast"

# create a new sce with only the tumor myoblasts
tumor_myoblast_sce <- filter_pb_sce[, myoblast_cells]
```

After filtering for our cell type of interest we should have a dataset with 6 columns, 1 for each group of `Tumor_Myoblast` cells in each of our 6 samples.

### Perform differential expression with `DESeq2`

Now we will use the `DESeq2` package to perform differential expression (DE) analysis on our pseudo-bulked SCE object.
From this point, we can proceed in the same way we would if we had a bulk RNA-seq dataset with 6 samples.
We will start with the unnormalized raw counts in the `counts` assay of the pseudo-bulked SCE and do the following with `DESeq2`:

- Create a `DESeqDataSet` object
- Normalize and log transform the counts data
- Estimate dispersions and shrink estimates
- Fit a negative binomial model and perform hypothesis testing using Wald statistics

You can also refer to our [materials from our previous workshops covering bulk RNA-seq](https://github.com/AlexsLemonade/training-modules/tree/master/RNA-seq#readme) for more information on using `DESeq`.

#### Create the `DESeqDataSet` object

To create the `DESeqDataSet` object we will need the unnormalized counts matrix, the metadata associated with the samples, and a design formula.
The first two items are already stored in our SCE object, so we can create a `DESeqDataSet` object directly from that object using the `DESeqDataSet()` function.
The design formula is used to indicate which columns of the metadata need to be considered in the DE comparison.
For our experiment we are comparing gene expression between different RMS subtypes.
The subtype information is stored in the `diagnosis_group` column of the `colData` in the pseudo-bulked SCE.

```{r deseq object, live=TRUE}
# set up the deseq object, group by diagnosis
deseq_object <- DESeq2::DESeqDataSet(tumor_myoblast_sce,
                                     design = ~ diagnosis_group)
```

The pseudo-bulked SCE object contains only one assay: the `counts` assay.
This is because `DESeq2` expects raw counts.
When we run `DESeq2` on our dataset, raw counts will first be normalized using size factors to account for differences in total sample counts.
Therefore we don't have to do any normalization on our own – we'll let `DESeq2` do all the work for us.

However, before we dive into DE analysis, we can do some initial exploration and visualization of our data to see if our samples separate by our known factor of interest, RMS subtype.
In particular, we can use principal component analysis (PCA) of our pseudo-bulked dataset to visualize any variation between samples.
If there is variation between RMS subtypes, we expect their respective samples to separate in PC space, likely indicating presence of differentially expressed genes.
We can evaluate this by plotting PC1 and PC2.

In order to create our PCA plot, we will first need to normalize our data to account for any technical variations across samples.
As a reminder, this is NOT required for running `DESeq2` analysis; we are just using it to visualize our data prior to DE analysis.

```{r normalize}
# estimate size factors first
deseq_object <- DESeq2::estimateSizeFactors(deseq_object)

# normalize and log transform to use for visualization
normalized_object <- DESeq2::rlog(deseq_object,
                                  blind = TRUE)
normalized_object
```

We now have a normalized and transformed object that can be directly input to the `DESeq2::plotPCA()` function, which will both calculate and plot the PC results.

```{r plotPCA, live=TRUE}
DESeq2::plotPCA(normalized_object, intgroup = "diagnosis_group")
```

As expected we see that samples group together based on RMS subtype and are separated along the PC1 axis, the PC contributing the highest amount of variation.

#### Run `DESeq`

We'll now use the convenience function `DESeq()` to perform our differential expression analysis.
This function calculates normalization factors, estimates gene-wise dispersions, fits a negative binomial model and performs hypothesis testing using Wald statistics.

```{r deseq, live=TRUE}
# run DESeq
deseq_object <- DESeq2::DESeq(deseq_object)
```

We can evaluate how well the model fit our data by looking at the dispersion estimates.
We expect to see the dispersion estimates decrease as means are increasing and follow the line of best fit.

```{r plot dispersion, live=TRUE}
plotDispEsts(deseq_object)
```

Now we can extract the results from the object, specifying the p-value threshold that we would like to use.

```{r results, live=TRUE}
# extract the results as a DataFrame
deseq_results <- DESeq2::results(deseq_object, alpha = 0.05)
```

But we aren't done yet!

The estimates of log2 fold change calculated by `DESeq()` are not corrected for expression level.
This means that when counts are small, we are likely to end up with some large fold change values that overestimate the true extent of the change between conditions.

We can correct this by applying a "shrinkage" procedure, which will adjust large values with small counts downward, while preserving values with larger counts, which are likely to be more accurate.

To do this, we will use the `lfcShrink()` function, but first we need to know the name and/or position of the "coefficient" that was calculated by `DESeq()`, which we can do with the `resultsNames()` function.

```{r coefficient, live=TRUE}
# identify position of coefficient
DESeq2::resultsNames(deseq_object)
```


```{r shrinkage}
# appyly logFC shrinkage using the default model
shrink_results <- DESeq2::lfcShrink(
  deseq_object,
  res = deseq_results,
  coef = 2,
  type = "apeglm"
)
head(shrink_results)
```

If you look at our `shrink_results` object, we see that the genes are labeled with the Ensembl gene identifiers, as those were the row names of the pseudo-bulked SCE we used as input to build our `DESeq2` object.
Although some of us may have all of the identifiers memorized by heart, it can be useful to have a human readable symbol in our results.
Before we save the results as a file, we will grab the gene symbols from the `rowData` of our original SCE object and add them as a new column.

```{r add gene symbol}
deseq_results <- shrink_results |>
  # directly add Ensembl id as a column
  # converting results into a data frame
  tibble::as_tibble(rownames = "ensembl_id")

# convert rowdata to data frame
sce_rowdata_df <- rowData(tumor_myoblast_sce) |>
  # create a column with rownames stored as ensembl id
  # use for joining with deseq results
  tibble::as_tibble(rownames = "ensembl_id")

# combine deseq results with rowdata by ensembl id
deseq_results <- deseq_results |>
  dplyr::left_join(sce_rowdata_df, by = "ensembl_id")

head(deseq_results)
```

We can save the new data frame that we have created with the Ensembl identifiers, gene symbols, and the `DESeq2` results as a tab separated (`tsv`) file.

```{r save deseq, live=TRUE}
# save our results as tsv
readr::write_tsv(deseq_results, deseq_output_file)
```

Next, we will take a look at how many genes are significant.
Here we will want to use the adjusted p-value, found in the `padj` column of the results, as this accounts for multiple test correction.

```{r significant results, live=TRUE}
# first look at the significant results
deseq_results_sig <- deseq_results |>
  # filter based on adjusted pvalue
  dplyr::filter(padj <= 0.05)

head(deseq_results_sig)
```


### Exploring the identified differentially expressed genes

Now that we have identified a set of genes that are differentially expressed in the tumor myoblasts between ARMS and ERMS subtypes, lets actually take a look at them and see if we can make some informative plots.
The first plot we'll make is a volcano plot using the [`EnhancedVolcano` package](https://github.com/kevinblighe/EnhancedVolcano).
This package automatically colors the points by cutoffs for both significance and fold change and labels many of the significant genes (subject to spacing).
`EnhancedVolcano` has many, many options, which is a good thing if you don't like all of its default settings.
Even better, it outputs a `ggplot2` object, so if we want to customize the plot further, we can use the same `ggplot2` commands we have used before.

```{r volcano}
EnhancedVolcano::EnhancedVolcano(deseq_results,
                x = 'log2FoldChange', # fold change statistic to plot
                y = 'pvalue', # significance values
                lab = deseq_results$gene_symbol, # labels for points
                pCutoff = 1e-05, # p value cutoff (default)
                FCcutoff = 1, # fold change cutoff (default)
                title = NULL, # no title
                subtitle = NULL, # or subtitle
                caption = NULL, # or caption
                drawConnectors = TRUE, # add some fun arrows
                labSize = 3  # smaller labels
                ) +
  # change the overall theme
  theme_bw() +
  # move the legend to the bottom
  theme(legend.position = "bottom")
```


We can also return back to the SCE object that we used to create our pseudo-bulked SCE and look at gene expression of some of the significant genes.
We can create UMAP plots as we did previously, but instead of labeling each cell with metadata, we can color cells by a specified gene's expression levels.
We will also use some of the `ggplot2` skills we picked up earlier, like `facet_grid()` to plot cells from different RMS subtypes separately.
This can help us validate the `DESeq2` results so that we can visualize gene expression changes across our cell type of interest on a single-cell level.

```{r expression umap, live=TRUE}
# filter to just myoblast cells and remove any NA's before plotting
myoblast_combined_sce <- rms_sce[, which(rms_sce$celltype_broad == "Tumor_Myoblast")]

# plot PTPRT (ENSG00000196090) expression in ARMS vs. ERMS
scater::plotReducedDim(myoblast_combined_sce,
                       dimred = "fastmnn_UMAP",
                       color_by = "ENSG00000196090", #PTPRT
                       point_size= 0.5,
                       point_alpha = 0.4,
                       other_fields = "diagnosis_group") +
  facet_grid(cols = vars(diagnosis_group))
```

In the above plot we only plotted the tumor myoblast cells that we used in our DE analysis.
However, we might be interested to see the expression of genes that are differentially expressed in other cell types present in our samples.

```{r celltype comparison}
# let's compare gene expression across some other cell types
# look at all tumor cells and pick one normal cell type
celltypes <- c("Tumor_Myoblast",
               "Tumor_Mesoderm",
               "Tumor_Myocyte",
               "Vascular Endothelium")

# subset to just celltypes that we are interested in
tumor_sce <- rms_sce[, which(rms_sce$celltype_broad %in% celltypes)]
```

Next we will look at a few DE genes that we identified, one up regulated gene and one down regulated gene, and compare their expression in myoblasts to other cell types in ARMS and ERMS samples.
We will use the `scater::plotExpression()` function to create a violin plot with RMS subtype on the x-axis and gene expression on the y-axis.
We can continue using `facet_grid()` to show separate panels for each cell type.
Because we want to show multiple genes here, we are going to add an additional option to `facet_grid()` to include multiple rows in our plot grid, one for each gene of interest.
One neat trick of the `scater::plotExpression()` function is that it actually creates a `Feature` column which corresponds to the features (in this case genes) being used in plotting.
We can then directly reference that `Feature` column when plotting, instead of using the `other_fields` option we used previously.

```{r multi-gene plot}
# pick a couple genes to look at
genes_to_plot <- c("ENSG00000196090", #PTPRT
                   "ENSG00000148935") #GAS2

# create a violin plot
scater::plotExpression(tumor_sce,
                       # a vector of genes to plot
                       features = genes_to_plot,
                       x = "diagnosis_group",
                       color_by = "diagnosis_group",
                       other_fields = "celltype_broad",
                       point_size = 0.1) +
  # each celltype is its own column
  facet_grid(cols = vars(celltype_broad),
             # each feature (gene) is its own row
             rows = vars(Feature)) +
  # change the font size of the facet labels
  theme(strip.text = element_text(size = 7)) +
  guides(color = guide_legend(
    title = "Subtype", # update the legend title
    # change the size of the legend colors
    override.aes = list(size = 3, alpha = 1))
    )
```

How do the expression of these genes change across cell types and RMS subtypes?

Go ahead and explore some genes on your own!
Feel free to plot any of the genes that are identified as significant, found in the DE results table, or your favorite gene.
Remember, you need to use the Ensembl gene identifier to refer to each gene.

```{r explore}
# now do some exploration of other genes on your own!
```

## Print session info

```{r session info}
sessionInfo()
```


    +
    ---
title: "Differential expression analysis for scRNA-seq data"
author: "Data Lab for ALSF"
date: 2023
output:
  html_notebook:
    toc: yes
    toc_float: yes
---

## Objectives

This notebook will demonstrate how to:

- Use pseudo-bulking to prepare scRNA-seq libraries for differential expression
- Perform differential expression with the `DESeq2` package
- Use `ggplot2` and `EnhancedVolcano` to visualize gene expression changes across cell types and samples

---

Just like bulk RNA-seq, it is likely that one of the goals when performing scRNA-seq will be to compare the gene expression of multiple samples to each other.
Unlike bulk RNA-seq analysis, scRNA-seq analysis allows us to identify and annotate cell types or subpopulations of cells present in each of our samples.
This means that we can account for differences in cell type composition across samples and specifically focus on cell types or populations of interest when performing differential expression (DE) analysis.
In this notebook, we will work with multiple samples to identify differentially expressed genes across cell types of interest using the [`DESeq2`](https://bioconductor.org/packages/release/bioc/html/DESeq2.html) package.

![Single-cell roadmap: Differential expression](diagrams/roadmap_differential_expression.png)

We will continue working with samples from the [`SCPCP000005` project](https://scpca.alexslemonade.org/projects/SCPCP000005), an investigation of pediatric solid tumors led by the Dyer and Chen labs at St. Jude Children's Research Hospital.
This particular dataset contains 10 different samples that have been integrated using `fastMNN`, following the same procedure we outlined in `02-dataset_integration.Rmd`.
These 10 samples represent two different types of rhabdomyosarcoma (RMS): embryonal rhabdomyosarcoma (ERMS) and alveolar rhabdomyosarcoma (ARMS).
These two subtypes are distinguished by the presence of the `PAX3/PAX7-FOXO1` fusion gene, which is present only in ARMS patients.
Additionally, cells found in ARMS tumors tend to have an increased mutational burden with cells in a more differentiated state compared to ERMS tumor cells ([Shern _et al._ 2014](https://doi.org/10.1158/2159-8290.CD-13-0639); [Stewart _et al._ 2018](https://doi.org/10.1016/j.ccell.2018.07.012)).
RMS tumors, regardless of subtype, are made up of cells typically associated with development of skeletal muscle: mesoderm, myoblasts, and myocytes ([Sebire and Malone 2003](https://doi.org/10.1136/jcp.56.6.412)).
[Patel _et al._ (2022)](https://doi.org/10.1016/j.devcel.2022.04.003) tested the hypothesis that cell types have distinct gene expression patterns in ARMS vs. ERMS samples.
Here we will look at a subset of the samples they sequenced and identify differentially expressed genes in tumor cells between ARMS and ERMS samples.

## Set up

```{r setup, message=FALSE}
# set seed for reproducibility
set.seed(2022)

# load libraries
library(ggplot2) # plotting functions
library(SingleCellExperiment)

# package used for differential expression analysis
library(DESeq2)
```

### Directories and files

We will start by reading in a `SingleCellExperiment` (SCE) object that contains both the uncorrected (merged but not integrated) and corrected (integrated) gene expression data for all 10 samples.

Prior to integration, all 10 samples went through the same filtering, normalization, and dimensionality reduction.
These 10 samples were then merged into one `SingleCellExperiment` object following the same steps outlined in `03-dataset_integration.Rmd`.
The merged object was then integrated with `fastMNN` to obtain a corrected gene expression assay and corrected reduced dimensionality results.
The final SCE object was stored in `data/rms/integrated/rms_all_sce.rds`.

We also have provided a metadata file, `data/rms/annotations/rms_sample_metadata.tsv`, that contains information from each sample, such as diagnosis, sex, age, etc.
Each row in this file corresponds to a sample found in the integrated SCE object.

To begin, let's set up our directories and files:

```{r filepaths}
# set up file paths
# data directory for RMS data
data_dir <- file.path("data", "rms")

# integrated file containing samples to use for DE analysis
integrated_sce_file <- file.path(
  data_dir,
  "integrated",
  "rms_all_sce.rds"
)

# sample metadata to set up DE analysis
sample_metadata_file <- file.path(
  data_dir,
  "annotations",
  "rms_sample_metadata.tsv"
)

# directory to store output
deseq_dir <- file.path("analysis", "rms", "deseq")
fs::dir_create(deseq_dir)

# results file to output from DE analysis
deseq_output_file <- file.path(
  deseq_dir,
  "rms_myoblast_deseq_results.tsv"
)

# output integrated sce object
output_sce_file <- file.path(
  data_dir,
  "integrated",
  "rms_subset_sce.rds"
)
```

We can go ahead and read in the SCE object and the metadata file.

```{r read files, live=TRUE}
# read in the SCE object that has already been integrated
integrated_sce <- readr::read_rds(integrated_sce_file)

# read in sample metadata file
sample_metadata <- readr::read_tsv(sample_metadata_file)
```

## Dataset exploration

Before we dive into differential expression, let's explore our integrated SCE object and the dataset a little more.

We'll start by looking at what's inside the object.
Here we should have both the original (uncorrected) data and the integrated (corrected) data for both the gene expression and the reduced dimensionality results.
How are those stored in our object?

```{r print sce, live=TRUE}
# print out entire object
integrated_sce
```


```{r print assay names, live=TRUE}
# look at the assay names in our object
assayNames(integrated_sce)
```

When we look at the assay names we should see that there are 3 matrices, `counts`, `logcounts`, and `fastmnn_corrected`.
The `counts` and `logcounts` assays correspond to the uncorrected gene expression data that has been merged but NOT integrated.
The `fastmnn_corrected` data contains the corrected gene expression data obtained from integration.
For this exercise we will not be using the `fastmnn_corrected` data (more on why not once we get to setting up the differential expression), but we need to be aware that it is present and be able to distinguish it from our uncorrected data.


```{r print reducedDim names, live=TRUE}
# look at the names of the dimension reductions
reducedDimNames(integrated_sce)
```

In the `reducedDim` slots you should see `PCA` and `UMAP`, which were both calculated from the combined data _before_ integration.
You should also see `fastmnn_PCA` and `fastmnn_UMAP` reduced dimensions, which correspond to the integrated results.

### Cell type annotations

Just like in the integration notebook, this dataset also contains the cell type annotations found in the `celltype_fine` and `celltype_broad` columns of the `colData`.
These cell types were originally assigned in [Patel _et al._ (2022)](https://doi.org/10.1016/j.devcel.2022.04.003).
We will use these cell type assignments to set up the DE analysis below, but they are not required for DE analysis itself.
It's important to note that DE analysis can be applied to any subpopulation of interest that is shared across samples besides just cell types.

Because we are going to be doing DE analysis between ARMS and ERMS samples, let's start by labeling cells in the integrated dataset based on their RMS subtype.
To do this we will need to be sure that the subtype is present in the `colData` of the integrated SCE object.
If it's not there, we need to add it in.

```{r coldata head, live=TRUE}
# look at the head of the coldata
head(colData(integrated_sce)) |>
  as.data.frame()
```

Uh oh, it looks like the RMS subtype is not found in the SCE object.
Fortunately we also have the sample metadata table that we read in earlier, which contains information about each of the samples present in the dataset.

```{r sample metadata, live=TRUE}
# print out sample metadata
head(sample_metadata)
```

Looking at this sample table, we see a column named `subdiagnosis` which accounts for the RMS subtype, ARMS or ERMS.
We also see other columns that contain information about each specific sample.

We can incorporate the information in this sample metadata table into the `colData` of the integrated SCE object.
This will allow us to match each of the samples in the SCE object with the RMS subtype and also allow us to use any of the columns in the sample metadata for plotting.

```{r modify coldata}
# add sample metadata to colData from the integrated SCE object
coldata_df <- colData(integrated_sce) |>
  # convert from DataFrame to data.frame
  as.data.frame() |>
  # merge with sample metadata
  dplyr::left_join(sample_metadata, by = c("sample" = "library_id")) |>
  # create new columns
  # cell_id is a combination of barcode and sample
  dplyr::mutate(
    cell_id = glue::glue("{sample}-{barcode}"),
    # simplify subdiagnosis
    diagnosis_group = forcats::fct_recode(
      subdiagnosis,
      "ARMS" = "Alveolar rhabdomyosarcoma",
      "ERMS" = "Embryonal rhabdomyosarcoma"
    )
  )

# add modified data frame back to SCE as DataFrame
colData(integrated_sce) <- DataFrame(
  coldata_df,
  row.names = coldata_df$cell_id
)
```

Now when we look at the `colData` of the SCE object we should see new columns, including the `diagnosis_group` column which indicates if each cell comes from an ERMS or ARMS sample.

```{r print new coldata, live=TRUE}
# take a look at the new modified colData
head(colData(integrated_sce)) |>
  as.data.frame()
```

### Plotting with annotations

We can now use that column to label any UMAP plots (or other plot types) that we make.
In the chunk below we will start by taking a look at our integration results and color our cells by RMS subtype.

**Reminder: You should always use the batch-corrected dimensionality reduction results for visualizing datasets containing multiple libraries or samples.**

```{r diagnosis group UMAP, live=TRUE}
# UMAP of all samples, separating by diagnosis group
scater::plotReducedDim(
  integrated_sce,
  dimred = "fastmnn_UMAP",
  color_by = "diagnosis_group",
  point_size = 0.5,
  point_alpha = 0.2
)
```

Interestingly, it looks like samples from the ARMS and ERMS subtypes tend to group with samples of the same subtype rather than all together.

In the integration notebook we also looked at the distribution of cell types after integration.
In that notebook, we discussed that cells of the same cell type are expected to integrate with other cells of the same type.
Is that the case with this dataset?

A word of caution when evaluating the cell type results for this dataset: The cell types for this dataset were assigned in a two stage process as described in [Patel _et al._ (2022)](https://doi.org/10.1016/j.devcel.2022.04.003).
The first stage assigned cells as tumor or non-tumor.
The next stage further classified tumor cells into one of three types of tumor cells: myoblast, myocyte, or mesoderm.
Some samples could not be further classified, so all of their tumor cells are denoted `Tumor`.
The samples which could be further classified have a mix of `Tumor_Mesoderm`, `Tumor_Myoblast`, and `Tumor_Myocyte`.

```{r celltype UMAP}
# UMAP of all samples labeled by cell type
scater::plotReducedDim(
  integrated_sce,
  dimred = "fastmnn_UMAP",
  # color each point by cell type
  color_by = "celltype_broad",
  point_size = 0.5,
  point_alpha = 0.4
) +
  # Modify the legend key with larger, easier to see points
  guides(color = guide_legend(override.aes = list(size = 3, alpha = 1)))
```

Unlike with the previous datasets we have seen where all cells of the same cell type always grouped together, this dataset shows some slightly different patterns and not all cells of the same cell type cluster together.
One reason is that tumor data can be heterogeneous and every tumor is unique.
Depending on the tumor type we may not expect every sample to integrate perfectly and more heterogeneous tumor types will be more difficult to integrate together.
In this particular case we are looking at two subtypes of RMS that have distinct mutation burdens and differentiation states, so it's likely that those differences contribute to how well they integrate.

To explore whether cells are grouping together both by cell type and by RMS subtype, we can create a plot that incorporates both pieces of metadata.
We will take advantage of the `facet_grid()` function from `ggplot2` to look at two variables in the `colData` at once - the cell type and the subdiagnosis.
In the below plot we will color our cells by cell type while also using `facet_grid()` so that cells from different subdiagnoses will be in their own plot panel.

```{r celltype subdiagnosis UMAP, live=TRUE}
# UMAP of all samples
# separating by diagnosis group and labeling cell type
scater::plotReducedDim(
  integrated_sce,
  dimred = "fastmnn_UMAP",
  # color each point by cell type
  color_by = "celltype_broad",
  point_size = 0.5,
  point_alpha = 0.4,
  # tell scater to use diagnosis_group for plotting
  other_fields = "diagnosis_group"
) +
  # include each diagnosis group as its own column
  facet_grid(cols = vars(diagnosis_group))
```

As expected, we see that cell types are separated, most likely due to different RMS subtypes.

We can also use a stacked barplot to look at the distribution of cell types across each sample, which will require a bit of wrangling first.

```{r celltype barplot}
# filter coldata to only include tumor cells
tumor_cells_df <- coldata_df |>
  # find rows where the cell type name contains the string "Tumor"
  dplyr::filter(stringr::str_detect(celltype_broad, "Tumor"))

# create a stacked barplot
ggplot(tumor_cells_df, aes(x = sample, fill = celltype_broad)) +
  geom_bar(position = "fill", color = "black", size = 0.2) +
  labs(
    x = "Sample",
    y = "Proportion of cells",
    fill = "Cell type"
  ) +
  scale_fill_brewer(palette = "Dark2") +
  theme_bw() +
  theme(axis.text.x = element_text(angle = 90, vjust = 0.5)) +
  # facet by diagnosis group
  facet_grid(
    cols = vars(diagnosis_group),
    # only show non-NA values on x-axis
    scales = "free_x",
    space = "free_x"
  )
```

Similar to the UMAP, this plot shows that ARMS and ERMS share a lot of the same cell types.

We also see that only 6 of these libraries have tumor cells that have been further classified into mesoderm, myoblast, and myocyte.
3 libraries contain cells that are only classified as tumor or non-tumor, and tumor cells are not further classified, and the remaining library is not even present in our plot because it was not assigned any cell types (all are `NA`).
We will continue our analysis only using the 6 libraries with fully classified cell types, removing the other 4 before we proceed with differential expression.

### Filtering samples

The reason we want to pare down our list of samples to consider is that we want to ensure that the cell types (or subpopulations) that we are interested in are present in all samples included in our DE analysis.
We want to remove any samples that do not contain our cell population(s) of interest as they have no counts to contribute to the DE analysis.

```{r subset sce}
# define samples to keep
library_ids <- c(
  "SCPCL000479",
  "SCPCL000480",
  "SCPCL000481",
  "SCPCL000484",
  "SCPCL000488",
  "SCPCL000491"
)

# subset sce to only contain samples of interest
samples_to_keep <- integrated_sce$sample %in% library_ids
rms_sce <- integrated_sce[, samples_to_keep]

# print out our new SCE
rms_sce
```

Before we move on, we'll remove the original integrated object from our environment to save some memory.

```{r remove sce}
rm(integrated_sce)
```

We will also save our new object in case we want to use it for other analysis later on.

```{r save sce}
# write RDS file with compression
readr::write_rds(rms_sce, file = output_sce_file, compress = "gz")
```

We now have an updated SCE object that contains 6 samples that were obtained from a mix of ARMS and ERMS patients.
We can then ask the question, do specific tumor cell types contain sets of differentially expressed genes between ARMS and ERMS samples?

We should make sure that we have enough biological replicates from each group to set up our experiment.
It is imperative to consider good experimental design and ensure that we have enough biological replicates (at least 3 for each group) when performing differential gene expression analysis.

If we look back at our stacked barplot we see that we picked 3 ARMS and 3 ERMS samples.
We can also see that the majority of cells are tumor cells, in particular the largest population of cells appears to be the `Tumor_Myoblast`.
For this example we will focus on identifying DE genes in these `Tumor_Myoblast` cells, but the principles applied below can be applied to any cell types or subpopulations of interest.

## Differential expression analysis

Now we are ready to start preparing for our DE analysis, where we will compare the gene expression of tumor myoblast cells between ARMS and ERMS samples.

Throughout the notebook we have been working with an integrated dataset that contains corrected gene expression data (`fastmnn_corrected` assay) and a corrected UMAP.
As a reminder, the uncorrected gene expression data, found in the `counts` and `logcounts` assays, correspond to data that has been merged (the first step we walked through prior to integration) into the same SCE but not yet integrated.
We do not want to use corrected gene expression values for differential expression; `DESeq2` expects the original raw counts as input so we will be using data found in the `counts` assay of the `SingleCellExperiment` object.

It is advised to only use the corrected values for any analyses being performed at the cell level, e.g., dimensionality reduction.
In contrast, it is not advised to use corrected values for any analyses that are gene-based, such as differential expression or marker gene detection, because within-batch and between-batch gene expression differences are no longer preserved.
The reason for this is two-fold – many of the DE models will expect uncorrected counts because they will account for between-sample variation within the model, and we want to ensure we are preserving variation that is present so as not to artificially inflate differences between populations.
See the [OSCA chapter on Using the corrected values](https://bioconductor.org/books/3.19/OSCA.multisample/using-corrected-values.html#using-corrected-values) for more insight.

### Pseudo-bulking

Before we can compare the gene expression profiles of myoblasts in ARMS vs. ERMS samples, we will need to "pseudo-bulk" the gene counts.
Pseudo-bulking creates a new counts matrix that contains the sum of the counts from all cells with a given label (e.g., cell type) for each sample ([Tung _et al._ 2017](https://doi.org/10.1038/srep39921)).
If we were to keep each cell's counts separate, they would be treated as replicates, leading to inflated statistics.
By pseudo-bulking first, we will now have one count for each gene for each sample and we can take advantage of well-established methods for differential expression with bulk RNA-seq.

Pseudo-bulking is implemented prior to differential expression analysis on single-cell data because it:

- Produces larger and less sparse counts, which allows us to use standard normalization and differential expression methods used by bulk RNA-seq.
- Collapses gene expression counts by sample, so that samples, rather than cells, represent replicates.
- Masks variance within a sample to emphasize variance across samples.
This can be both good and bad!
Masking intra-sample variation means you might not identify genes where average expression doesn't change between samples but the degree of cell-to-cell variation does.

Before we apply pseudo-bulking to our dataset, let's look at a simple example of how pseudo-bulking works.
We'll start by creating a fake matrix of counts.

```{r create matrix}
# create an example counts matrix
counts_mtx <- matrix(
  1:12,
  ncol = 4,
  dimnames = list(
    c("geneA", "geneB", "geneC"),
    c("A-cell1", "A-cell2", "B-cell1", "B-cell2")
  )
)
counts_mtx
```

Next we will create a pseudo-bulked version of this matrix with only 2 columns: 1 for group `A` and 1 for group `B`.
To do this we will use the `DelayedArray::colsum()` function, which allows us to sum the counts for each row across groups of columns.

```{r pseudobulk matrix, live=TRUE}
# define the group that each column belongs to
groups <- c("A", "A", "B", "B")

# sum counts across cells (columns) by group label
pb_counts <- DelayedArray::colsum(counts_mtx, groups)
pb_counts
```

Looking at this output, you should see that the original 4 columns have been condensed to only 2 columns: 1 column to represent all cells from group `A`, and 1 column to represent all cells from group `B`.

Now the actual pseudo-bulking for our dataset!

We will use the [`scuttle::aggregateAcrossCells()` function](https://rdrr.io/github/LTLA/scuttle/man/aggregateAcrossCells.html) to pseudo-bulk our dataset.
This function takes as input an SCE object and the grouping assignments for each cell.
The output will be an SCE object that contains only the pseudo-bulked counts for all genes across all specified groups, rather than across all cells.
We can then subset this SCE to just include our cell type of interest (tumor myoblasts) for input to the DE analysis.

We can pseudo-bulk using any grouping that we are interested in.
For right now, we are interested in looking at gene expression across cell types, so we want to group the pseudo-bulked counts matrix by both cell type and original sample.

```{r pseudobulk sce}
# first subset the coldata
# to only have the columns we care about for pseudo-bulking
pb_groups <- colData(rms_sce)[, c("celltype_broad", "sample")]

# create a new SCE object that contains
# the pseudo-bulked counts across the provided groups
pb_sce <- scuttle::aggregateAcrossCells(
  rms_sce,
  id = pb_groups
)

# column names aren't automatically added to the pseudo-bulked sce,
# so let's add them in
colnames(pb_sce) <- glue::glue(
  "{pb_sce$celltype_broad}_{pb_sce$sample}"
)

pb_sce
```

How does the new pseudo-bulked `SingleCellExperiment` look different?
How many columns does it have?

Let's take a look at what the `colData` looks like in the pseudo-bulked SCE object.

```{r pseudobulk colData, live=TRUE}
# note the new column with number of cells per group
head(colData(pb_sce)) |>
  as.data.frame()
```

You should see that columns such as `sum`, `detected`, `subsets_mito_sum`, and other columns that typically contain per cell QC statistics now contain `NA` rather than numeric values.
This is because these values were initially calculated on a per cell level (we did this using `scuttle::addPerCellQCMetrics()`), but we no longer have a single column per cell.
Instead, each column now represents a _group_ of cells, in this case comprised of cells of a given cell type and sample combination.
Therefore, the values that we calculated on a per-cell level are no longer applicable to this pseudo-bulked SCE object.

You should also see a new column that wasn't present previously, the `ncells` column.
This column was added during pseudo-bulking and indicates the total number of cells that were summed together to form each column of the SCE object.

Before we proceed we will want to filter out any columns that have a low number of cells.
A low number of cells will usually result in small counts that can cause issues with the statistical approximations made during differential expression analysis.
This is equivalent to filtering out any libraries in bulk RNA-seq analysis that have low library sizes.

We can set a threshold for the number of cells required to continue with our analysis and remove any groups that do not meet the minimum threshold.
Here we will use 10, but the threshold you use for your dataset can vary depending on the composition of cell types.

```{r filter pseudobulk, live=TRUE}
# remove any groups with fewer than 10 cells
filter_pb_sce <- pb_sce[, pb_sce$ncells >= 10]
```

We can then take a look and see how many cell type-sample columns we removed, if any.

```{r print dim, live=TRUE}
# print out dimensions of unfiltered pseudobulk sce
dim(pb_sce)

# dimensions of filtered pseudobulk sce
dim(filter_pb_sce)
```

It looks like we only got rid of one group.
We can do a quick check to see which group was removed by finding which column is no longer present in the filtered object.

```{r removed columns, live=TRUE}
# find removed columns
removed_cols <- !(colnames(pb_sce) %in% colnames(filter_pb_sce))

# print out missing columns
colnames(pb_sce)[removed_cols]
```

The last step we want to do to prepare our dataset for DE is to subset the pseudo-bulked SCE object to contain only the cell type that we are interested in comparing across the two RMS subtypes.
As mentioned previously, we are specifically interested in the `Tumor_Myoblast` cell type.

```{r filter celltype}
# logical vector indicating if cells are tumor myoblast or not
myoblast_cells <- filter_pb_sce$celltype_broad == "Tumor_Myoblast"

# create a new sce with only the tumor myoblasts
tumor_myoblast_sce <- filter_pb_sce[, myoblast_cells]
```

After filtering for our cell type of interest we should have a dataset with 6 columns, 1 for each group of `Tumor_Myoblast` cells in each of our 6 samples.

### Perform differential expression with `DESeq2`

Now we will use the `DESeq2` package to perform differential expression (DE) analysis on our pseudo-bulked SCE object.
From this point, we can proceed in the same way we would if we had a bulk RNA-seq dataset with 6 samples.
We will start with the unnormalized raw counts in the `counts` assay of the pseudo-bulked SCE and do the following with `DESeq2`:

- Create a `DESeqDataSet` object
- Normalize and log transform the counts data
- Estimate dispersions and shrink estimates
- Fit a negative binomial model and perform hypothesis testing using Wald statistics

You can also refer to our [materials from our previous workshops covering bulk RNA-seq](https://github.com/AlexsLemonade/training-modules/tree/master/RNA-seq#readme) for more information on using `DESeq`.

#### Create the `DESeqDataSet` object

To create the `DESeqDataSet` object we will need the unnormalized counts matrix, the metadata associated with the samples, and a design formula.
The first two items are already stored in our SCE object, so we can create a `DESeqDataSet` object directly from that object using the `DESeqDataSet()` function.
The design formula is used to indicate which columns of the metadata need to be considered in the DE comparison.
For our experiment we are comparing gene expression between different RMS subtypes.
The subtype information is stored in the `diagnosis_group` column of the `colData` in the pseudo-bulked SCE.

```{r deseq object, live=TRUE}
# set up the deseq object, group by diagnosis
deseq_object <- DESeq2::DESeqDataSet(
  tumor_myoblast_sce,
  design = ~diagnosis_group
)
```

The pseudo-bulked SCE object contains only one assay: the `counts` assay.
This is because `DESeq2` expects raw counts.
When we run `DESeq2` on our dataset, raw counts will first be normalized using size factors to account for differences in total sample counts.
Therefore we don't have to do any normalization on our own – we'll let `DESeq2` do all the work for us.

However, before we dive into DE analysis, we can do some initial exploration and visualization of our data to see if our samples separate by our known factor of interest, RMS subtype.
In particular, we can use principal component analysis (PCA) of our pseudo-bulked dataset to visualize any variation between samples.
If there is variation between RMS subtypes, we expect their respective samples to separate in PC space, likely indicating presence of differentially expressed genes.
We can evaluate this by plotting PC1 and PC2.

In order to create our PCA plot, we will first need to normalize our data to account for any technical variations across samples.
As a reminder, this is NOT required for running `DESeq2` analysis; we are just using it to visualize our data prior to DE analysis.

```{r normalize}
# estimate size factors first
deseq_object <- DESeq2::estimateSizeFactors(deseq_object)

# normalize and log transform to use for visualization
normalized_object <- DESeq2::rlog(
  deseq_object,
  blind = TRUE
)
normalized_object
```

We now have a normalized and transformed object that can be directly input to the `DESeq2::plotPCA()` function, which will both calculate and plot the PC results.

```{r plotPCA, live=TRUE}
DESeq2::plotPCA(normalized_object, intgroup = "diagnosis_group")
```

As expected we see that samples group together based on RMS subtype and are separated along the PC1 axis, the PC contributing the highest amount of variation.

#### Run `DESeq`

We'll now use the convenience function `DESeq()` to perform our differential expression analysis.
This function calculates normalization factors, estimates gene-wise dispersions, fits a negative binomial model and performs hypothesis testing using Wald statistics.

```{r deseq, live=TRUE}
# run DESeq
deseq_object <- DESeq2::DESeq(deseq_object)
```

We can evaluate how well the model fit our data by looking at the dispersion estimates.
We expect to see the dispersion estimates decrease as means are increasing and follow the line of best fit.

```{r plot dispersion, live=TRUE}
plotDispEsts(deseq_object)
```

Now we can extract the results from the object, specifying the p-value threshold that we would like to use.

```{r results, live=TRUE}
# extract the results as a DataFrame
deseq_results <- DESeq2::results(deseq_object, alpha = 0.05)
```

But we aren't done yet!

The estimates of log2 fold change calculated by `DESeq()` are not corrected for expression level.
This means that when counts are small, we are likely to end up with some large fold change values that overestimate the true extent of the change between conditions.

We can correct this by applying a "shrinkage" procedure, which will adjust large values with small counts downward, while preserving values with larger counts, which are likely to be more accurate.

To do this, we will use the `lfcShrink()` function, but first we need to know the name and/or position of the "coefficient" that was calculated by `DESeq()`, which we can do with the `resultsNames()` function.

```{r coefficient, live=TRUE}
# identify position of coefficient
DESeq2::resultsNames(deseq_object)
```


```{r shrinkage}
# appyly logFC shrinkage using the default model
shrink_results <- DESeq2::lfcShrink(
  deseq_object,
  res = deseq_results,
  coef = 2,
  type = "apeglm"
)
head(shrink_results)
```

If you look at our `shrink_results` object, we see that the genes are labeled with the Ensembl gene identifiers, as those were the row names of the pseudo-bulked SCE we used as input to build our `DESeq2` object.
Although some of us may have all of the identifiers memorized by heart, it can be useful to have a human readable symbol in our results.
Before we save the results as a file, we will grab the gene symbols from the `rowData` of our original SCE object and add them as a new column.

```{r add gene symbol}
deseq_results <- shrink_results |>
  # directly add Ensembl id as a column
  # converting results into a data frame
  tibble::as_tibble(rownames = "ensembl_id")

# convert rowdata to data frame
sce_rowdata_df <- rowData(tumor_myoblast_sce) |>
  # create a column with rownames stored as ensembl id
  # use for joining with deseq results
  tibble::as_tibble(rownames = "ensembl_id")

# combine deseq results with rowdata by ensembl id
deseq_results <- deseq_results |>
  dplyr::left_join(sce_rowdata_df, by = "ensembl_id")

head(deseq_results)
```

We can save the new data frame that we have created with the Ensembl identifiers, gene symbols, and the `DESeq2` results as a tab separated (`tsv`) file.

```{r save deseq, live=TRUE}
# save our results as tsv
readr::write_tsv(deseq_results, deseq_output_file)
```

Next, we will take a look at how many genes are significant.
Here we will want to use the adjusted p-value, found in the `padj` column of the results, as this accounts for multiple test correction.

```{r significant results, live=TRUE}
# first look at the significant results
deseq_results_sig <- deseq_results |>
  # filter based on adjusted pvalue
  dplyr::filter(padj <= 0.05)

head(deseq_results_sig)
```


### Exploring the identified differentially expressed genes

Now that we have identified a set of genes that are differentially expressed in the tumor myoblasts between ARMS and ERMS subtypes, lets actually take a look at them and see if we can make some informative plots.
The first plot we'll make is a volcano plot using the [`EnhancedVolcano` package](https://github.com/kevinblighe/EnhancedVolcano).
This package automatically colors the points by cutoffs for both significance and fold change and labels many of the significant genes (subject to spacing).
`EnhancedVolcano` has many, many options, which is a good thing if you don't like all of its default settings.
Even better, it outputs a `ggplot2` object, so if we want to customize the plot further, we can use the same `ggplot2` commands we have used before.

```{r volcano}
EnhancedVolcano::EnhancedVolcano(
  deseq_results,
  x = "log2FoldChange", # fold change statistic to plot
  y = "pvalue", # significance values
  lab = deseq_results$gene_symbol, # labels for points
  pCutoff = 1e-05, # p value cutoff (default)
  FCcutoff = 1, # fold change cutoff (default)
  title = NULL, # no title
  subtitle = NULL, # or subtitle
  caption = NULL, # or caption
  drawConnectors = TRUE, # add some fun arrows
  labSize = 3 # smaller labels
) +
  # change the overall theme
  theme_bw() +
  # move the legend to the bottom
  theme(legend.position = "bottom")
```


We can also return back to the SCE object that we used to create our pseudo-bulked SCE and look at gene expression of some of the significant genes.
We can create UMAP plots as we did previously, but instead of labeling each cell with metadata, we can color cells by a specified gene's expression levels.
We will also use some of the `ggplot2` skills we picked up earlier, like `facet_grid()` to plot cells from different RMS subtypes separately.
This can help us validate the `DESeq2` results so that we can visualize gene expression changes across our cell type of interest on a single-cell level.

```{r expression umap, live=TRUE}
# filter to just myoblast cells and remove any NA's before plotting
myoblast_combined_sce <- rms_sce[, which(rms_sce$celltype_broad == "Tumor_Myoblast")]

# plot PTPRT (ENSG00000196090) expression in ARMS vs. ERMS
scater::plotReducedDim(
  myoblast_combined_sce,
  dimred = "fastmnn_UMAP",
  color_by = "ENSG00000196090", # PTPRT
  point_size = 0.5,
  point_alpha = 0.4,
  other_fields = "diagnosis_group"
) +
  facet_grid(cols = vars(diagnosis_group))
```

In the above plot we only plotted the tumor myoblast cells that we used in our DE analysis.
However, we might be interested to see the expression of genes that are differentially expressed in other cell types present in our samples.

```{r celltype comparison}
# let's compare gene expression across some other cell types
# look at all tumor cells and pick one normal cell type
celltypes <- c(
  "Tumor_Myoblast",
  "Tumor_Mesoderm",
  "Tumor_Myocyte",
  "Vascular Endothelium"
)

# subset to just celltypes that we are interested in
tumor_sce <- rms_sce[, which(rms_sce$celltype_broad %in% celltypes)]
```

Next we will look at a few DE genes that we identified, one up regulated gene and one down regulated gene, and compare their expression in myoblasts to other cell types in ARMS and ERMS samples.
We will use the `scater::plotExpression()` function to create a violin plot with RMS subtype on the x-axis and gene expression on the y-axis.
We can continue using `facet_grid()` to show separate panels for each cell type.
Because we want to show multiple genes here, we are going to add an additional option to `facet_grid()` to include multiple rows in our plot grid, one for each gene of interest.
One neat trick of the `scater::plotExpression()` function is that it actually creates a `Feature` column which corresponds to the features (in this case genes) being used in plotting.
We can then directly reference that `Feature` column when plotting, instead of using the `other_fields` option we used previously.

```{r multi-gene plot}
# pick a couple genes to look at
genes_to_plot <- c(
  "ENSG00000196090", # PTPRT
  "ENSG00000148935"
) # GAS2

# create a violin plot
scater::plotExpression(
  tumor_sce,
  # a vector of genes to plot
  features = genes_to_plot,
  x = "diagnosis_group",
  color_by = "diagnosis_group",
  other_fields = "celltype_broad",
  point_size = 0.1
) +
  # each celltype is its own column
  facet_grid(
    cols = vars(celltype_broad),
    # each feature (gene) is its own row
    rows = vars(Feature)
  ) +
  # change the font size of the facet labels
  theme(strip.text = element_text(size = 7)) +
  guides(
    color = guide_legend(
      # update the legend title
      title = "Subtype",
      # change the size of the legend colors
      override.aes = list(size = 3, alpha = 1)
    )
  )
```

How do the expression of these genes change across cell types and RMS subtypes?

Go ahead and explore some genes on your own!
Feel free to plot any of the genes that are identified as significant, found in the DE results table, or your favorite gene.
Remember, you need to use the Ensembl gene identifier to refer to each gene.

```{r explore}
# now do some exploration of other genes on your own!
```

## Print session info

```{r session info}
sessionInfo()
```


    diff --git a/scRNA-seq-advanced/04-gene_set_enrichment_analysis-live.Rmd b/scRNA-seq-advanced/04-gene_set_enrichment_analysis-live.Rmd index f022ca51..504d97f4 100644 --- a/scRNA-seq-advanced/04-gene_set_enrichment_analysis-live.Rmd +++ b/scRNA-seq-advanced/04-gene_set_enrichment_analysis-live.Rmd @@ -55,12 +55,14 @@ library(msigdbr) #### Directories -```{r create_dir, live = TRUE} +```{r create_dir} # We'll use the differential expression results as GSEA input +rms_analysis_dir <- file.path("analysis", "rms") # We'll create a directory to specifically hold the pathway results if it doesn't # exist yet - +results_dir <- file.path(rms_analysis_dir, "pathway-analysis") +fs::dir_create(results_dir) ``` #### Input files @@ -134,7 +136,7 @@ The enrichment score for a pathway is the running sum's maximum deviation from z GSEA also assesses statistical significance of the scores for each pathway through permutation testing. As a result, each input pathway will have a p-value associated with it that is then corrected for multiple hypothesis testing ([Subramanian _et al._ 2005](https://doi.org/10.1073/pnas.0506580102); [Yu](http://yulab-smu.top/clusterProfiler-book/chapter2.html#gene-set-enrichment-analysis)). -The implementation of GSEA we use in here examples requires a gene list ordered by some statistic and input gene sets. +The implementation of GSEA we use here requires a gene list ordered by some statistic and input gene sets. When you use previously computed gene-level statistics with GSEA, it is called GSEA pre-ranked. ## DESeq2 results @@ -164,7 +166,7 @@ You can see an example in this Harvard Chan Bioinformatics Core Training materia One good thing about Ensembl gene identifiers is that they are less likely to be duplicated than, for example, gene symbols. (Multiple Ensembl gene identifiers can map to the same symbol.) -The GSEA approach requires on discriminating between genes that are in a gene set and those that are not. +The GSEA approach requires discriminating between genes that are in a gene set and those that are not. Practically speaking, gene sets are just collections of gene identifiers! When the function we use for GSEA pre-ranked gets a list with duplicated gene identifiers, it can produce unexpected results. So, let's check for duplicates in the data frame of DESeq2 results. @@ -227,10 +229,11 @@ Normalized enrichment scores (NES) are enrichment scores that are scaled to make Pathways with significant, highly positive NES are enriched in ERMS myoblasts, whereas pathways with significant, highly negative NES are enriched in ARMS myoblasts. -Let's write these results to file. +Let's write these results to a file. ```{r write_gsea} -gsea_results@result |> readr::write_tsv(output_file) +gsea_results@result |> + readr::write_tsv(output_file) ``` ### Visualizing GSEA results diff --git a/scRNA-seq-advanced/04-gene_set_enrichment_analysis.Rmd b/scRNA-seq-advanced/04-gene_set_enrichment_analysis.Rmd index 7c43a03b..513c4ea7 100644 --- a/scRNA-seq-advanced/04-gene_set_enrichment_analysis.Rmd +++ b/scRNA-seq-advanced/04-gene_set_enrichment_analysis.Rmd @@ -55,7 +55,7 @@ library(msigdbr) #### Directories -```{r create_dir, live = TRUE} +```{r create_dir} # We'll use the differential expression results as GSEA input rms_analysis_dir <- file.path("analysis", "rms") @@ -69,9 +69,11 @@ fs::dir_create(results_dir) ```{r input_files} -input_file <- file.path(rms_analysis_dir, - "deseq", - "rms_myoblast_deseq_results.tsv") +input_file <- file.path( + rms_analysis_dir, + "deseq", + "rms_myoblast_deseq_results.tsv" +) ``` #### Output files @@ -79,8 +81,10 @@ input_file <- file.path(rms_analysis_dir, We'll save our table of GSEA results as a TSV. ```{r output_files} -output_file <- file.path(results_dir, - "rms_myoblast_gsea_results.tsv") +output_file <- file.path( + results_dir, + "rms_myoblast_gsea_results.tsv" +) ``` ## Gene sets @@ -113,11 +117,13 @@ Here's an excerpt of the [collection description](https://www.gsea-msigdb.org/gs Notably, there are only 50 gene sets included in this collection. The fewer gene sets we test, the lower our multiple hypothesis testing burden. -We can retrieve only the Hallmark gene sets by specifying `category = "H"` to the `msigdbr()` function. +We can retrieve only the Hallmark gene sets by specifying `collection = "H"` to the `msigdbr()` function. ```{r immunologic_sets, live = TRUE} -hs_hallmarks_df <- msigdbr(species = "Homo sapiens", - category = "H") +hs_hallmarks_df <- msigdbr( + species = "Homo sapiens", + collection = "H" +) ``` ## Gene Set Enrichment Analysis @@ -137,7 +143,7 @@ The enrichment score for a pathway is the running sum's maximum deviation from z GSEA also assesses statistical significance of the scores for each pathway through permutation testing. As a result, each input pathway will have a p-value associated with it that is then corrected for multiple hypothesis testing ([Subramanian _et al._ 2005](https://doi.org/10.1073/pnas.0506580102); [Yu](http://yulab-smu.top/clusterProfiler-book/chapter2.html#gene-set-enrichment-analysis)). -The implementation of GSEA we use in here examples requires a gene list ordered by some statistic and input gene sets. +The implementation of GSEA we use here requires a gene list ordered by some statistic and input gene sets. When you use previously computed gene-level statistics with GSEA, it is called GSEA pre-ranked. ## DESeq2 results @@ -167,7 +173,7 @@ You can see an example in this Harvard Chan Bioinformatics Core Training materia One good thing about Ensembl gene identifiers is that they are less likely to be duplicated than, for example, gene symbols. (Multiple Ensembl gene identifiers can map to the same symbol.) -The GSEA approach requires on discriminating between genes that are in a gene set and those that are not. +The GSEA approach requires discriminating between genes that are in a gene set and those that are not. Practically speaking, gene sets are just collections of gene identifiers! When the function we use for GSEA pre-ranked gets a list with duplicated gene identifiers, it can produce unexpected results. So, let's check for duplicates in the data frame of DESeq2 results. @@ -211,20 +217,23 @@ Now for the analysis! We can use the `GSEA()` function to perform GSEA with any generic set of gene sets, but there are several functions for using specific, commonly used gene sets (e.g., `gseKEGG()`). ```{r run_gsea} -gsea_results <- GSEA(geneList = lfc_vector, # ordered ranked gene list - minGSSize = 25, # minimum gene set size - maxGSSize = 500, # maximum gene set set - pvalueCutoff = 0.05, - pAdjustMethod = "BH", # correction for multiple hypothesis testing - TERM2GENE = dplyr::select(hs_hallmarks_df, - gs_name, - ensembl_gene)) # pass the correct identifier column +gsea_results <- GSEA( + geneList = lfc_vector, # ordered ranked gene list + minGSSize = 25, # minimum gene set size + maxGSSize = 500, # maximum gene set set + pvalueCutoff = 0.05, + pAdjustMethod = "BH", # correction for multiple hypothesis testing + TERM2GENE = dplyr::select(hs_hallmarks_df, + gs_name, + ensembl_gene) # pass the correct identifier column +) ``` Let's take a look at the GSEA results. ```{r view_gsea, live = TRUE, eval = FALSE} -View(gsea_results@result |> - dplyr::arrange(dplyr::desc(NES)) +View( + gsea_results@result |> + dplyr::arrange(dplyr::desc(NES)) ) ``` @@ -232,10 +241,11 @@ Normalized enrichment scores (NES) are enrichment scores that are scaled to make Pathways with significant, highly positive NES are enriched in ERMS myoblasts, whereas pathways with significant, highly negative NES are enriched in ARMS myoblasts. -Let's write these results to file. +Let's write these results to a file. ```{r write_gsea} -gsea_results@result |> readr::write_tsv(output_file) +gsea_results@result |> + readr::write_tsv(output_file) ``` ### Visualizing GSEA results @@ -248,10 +258,12 @@ Let's take a look at 3 different pathways -- one with a highly positive NES, one Let's take look at a pathway with a highly positive NES (`HALLMARK_MYC_TARGETS_V2`) using a GSEA plot. ```{r highly_pos} -enrichplot::gseaplot(gsea_results, - geneSetID = "HALLMARK_MYC_TARGETS_V2", - title = "HALLMARK_MYC_TARGETS_V2", - color.line = "#0066FF") +enrichplot::gseaplot( + gsea_results, + geneSetID = "HALLMARK_MYC_TARGETS_V2", + title = "HALLMARK_MYC_TARGETS_V2", + color.line = "#0066FF" +) ``` Notice how the genes that are in the gene set, indicated by the black bars, tend to be on the left side of the graph indicating that they have positive gene-level scores. @@ -261,10 +273,12 @@ Notice how the genes that are in the gene set, indicated by the black bars, tend The gene set `HALLMARK_MYOGENESIS` had a highly negative NES. ```{r highly_neg} -enrichplot::gseaplot(gsea_results, - geneSetID = "HALLMARK_MYOGENESIS", - title = "HALLMARK_MYOGENESIS", - color.line = "#0066FF") +enrichplot::gseaplot( + gsea_results, + geneSetID = "HALLMARK_MYOGENESIS", + title = "HALLMARK_MYOGENESIS", + color.line = "#0066FF" +) ``` This gene set shows the opposite pattern -- genes in the pathway tend to be on the right side of the graph. @@ -275,10 +289,12 @@ The `@results` slot will only show gene sets that pass the `pvalueCutoff` thresh Let's look at `HALLMARK_P53_PATHWAY`, which was not in the results we viewed earlier. ```{r p53, live = TRUE} -enrichplot::gseaplot(gsea_results, - geneSetID = "HALLMARK_P53_PATHWAY", - title = "HALLMARK_P53_PATHWAY", - color.line = "#0066FF") +enrichplot::gseaplot( + gsea_results, + geneSetID = "HALLMARK_P53_PATHWAY", + title = "HALLMARK_P53_PATHWAY", + color.line = "#0066FF" +) ``` Genes in the pathway are distributed more evenly throughout the ranked list, resulting in a more "middling" score. diff --git a/scRNA-seq-advanced/04-gene_set_enrichment_analysis.nb.html b/scRNA-seq-advanced/04-gene_set_enrichment_analysis.nb.html index 307ca33d..07e65fd9 100644 --- a/scRNA-seq-advanced/04-gene_set_enrichment_analysis.nb.html +++ b/scRNA-seq-advanced/04-gene_set_enrichment_analysis.nb.html @@ -3090,10 +3090,9 @@

    Gene Set Enrichment Analysis

    input pathway will have a p-value associated with it that is then corrected for multiple hypothesis testing (Subramanian et al. 2005; Yu).

    -

    The implementation of GSEA we use in here examples requires a gene -list ordered by some statistic and input gene sets. When you use -previously computed gene-level statistics with GSEA, it is called GSEA -pre-ranked.

    +

    The implementation of GSEA we use here requires a gene list ordered +by some statistic and input gene sets. When you use previously computed +gene-level statistics with GSEA, it is called GSEA pre-ranked.

    DESeq2 results

    @@ -3152,12 +3151,12 @@

    DESeq2 results

    One good thing about Ensembl gene identifiers is that they are less likely to be duplicated than, for example, gene symbols. (Multiple Ensembl gene identifiers can map to the same symbol.)

    -

    The GSEA approach requires on discriminating between genes that are -in a gene set and those that are not. Practically speaking, gene sets -are just collections of gene identifiers! When the function we use for -GSEA pre-ranked gets a list with duplicated gene identifiers, it can -produce unexpected results. So, let’s check for duplicates in the data -frame of DESeq2 results.

    +

    The GSEA approach requires discriminating between genes that are in a +gene set and those that are not. Practically speaking, gene sets are +just collections of gene identifiers! When the function we use for GSEA +pre-ranked gets a list with duplicated gene identifiers, it can produce +unexpected results. So, let’s check for duplicates in the data frame of +DESeq2 results.

    @@ -3258,9 +3257,10 @@

    Run GSEA

    Let’s take a look at the GSEA results.

    - -
    View(gsea_results@result |>
    -       dplyr::arrange(dplyr::desc(NES))
    +
    +
    View(
    +  gsea_results@result |>
    +    dplyr::arrange(dplyr::desc(NES))
     )
    @@ -3271,11 +3271,12 @@

    Run GSEA

    Pathways with significant, highly positive NES are enriched in ERMS myoblasts, whereas pathways with significant, highly negative NES are enriched in ARMS myoblasts.

    -

    Let’s write these results to file.

    +

    Let’s write these results to a file.

    - -
    gsea_results@result |> readr::write_tsv(output_file)
    + +
    gsea_results@result |> 
    +  readr::write_tsv(output_file)
    @@ -3428,7 +3429,7 @@

    Session Info

    -
    ---
title: "Pathway analysis: Gene Set Enrichment Analysis (GSEA)"
output:
  html_notebook:
    toc: true
    toc_float: true
author: CCDL for ALSF
date: 2024
---

## Objectives

This notebook will demonstrate how to:

- Prepare tabular data of gene-level statistics for use with Gene Set Enrichment Analysis (GSEA)
- Access [Molecular Signatures Database gene set collections](https://www.gsea-msigdb.org/gsea/msigdb/collections.jsp) via the `msigdbr` package
- Perform GSEA with the `clusterProfiler` package
- Visualize GSEA results with the `enrichplot` package

---

In this notebook, we'll analyze the differential expression results from the last notebook.

GSEA is a functional class scoring (FCS) approach to pathway analysis that was first introduced in [Subramanian _et al._ (2005)](https://doi.org/10.1073/pnas.0506580102).
The rationale behind FCS approaches is that small changes in individual genes that participate in the same biological process or pathway can be significant and of biological interest.

There are 3 general steps in FCS methods ([Khatri _et al._ 2012](https://doi.org/10.1371/journal.pcbi.1002375)):

1. Calculate a gene-level statistic (here, we'll use the summary log fold changes in our DESeq2 results)
2. Aggregate gene-level statistics into a pathway-level statistic
3. Assess the statistical significance of the pathway-level statistic

#### Other resources

* For another example using `clusterProfiler` for GSEA, see [_Intro to DGE: Functional Analysis._ from Harvard Chan Bioinformatics Core Training.](https://hbctraining.github.io/Training-modules/DGE-functional-analysis/lessons/02_functional_analysis.html)
* The way we'll use `clusterProfiler` here uses `fgsea` (Fast Gene Set Enrichment Analysis) under the hood.
You can read more about `fgsea` in [Korotkevich _et al._ (2021)](https://doi.org/10.1101/060012).
* See the [refine.bio examples for "Gene set enrichment analysis - RNA-seq"](https://alexslemonade.github.io/refinebio-examples/03-rnaseq/pathway-analysis_rnaseq_02_gsea.html) from which this material has been adapted.

## Set up

### Libraries

```{r setup}
# set seed for reproducibility
set.seed(2025)

# Package to run GSEA
library(clusterProfiler)
# Package that contains the MSigDB gene sets in tidy format
library(msigdbr)
```

### Directories and Files

#### Directories

```{r create_dir, live = TRUE}
# We'll use the differential expression results as GSEA input
rms_analysis_dir <- file.path("analysis", "rms")

# We'll create a directory to specifically hold the pathway results if it doesn't
# exist yet
results_dir <- file.path(rms_analysis_dir, "pathway-analysis")
fs::dir_create(results_dir)
```

#### Input files


```{r input_files}
input_file <- file.path(rms_analysis_dir,
                        "deseq",
                        "rms_myoblast_deseq_results.tsv")
```

#### Output files

We'll save our table of GSEA results as a TSV.

```{r output_files}
output_file <- file.path(results_dir,
                         "rms_myoblast_gsea_results.tsv")
```

## Gene sets

We will use gene sets from the [Molecular Signatures Database (MSigDB)](https://www.gsea-msigdb.org/gsea/msigdb/index.jsp) from the Broad Institute ([Subramanian, Tamayo *et al.* 2005](https://doi.org/10.1073/pnas.0506580102)).
The [`msigdbr`](https://cran.r-project.org/web/packages/msigdbr/index.html) package contains MSigDB datasets already in the tidy format required by `clusterProfiler` and supports multiple organisms.

Let's take a look at what organisms the package supports.

```{r show_species}
msigdbr_species()
```

MSigDB contains 8 different gene set collections.

    H: hallmark gene sets
    C1: positional gene sets
    C2: curated gene sets
    C3: motif gene sets
    C4: computational gene sets
    C5: GO gene sets
    C6: oncogenic signatures
    C7: immunologic signatures

We'll use the Hallmark collection for GSEA.
Here's an excerpt of the [collection description](https://www.gsea-msigdb.org/gsea/msigdb/collection_details.jsp#H):

> Hallmark gene sets summarize and represent specific well-defined biological states or processes and display coherent expression. These gene sets were generated by a computational methodology based on identifying gene set overlaps and retaining genes that display coordinate expression. The hallmarks reduce noise and redundancy and provide a better delineated biological space for GSEA.

Notably, there are only 50 gene sets included in this collection.
The fewer gene sets we test, the lower our multiple hypothesis testing burden.

We can retrieve only the Hallmark gene sets by specifying `category = "H"` to the `msigdbr()` function.

```{r immunologic_sets, live = TRUE}
hs_hallmarks_df <- msigdbr(species = "Homo sapiens",
                           category = "H")
```

## Gene Set Enrichment Analysis

_Adapted from [refine.bio examples](https://github.com/AlexsLemonade/refinebio-examples/blob/33cdeff66d57f9fe8ee4fcb5156aea4ac2dce07f/03-rnaseq/pathway-analysis_rnaseq_02_gsea.Rmd)_

![](diagrams/subramanian_fig1.jpg)

**Figure 1. [Subramanian _et al._ (2005)](https://doi.org/10.1073/pnas.0506580102).**

GSEA calculates a pathway-level metric, called an enrichment score (sometimes abbreviated as ES), by ranking genes by a gene-level statistic.
This score reflects whether or not a gene set or pathway is over-represented at the top or bottom of the gene rankings ([Subramanian _et al._ 2005](https://doi.org/10.1073/pnas.0506580102); [Yu](http://yulab-smu.top/clusterProfiler-book/chapter2.html#gene-set-enrichment-analysis))

Specifically, all genes are ranked from most positive to most negative based on their statistic and a running sum is calculated:
Starting with the most highly ranked genes, the running sum increases for each gene in the pathway and decreases for each gene not in the pathway.
The enrichment score for a pathway is the running sum's maximum deviation from zero.
GSEA also assesses statistical significance of the scores for each pathway through permutation testing.
As a result, each input pathway will have a p-value associated with it that is then corrected for multiple hypothesis testing ([Subramanian _et al._ 2005](https://doi.org/10.1073/pnas.0506580102); [Yu](http://yulab-smu.top/clusterProfiler-book/chapter2.html#gene-set-enrichment-analysis)).

The implementation of GSEA we use in here examples requires a gene list ordered by some statistic and input gene sets.
When you use previously computed gene-level statistics with GSEA, it is called GSEA pre-ranked.

## DESeq2 results

```{r read_in_markers, live = TRUE}
deseq_df <- readr::read_tsv(input_file)
```

```{r deseq_head}
head(deseq_df)
```

This data frame uses Ensembl gene identifiers.
We'll need to make sure our gene sets use the same identifiers.
Let's take a look at the first few rows of the data frame that contains the hallmark gene sets.

```{r hallmark_head, live = TRUE}
head(hs_hallmarks_df)
```

We can see that the gene sets from `msigdbr` have Ensembl gene identifiers associated with them, so we don't need to do any conversion.
However, we'll need to pass the correct column to the function that runs GSEA.

If we needed to do gene identifier conversion, we would likely use the `AnnotationDbi` package.
You can see an example in this Harvard Chan Bioinformatics Core Training material: <https://hbctraining.github.io/DGE_workshop_salmon_online/lessons/AnnotationDbi_lesson.html>

One good thing about Ensembl gene identifiers is that they are less likely to be duplicated than, for example, gene symbols.
(Multiple Ensembl gene identifiers can map to the same symbol.)

The GSEA approach requires on discriminating between genes that are in a gene set and those that are not.
Practically speaking, gene sets are just collections of gene identifiers!
When the function we use for GSEA pre-ranked gets a list with duplicated gene identifiers, it can produce unexpected results.
So, let's check for duplicates in the data frame of DESeq2 results.

```{r check_duplicates, live = TRUE}
any(duplicated(deseq_df$ensembl_id))
```

There are no duplicates for us to worry about!

### Pre-ranked list

The `GSEA()` function takes a pre-ranked (sorted) named vector of statistics, where the names in the vector are gene identifiers.
This is step 1 -- gene-level statistics.

```{r lfc_vector}
lfc_vector <- deseq_df |>
  # Extract a vector of `log2FoldChange` named by `ensembl_id`
  dplyr::pull(log2FoldChange, name = ensembl_id)
lfc_vector <- sort(lfc_vector, decreasing = TRUE)
```

Let's look at the top ranked values.

```{r head_lfc, live = TRUE}
# Look at first entries of the log fold change vector
head(lfc_vector)
```

And the bottom of the list.

```{r tail_lfc, live = TRUE}
# Look at the last entries of the log fold change vector
tail(lfc_vector)
```

## Run GSEA

Now for the analysis!

We can use the `GSEA()` function to perform GSEA with any generic set of gene sets, but there are several functions for using specific, commonly used gene sets (e.g., `gseKEGG()`).

```{r run_gsea}
gsea_results <- GSEA(geneList = lfc_vector,  # ordered ranked gene list
                     minGSSize = 25,  # minimum gene set size
                     maxGSSize = 500,  # maximum gene set set
                     pvalueCutoff = 0.05,
                     pAdjustMethod = "BH",  # correction for multiple hypothesis testing
                     TERM2GENE = dplyr::select(hs_hallmarks_df,
                                               gs_name,
                                               ensembl_gene))  # pass the correct identifier column
```
Let's take a look at the GSEA results.

```{r view_gsea, live = TRUE, eval = FALSE}
View(gsea_results@result |>
       dplyr::arrange(dplyr::desc(NES))
)
```

Normalized enrichment scores (NES) are enrichment scores that are scaled to make gene sets that contain different number of genes comparable.

Pathways with significant, highly positive NES are enriched in ERMS myoblasts, whereas pathways with significant, highly negative NES are enriched in ARMS myoblasts.

Let's write these results to file.

```{r write_gsea}
gsea_results@result |> readr::write_tsv(output_file)
```

### Visualizing GSEA results

We can visualize GSEA results for individual pathways or gene sets using `enrichplot::gseaplot()`.
Let's take a look at 3 different pathways -- one with a highly positive NES, one with a highly negative NES, and one that was not a significant result -- to get more insight into how ES are calculated.

#### Highly Positive NES

Let's take look at a pathway with a highly positive NES (`HALLMARK_MYC_TARGETS_V2`) using a GSEA plot.

```{r highly_pos}
enrichplot::gseaplot(gsea_results,
                     geneSetID = "HALLMARK_MYC_TARGETS_V2",
                     title = "HALLMARK_MYC_TARGETS_V2",
                     color.line = "#0066FF")
```

Notice how the genes that are in the gene set, indicated by the black bars, tend to be on the left side of the graph indicating that they have positive gene-level scores.

#### Highly Negative NES

The gene set `HALLMARK_MYOGENESIS` had a highly negative NES.

```{r highly_neg}
enrichplot::gseaplot(gsea_results,
                     geneSetID = "HALLMARK_MYOGENESIS",
                     title = "HALLMARK_MYOGENESIS",
                     color.line = "#0066FF")
```

This gene set shows the opposite pattern -- genes in the pathway tend to be on the right side of the graph.

#### A non-significant result

The `@results` slot will only show gene sets that pass the `pvalueCutoff` threshold we supplied to `GSEA()`, but we can plot any gene set so long as we know its name.
Let's look at `HALLMARK_P53_PATHWAY`, which was not in the results we viewed earlier.

```{r p53, live = TRUE}
enrichplot::gseaplot(gsea_results,
                     geneSetID = "HALLMARK_P53_PATHWAY",
                     title = "HALLMARK_P53_PATHWAY",
                     color.line = "#0066FF")
```

Genes in the pathway are distributed more evenly throughout the ranked list, resulting in a more "middling" score.

*Note: The plots returned by `enrichplot::gseaplot` are ggplots, so we could use `ggplot2::ggsave()` to save them to file if we wanted to.*

## Session Info

```{r session_info}
sessionInfo()
```

    +
    ---
title: "Pathway analysis: Gene Set Enrichment Analysis (GSEA)"
output:
  html_notebook:
    toc: true
    toc_float: true
author: CCDL for ALSF
date: 2024
---

## Objectives

This notebook will demonstrate how to:

- Prepare tabular data of gene-level statistics for use with Gene Set Enrichment Analysis (GSEA)
- Access [Molecular Signatures Database gene set collections](https://www.gsea-msigdb.org/gsea/msigdb/collections.jsp) via the `msigdbr` package
- Perform GSEA with the `clusterProfiler` package
- Visualize GSEA results with the `enrichplot` package

---

In this notebook, we'll analyze the differential expression results from the last notebook.

GSEA is a functional class scoring (FCS) approach to pathway analysis that was first introduced in [Subramanian _et al._ (2005)](https://doi.org/10.1073/pnas.0506580102).
The rationale behind FCS approaches is that small changes in individual genes that participate in the same biological process or pathway can be significant and of biological interest.

There are 3 general steps in FCS methods ([Khatri _et al._ 2012](https://doi.org/10.1371/journal.pcbi.1002375)):

1. Calculate a gene-level statistic (here, we'll use the summary log fold changes in our DESeq2 results)
2. Aggregate gene-level statistics into a pathway-level statistic
3. Assess the statistical significance of the pathway-level statistic

#### Other resources

* For another example using `clusterProfiler` for GSEA, see [_Intro to DGE: Functional Analysis._ from Harvard Chan Bioinformatics Core Training.](https://hbctraining.github.io/Training-modules/DGE-functional-analysis/lessons/02_functional_analysis.html)
* The way we'll use `clusterProfiler` here uses `fgsea` (Fast Gene Set Enrichment Analysis) under the hood.
You can read more about `fgsea` in [Korotkevich _et al._ (2021)](https://doi.org/10.1101/060012).
* See the [refine.bio examples for "Gene set enrichment analysis - RNA-seq"](https://alexslemonade.github.io/refinebio-examples/03-rnaseq/pathway-analysis_rnaseq_02_gsea.html) from which this material has been adapted.

## Set up

### Libraries

```{r setup}
# set seed for reproducibility
set.seed(2025)

# Package to run GSEA
library(clusterProfiler)
# Package that contains the MSigDB gene sets in tidy format
library(msigdbr)
```

### Directories and Files

#### Directories

```{r create_dir}
# We'll use the differential expression results as GSEA input
rms_analysis_dir <- file.path("analysis", "rms")

# We'll create a directory to specifically hold the pathway results if it doesn't
# exist yet
results_dir <- file.path(rms_analysis_dir, "pathway-analysis")
fs::dir_create(results_dir)
```

#### Input files


```{r input_files}
input_file <- file.path(rms_analysis_dir,
                        "deseq",
                        "rms_myoblast_deseq_results.tsv")
```

#### Output files

We'll save our table of GSEA results as a TSV.

```{r output_files}
output_file <- file.path(results_dir,
                         "rms_myoblast_gsea_results.tsv")
```

## Gene sets

We will use gene sets from the [Molecular Signatures Database (MSigDB)](https://www.gsea-msigdb.org/gsea/msigdb/index.jsp) from the Broad Institute ([Subramanian, Tamayo *et al.* 2005](https://doi.org/10.1073/pnas.0506580102)).
The [`msigdbr`](https://cran.r-project.org/web/packages/msigdbr/index.html) package contains MSigDB datasets already in the tidy format required by `clusterProfiler` and supports multiple organisms.

Let's take a look at what organisms the package supports.

```{r show_species}
msigdbr_species()
```

MSigDB contains 8 different gene set collections.

    H: hallmark gene sets
    C1: positional gene sets
    C2: curated gene sets
    C3: motif gene sets
    C4: computational gene sets
    C5: GO gene sets
    C6: oncogenic signatures
    C7: immunologic signatures

We'll use the Hallmark collection for GSEA.
Here's an excerpt of the [collection description](https://www.gsea-msigdb.org/gsea/msigdb/collection_details.jsp#H):

> Hallmark gene sets summarize and represent specific well-defined biological states or processes and display coherent expression. These gene sets were generated by a computational methodology based on identifying gene set overlaps and retaining genes that display coordinate expression. The hallmarks reduce noise and redundancy and provide a better delineated biological space for GSEA.

Notably, there are only 50 gene sets included in this collection.
The fewer gene sets we test, the lower our multiple hypothesis testing burden.

We can retrieve only the Hallmark gene sets by specifying `category = "H"` to the `msigdbr()` function.

```{r immunologic_sets, live = TRUE}
hs_hallmarks_df <- msigdbr(species = "Homo sapiens",
                           category = "H")
```

## Gene Set Enrichment Analysis

_Adapted from [refine.bio examples](https://github.com/AlexsLemonade/refinebio-examples/blob/33cdeff66d57f9fe8ee4fcb5156aea4ac2dce07f/03-rnaseq/pathway-analysis_rnaseq_02_gsea.Rmd)_

![](diagrams/subramanian_fig1.jpg)

**Figure 1. [Subramanian _et al._ (2005)](https://doi.org/10.1073/pnas.0506580102).**

GSEA calculates a pathway-level metric, called an enrichment score (sometimes abbreviated as ES), by ranking genes by a gene-level statistic.
This score reflects whether or not a gene set or pathway is over-represented at the top or bottom of the gene rankings ([Subramanian _et al._ 2005](https://doi.org/10.1073/pnas.0506580102); [Yu](http://yulab-smu.top/clusterProfiler-book/chapter2.html#gene-set-enrichment-analysis))

Specifically, all genes are ranked from most positive to most negative based on their statistic and a running sum is calculated:
Starting with the most highly ranked genes, the running sum increases for each gene in the pathway and decreases for each gene not in the pathway.
The enrichment score for a pathway is the running sum's maximum deviation from zero.
GSEA also assesses statistical significance of the scores for each pathway through permutation testing.
As a result, each input pathway will have a p-value associated with it that is then corrected for multiple hypothesis testing ([Subramanian _et al._ 2005](https://doi.org/10.1073/pnas.0506580102); [Yu](http://yulab-smu.top/clusterProfiler-book/chapter2.html#gene-set-enrichment-analysis)).

The implementation of GSEA we use here requires a gene list ordered by some statistic and input gene sets.
When you use previously computed gene-level statistics with GSEA, it is called GSEA pre-ranked.

## DESeq2 results

```{r read_in_markers, live = TRUE}
deseq_df <- readr::read_tsv(input_file)
```

```{r deseq_head}
head(deseq_df)
```

This data frame uses Ensembl gene identifiers.
We'll need to make sure our gene sets use the same identifiers.
Let's take a look at the first few rows of the data frame that contains the hallmark gene sets.

```{r hallmark_head, live = TRUE}
head(hs_hallmarks_df)
```

We can see that the gene sets from `msigdbr` have Ensembl gene identifiers associated with them, so we don't need to do any conversion.
However, we'll need to pass the correct column to the function that runs GSEA.

If we needed to do gene identifier conversion, we would likely use the `AnnotationDbi` package.
You can see an example in this Harvard Chan Bioinformatics Core Training material: <https://hbctraining.github.io/DGE_workshop_salmon_online/lessons/AnnotationDbi_lesson.html>

One good thing about Ensembl gene identifiers is that they are less likely to be duplicated than, for example, gene symbols.
(Multiple Ensembl gene identifiers can map to the same symbol.)

The GSEA approach requires discriminating between genes that are in a gene set and those that are not.
Practically speaking, gene sets are just collections of gene identifiers!
When the function we use for GSEA pre-ranked gets a list with duplicated gene identifiers, it can produce unexpected results.
So, let's check for duplicates in the data frame of DESeq2 results.

```{r check_duplicates, live = TRUE}
any(duplicated(deseq_df$ensembl_id))
```

There are no duplicates for us to worry about!

### Pre-ranked list

The `GSEA()` function takes a pre-ranked (sorted) named vector of statistics, where the names in the vector are gene identifiers.
This is step 1 -- gene-level statistics.

```{r lfc_vector}
lfc_vector <- deseq_df |>
  # Extract a vector of `log2FoldChange` named by `ensembl_id`
  dplyr::pull(log2FoldChange, name = ensembl_id)
lfc_vector <- sort(lfc_vector, decreasing = TRUE)
```

Let's look at the top ranked values.

```{r head_lfc, live = TRUE}
# Look at first entries of the log fold change vector
head(lfc_vector)
```

And the bottom of the list.

```{r tail_lfc, live = TRUE}
# Look at the last entries of the log fold change vector
tail(lfc_vector)
```

## Run GSEA

Now for the analysis!

We can use the `GSEA()` function to perform GSEA with any generic set of gene sets, but there are several functions for using specific, commonly used gene sets (e.g., `gseKEGG()`).

```{r run_gsea}
gsea_results <- GSEA(geneList = lfc_vector,  # ordered ranked gene list
                     minGSSize = 25,  # minimum gene set size
                     maxGSSize = 500,  # maximum gene set set
                     pvalueCutoff = 0.05,
                     pAdjustMethod = "BH",  # correction for multiple hypothesis testing
                     TERM2GENE = dplyr::select(hs_hallmarks_df,
                                               gs_name,
                                               ensembl_gene))  # pass the correct identifier column
```
Let's take a look at the GSEA results.

```{r view_gsea, live = TRUE, eval = FALSE}
View(
  gsea_results@result |>
    dplyr::arrange(dplyr::desc(NES))
)
```

Normalized enrichment scores (NES) are enrichment scores that are scaled to make gene sets that contain different number of genes comparable.

Pathways with significant, highly positive NES are enriched in ERMS myoblasts, whereas pathways with significant, highly negative NES are enriched in ARMS myoblasts.

Let's write these results to a file.

```{r write_gsea}
gsea_results@result |> 
  readr::write_tsv(output_file)
```

### Visualizing GSEA results

We can visualize GSEA results for individual pathways or gene sets using `enrichplot::gseaplot()`.
Let's take a look at 3 different pathways -- one with a highly positive NES, one with a highly negative NES, and one that was not a significant result -- to get more insight into how ES are calculated.

#### Highly Positive NES

Let's take look at a pathway with a highly positive NES (`HALLMARK_MYC_TARGETS_V2`) using a GSEA plot.

```{r highly_pos}
enrichplot::gseaplot(gsea_results,
                     geneSetID = "HALLMARK_MYC_TARGETS_V2",
                     title = "HALLMARK_MYC_TARGETS_V2",
                     color.line = "#0066FF")
```

Notice how the genes that are in the gene set, indicated by the black bars, tend to be on the left side of the graph indicating that they have positive gene-level scores.

#### Highly Negative NES

The gene set `HALLMARK_MYOGENESIS` had a highly negative NES.

```{r highly_neg}
enrichplot::gseaplot(gsea_results,
                     geneSetID = "HALLMARK_MYOGENESIS",
                     title = "HALLMARK_MYOGENESIS",
                     color.line = "#0066FF")
```

This gene set shows the opposite pattern -- genes in the pathway tend to be on the right side of the graph.

#### A non-significant result

The `@results` slot will only show gene sets that pass the `pvalueCutoff` threshold we supplied to `GSEA()`, but we can plot any gene set so long as we know its name.
Let's look at `HALLMARK_P53_PATHWAY`, which was not in the results we viewed earlier.

```{r p53, live = TRUE}
enrichplot::gseaplot(gsea_results,
                     geneSetID = "HALLMARK_P53_PATHWAY",
                     title = "HALLMARK_P53_PATHWAY",
                     color.line = "#0066FF")
```

Genes in the pathway are distributed more evenly throughout the ranked list, resulting in a more "middling" score.

*Note: The plots returned by `enrichplot::gseaplot` are ggplots, so we could use `ggplot2::ggsave()` to save them to file if we wanted to.*

## Session Info

```{r session_info}
sessionInfo()
```

    diff --git a/scRNA-seq-advanced/05-aucell-live.Rmd b/scRNA-seq-advanced/05-aucell-live.Rmd index 67e7a408..f6280b01 100644 --- a/scRNA-seq-advanced/05-aucell-live.Rmd +++ b/scRNA-seq-advanced/05-aucell-live.Rmd @@ -18,21 +18,21 @@ date: 2024 --- -In this notebook, we'll demonstrate how to use the AUCell method, introduced in [Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463). +In this notebook, we'll demonstrate how to use the `AUCell` method, introduced in [Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463). -We can use AUCell when we are interested in a gene set's relative expression or activity in an individual cell. +We can use `AUCell` when we are interested in a gene set's relative expression or activity in an individual cell. Gene sets can come from a curated collection of prior knowledge, like the Hallmark collection we used in the last notebook, or we can use our own custom gene sets (e.g., a set of marker genes for a cell type of interest). -A nice feature of AUCell is that it is based on ranking genes from highest to lowest expression value in an individual cell, which is helpful in the following ways ([AUCell vignette](https://bioconductor.org/packages/release/bioc/vignettes/AUCell/inst/doc/AUCell.html)): +A nice feature of `AUCell` is that it is based on ranking genes from highest to lowest expression value in an individual cell, which is helpful in the following ways ([`AUCell` vignette](https://bioconductor.org/packages/release/bioc/vignettes/AUCell/inst/doc/AUCell.html)): - It can take a number of different values as input (e.g., raw counts, TPM) - It compensates for differences in library size, where something like averaging raw count values of genes in a gene set would not - It scales to larger datasets, since creating rankings is not as resource-intensive as something like permutation testing, and we could split up the object into subsets of cells if needed -AUCell calculates the area under the recovery curve (AUC), which "represents the proportion of expressed genes in the signature and their relative expression value compared to the other genes within the cell" ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)). +`AUCell` calculates the area under the recovery curve (AUC), which "represents the proportion of expressed genes in the signature and their relative expression value compared to the other genes within the cell" ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)). We will visualize some recovery curves in the notebook to give you a better intuition about the AUC and its meaning. -The AUC values we get out of AUCell can be used in a number of ways ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)): +The AUC values we get out of `AUCell` can be used in a number of ways ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)): - As continuous values we can use for visualization or clustering - For binary assignment (i.e., "on" and "off" or "expressed" and "not expressed") if we pick a threshold either automatically using built-in functionality or manually by inspecting the distribution of scores ourselves @@ -82,7 +82,7 @@ sce_file <- fs::path(processed_dir, "SCPCL000822_processed.rds") ``` -We will save the AUCell results as a table in the analysis directory. +We will save the `AUCell` results as a table in the analysis directory. ```{r setup_output_files, live = TRUE} @@ -98,7 +98,7 @@ source(fs::path("util", "aucell_functions.R")) ``` This loads one custom function, called `plot_recovery_curve()`, into our environment. -This function is adapted from [the AUCell vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd#L295-L316). +This function is adapted from [the `AUCell` vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd#L295-L316). ## Set up gene sets @@ -146,21 +146,33 @@ ewing_gene_set_collection <- ewing_gene_set_names |> GeneSetCollection() ``` -## Read in and prepare SingleCellExperiment +## Read in and prepare the `SingleCellExperiment` ```{r read_in_sce, live = TRUE} ``` +Our object includes counts for all genes that were present in the index when quantifying gene expression. +There are a number of genes that are present in the object but not detected in any of the cells. +We don't want genes that are not found in our data set to impact our rankings, so let's remove them. + +```{r, filter_sce} +# remove genes that are not detected in any of the cells from the SCE object +genes_to_keep <- rowData(sce)$detected > 0 +sce <- sce[genes_to_keep, ] +``` + + The `AUCell` functions takes an expression matrix with genes as rows and cells as column. We can extract a counts matrix in sparse format for use with `AUCell`. + ```{r counts_matrix} # Extract counts matrix counts_matrix <- counts(sce) ``` -There may be genes in our gene set that do not appear in the SingleCellExperiment object. +There may be genes in our gene set that do not appear in the `SingleCellExperiment` object. We can remove them using the `subsetGeneSets()` function. ```{r subset_gene_sets, live = TRUE} @@ -168,9 +180,9 @@ We can remove them using the `subsetGeneSets()` function. ``` -## AUCell +## `AUCell` -AUCell relies on ranking genes from highest to lowest expression value to calculate the AUC. +`AUCell` relies on ranking genes from highest to lowest expression value to calculate the AUC. The AUC is the area under the recovery curve, which captures the number of genes in a gene set that are present in the rankings above some threshold (i.e., it is the area under the curve to the left of this gene rank). By default, the top 5% of genes are used as the threshold. @@ -185,16 +197,16 @@ To make our rankings – and therefore results – reproducible, we will set a s ### Cell ranking -The first step in AUCell is to rank genes for each cell from highest to lowest expression value. -We can do this using the `AUCell_buildRankings()` function, which will output a visualization showing the distribution of the number of genes detected in the cells in our SingleCellExperiment object. +The first step in `AUCell` is to rank genes for each cell from highest to lowest expression value. +We can do this using the `AUCell_buildRankings()` function, which will output a visualization showing the distribution of the number of genes detected in the cells in our `SingleCellExperiment` object. ```{r cell_rankings, live = TRUE} ``` -The AUCell authors recommend making sure most cells have at least the number of genes we will use as the max rank to calculate the AUC. +The `AUCell` authors recommend making sure most cells have at least the number of genes we will use as the max rank to calculate the AUC. -The AUC max rank value tells AUCell the cutoff in the gene rankings to use for calculating AUC; we will visualize this curve and max rank in just a moment. +The AUC max rank value tells `AUCell` the cutoff in the gene rankings to use for calculating AUC; we will visualize this curve and max rank in just a moment. If we picked a max rank higher than the number of genes detected in most cells, the non-detected genes that are randomly ordered would play an outsized role in our AUC values. By default, the max rank is the top 5% highest expressed genes. @@ -222,10 +234,10 @@ We can use a function called `ceiling()` to round this and save it to a variable ### Plotting AUC -The AUC values we get out of AUCell are the area under a recovery curve and estimate the proportion of genes in the gene set that are highly expressed (i.e., highly ranked). +The AUC values we get out of `AUCell` are the area under a recovery curve and estimate the proportion of genes in the gene set that are highly expressed (i.e., highly ranked). Let's plot the recovery curve for a cell with high AUC and a cell with low AUC to get a better intuition about AUC values. -Earlier, we loaded a custom function we adapted from [the AUCell vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd) called `plot_recovery_curve()` with `source()`. +Earlier, we loaded a custom function we adapted from [the `AUCell` vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd) called `plot_recovery_curve()` with `source()`. First, we'll start with a cell with a high AUC. We picked this barcode ahead of time when we wrote the notebook. @@ -286,7 +298,7 @@ head(auc_df) ### Assignments -AUCell can assign cells as having an active gene set or not by picking a threshold automatically. +`AUCell` can assign cells as having an active gene set or not by picking a threshold automatically. We'll explore these in a later plot, but for now, let's calculate the threshold and assign cells with `AUCell_exploreThresholds()`. ```{r auc_assignments, live = TRUE} @@ -367,7 +379,7 @@ auc_plotting_df |> #### Adding AUC to `colData` -We can also add the AUC values back into the SingleCellExperiment for convenience, e.g., for plotting. +We can also add the AUC values back into the `SingleCellExperiment` for convenience, e.g., for plotting. We'll add it to the existing `colData`. First, let's rename the gene set columns to something more easily typed. diff --git a/scRNA-seq-advanced/05-aucell.Rmd b/scRNA-seq-advanced/05-aucell.Rmd index f74fc0fd..55d3891b 100644 --- a/scRNA-seq-advanced/05-aucell.Rmd +++ b/scRNA-seq-advanced/05-aucell.Rmd @@ -18,21 +18,21 @@ date: 2024 --- -In this notebook, we'll demonstrate how to use the AUCell method, introduced in [Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463). +In this notebook, we'll demonstrate how to use the `AUCell` method, introduced in [Aibar _et al_. 2017](https://doi.org/10.1038/nmeth.4463). -We can use AUCell when we are interested in a gene set's relative expression or activity in an individual cell. +We can use `AUCell` when we are interested in a gene set's relative expression or activity in an individual cell. Gene sets can come from a curated collection of prior knowledge, like the Hallmark collection we used in the last notebook, or we can use our own custom gene sets (e.g., a set of marker genes for a cell type of interest). -A nice feature of AUCell is that it is based on ranking genes from highest to lowest expression value in an individual cell, which is helpful in the following ways ([AUCell vignette](https://bioconductor.org/packages/release/bioc/vignettes/AUCell/inst/doc/AUCell.html)): +A nice feature of `AUCell` is that it is based on ranking genes from highest to lowest expression value in an individual cell, which is helpful in the following ways ([`AUCell` vignette](https://bioconductor.org/packages/release/bioc/vignettes/AUCell/inst/doc/AUCell.html)): - It can take a number of different values as input (e.g., raw counts, TPM) - It compensates for differences in library size, where something like averaging raw count values of genes in a gene set would not - It scales to larger datasets, since creating rankings is not as resource-intensive as something like permutation testing, and we could split up the object into subsets of cells if needed -AUCell calculates the area under the recovery curve (AUC), which "represents the proportion of expressed genes in the signature and their relative expression value compared to the other genes within the cell" ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)). +`AUCell` calculates the area under the recovery curve (AUC), which "represents the proportion of expressed genes in the signature and their relative expression value compared to the other genes within the cell" ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)). We will visualize some recovery curves in the notebook to give you a better intuition about the AUC and its meaning. -The AUC values we get out of AUCell can be used in a number of ways ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)): +The AUC values we get out of `AUCell` can be used in a number of ways ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)): - As continuous values we can use for visualization or clustering - For binary assignment (i.e., "on" and "off" or "expressed" and "not expressed") if we pick a threshold either automatically using built-in functionality or manually by inspecting the distribution of scores ourselves @@ -77,16 +77,20 @@ fs::dir_create(analysis_dir) The input will be a `SingleCellExperiment` for an individual Ewing sarcoma library. ```{r setup_input_files} -sce_file <- fs::path(processed_dir, - "SCPCS000490", - "SCPCL000822_processed.rds") +sce_file <- fs::path( + processed_dir, + "SCPCS000490", + "SCPCL000822_processed.rds" +) ``` -We will save the AUCell results as a table in the analysis directory. +We will save the `AUCell` results as a table in the analysis directory. ```{r setup_output_files, live = TRUE} -output_file <- fs::path(analysis_dir, - "ewing_sarcoma_aucell_results.tsv") +output_file <- fs::path( + analysis_dir, + "ewing_sarcoma_aucell_results.tsv" +) ``` @@ -99,7 +103,7 @@ source(fs::path("util", "aucell_functions.R")) ``` This loads one custom function, called `plot_recovery_curve()`, into our environment. -This function is adapted from [the AUCell vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd#L295-L316). +This function is adapted from [the `AUCell` vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd#L295-L316). ## Set up gene sets @@ -112,8 +116,10 @@ We would expect both of these gene sets to have high expression in tumor cells. ```{r genesets} # Create a named vector with the relevant gene set names -ewing_gene_set_names <- c(zhang = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION", - riggi = "RIGGI_EWING_SARCOMA_PROGENITOR_UP") +ewing_gene_set_names <- c( + zhang = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION", + riggi = "RIGGI_EWING_SARCOMA_PROGENITOR_UP" +) ewing_gene_set_names ``` @@ -122,60 +128,68 @@ These gene sets come from the C2 gene set collection from MSigDB. Let's retrieve them using `msigdbr()`. ```{r extract_genesets, live = TRUE} -ewing_gene_sets_df <- msigdbr(species = "Homo sapiens", - category = "C2", - subcategory = "CGP") |> - dplyr::filter(gs_name %in% ewing_gene_set_names) +ewing_gene_sets_df <- msigdbr( + species = "Homo sapiens", + collection = "C2", + subcollection = "CGP" +) |> + dplyr::filter(gs_name %in% ewing_gene_set_names) |> + dplyr::select(gs_name, ensembl_gene) ``` -`AUCell` uses gene sets in a particular format that comes from the `GSEABase` package. -We need to create a `GeneSetCollection`. +We can provide these gene sets as a named list to `AUCell`, so let's create that now. -```{r gene_set_collection} -ewing_gene_set_collection <- ewing_gene_set_names |> - purrr::map( - # For each gene set - \(gene_set_name) { - ewing_gene_sets_df |> - # Subset to the rows in that gene set - dplyr::filter(gs_name == gene_set_name) |> - # Grab the Ensembl gene identifiers - dplyr::pull(ensembl_gene) |> - # Create a GeneSet object - GeneSet(setName = gene_set_name, - geneIdType = ENSEMBLIdentifier()) - } - ) |> - # Turn the list of GeneSet objects into a GeneSet collection - GeneSetCollection() +```{r create_gene_list} +ewing_gene_set_list <- split( + ewing_gene_sets_df$ensembl_gene, + ewing_gene_sets_df$gs_name +) + +# briefly, show the resulting list +purrr::map(ewing_gene_set_list, head) ``` -## Read in and prepare SingleCellExperiment +## Read in and prepare the `SingleCellExperiment` ```{r read_in_sce, live = TRUE} sce <- readr::read_rds(sce_file) ``` +Our object includes counts for all genes that were present in the index when quantifying gene expression. +There are a number of genes that are present in the object but not detected in any of the cells. +We don't want genes that are not found in our data set to impact our rankings, so let's remove them. + +```{r, filter_sce} +# remove genes that are not detected in any of the cells from the SCE object +genes_to_keep <- rowData(sce)$detected > 0 +sce <- sce[genes_to_keep, ] +``` + + The `AUCell` functions takes an expression matrix with genes as rows and cells as column. We can extract a counts matrix in sparse format for use with `AUCell`. + ```{r counts_matrix} # Extract counts matrix counts_matrix <- counts(sce) ``` -There may be genes in our gene set that do not appear in the SingleCellExperiment object. -We can remove them using the `subsetGeneSets()` function. +There may be genes in our gene set that do not appear in the `SingleCellExperiment` object which we should remove before proceeding. ```{r subset_gene_sets, live = TRUE} -# Remove genes from gene sets if they are not in the SCE -ewing_gene_set_collection <- subsetGeneSets(ewing_gene_set_collection, - rownames(counts_matrix)) +# Remove genes from list if they are not in the SCE +ewing_gene_set_list <- ewing_gene_set_list |> + purrr::map( + \(gene_list) { + intersect(gene_list, rownames(sce)) # keep only the intersection with our genes + } +) ``` -## AUCell +## `AUCell` -AUCell relies on ranking genes from highest to lowest expression value to calculate the AUC. +`AUCell` relies on ranking genes from highest to lowest expression value to calculate the AUC. The AUC is the area under the recovery curve, which captures the number of genes in a gene set that are present in the rankings above some threshold (i.e., it is the area under the curve to the left of this gene rank). By default, the top 5% of genes are used as the threshold. @@ -190,16 +204,16 @@ set.seed(2024) ### Cell ranking -The first step in AUCell is to rank genes for each cell from highest to lowest expression value. -We can do this using the `AUCell_buildRankings()` function, which will output a visualization showing the distribution of the number of genes detected in the cells in our SingleCellExperiment object. +The first step in `AUCell` is to rank genes for each cell from highest to lowest expression value. +We can do this using the `AUCell_buildRankings()` function, which will output a visualization showing the distribution of the number of genes detected in the cells in our `SingleCellExperiment` object. ```{r cell_rankings, live = TRUE} cell_rankings <- AUCell_buildRankings(counts_matrix) ``` -The AUCell authors recommend making sure most cells have at least the number of genes we will use as the max rank to calculate the AUC. +The `AUCell` authors recommend making sure most cells have at least the number of genes we will use as the max rank to calculate the AUC. -The AUC max rank value tells AUCell the cutoff in the gene rankings to use for calculating AUC; we will visualize this curve and max rank in just a moment. +The AUC max rank value tells `AUCell` the cutoff in the gene rankings to use for calculating AUC; we will visualize this curve and max rank in just a moment. If we picked a max rank higher than the number of genes detected in most cells, the non-detected genes that are randomly ordered would play an outsized role in our AUC values. By default, the max rank is the top 5% highest expressed genes. @@ -227,20 +241,22 @@ auc_max_rank <- ceiling(nrow(cell_rankings) * 0.01) ### Plotting AUC -The AUC values we get out of AUCell are the area under a recovery curve and estimate the proportion of genes in the gene set that are highly expressed (i.e., highly ranked). +The AUC values we get out of `AUCell` are the area under a recovery curve and estimate the proportion of genes in the gene set that are highly expressed (i.e., highly ranked). Let's plot the recovery curve for a cell with high AUC and a cell with low AUC to get a better intuition about AUC values. -Earlier, we loaded a custom function we adapted from [the AUCell vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd) called `plot_recovery_curve()` with `source()`. +Earlier, we loaded a custom function we adapted from [the `AUCell` vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd) called `plot_recovery_curve()` with `source()`. First, we'll start with a cell with a high AUC. We picked this barcode ahead of time when we wrote the notebook. ```{r high_recovery_curve} -plot_recovery_curve(cell_rankings, - ewing_gene_set_collection, - gene_set_name = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION", - barcode = "CTGAGCGGTCTTTATC", - auc_max_rank = auc_max_rank) # 1% threshold +plot_recovery_curve( + cell_rankings, + ewing_gene_set_list, + gene_set_name = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION", + barcode = "CTGAGCGGTCTTTATC", + auc_max_rank = auc_max_rank # 1% threshold +) ``` The x-axis is the gene ranks for all genes. @@ -250,11 +266,13 @@ The AUC is the area under this recovery curve at the max rank threshold chosen f Now, let's look at an example with a low AUC. ```{r low_recovery_curve} -plot_recovery_curve(cell_rankings, - ewing_gene_set_collection, - gene_set_name = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION", - barcode = "AGATAGAGTCACAATC", - auc_max_rank = auc_max_rank) # 1% threshold +plot_recovery_curve( + cell_rankings, + ewing_gene_set_list, + gene_set_name = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION", + barcode = "AGATAGAGTCACAATC", + auc_max_rank = auc_max_rank # 1% threshold +) ``` Far fewer genes in the gene set are ranked above the threshold, yielding a lower AUC value. @@ -264,9 +282,11 @@ Far fewer genes in the gene set are ranked above the threshold, yielding a lower Once we have the rankings, we can calculate the AUC scores for both gene sets in all cells with the `AUCell_calcAUC()` function. ```{r calc_auc, live = TRUE} -cell_auc <- AUCell_calcAUC(geneSets = ewing_gene_set_collection, - rankings = cell_rankings, - aucMaxRank = auc_max_rank) +cell_auc <- AUCell_calcAUC( + geneSets = ewing_gene_set_list, + rankings = cell_rankings, + aucMaxRank = auc_max_rank +) ``` This function returns an `aucellResults` object. @@ -293,22 +313,26 @@ head(auc_df) ### Assignments -AUCell can assign cells as having an active gene set or not by picking a threshold automatically. +`AUCell` can assign cells as having an active gene set or not by picking a threshold automatically. We'll explore these in a later plot, but for now, let's calculate the threshold and assign cells with `AUCell_exploreThresholds()`. ```{r auc_assignments, live = TRUE} -auc_assignments <- AUCell_exploreThresholds(cell_auc, - plotHist = FALSE, - assignCells = TRUE) +auc_assignments <- AUCell_exploreThresholds( + cell_auc, + plotHist = FALSE, + assignCells = TRUE +) ``` We're going to plot the distribution of AUC values with `ggplot2`, so we will want the AUC values in a longer format. ```{r auc_plotting_df} auc_plotting_df <- auc_df |> - tidyr::pivot_longer(!barcodes, - names_to = "gene_set", - values_to = "auc") |> + tidyr::pivot_longer( + !barcodes, + names_to = "gene_set", + values_to = "auc" + ) |> dplyr::mutate( # Create a new logical column called assigned assigned = dplyr::case_when( @@ -376,7 +400,7 @@ auc_plotting_df |> #### Adding AUC to `colData` -We can also add the AUC values back into the SingleCellExperiment for convenience, e.g., for plotting. +We can also add the AUC values back into the `SingleCellExperiment` for convenience, e.g., for plotting. We'll add it to the existing `colData`. First, let's rename the gene set columns to something more easily typed. @@ -384,9 +408,10 @@ First, let's rename the gene set columns to something more easily typed. ```{r rename_gene_set} auc_df <- auc_df |> # Use shorter names - dplyr::rename(zhang_auc = ewing_gene_set_names[["zhang"]], - riggi_auc = ewing_gene_set_names[["riggi"]]) - + dplyr::rename( + zhang_auc = ewing_gene_set_names[["zhang"]], + riggi_auc = ewing_gene_set_names[["riggi"]] + ) ``` And join it to the existing `colData`. @@ -419,18 +444,26 @@ We can use the `plotUMAP()` function from the `scater` package to plot a UMAP wi ```{r plot_umap_zhang} scater::plotUMAP(sce, colour_by = "zhang_auc") + # Use the gene set name, replacing underscores with spaces - ggplot2::ggtitle(stringr::str_replace_all(ewing_gene_set_names[["zhang"]], - "\\_", - " ")) + ggplot2::ggtitle( + stringr::str_replace_all( + ewing_gene_set_names[["zhang"]], + "\\_", + " " + ) + ) ``` Let's color the points by the AUC values for the other gene set. ```{r plot_umap_riggi, live = TRUE} scater::plotUMAP(sce, colour_by = "riggi_auc") + - ggplot2::ggtitle(stringr::str_replace_all(ewing_gene_set_names[["riggi"]], - "\\_", - " ")) + ggplot2::ggtitle( + stringr::str_replace_all( + ewing_gene_set_names[["riggi"]], + "\\_", + " " + ) + ) ``` We would want to do something more formal to confirm, but it seems like the same cells have high AUC values for both gene sets! diff --git a/scRNA-seq-advanced/05-aucell.nb.html b/scRNA-seq-advanced/05-aucell.nb.html index b10258db..7300e6c4 100644 --- a/scRNA-seq-advanced/05-aucell.nb.html +++ b/scRNA-seq-advanced/05-aucell.nb.html @@ -2898,17 +2898,17 @@

    Objectives

    plotting
    -

    In this notebook, we’ll demonstrate how to use the AUCell method, -introduced in Aibar et -al. 2017..

    -

    We can use AUCell when we are interested in a gene set’s relative -expression or activity in an individual cell. Gene sets can come from a -curated collection of prior knowledge, like the Hallmark collection we -used in the last notebook, or we can use our own custom gene sets (e.g., -a set of marker genes for a cell type of interest).

    -

    A nice feature of AUCell is that it is based on ranking genes from -highest to lowest expression value in an individual cell, which is -helpful in the following ways (AUCell +

    In this notebook, we’ll demonstrate how to use the +AUCell method, introduced in Aibar et al. +2017..

    +

    We can use AUCell when we are interested in a gene set’s +relative expression or activity in an individual cell. Gene sets can +come from a curated collection of prior knowledge, like the Hallmark +collection we used in the last notebook, or we can use our own custom +gene sets (e.g., a set of marker genes for a cell type of interest).

    +

    A nice feature of AUCell is that it is based on ranking +genes from highest to lowest expression value in an individual cell, +which is helpful in the following ways (AUCell vignette):

    • It can take a number of different values as input (e.g., raw counts, @@ -2919,15 +2919,15 @@

      Objectives

      resource-intensive as something like permutation testing, and we could split up the object into subsets of cells if needed
    -

    AUCell calculates the area under the recovery curve (AUC), which -“represents the proportion of expressed genes in the signature and their -relative expression value compared to the other genes within the cell” -(Aibar et al. +

    AUCell calculates the area under the recovery curve +(AUC), which “represents the proportion of expressed genes in the +signature and their relative expression value compared to the other +genes within the cell” (Aibar et al. 2017.). We will visualize some recovery curves in the notebook to give you a better intuition about the AUC and its meaning.

    -

    The AUC values we get out of AUCell can be used in a number of ways -(Aibar et al. -2017.):

    +

    The AUC values we get out of AUCell can be used in a +number of ways (Aibar +et al. 2017.):

    • As continuous values we can use for visualization or clustering
    • For binary assignment (i.e., “on” and “off” or “expressed” and “not @@ -3022,8 +3022,8 @@

      Files

      -

      We will save the AUCell results as a table in the analysis -directory.

      +

      We will save the AUCell results as a table in the +analysis directory.

      @@ -3048,7 +3048,7 @@

      Functions

      This loads one custom function, called plot_recovery_curve(), into our environment. This function is adapted from the -AUCell vignette.

      +AUCell vignette.

    @@ -3121,8 +3121,8 @@

    Set up gene sets

    -
    -

    Read in and prepare SingleCellExperiment

    +
    +

    Read in and prepare the SingleCellExperiment

    @@ -3130,6 +3130,20 @@

    Read in and prepare SingleCellExperiment

    +

    Our object includes counts for all genes that were present in the +index when quantifying gene expression. There are a number of genes that +are present in the object but not detected in any of the cells. We don’t +want genes that are not found in our data set to impact our rankings, so +let’s remove them.

    + + + +
    # remove genes that are not detected in any of the cells from the SCE object
    +genes_to_keep <- rowData(sce)$detected > 0
    +sce <- sce[genes_to_keep, ]
    + + +

    The AUCell functions takes an expression matrix with genes as rows and cells as column. We can extract a counts matrix in sparse format for use with AUCell.

    @@ -3142,7 +3156,7 @@

    Read in and prepare SingleCellExperiment

    There may be genes in our gene set that do not appear in the -SingleCellExperiment object. We can remove them using the +SingleCellExperiment object. We can remove them using the subsetGeneSets() function.

    @@ -3155,13 +3169,13 @@

    Read in and prepare SingleCellExperiment

    -

    AUCell

    -

    AUCell relies on ranking genes from highest to lowest expression -value to calculate the AUC. The AUC is the area under the recovery -curve, which captures the number of genes in a gene set that are present -in the rankings above some threshold (i.e., it is the area under the -curve to the left of this gene rank). By default, the top 5% of genes -are used as the threshold.

    +

    AUCell

    +

    AUCell relies on ranking genes from highest to lowest +expression value to calculate the AUC. The AUC is the area under the +recovery curve, which captures the number of genes in a gene set that +are present in the rankings above some threshold (i.e., it is the area +under the curve to the left of this gene rank). By default, the top 5% +of genes are used as the threshold.

    Some genes will not be detected (i.e., have 0 counts). Genes can also have the same expression level (i.e., ties). These undetected genes and ties will be randomly ordered in our ranking. To make our rankings – and @@ -3175,11 +3189,11 @@

    AUCell

    Cell ranking

    -

    The first step in AUCell is to rank genes for each cell from highest -to lowest expression value. We can do this using the +

    The first step in AUCell is to rank genes for each cell +from highest to lowest expression value. We can do this using the AUCell_buildRankings() function, which will output a visualization showing the distribution of the number of genes detected -in the cells in our SingleCellExperiment object.

    +in the cells in our SingleCellExperiment object.

    @@ -3198,13 +3212,14 @@

    Cell ranking

    -

    The AUCell authors recommend making sure most cells have at least the -number of genes we will use as the max rank to calculate the AUC.

    -

    The AUC max rank value tells AUCell the cutoff in the gene rankings -to use for calculating AUC; we will visualize this curve and max rank in -just a moment. If we picked a max rank higher than the number of genes -detected in most cells, the non-detected genes that are randomly ordered -would play an outsized role in our AUC values.

    +

    The AUCell authors recommend making sure most cells have +at least the number of genes we will use as the max rank to calculate +the AUC.

    +

    The AUC max rank value tells AUCell the cutoff in the +gene rankings to use for calculating AUC; we will visualize this curve +and max rank in just a moment. If we picked a max rank higher than the +number of genes detected in most cells, the non-detected genes that are +randomly ordered would play an outsized role in our AUC values.

    By default, the max rank is the top 5% highest expressed genes. We can calculate the default max rank by taking into account the number of genes.

    @@ -3213,8 +3228,8 @@

    Cell ranking

    nrow(cell_rankings) * 0.05
    - -
    [1] 3015.95
    + +
    [1] 1745.7
    @@ -3227,8 +3242,8 @@

    Cell ranking

    nrow(cell_rankings) * 0.01
    - -
    [1] 603.19
    + +
    [1] 349.14
    @@ -3245,14 +3260,14 @@

    Cell ranking

    Plotting AUC

    -

    The AUC values we get out of AUCell are the area under a recovery -curve and estimate the proportion of genes in the gene set that are -highly expressed (i.e., highly ranked).

    +

    The AUC values we get out of AUCell are the area under a +recovery curve and estimate the proportion of genes in the gene set that +are highly expressed (i.e., highly ranked).

    Let’s plot the recovery curve for a cell with high AUC and a cell with low AUC to get a better intuition about AUC values. Earlier, we loaded a custom function we adapted from the -AUCell vignette called plot_recovery_curve() with -source().

    +AUCell vignette called +plot_recovery_curve() with source().

    First, we’ll start with a cell with a high AUC. We picked this barcode ahead of time when we wrote the notebook.

    @@ -3265,7 +3280,7 @@

    Plotting AUC

    auc_max_rank = auc_max_rank) # 1% threshold
    -

    +

    @@ -3285,7 +3300,7 @@

    Plotting AUC

    auc_max_rank = auc_max_rank) # 1% threshold
    -

    +

    @@ -3312,7 +3327,7 @@

    Calculating the AUC

    str(cell_auc)
    - +
    Formal class 'aucellResults' [package "AUCell"] with 6 slots
       ..@ nGenesDetected : num(0) 
       ..@ colData        :Formal class 'DFrame' [package "S4Vectors"] with 6 slots
    @@ -3325,7 +3340,7 @@ 

    Calculating the AUC

    ..@ assays :Formal class 'SimpleAssays' [package "SummarizedExperiment"] with 1 slot .. .. ..@ data:Formal class 'SimpleList' [package "S4Vectors"] with 4 slots .. .. .. .. ..@ listData :List of 1 - .. .. .. .. .. ..$ AUC: num [1:2, 1:4277] 0.1103 0.1206 0.0352 0.0538 0.1671 ... + .. .. .. .. .. ..$ AUC: num [1:2, 1:4277] 0.0912 0.1544 0.0179 0.051 0.1426 ... .. .. .. .. .. .. ..- attr(*, "dimnames")=List of 2 .. .. .. .. .. .. .. ..$ gene sets: chr [1:2] "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION" "RIGGI_EWING_SARCOMA_PROGENITOR_UP" .. .. .. .. .. .. .. ..$ cells : chr [1:4277] "AAGCATCTCGTTGCCT" "CTGTATTTCCAAGAGG" "CAGCAGCTCCTCAGAA" "AGGTCTAAGGGACTGT" ... @@ -3363,7 +3378,7 @@

    Calculating the AUC

    @@ -3371,9 +3386,9 @@

    Calculating the AUC

    Assignments

    -

    AUCell can assign cells as having an active gene set or not by -picking a threshold automatically. We’ll explore these in a later plot, -but for now, let’s calculate the threshold and assign cells with +

    AUCell can assign cells as having an active gene set or +not by picking a threshold automatically. We’ll explore these in a later +plot, but for now, let’s calculate the threshold and assign cells with AUCell_exploreThresholds().

    @@ -3414,7 +3429,7 @@

    Assignments

    @@ -3435,7 +3450,7 @@

    Assignments

    @@ -3465,7 +3480,7 @@

    Assignments

    ggplot2::theme_bw()
    -

    +

    @@ -3486,9 +3501,9 @@

    Assignments

    UMAPs

    Adding AUC to colData

    -

    We can also add the AUC values back into the SingleCellExperiment for -convenience, e.g., for plotting. We’ll add it to the existing -colData.

    +

    We can also add the AUC values back into the +SingleCellExperiment for convenience, e.g., for plotting. +We’ll add it to the existing colData.

    First, let’s rename the gene set columns to something more easily typed.

    @@ -3544,7 +3559,7 @@

    Plotting UMAPs

    " ")) -

    +

    @@ -3558,7 +3573,7 @@

    Plotting UMAPs

    " ")) -

    +

    @@ -3665,7 +3680,7 @@

    Session Info

    -
    ---
title: "Pathway Analysis: AUCell"
output:
  html_notebook:
    toc: true
    toc_float: true
author: CCDL for ALSF
date: 2024
---

*Adapted from [the AUCell vignette](https://bioconductor.org/packages/release/bioc/vignettes/AUCell/inst/doc/AUCell.html) and [the `cell-type-ewings` module](https://github.com/AlexsLemonade/OpenScPCA-analysis/tree/main/analyses/cell-type-ewings) that is part of the Open Single-cell Pediatric Cancer Atlas project.*

## Objectives

- Introduce the `AUCell` R package
- Illustrate how AUC values are calculated
- Demonstrate how AUC values can be used for cell assignment and plotting

---

In this notebook, we'll demonstrate how to use the AUCell method, introduced in [Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463).

We can use AUCell when we are interested in a gene set's relative expression or activity in an individual cell.
Gene sets can come from a curated collection of prior knowledge, like the Hallmark collection we used in the last notebook, or we can use our own custom gene sets (e.g., a set of marker genes for a cell type of interest).

A nice feature of AUCell is that it is based on ranking genes from highest to lowest expression value in an individual cell, which is helpful in the following ways ([AUCell vignette](https://bioconductor.org/packages/release/bioc/vignettes/AUCell/inst/doc/AUCell.html)):

- It can take a number of different values as input (e.g., raw counts, TPM) 
- It compensates for differences in library size, where something like averaging raw count values of genes in a gene set would not 
- It scales to larger datasets, since creating rankings is not as resource-intensive as something like permutation testing, and we could split up the object into subsets of cells if needed

AUCell calculates the area under the recovery curve (AUC), which "represents the proportion of expressed genes in the signature and their relative expression value compared to the other genes within the cell" ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)).
We will visualize some recovery curves in the notebook to give you a better intuition about the AUC and its meaning.

The AUC values we get out of AUCell can be used in a number of ways ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)):

- As continuous values we can use for visualization or clustering
- For binary assignment (i.e., "on" and "off" or "expressed" and "not expressed") if we pick a threshold either automatically using built-in functionality or manually by inspecting the distribution of scores ourselves

We will use an snRNA-seq of a Ewing sarcoma sample from the [`SCPCP000015` project](https://scpca.alexslemonade.org/projects/SCPCP000015) on the Single-cell Pediatric Cancer Atlas Portal and two relevant gene sets from the Molecular Signatures Database (MSigDB) to demonstrate this method.

## Set up

### Libraries

```{r setup}
# We will be loading a SingleCellExperiment object into our environment but don't need to see the startup messages
suppressPackageStartupMessages({
  library(SingleCellExperiment)
})

# Library we'll use for the gene set analysis itself
library(AUCell)

# Libraries for accessing and working with gene sets
library(GSEABase)
library(msigdbr)
```

### Directories and files

#### Directories

```{r setup_directories}
# Input data 
ewing_data_dir <- fs::path("data", "ewing-sarcoma")
processed_dir <- fs::path(ewing_data_dir, "processed")

# Directory for holding pathway analysis results
analysis_dir <- fs::path("analysis", "ewing-sarcoma", "pathway-analysis")
# Create if it doesn't exist yet
fs::dir_create(analysis_dir)
```

#### Files

The input will be a `SingleCellExperiment` for an individual Ewing sarcoma library.

```{r setup_input_files}
sce_file <- fs::path(processed_dir, 
                     "SCPCS000490", 
                     "SCPCL000822_processed.rds")
```

We will save the AUCell results as a table in the analysis directory.

```{r setup_output_files, live = TRUE}
output_file <- fs::path(analysis_dir,
                        "ewing_sarcoma_aucell_results.tsv")
```


### Functions

The `source()` function allows us to load in custom functions we saved in an `.R` file.

```{r source_functions}
source(fs::path("util", "aucell_functions.R"))
```

This loads one custom function, called `plot_recovery_curve()`, into our environment.
This function is adapted from [the AUCell vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd#L295-L316).

## Set up gene sets

We are going to use two gene sets pertaining to Ewing sarcoma.

* [`ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION`](https://www.gsea-msigdb.org/gsea/msigdb/geneset_page.jsp?geneSetName=ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION), which are genes that were highly expressed in a rhabdomyosarcoma cell line engineered to express the EWSR1-FLI1 fusion.
* [`RIGGI_EWING_SARCOMA_PROGENITOR_UP`](https://www.gsea-msigdb.org/gsea/msigdb/cards/RIGGI_EWING_SARCOMA_PROGENITOR_UP), which are genes that were highly expressed in mesenchymal stem cells engineered to express the EWS-FLI1 fusion protein.

We would expect both of these gene sets to have high expression in tumor cells.

```{r genesets}
# Create a named vector with the relevant gene set names
ewing_gene_set_names <- c(zhang = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION",
                          riggi = "RIGGI_EWING_SARCOMA_PROGENITOR_UP")

ewing_gene_set_names
```

These gene sets come from the C2 gene set collection from MSigDB.
Let's retrieve them using `msigdbr()`.

```{r extract_genesets, live = TRUE}
ewing_gene_sets_df <- msigdbr(species = "Homo sapiens",
                              category = "C2",
                              subcategory = "CGP") |>
  dplyr::filter(gs_name %in% ewing_gene_set_names)
```

`AUCell` uses gene sets in a particular format that comes from the `GSEABase` package.
We need to create a `GeneSetCollection`.

```{r gene_set_collection}
ewing_gene_set_collection <- ewing_gene_set_names |>
  purrr::map(
    # For each gene set
    \(gene_set_name) {
      ewing_gene_sets_df |>
        # Subset to the rows in that gene set
        dplyr::filter(gs_name == gene_set_name) |>
        # Grab the Ensembl gene identifiers
        dplyr::pull(ensembl_gene) |>
        # Create a GeneSet object
        GeneSet(setName = gene_set_name,
                geneIdType = ENSEMBLIdentifier())
    }
  ) |>
  # Turn the list of GeneSet objects into a GeneSet collection
  GeneSetCollection()
```

## Read in and prepare SingleCellExperiment

```{r read_in_sce, live = TRUE}
sce <- readr::read_rds(sce_file)
```

The `AUCell` functions takes an expression matrix with genes as rows and cells as column.
We can extract a counts matrix in sparse format for use with `AUCell`.

```{r counts_matrix}
# Extract counts matrix
counts_matrix <- counts(sce)
```

There may be genes in our gene set that do not appear in the SingleCellExperiment object.
We can remove them using the `subsetGeneSets()` function.

```{r subset_gene_sets, live = TRUE}
# Remove genes from gene sets if they are not in the SCE
ewing_gene_set_collection <- subsetGeneSets(ewing_gene_set_collection,
                                            rownames(counts_matrix))
```

## AUCell

AUCell relies on ranking genes from highest to lowest expression value to calculate the AUC.
The AUC is the area under the recovery curve, which captures the number of genes in a gene set that are present in the rankings above some threshold (i.e., it is the area under the curve to the left of this gene rank).
By default, the top 5% of genes are used as the threshold.

Some genes will not be detected (i.e., have 0 counts).
Genes can also have the same expression level (i.e., ties).
These undetected genes and ties will be randomly ordered in our ranking.
To make our rankings – and therefore results – reproducible, we will set a seed.

```{r set_seed, live = TRUE}
set.seed(2024)
```

### Cell ranking

The first step in AUCell is to rank genes for each cell from highest to lowest expression value.
We can do this using the `AUCell_buildRankings()` function, which will output a visualization showing the distribution of the number of genes detected in the cells in our SingleCellExperiment object.

```{r cell_rankings, live = TRUE}
cell_rankings <- AUCell_buildRankings(counts_matrix)
```

The AUCell authors recommend making sure most cells have at least the number of genes we will use as the max rank to calculate the AUC.

The AUC max rank value tells AUCell the cutoff in the gene rankings to use for calculating AUC; we will visualize this curve and max rank in just a moment.
If we picked a max rank higher than the number of genes detected in most cells, the non-detected genes that are randomly ordered would play an outsized role in our AUC values.

By default, the max rank is the top 5% highest expressed genes.
We can calculate the default max rank by taking into account the number of genes.

```{r explore_auc_max_rank}
nrow(cell_rankings) * 0.05
```

This number is probably too high, given the distribution of the number of genes detected by cell we visualized with `AUCell_buildRankings()`.

What if we chose a 1% threshold?

```{r lower_max_rank, live = TRUE}
nrow(cell_rankings) * 0.01
```

That is probably a more reasonable choice for this dataset.

We can use a function called `ceiling()` to round this and save it to a variable for later use.

```{r auc_max_rank, live = TRUE}
auc_max_rank <- ceiling(nrow(cell_rankings) * 0.01)
```

### Plotting AUC

The AUC values we get out of AUCell are the area under a recovery curve and estimate the proportion of genes in the gene set that are highly expressed (i.e., highly ranked).

Let's plot the recovery curve for a cell with high AUC and a cell with low AUC to get a better intuition about AUC values.
Earlier, we loaded a custom function we adapted from [the AUCell vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd) called `plot_recovery_curve()` with `source()`.

First, we'll start with a cell with a high AUC.
We picked this barcode ahead of time when we wrote the notebook.

```{r high_recovery_curve}
plot_recovery_curve(cell_rankings,
                    ewing_gene_set_collection,
                    gene_set_name = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION",
                    barcode = "CTGAGCGGTCTTTATC",
                    auc_max_rank = auc_max_rank)  # 1% threshold 
```

The x-axis is the gene ranks for all genes.
The y-axis is the number of genes in the signature at a given point in the gene ranking – the line will rise when a gene in the gene set is encountered in the ranking from highest to lowest.
The AUC is the area under this recovery curve at the max rank threshold chosen for this dataset.

Now, let's look at an example with a low AUC.

```{r low_recovery_curve}
plot_recovery_curve(cell_rankings,
                    ewing_gene_set_collection,
                    gene_set_name = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION",
                    barcode = "AGATAGAGTCACAATC",
                    auc_max_rank = auc_max_rank)  # 1% threshold
```

Far fewer genes in the gene set are ranked above the threshold, yielding a lower AUC value.

### Calculating the AUC

Once we have the rankings, we can calculate the AUC scores for both gene sets in all cells with the `AUCell_calcAUC()` function.

```{r calc_auc, live = TRUE}
cell_auc <- AUCell_calcAUC(geneSets = ewing_gene_set_collection, 
                           rankings = cell_rankings,
                           aucMaxRank = auc_max_rank)
```

This function returns an `aucellResults` object.

```{r check_str, live = TRUE}
str(cell_auc)
```

It can be much more convenient to work with this in a tabular format.

```{r auc_to_table}
# Extract AUC
auc_df <- cell_auc@assays@data$AUC |>
  # Transpose
  t() |>
  # Convert to data frame
  as.data.frame() |>
  # Make the barcodes a column
  tibble::rownames_to_column("barcodes") 

# Look at first few rows
head(auc_df)
```

### Assignments

AUCell can assign cells as having an active gene set or not by picking a threshold automatically.
We'll explore these in a later plot, but for now, let's calculate the threshold and assign cells with `AUCell_exploreThresholds()`.

```{r auc_assignments, live = TRUE}
auc_assignments <- AUCell_exploreThresholds(cell_auc, 
                                            plotHist = FALSE, 
                                            assignCells = TRUE)
```

We're going to plot the distribution of AUC values with `ggplot2`, so we will want the AUC values in a longer format.

```{r auc_plotting_df}
auc_plotting_df <- auc_df |>
  tidyr::pivot_longer(!barcodes,
                      names_to = "gene_set",
                      values_to = "auc") |>
  dplyr::mutate(
    # Create a new logical column called assigned
    assigned = dplyr::case_when(
      # For Zhang gene set rows, set to TRUE when the barcode is in the 
      # assignment list
      gene_set == ewing_gene_set_names[["zhang"]] & 
        barcodes %in% auc_assignments[[ewing_gene_set_names[["zhang"]]]]$assignment ~ TRUE,
      # For Riggi gene set rows, set to TRUE when the barcode is in the 
      # assignment list
      gene_set == ewing_gene_set_names[["riggi"]] & 
        barcodes %in% auc_assignments[[ewing_gene_set_names[["riggi"]]]]$assignment ~ TRUE,
      # Otherwise, set to FALSE
      .default = FALSE
    )
  )

auc_plotting_df
```

To draw vertical lines representing the automatically chosen threshold, we can create a separate data frame.

```{r auc_threshold_df}
auc_threshold_df <- data.frame(
  gene_set = ewing_gene_set_names,
  # Grab thresholds associated with each gene set from assignements object
  threshold = c(auc_assignments[[ewing_gene_set_names["zhang"]]]$aucThr$selected, 
                auc_assignments[[ewing_gene_set_names["riggi"]]]$aucThr$selected)
)

auc_threshold_df
```

Now let's make a density plot, plotting the density of the assigned and unassigned cells separately and drawing a vertical line for the threshold.

```{r auc_density_plot}
auc_plotting_df |>
  ggplot2::ggplot(
    ggplot2::aes(
      x = auc,  # AUC values
      color = assigned,  # Group by assignment
      fill = assigned,   # Group by assignment
    )
  ) +
  ggplot2::geom_density(alpha = 0.2) +
  # Draw a vertical dotted line showing the threshold for each gene set
  ggplot2::geom_vline(data = auc_threshold_df,
                      mapping = ggplot2::aes(xintercept = threshold),
                      lty = 2) +
  # Plot each gene set in its own facet
  ggplot2::facet_grid(cols = ggplot2::vars(gene_set)) +
  # Use a built-in theme
  ggplot2::theme_bw()
```

For these particular gene sets, the AUC values appear to be bimodally distributed, and we can easily identify cells where the genes are highly expressed.

Let's write this table to the output file.

```{r save_auc}
auc_plotting_df |> 
  readr::write_tsv(output_file)
```

### UMAPs

#### Adding AUC to `colData`

We can also add the AUC values back into the SingleCellExperiment for convenience, e.g., for plotting.
We'll add it to the existing `colData`.

First, let's rename the gene set columns to something more easily typed.

```{r rename_gene_set}
auc_df <- auc_df |>
  # Use shorter names
  dplyr::rename(zhang_auc = ewing_gene_set_names[["zhang"]],
                riggi_auc = ewing_gene_set_names[["riggi"]])

```

And join it to the existing `colData`.

```{r coldata, live = TRUE}
# Extract the existing colData, and left join it with the AUC values by the
# barcodes
coldata_df <- colData(sce) |>
  as.data.frame() |>
  dplyr::left_join(
    auc_df,
    by = "barcodes"
  )
```

Now, we're ready to add it back to the object.

```{r add_back_colData, live = TRUE}
# We need to save this as a DataFrame
colData(sce) <- DataFrame(
  coldata_df,
  row.names = colData(sce)$barcodes
)
```

#### Plotting UMAPs

We can use the `plotUMAP()` function from the `scater` package to plot a UMAP with the points colored by the AUC value

```{r plot_umap_zhang}
scater::plotUMAP(sce, colour_by = "zhang_auc") +
  # Use the gene set name, replacing underscores with spaces
  ggplot2::ggtitle(stringr::str_replace_all(ewing_gene_set_names[["zhang"]], 
                                            "\\_", 
                                            " "))
```

Let's color the points by the AUC values for the other gene set.

```{r plot_umap_riggi, live = TRUE}
scater::plotUMAP(sce, colour_by = "riggi_auc") + 
  ggplot2::ggtitle(stringr::str_replace_all(ewing_gene_set_names[["riggi"]], 
                                            "\\_", 
                                            " "))
```

We would want to do something more formal to confirm, but it seems like the same cells have high AUC values for both gene sets!

## Session Info

```{r session_info}
sessionInfo()
```

    +
    ---
title: "Pathway Analysis: AUCell"
output:
  html_notebook:
    toc: true
    toc_float: true
author: CCDL for ALSF
date: 2024
---

*Adapted from [the AUCell vignette](https://bioconductor.org/packages/release/bioc/vignettes/AUCell/inst/doc/AUCell.html) and [the `cell-type-ewings` module](https://github.com/AlexsLemonade/OpenScPCA-analysis/tree/main/analyses/cell-type-ewings) that is part of the Open Single-cell Pediatric Cancer Atlas project.*

## Objectives

- Introduce the `AUCell` R package
- Illustrate how AUC values are calculated
- Demonstrate how AUC values can be used for cell assignment and plotting

---

In this notebook, we'll demonstrate how to use the `AUCell` method, introduced in [Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463).

We can use `AUCell` when we are interested in a gene set's relative expression or activity in an individual cell.
Gene sets can come from a curated collection of prior knowledge, like the Hallmark collection we used in the last notebook, or we can use our own custom gene sets (e.g., a set of marker genes for a cell type of interest).

A nice feature of `AUCell` is that it is based on ranking genes from highest to lowest expression value in an individual cell, which is helpful in the following ways ([`AUCell` vignette](https://bioconductor.org/packages/release/bioc/vignettes/AUCell/inst/doc/AUCell.html)):

- It can take a number of different values as input (e.g., raw counts, TPM) 
- It compensates for differences in library size, where something like averaging raw count values of genes in a gene set would not 
- It scales to larger datasets, since creating rankings is not as resource-intensive as something like permutation testing, and we could split up the object into subsets of cells if needed

`AUCell` calculates the area under the recovery curve (AUC), which "represents the proportion of expressed genes in the signature and their relative expression value compared to the other genes within the cell" ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)).
We will visualize some recovery curves in the notebook to give you a better intuition about the AUC and its meaning.

The AUC values we get out of `AUCell` can be used in a number of ways ([Aibar _et al_. 2017.](https://doi.org/10.1038/nmeth.4463)):

- As continuous values we can use for visualization or clustering
- For binary assignment (i.e., "on" and "off" or "expressed" and "not expressed") if we pick a threshold either automatically using built-in functionality or manually by inspecting the distribution of scores ourselves

We will use an snRNA-seq of a Ewing sarcoma sample from the [`SCPCP000015` project](https://scpca.alexslemonade.org/projects/SCPCP000015) on the Single-cell Pediatric Cancer Atlas Portal and two relevant gene sets from the Molecular Signatures Database (MSigDB) to demonstrate this method.

## Set up

### Libraries

```{r setup}
# We will be loading a SingleCellExperiment object into our environment but don't need to see the startup messages
suppressPackageStartupMessages({
  library(SingleCellExperiment)
})

# Library we'll use for the gene set analysis itself
library(AUCell)

# Libraries for accessing and working with gene sets
library(GSEABase)
library(msigdbr)
```

### Directories and files

#### Directories

```{r setup_directories}
# Input data 
ewing_data_dir <- fs::path("data", "ewing-sarcoma")
processed_dir <- fs::path(ewing_data_dir, "processed")

# Directory for holding pathway analysis results
analysis_dir <- fs::path("analysis", "ewing-sarcoma", "pathway-analysis")
# Create if it doesn't exist yet
fs::dir_create(analysis_dir)
```

#### Files

The input will be a `SingleCellExperiment` for an individual Ewing sarcoma library.

```{r setup_input_files}
sce_file <- fs::path(processed_dir, 
                     "SCPCS000490", 
                     "SCPCL000822_processed.rds")
```

We will save the `AUCell` results as a table in the analysis directory.

```{r setup_output_files, live = TRUE}
output_file <- fs::path(analysis_dir,
                        "ewing_sarcoma_aucell_results.tsv")
```


### Functions

The `source()` function allows us to load in custom functions we saved in an `.R` file.

```{r source_functions}
source(fs::path("util", "aucell_functions.R"))
```

This loads one custom function, called `plot_recovery_curve()`, into our environment.
This function is adapted from [the `AUCell` vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd#L295-L316).

## Set up gene sets

We are going to use two gene sets pertaining to Ewing sarcoma.

* [`ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION`](https://www.gsea-msigdb.org/gsea/msigdb/geneset_page.jsp?geneSetName=ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION), which are genes that were highly expressed in a rhabdomyosarcoma cell line engineered to express the EWSR1-FLI1 fusion.
* [`RIGGI_EWING_SARCOMA_PROGENITOR_UP`](https://www.gsea-msigdb.org/gsea/msigdb/cards/RIGGI_EWING_SARCOMA_PROGENITOR_UP), which are genes that were highly expressed in mesenchymal stem cells engineered to express the EWS-FLI1 fusion protein.

We would expect both of these gene sets to have high expression in tumor cells.

```{r genesets}
# Create a named vector with the relevant gene set names
ewing_gene_set_names <- c(zhang = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION",
                          riggi = "RIGGI_EWING_SARCOMA_PROGENITOR_UP")

ewing_gene_set_names
```

These gene sets come from the C2 gene set collection from MSigDB.
Let's retrieve them using `msigdbr()`.

```{r extract_genesets, live = TRUE}
ewing_gene_sets_df <- msigdbr(species = "Homo sapiens",
                              category = "C2",
                              subcategory = "CGP") |>
  dplyr::filter(gs_name %in% ewing_gene_set_names)
```

`AUCell` uses gene sets in a particular format that comes from the `GSEABase` package.
We need to create a `GeneSetCollection`.

```{r gene_set_collection}
ewing_gene_set_collection <- ewing_gene_set_names |>
  purrr::map(
    # For each gene set
    \(gene_set_name) {
      ewing_gene_sets_df |>
        # Subset to the rows in that gene set
        dplyr::filter(gs_name == gene_set_name) |>
        # Grab the Ensembl gene identifiers
        dplyr::pull(ensembl_gene) |>
        # Create a GeneSet object
        GeneSet(setName = gene_set_name,
                geneIdType = ENSEMBLIdentifier())
    }
  ) |>
  # Turn the list of GeneSet objects into a GeneSet collection
  GeneSetCollection()
```

## Read in and prepare the `SingleCellExperiment`

```{r read_in_sce, live = TRUE}
sce <- readr::read_rds(sce_file)
```

Our object includes counts for all genes that were present in the index when quantifying gene expression.
There are a number of genes that are present in the object but not detected in any of the cells. 
We don't want genes that are not found in our data set to impact our rankings, so let's remove them. 

```{r, filter_sce}
# remove genes that are not detected in any of the cells from the SCE object
genes_to_keep <- rowData(sce)$detected > 0
sce <- sce[genes_to_keep, ]
```


The `AUCell` functions takes an expression matrix with genes as rows and cells as column.
We can extract a counts matrix in sparse format for use with `AUCell`.


```{r counts_matrix}
# Extract counts matrix
counts_matrix <- counts(sce)
```

There may be genes in our gene set that do not appear in the `SingleCellExperiment` object.
We can remove them using the `subsetGeneSets()` function.

```{r subset_gene_sets, live = TRUE}
# Remove genes from gene sets if they are not in the SCE
ewing_gene_set_collection <- subsetGeneSets(ewing_gene_set_collection,
                                            rownames(counts_matrix))
```

## `AUCell`

`AUCell` relies on ranking genes from highest to lowest expression value to calculate the AUC.
The AUC is the area under the recovery curve, which captures the number of genes in a gene set that are present in the rankings above some threshold (i.e., it is the area under the curve to the left of this gene rank).
By default, the top 5% of genes are used as the threshold.

Some genes will not be detected (i.e., have 0 counts).
Genes can also have the same expression level (i.e., ties).
These undetected genes and ties will be randomly ordered in our ranking.
To make our rankings – and therefore results – reproducible, we will set a seed.

```{r set_seed, live = TRUE}
set.seed(2024)
```

### Cell ranking

The first step in `AUCell` is to rank genes for each cell from highest to lowest expression value.
We can do this using the `AUCell_buildRankings()` function, which will output a visualization showing the distribution of the number of genes detected in the cells in our `SingleCellExperiment` object.

```{r cell_rankings, live = TRUE}
cell_rankings <- AUCell_buildRankings(counts_matrix)
```

The `AUCell` authors recommend making sure most cells have at least the number of genes we will use as the max rank to calculate the AUC.

The AUC max rank value tells `AUCell` the cutoff in the gene rankings to use for calculating AUC; we will visualize this curve and max rank in just a moment.
If we picked a max rank higher than the number of genes detected in most cells, the non-detected genes that are randomly ordered would play an outsized role in our AUC values.

By default, the max rank is the top 5% highest expressed genes.
We can calculate the default max rank by taking into account the number of genes.

```{r explore_auc_max_rank}
nrow(cell_rankings) * 0.05
```

This number is probably too high, given the distribution of the number of genes detected by cell we visualized with `AUCell_buildRankings()`.

What if we chose a 1% threshold?

```{r lower_max_rank, live = TRUE}
nrow(cell_rankings) * 0.01
```

That is probably a more reasonable choice for this dataset.

We can use a function called `ceiling()` to round this and save it to a variable for later use.

```{r auc_max_rank, live = TRUE}
auc_max_rank <- ceiling(nrow(cell_rankings) * 0.01)
```

### Plotting AUC

The AUC values we get out of `AUCell` are the area under a recovery curve and estimate the proportion of genes in the gene set that are highly expressed (i.e., highly ranked).

Let's plot the recovery curve for a cell with high AUC and a cell with low AUC to get a better intuition about AUC values.
Earlier, we loaded a custom function we adapted from [the `AUCell` vignette](https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd) called `plot_recovery_curve()` with `source()`.

First, we'll start with a cell with a high AUC.
We picked this barcode ahead of time when we wrote the notebook.

```{r high_recovery_curve}
plot_recovery_curve(cell_rankings,
                    ewing_gene_set_collection,
                    gene_set_name = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION",
                    barcode = "CTGAGCGGTCTTTATC",
                    auc_max_rank = auc_max_rank)  # 1% threshold 
```

The x-axis is the gene ranks for all genes.
The y-axis is the number of genes in the signature at a given point in the gene ranking – the line will rise when a gene in the gene set is encountered in the ranking from highest to lowest.
The AUC is the area under this recovery curve at the max rank threshold chosen for this dataset.

Now, let's look at an example with a low AUC.

```{r low_recovery_curve}
plot_recovery_curve(cell_rankings,
                    ewing_gene_set_collection,
                    gene_set_name = "ZHANG_TARGETS_OF_EWSR1_FLI1_FUSION",
                    barcode = "AGATAGAGTCACAATC",
                    auc_max_rank = auc_max_rank)  # 1% threshold
```

Far fewer genes in the gene set are ranked above the threshold, yielding a lower AUC value.

### Calculating the AUC

Once we have the rankings, we can calculate the AUC scores for both gene sets in all cells with the `AUCell_calcAUC()` function.

```{r calc_auc, live = TRUE}
cell_auc <- AUCell_calcAUC(geneSets = ewing_gene_set_collection, 
                           rankings = cell_rankings,
                           aucMaxRank = auc_max_rank)
```

This function returns an `aucellResults` object.

```{r check_str, live = TRUE}
str(cell_auc)
```

It can be much more convenient to work with this in a tabular format.

```{r auc_to_table}
# Extract AUC
auc_df <- cell_auc@assays@data$AUC |>
  # Transpose
  t() |>
  # Convert to data frame
  as.data.frame() |>
  # Make the barcodes a column
  tibble::rownames_to_column("barcodes") 

# Look at first few rows
head(auc_df)
```

### Assignments

`AUCell` can assign cells as having an active gene set or not by picking a threshold automatically.
We'll explore these in a later plot, but for now, let's calculate the threshold and assign cells with `AUCell_exploreThresholds()`.

```{r auc_assignments, live = TRUE}
auc_assignments <- AUCell_exploreThresholds(cell_auc, 
                                            plotHist = FALSE, 
                                            assignCells = TRUE)
```

We're going to plot the distribution of AUC values with `ggplot2`, so we will want the AUC values in a longer format.

```{r auc_plotting_df}
auc_plotting_df <- auc_df |>
  tidyr::pivot_longer(!barcodes,
                      names_to = "gene_set",
                      values_to = "auc") |>
  dplyr::mutate(
    # Create a new logical column called assigned
    assigned = dplyr::case_when(
      # For Zhang gene set rows, set to TRUE when the barcode is in the 
      # assignment list
      gene_set == ewing_gene_set_names[["zhang"]] & 
        barcodes %in% auc_assignments[[ewing_gene_set_names[["zhang"]]]]$assignment ~ TRUE,
      # For Riggi gene set rows, set to TRUE when the barcode is in the 
      # assignment list
      gene_set == ewing_gene_set_names[["riggi"]] & 
        barcodes %in% auc_assignments[[ewing_gene_set_names[["riggi"]]]]$assignment ~ TRUE,
      # Otherwise, set to FALSE
      .default = FALSE
    )
  )

auc_plotting_df
```

To draw vertical lines representing the automatically chosen threshold, we can create a separate data frame.

```{r auc_threshold_df}
auc_threshold_df <- data.frame(
  gene_set = ewing_gene_set_names,
  # Grab thresholds associated with each gene set from assignements object
  threshold = c(auc_assignments[[ewing_gene_set_names["zhang"]]]$aucThr$selected, 
                auc_assignments[[ewing_gene_set_names["riggi"]]]$aucThr$selected)
)

auc_threshold_df
```

Now let's make a density plot, plotting the density of the assigned and unassigned cells separately and drawing a vertical line for the threshold.

```{r auc_density_plot}
auc_plotting_df |>
  ggplot2::ggplot(
    ggplot2::aes(
      x = auc,  # AUC values
      color = assigned,  # Group by assignment
      fill = assigned,   # Group by assignment
    )
  ) +
  ggplot2::geom_density(alpha = 0.2) +
  # Draw a vertical dotted line showing the threshold for each gene set
  ggplot2::geom_vline(data = auc_threshold_df,
                      mapping = ggplot2::aes(xintercept = threshold),
                      lty = 2) +
  # Plot each gene set in its own facet
  ggplot2::facet_grid(cols = ggplot2::vars(gene_set)) +
  # Use a built-in theme
  ggplot2::theme_bw()
```

For these particular gene sets, the AUC values appear to be bimodally distributed, and we can easily identify cells where the genes are highly expressed.

Let's write this table to the output file.

```{r save_auc}
auc_plotting_df |> 
  readr::write_tsv(output_file)
```

### UMAPs

#### Adding AUC to `colData`

We can also add the AUC values back into the `SingleCellExperiment` for convenience, e.g., for plotting.
We'll add it to the existing `colData`.

First, let's rename the gene set columns to something more easily typed.

```{r rename_gene_set}
auc_df <- auc_df |>
  # Use shorter names
  dplyr::rename(zhang_auc = ewing_gene_set_names[["zhang"]],
                riggi_auc = ewing_gene_set_names[["riggi"]])

```

And join it to the existing `colData`.

```{r coldata, live = TRUE}
# Extract the existing colData, and left join it with the AUC values by the
# barcodes
coldata_df <- colData(sce) |>
  as.data.frame() |>
  dplyr::left_join(
    auc_df,
    by = "barcodes"
  )
```

Now, we're ready to add it back to the object.

```{r add_back_colData, live = TRUE}
# We need to save this as a DataFrame
colData(sce) <- DataFrame(
  coldata_df,
  row.names = colData(sce)$barcodes
)
```

#### Plotting UMAPs

We can use the `plotUMAP()` function from the `scater` package to plot a UMAP with the points colored by the AUC value

```{r plot_umap_zhang}
scater::plotUMAP(sce, colour_by = "zhang_auc") +
  # Use the gene set name, replacing underscores with spaces
  ggplot2::ggtitle(stringr::str_replace_all(ewing_gene_set_names[["zhang"]], 
                                            "\\_", 
                                            " "))
```

Let's color the points by the AUC values for the other gene set.

```{r plot_umap_riggi, live = TRUE}
scater::plotUMAP(sce, colour_by = "riggi_auc") + 
  ggplot2::ggtitle(stringr::str_replace_all(ewing_gene_set_names[["riggi"]], 
                                            "\\_", 
                                            " "))
```

We would want to do something more formal to confirm, but it seems like the same cells have high AUC values for both gene sets!

## Session Info

```{r session_info}
sessionInfo()
```

    diff --git a/scRNA-seq-advanced/diagrams/roadmap_multi_integrate.png b/scRNA-seq-advanced/diagrams/roadmap_multi_integrate.png index 6fc50c84..cc519690 100644 Binary files a/scRNA-seq-advanced/diagrams/roadmap_multi_integrate.png and b/scRNA-seq-advanced/diagrams/roadmap_multi_integrate.png differ diff --git a/scRNA-seq-advanced/diagrams/technical_merge_sce.png b/scRNA-seq-advanced/diagrams/technical_merge_sce.png index 48ab0948..03cc4332 100644 Binary files a/scRNA-seq-advanced/diagrams/technical_merge_sce.png and b/scRNA-seq-advanced/diagrams/technical_merge_sce.png differ diff --git a/scRNA-seq-advanced/exercise_02-integration.Rmd b/scRNA-seq-advanced/exercise_02-integration.Rmd index 62c67cfe..b4596300 100644 --- a/scRNA-seq-advanced/exercise_02-integration.Rmd +++ b/scRNA-seq-advanced/exercise_02-integration.Rmd @@ -274,9 +274,10 @@ We will use the default clustering parameters for graph-based clustering to star Remember that the clustering results depend only the expression data that was used to generate the PC matrix – the cell labels are not used in the clustering algorithms. ```{r cluster unintegrated} -merged_sce$cluster_unintegrated <- bluster::clusterRows( - reducedDim(merged_sce, "PCA"), - bluster::NNGraphParam() +merged_sce$cluster_unintegrated <- scran::clusterCells( + merged_sce, + use.dimred = "PCA", + BLUSPARAM = bluster::NNGraphParam() ) ``` diff --git a/scRNA-seq-advanced/exercise_04-scrna_pathway.Rmd b/scRNA-seq-advanced/exercise_04-scrna_pathway.Rmd index d1c6266a..8577bf32 100644 --- a/scRNA-seq-advanced/exercise_04-scrna_pathway.Rmd +++ b/scRNA-seq-advanced/exercise_04-scrna_pathway.Rmd @@ -8,8 +8,8 @@ output: toc_float: true --- -In this notebook, we will use AUCell with a custom gene set of tumor marker genes for Ewing sarcoma, as well as a collection from the Molecular Signatures Database (MSigDB) of your choosing. -Because AUCell will assign cells as expressing (or not expressing) a gene set, it can be used as part of a strategy for cell typing or identifying malignant cells when a high quality marker gene set is available. +In this notebook, we will use `AUCell` with a custom gene set of tumor marker genes for Ewing sarcoma, as well as a collection from the Molecular Signatures Database (MSigDB) of your choosing. +Because `AUCell` will assign cells as expressing (or not expressing) a gene set, it can be used as part of a strategy for cell typing or identifying malignant cells when a high quality marker gene set is available. Once tumor cells are identified using an automatically selected threshold, we can visualize that information or even use it in tandem with AUC values from other gene sets. Our goal in this notebook is to explore gene sets from MSigDB that may have different relative expression (i.e., AUC values) between malignant and non-malignant cells. @@ -19,8 +19,8 @@ In practice, if we were to publish these results, we would want to explicitly te In this notebook, you will: - Part A: Read in and prepare the data -- Part B: Run AUCell with tumor marker genes -- Part C: Run AUCell with an MSigDB collection +- Part B: Run `AUCell` with tumor marker genes +- Part C: Run `AUCell` with an MSigDB collection - Part D: Visualize the results There are multiple Ewing sarcoma samples you can choose for your analysis, available in the following directory: @@ -52,7 +52,7 @@ library(msigdbr) ### Set a seed -Some genes sharing expression values or that are not detected will be randomly ordered in the AUCell rankings, so we need to set a seed using `set.seed()`. +Some genes sharing expression values or that are not detected will be randomly ordered in the `AUCell` rankings, so we need to set a seed using `set.seed()`. ```{r set_seed, solution = TRUE} @@ -62,7 +62,7 @@ Some genes sharing expression values or that are not detected will be randomly o #### Directories -Set up the directories to read in a Ewing sarcoma SingleCellExperiment and output pathway analysis results. +Set up the directories to read in a Ewing sarcoma `SingleCellExperiment` and output pathway analysis results. ```{r set_up_directories, solution = TRUE} # Input data @@ -118,12 +118,12 @@ marker_genes_df You can use the information in the `source` column to review the publication a marker gene's inclusion is supported by. -Now we need to get the gene set ready for use with AUCell. +Now we need to get the gene set ready for use with `AUCell`. You're going to save the gene set to a vector called `ensg_tumor_markers`. Here are some things to keep in mind when preparing the gene set: - You are only interested in the _tumor_ gene set -- The SingleCellExperiment object uses Ensembl gene identifiers +- The `SingleCellExperiment` object uses Ensembl gene identifiers - You'll want to remove any duplicate genes in the gene set ```{r format_marker_genes, solution = TRUE} @@ -131,7 +131,7 @@ Here are some things to keep in mind when preparing the gene set: ``` -Next, convert the vector of Ensembl ids to a `GeneSet` object for use with AUCell: +Next, convert the vector of Ensembl ids to a `GeneSet` object for use with `AUCell`: ```{r GeneSet} ensg_tumor_markers <- GeneSet(ensg_tumor_markers, @@ -142,7 +142,7 @@ ensg_tumor_markers <- GeneSet(ensg_tumor_markers, ### Prepare the single-nuclei data Now that we have our gene set prepared, let's prepare the data. -First, we will read in the SingleCellExperiment from the file we chose earlier. +First, we will read in the `SingleCellExperiment` from the file we chose earlier. ```{r read_in_sce} sce <- readr::read_rds(sce_file) @@ -154,7 +154,7 @@ We will mostly be using the raw counts, so save the counts matrix to a separate ``` -## Part B: Run AUCell with tumor marker genes +## Part B: Run `AUCell` with tumor marker genes Calculate the gene rankings for individual cells. @@ -182,7 +182,7 @@ Save the output to `cell_auc`. ``` Automatically calculate a threshold for the AUC values using `AUCell_exploreThresholds()`. -You'll want to set `assignCells = TRUE`, which tells AUCell to assign tumor cell labels to cells above the automatically chosen threshold. +You'll want to set `assignCells = TRUE`, which tells `AUCell` to assign tumor cell labels to cells above the automatically chosen threshold. Save the output to `auc_assignments`. @@ -192,7 +192,7 @@ Save the output to `auc_assignments`. ### Wrangle data for plotting -We will create a data frame that holds the barcodes, AUC values, and a column called `tumor_cell` that indicates whether or not AUCell classified the cell as a tumor cell using the AUC values for the marker gene set. +We will create a data frame that holds the barcodes, AUC values, and a column called `tumor_cell` that indicates whether or not `AUCell` classified the cell as a tumor cell using the AUC values for the marker gene set. ```{r auc_df} auc_df <- cell_auc@assays@data$AUC |> # start with the internal AUC table @@ -226,11 +226,11 @@ Add a vertical line representing the threshold that was automatically selected a ``` -## Part C: Run AUCell with an MSigDB collection +## Part C: Run `AUCell` with an MSigDB collection -Now, let's run AUCell using a collection from the Molecular Signatures Database (MSigDB). +Now, let's run `AUCell` using a collection from the Molecular Signatures Database (MSigDB). -We can use the results to explore if there are gene sets that have different expression in tumor cells vs. other cells in the sample, using the labels from AUCell. +We can use the results to explore if there are gene sets that have different expression in tumor cells vs. other cells in the sample, using the labels from `AUCell`. Let's look at what collections are available as part of the `msigdbr` package: @@ -243,13 +243,13 @@ It may be helpful to cross-reference this with the MSigDB website: @@ -270,7 +270,7 @@ collection_list <- unique(collection_df$gs_name) |> GeneSetCollection() ``` -AUCell provides a wrapper function that runs the ranking and AUC calculation steps called `AUCell_run()`. +`AUCell` provides a wrapper function that runs the ranking and AUC calculation steps called `AUCell_run()`. Because we're using the same sample as we did with the tumor marker genes, we can use the same AUC max rank value we used earlier (`auc_max_rank`). You may need to change the names of the variables passed to the `exprsMat` and `aucMaxRank` arguments below depending on what you used earlier! @@ -315,11 +315,11 @@ Write this data frame to the output file you saved as a variable earlier. ### Plot UMAPs -We can use the data in `auc_df` to make UMAP visualizations once we add it to the SingleCellExperiment object. +We can use the data in `auc_df` to make UMAP visualizations once we add it to the `SingleCellExperiment` object. #### Add AUC values to `colData` -To prepare for use with `scater::plotUMAP`, let's add the `auc_df` to the `colData` of the SingleCellExperiment. +To prepare for use with `scater::plotUMAP`, let's add the `auc_df` to the `colData` of the `SingleCellExperiment` object. Don't forget to provide the `row.names` argument when converting back to a `DataFrame`. ```{r add_to_col_data, solution = TRUE} @@ -331,7 +331,7 @@ Don't forget to provide the `row.names` argument when converting back to a `Data #### Plot tumor cell assignments (UMAP) -First, plot a UMAP, coloring cells by whether or not they are a tumor cell according to AUCell. +First, plot a UMAP, coloring cells by whether or not they are a tumor cell according to `AUCell`. ```{r plot_tumor_cell, solution = TRUE} diff --git a/scRNA-seq-advanced/exercise_05-cluster_evaluation.Rmd b/scRNA-seq-advanced/exercise_05-cluster_evaluation.Rmd index 44b54e73..ec2a192b 100644 --- a/scRNA-seq-advanced/exercise_05-cluster_evaluation.Rmd +++ b/scRNA-seq-advanced/exercise_05-cluster_evaluation.Rmd @@ -29,6 +29,7 @@ These quantities are explained more in depth when they are introduced, but you c - The `bluster` package vignette on cluster evaluation: - The "Clustering Redux" chapter in _Orchestrating Single Cell Analysis_: +We'll specifically run `bluster` using the [`scran::clusterCells()`](https://rdrr.io/github/MarioniLab/scran/man/clusterCells.html) function. Before we dive in, it's important to bear some language caveats in mind: We sometimes use phrasing like "the best clusters" or "the optimal clusters," but the truth is, it's quite hard (if even possible) to know which clustering assignments are truly _the best._ @@ -149,15 +150,6 @@ We will begin by calculating clusters using different values of the `resolution` Then, we'll compare results to one another with the ultimate goal of identifying a reasonable `resolution` parameter to achieve reliable clusters. To specifically explore the effect of different `resolution` parameters, we won't vary any other parameters; we'll also use 20 nearest neighbors and Jaccard weighting for all clusterings. -To begin, extract the PCA matrix from the SCE using the `reducedDim()` function, and save it to a variable called `pca_matrix`. -You'll need to provide two arguments to this function: The SCE object, and the name of the reduced dimension we'd like to pull out; here, it's `"PCA"`. - -We'll need this matrix to both perform and evaluate clusters. - -```{r extract pca matrix, solution = TRUE} - -``` - We'll define our vector of values to try out for the `resolution` parameter. For the `modularity` objective function, `resolution` values around 1 tend to perform well, so we'll explore a range of values around 1: @@ -166,9 +158,12 @@ res_params <- seq(0.25, 1.5, 0.25) res_params ``` -Now, we're going to use `purrr::map()` to perform clustering for each of these values using `bluster::clusterRows()`. +Now, we're going to use `purrr::map()` to perform clustering for each of these values using `scran::clusterCells()` which uses `bluster` to perform clustering. +The `scran::clusterCells()` function takes an SCE object and a specification of which assay or reduced dimension to use for clustering - here, we want to use the PCA reduced dimension, which is a very standard choice for single-cell clustering. +The other argument that `scran::clusterCells()` takes is `BLUSPARAM`, which will specify the algorithm and any additional algorithm parameters for `bluster` to use. +This argument is made with a special `bluster` function, for example `bluster::KMeansParam()` or `bluster::NNGraphParam()` which respectively specify k-means clustering and graph-based clustering. -The result will be a list of vectors containing cluster assignments for each cell. +When we run `purrr::map()` over our different resolution parameters, we'll end up with a list of vectors containing cluster assignments for each cell. Before we do this, we'll name our `res_params` vector with the `resolution` values themselves. This will ensure the output from `purrr::map()` is also named according to these resolution parameters, which will help us keep track of which clustering is which! @@ -178,7 +173,7 @@ names(res_params) <- res_params ``` Time to cluster! -To make this code a bit easier to follow, we'll define our clustering parameters first using `bluster::NNGraphParam()`, and then we'll call `bluster::clusterRows()`. +To make this code a bit easier to follow, we'll define our clustering parameters first using `bluster::NNGraphParam()` for graph-based clustering, and then we'll call `scran::clusterCells()`. ```{r perform clustering} cluster_list <- res_params |> @@ -200,13 +195,17 @@ cluster_list <- res_params |> ) # Perform clustering - bluster::clusterRows(pca_matrix, cluster_params) + scran::clusterCells( + sce, + use.dimred = "PCA", # specify to use the PCA matrix for clustering + BLUSPARAM = cluster_params + ) } ) ``` How many clusters were created for each `resolution` parameter? -The cluster vector created by `bluster::clusterRows()` is a factor, so we can answer this by finding the length of each clustering's levels using `purrr::map()`. +The cluster vector created by `scran::clusterCells()` is a factor, so we can answer this by finding the length of each clustering's levels using `purrr::map()`. In the chunk below, use `purrr::map()` to get, for each clustering, the `length()` of its `levels()`. @@ -270,6 +269,12 @@ How does one choose among these potential results? In this section, we'll walk through several metrics to evaluate cluster quality and their interpretations. +Part of evaluating these clusters will involve working directly with the PCA matrix we clustered, so we'll go ahead and pull that out into its own variable called `pca_matrix` for convenience: + +```{r extract pca matrix, solution = TRUE} + +``` + ### Silhouette width @@ -391,7 +396,7 @@ This function returns a capital-D `DataFrame` with one row per cell and the foll In the chunk below, use `purrr::map()` to calculate the neighborhood purity on each vector of clusters. Just like we did for calculating silhouette width, you'll want to coerce the `bluster::neighborPurity()` output into a small-d `data.frame` with row names moved into a new column `barcodes`. -_Unlike_ for silhouette width though, the `bluster::neighborhoodPurity()` function does not automatically save the cluster assignments to the output data frame, so you'll also want to add a `dplyr::mutate()` statement to ensure there is a `cluster` column in the output. +_Unlike_ for silhouette width though, the `bluster::neighborPurity()` function does not automatically save the cluster assignments to the output data frame, so you'll also want to add a `dplyr::mutate()` statement to ensure there is a `cluster` column in the output. Finally, after the `purrr:map()` statement, bind (hint!) all rows together, specifying `.id = "resolution"`, to create one data frame with all results. Save this result to `cluster_purity_df`, and print the data frame once you have created it (again, remember that R Markdown will only print 1000 rows, even when there are more!) diff --git a/scRNA-seq-advanced/util/aucell_functions.R b/scRNA-seq-advanced/util/aucell_functions.R index 16f3482f..e4bbfa98 100644 --- a/scRNA-seq-advanced/util/aucell_functions.R +++ b/scRNA-seq-advanced/util/aucell_functions.R @@ -3,8 +3,8 @@ #' Adapted from https://github.com/aertslab/AUCell/blob/91753b327a39dc7a4bbed46408ec2271c485f2f0/vignettes/AUCell.Rmd#L295-L316 #' #' @param cell_rankings Cell rankings; the output of AUCell::AUCell_buildRankings() -#' @param gene_set_collection A GSEABase GeneSetCollection object -#' @param gene_set_name The name of the gene set from the GeneSetCollection +#' @param gene_set_list A list of gene sets +#' @param gene_set_name The name of the gene set the gene_set_list #' that you would like to plot a recovery curve for #' @param barcode The cell barcode that you would like to plot a recovery #' curve for @@ -14,7 +14,7 @@ #' #' @return Outputs a recovery curve plot with the plot_recovery_curve <- function(cell_rankings, - gene_set_collection, + gene_set_list, gene_set_name, barcode, auc_max_rank, @@ -24,8 +24,8 @@ plot_recovery_curve <- function(cell_rankings, # Pull out the gene set and identify where in the cell ranks those genes # lie - gene_set <- gene_set_collection[[gene_set_name]] - gene_set_ranks <- cell_rankings[geneIds(gene_set), ] + gene_set <- gene_set_list[[gene_set_name]] + gene_set_ranks <- cell_rankings[gene_set, ] # Index of the cell with the barcode barcode_index <- which(colnames(gene_set_ranks) == barcode) @@ -35,7 +35,7 @@ plot_recovery_curve <- function(cell_rankings, plot(aucCurve, type="s", col="darkblue", lwd=1, xlab="Gene rank", ylab="# genes in the gene set", - xlim=c(0, auc_max_rank*3), ylim=c(0, nGenes(gene_set)), + xlim=c(0, auc_max_rank*3), ylim=c(0, length(gene_set)), main="Recovery curve", sub=paste("Cell:", colnames(gene_set_ranks)[barcode_index])) aucShade <- aucCurve[which(aucCurve[,1] < auc_max_rank),] diff --git a/scRNA-seq/06-celltype_annotation.Rmd b/scRNA-seq/06-celltype_annotation.Rmd index 82219b58..76bb8a61 100644 --- a/scRNA-seq/06-celltype_annotation.Rmd +++ b/scRNA-seq/06-celltype_annotation.Rmd @@ -61,24 +61,32 @@ We aren't planning any significant modifications of the underlying data, so we w ```{r filepaths, live=TRUE} # directory for the input data -data_dir <- file.path("data", - "PBMC-TotalSeqB", - "normalized") +data_dir <- file.path( + "data", + "PBMC-TotalSeqB", + "normalized" +) # the input file itself -sce_file <- file.path(data_dir, - "PBMC_TotalSeqB_normalized_sce.rds") +sce_file <- file.path( + data_dir, + "PBMC_TotalSeqB_normalized_sce.rds" +) # A directory to store outputs -analysis_dir <- file.path("analysis", - "PBMC-TotalSeqB") +analysis_dir <- file.path( + "analysis", + "PBMC-TotalSeqB" +) # Create directory if it doesn't exist fs::dir_create(analysis_dir) # output table path -cellinfo_file <- file.path(analysis_dir, - "PBMC_TotalSeqB_cellinfo.tsv") +cellinfo_file <- file.path( + analysis_dir, + "PBMC_TotalSeqB_cellinfo.tsv" +) ``` @@ -198,14 +206,12 @@ Let's plot the ADT results for those two markers as well below: ```{r plot CD4, live=TRUE} # plot CD4 marker -scater::plotUMAP(sce, - color_by = "CD4") +scater::plotUMAP(sce, color_by = "CD4") ``` ```{r plot CD8, live=TRUE} # plot CD8 marker -scater::plotUMAP(sce, - color_by = "CD8") +scater::plotUMAP(sce, color_by = "CD8") ``` @@ -306,8 +312,7 @@ Creating and assigning values to this column can be done with the `$` shortcut, ```{r plot thresholds} sce$threshold_celltype <- adt_df$celltype -scater::plotUMAP(sce, - color_by = "threshold_celltype") + +scater::plotUMAP(sce, color_by = "threshold_celltype") + guides(color = guide_legend(title = "Cell type")) ``` @@ -490,12 +495,12 @@ We will also take this time to dive a bit deeper into the steps that `SingleR` p As mentioned, the first step is training the model, during which we identify the genes that will be used for the correlation analysis later. While this step is not particularly slow, if we were classifying multiple samples, we would not want to have to repeat it for every sample. -To do the training, we will use the `trainSingleR()` function. +To do the training, we will use the `SingleR::trainSingleR()` function. For this we will start with our reference and the labels we want to train the model with. We can then specify the method used to select the genes that will be used for classification. The default method is `"de"`, which performs a differential expression analysis for each pair of labels, but we could also use `"sd"` to select the genes which are most variable across labels, or `"all"` to use all genes. -If we want to get really fancy, we could even provide a specific list of genes to use. +If we wanted to get really fancy, we could even provide a specific list of genes to use with the `restrict` argument. We should note here that the reference dataset for `SingleR` does not need to be from a compendium like `celldex`! If you have any well-classified dataset that you want to use as a reference, you can, as long as you can create a gene by sample expression matrix and a vector of cell types for each sample. @@ -503,7 +508,8 @@ You will want to ensure that the cell types you expect to see in your sample are You can even use a previously-annotated `SingleCellExperiment` as a reference for a new dataset. For more details about custom references, see the [OSCA chapter on cell type annotation](http://bioconductor.org/books/3.19/OSCA.basic/cell-type-annotation.html#using-custom-references) -We do want to be sure that the genes selected for the model will be among those present in our SCE object, so we will use the `restrict` argument with a vector of the genes in our SCE. +We do want to be sure that the genes included in the model will be among those present in our SCE object, so we will use the `test.genes` argument with a vector of the genes in our SCE. +Otherwise, the function will assume that both the reference and test data set have the same genes in the same order (which they don't!), and the next step of actually classifying cell types wouldn't work. This step would happen automatically with the `SingleR::SingleR()` function, but we need to add it manually for this use case. @@ -512,10 +518,10 @@ This step would happen automatically with the `SingleR::SingleR()` function, but singler_finemodel <- SingleR::trainSingleR( monaco_ref, # reference dataset labels = monaco_ref$label.fine, # labels for training dataset + # consider only genes in the sce object + test.genes = rownames(sce), # use DE to select genes (default) genes = "de", - # only use genes in the sce object - restrict = rownames(sce), # parallel processing BPPARAM = BiocParallel::MulticoreParam(4) ) @@ -611,8 +617,10 @@ Now that we have that set up, we can plot using our collapsed and ordered cell t ```{r plot collapsed, live=TRUE} sce$celltype_collapsed <- collapsed_labels -scater::plotUMAP(sce, - color_by = "celltype_collapsed") +scater::plotUMAP( + sce, + color_by = "celltype_collapsed" +) ``` diff --git a/scripts/link-data.sh b/scripts/link-data.sh index aaed38f2..7fde6ea4 100644 --- a/scripts/link-data.sh +++ b/scripts/link-data.sh @@ -63,6 +63,8 @@ link_locs=( RNA-seq/data/gastric-cancer/salmon_quant/SRR585577 RNA-seq/QC/gastric-cancer/fastp/SRR585571 RNA-seq/QC/gastric-cancer/fastqc/SRR585571 + RNA-seq/QC/gastric-cancer/fastp/SRR585574 + RNA-seq/QC/gastric-cancer/fastqc/SRR585574 RNA-seq/data/NB-cell/NB-cell_metadata.tsv RNA-seq/data/NB-cell/salmon_quant RNA-seq/data/leukemia/SRP049821_metadata.tsv @@ -86,6 +88,7 @@ link_locs=( scRNA-seq-advanced/analysis/mouse-liver/markers scRNA-seq-advanced/data/PBMC-TotalSeqB/raw_feature_bc_matrix scRNA-seq-advanced/data/PBMC-TotalSeqB/normalized/PBMC_TotalSeqB_normalized_sce.rds + scRNA-seq-advanced/data/glioblastoma-10x/filtered_feature_bc_matrix scRNA-seq-advanced/data/glioblastoma-10x/raw_feature_bc_matrix scRNA-seq-advanced/data/ewing-sarcoma/annotations/ewing_sarcoma_sample_metadata.tsv scRNA-seq-advanced/data/ewing-sarcoma/processed diff --git a/scripts/make-live.R b/scripts/make-live.R index 14cee77a..322c2819 100644 --- a/scripts/make-live.R +++ b/scripts/make-live.R @@ -14,11 +14,15 @@ library(optparse) # Set up optparse options option_list <- list( make_option( - opt_str = "--notebook", type = "character", - help = "The notebook file to process."), + opt_str = "--notebook", + type = "character", + help = "The notebook file to process." + ), make_option( - opt_str = "--render", type = "character", - default = "TRUE", help = "Needs a 'TRUE/FALSE' to determine whether the markdown::render() step will be run." + opt_str = "--render", + type = "character", + default = "TRUE", + help = "Needs a 'TRUE/FALSE' to determine whether the markdown::render() step will be run." ) ) @@ -26,7 +30,7 @@ option_list <- list( opt <- parse_args(OptionParser(option_list = option_list)) # Check that render is TRUE or FALSE (or an obvious variant) -if (! tolower(opt$render) %in% c("true", "false", "t", "f")){ +if (!tolower(opt$render) %in% c("true", "false", "t", "f")) { stop("`--render` option must be TRUE or FALSE") } @@ -34,13 +38,17 @@ if (! tolower(opt$render) %in% c("true", "false", "t", "f")){ render <- as.logical(opt$render) # Check that the file is an Rmarkdown file: -if (! stringr::str_detect(opt$notebook, "\\.Rmd$")){ +if (!stringr::str_detect(opt$notebook, "\\.Rmd$")) { stop(opt$notebook, " is not a `.Rmd` notebook") } # Install exrcise package if needed. -if (!"exrcise" %in% installed.packages()){ - remotes::install_github("AlexsLemonade/exrcise", dependencies = TRUE, upgrade = "never") +if (!"exrcise" %in% installed.packages()) { + remotes::install_github( + "AlexsLemonade/exrcise", + dependencies = TRUE, + upgrade = "never" + ) } diff --git a/scripts/render-live.sh b/scripts/render-live.sh index 88c54a40..95502bc1 100644 --- a/scripts/render-live.sh +++ b/scripts/render-live.sh @@ -37,6 +37,7 @@ files=( scRNA-seq-advanced/03-differential_expression.Rmd scRNA-seq-advanced/04-gene_set_enrichment_analysis.Rmd scRNA-seq-advanced/05-aucell.Rmd + spatial/01-spatial_intro.Rmd # machine-learning/01-openpbta_heatmap.Rmd # machine-learning/02-openpbta_consensus_clustering.Rmd # machine-learning/03-openpbta_PLIER.Rmd diff --git a/spatial/.gitignore b/spatial/.gitignore new file mode 100644 index 00000000..a5e65c23 --- /dev/null +++ b/spatial/.gitignore @@ -0,0 +1,4 @@ +# ignore data directory +data + + diff --git a/spatial/01-spatial_intro.Rmd b/spatial/01-spatial_intro.Rmd new file mode 100644 index 00000000..d6e67cfd --- /dev/null +++ b/spatial/01-spatial_intro.Rmd @@ -0,0 +1,426 @@ +--- +title: "Importing, processing, and exploring Visium data" +author: Data Lab for ALSF +date: 2026 +output: + html_notebook: + toc: true + toc_float: true +--- + +## Objectives + +- Read Visium data into R +- Filter to spots overlapping tissue +- Calculate quality control measures on spatial transcriptomic data +- Remove likely low-quality spots with `SpotSweeper()` +- Normalize spatial expression data +- Visualize spatial transcriptomic data + +## Introduction + +In this notebook, we'll learn some basic import, processing, and visualization of a Visium data set, starting with the Space Ranger output. + +The data we'll use in this notebook is from an anaplastic Wilms Tumor sample in the [Single-cell Pediatric Cancer Atlas](https://scpca.alexslemonade.org/projects/SCPCP000006). +This sample was processed using first-generation 3' Visium technology, and it was quantified with `Space Ranger 1.3.1`. + +## Set up + +To begin, we'll load the core library for spatial transcriptomic objects in Bioconductor, `SpatialExperiment`. +We won't be doing any calculations involving random numbers here, so we don't need to set a seed. + +```{r libraries} +# Load libraries + +# The main class we use for spatial transcriptomic data +library(SpatialExperiment) + +# To support plotting +library(patchwork) + +# Set ggplot2 theme +ggplot2::theme_set(ggplot2::theme_bw()) +``` + +### Directories and files + +Next we'll define paths and files. + +```{r inputs, live=TRUE} +# define sample id we'll be analyzing +sample_id <- "SCPCS000190" + +# main data directory for this sample +data_dir <- file.path("data/wilms-tumor", sample_id) + +# Path to Space Ranger output +raw_spaceranger_dir <- file.path(data_dir, "outs") + +# reference data directory +ref_dir <- file.path("data/reference") + +# Path to mitochondrial genes table +mito_file <- file.path(ref_dir, "hs_mitochondrial_genes.tsv") +``` + + +```{r outputs} +# Directory and file to save output +normalized_dir <- file.path(data_dir, "normalized") + +# create the directory if it does not exist +fs::dir_create(normalized_dir) + +# output RDS file for normalized Spatial Experiment (SPE) object +output_spe_file <- file.path( + normalized_dir, + "SCPCS000190_normalized.rds" +) +``` + + + +## Reading Space Ranger data + +We'll use the `VisiumIO` package to read this data in. +The directory structure typically looks something like this, with differences for different technology versions. + +```markdown +_add file tree here with standard visium output_ +``` + +Let's see what we have: + +```{r dir-spaceranger, live = TRUE} +dir(raw_spaceranger_dir) +``` + +The filtered/raw matrix directories are analogous to we'd get from `Cell Ranger` for single-cell sequencing, but the `spatial` directory contains spatial information, including images, that augment the data with its full spatial context. +Without this directory, we could still read in the filtered/raw and make an `SCE`, but you wouldn't have the spatial info. + +Along those lines, let's have a closer look at what's in the `spatial` directory: + +```{r dir-spatial, live = TRUE} +dir( + file.path(raw_spaceranger_dir, "spatial") +) +``` + +Placeholder to describe each image file: + +- `"aligned_fiducials.jpg"` +- `"detected_tissue_image.jpg"` +- `"tissue_hires_image.png"` +- `"tissue_lowres_image.png"` +- `"tissue_positions_list.csv"` +- `"scalefactors_json.json"` + +It's worth mentioning that this is a slightly older technology than the current Visium `CytAssist` image, which means there is no specific `Cytassist` image. +This won't affect our analysis, but worth noting when reading in since by default the functions assume that image is present. + + +We'll import the raw, not filtered, version of the data here so that we can see full processing steps and save it to a variable called `spe`, using the package `VisiumIO`. + +```{r import-visium, live = TRUE} +# path to quantified RNA 10X output +rna_dir <- file.path(raw_spaceranger_dir, "raw_feature_bc_matrix") + +# path to spatial 10X output +spatial_dir <- file.path(raw_spaceranger_dir, "spatial") + +spe <- VisiumIO::import( + VisiumIO::TENxVisium( + # path that contains the sequencing data + resources = rna_dir, + # path that contains the spatial data + spatialResource = spatial_dir, + # which image(s) to import - use lowres to save some memory/space + # we also need to override the default since it assumes there is a CytAssist image + images = "lowres" + ) +) + +# print the spe +spe +``` + +Let's get to know our `SPE` object! +It's essentially an `SCE`, but with a few more bells and whistles. +This means there are some functions we'll use to explore it that you might recognize, but there will also be some new ones only used for `SPE` objects. + +First, note that we have 4992 spots - this is not a random value. +It's the number of spots on a Visium 6.5 x 6.5 slide. + +Like `SCE`s, we have slots like `assays` for count matrices, `rowData` for feature metadata, and `colData` for _spot_ metadata - recall, our experimental units are not individual cells in spatial transcriptomics. +Let's take a tour: + +```{r spe-rowdata} +rowData(spe) +``` + +```{r spe-assays} +counts(spe)[1:10, 1:10] +``` +Again, for the colData`, we're looking at spots and not cells. +With this technology, where each spot is 55 µm, we can expect there's somewhere between 1-10 cells per spot. + +```{r spe-coldata, live = TRUE} +colData(spe) +``` +Check out that `in_tissue` column. +It contains 0/1 because we read in the raw `Space Ranger` output, which includes all spots including those which don't actually overlap tissue. +Ultimately, we won't want to analyze the spots that are not on top of tissue, so we'll have to deal with that. + +Unlike `SCE`s, we have spatial information in a dedicated `spatialCoords` slot which contains the x/y coordinates on the slide: +```{r spe-spatialcoords, live = TRUE} +# handy function from our code package SpatialExperiment +# literally is giving x/y coordinates +spatialCoords(spe) |> head() +``` + +We also have a slot that just holds the images, `imgData`, although looking at it directly isn't very interesting. +But, this stored information will help us make figures! + +```{r spe-imgdata, live = TRUE} +imgData(spe) +``` + +### Introduction to spatial data visualization + +Before we get further, let's go ahead and actually look at our data. +Because the spatial and image information is contained in the object, we can plot directly from this object using the viz package [`ggspavis`](https://bioconductor.posit.co/packages/release/bioc/vignettes/ggspavis/inst/doc/ggspavis_overview.html). + + +```{r plot-visium, live = TRUE} +#| fig.width: 7 + +ggspavis::plotVisium(spe) +``` + +Here, we see the spots overlaid on the slide as well as slide boundaries. +You'll notice that every single spot is shown, including ones that don't overlap tissue directly - indeed, all 4992 spots are present in the `SPE` right now! + +By default this shows the spots, but we can hide them and zoom into the tissue on the slide. +This is a helpful way to pop up the H&E for side-by-side comparisons with other plots you might make. + +```{r plot-visium-zoom, live = TRUE} +#| fig.width: 5 + +ggspavis::plotVisium( + spe, + spots = FALSE, # don't show the spots + zoom = TRUE # zoom into the slide area +) +``` + +TODO: save this text for when we actually plot genes? +Let's take a moment to chat about Wilms Tumor - these tumors in the developing kidney are often composed of a couple histologic compartments, blastemal, stromal (with mesenchymal biology), and sometimes epithelial components. +In this tissue section, we can see two major components: the bluer indicating densely cellular, ECM-poor regions consistent with blastema, and paler pink regions consistent with stromal tissue. + + +There's another function in `ggspavis` called `plotCoords` which hides the H&E to just show the spots; this plot is not currently very compelling without any colors, but we'll add those soon! + +```{r plot-coords, live = TRUE} +#| fig.width: 7 + +# plot just the spots +ggspavis::plotCoords(spe) +``` + +The spot layout looks a bit different. +By default, this function will only plot spots that overlay tissue, even if those spots are still in the `SPE`. + +We can override this if we want, but really what we want to do is actually remove those spots since they are entirely uninformative - we'll do this in the next section. + + +## Filtering empty spots + +We only care about spots on top of tissue, so let's begin by removing the `in_tissue = 0` spots. +Note that if we had read in the `filtered_feature_bc_matrix` instead, the data would already be filtered to only `1` in this column, but we need to take this step because we read in the `raw`. +We can make a rough analogy, that this analysis step is like filtering empty droplets to retain only droplets with cells in scRNA-seq, but here we're retaining only spots over tissue. + +We can visualize which spots those are, and we'll do it over the H&E to clearly see the relationship. +We'll use the `annotate` argument, but `plotVisium` sees that this column is an integer and forces it to use a continuous color scale; we'll go ahead and make it a factor version of it to plot with. + +```{r plot_in_tissue} +#| fig.width: 7 + +# make a factor version of this column to plot with, specifying "No" and "Yes" plot labels +spe$in_tissue_factor <- factor(spe$in_tissue, levels = c(0, 1), labels = c("No", "Yes")) + +ggspavis::plotVisium( + spe, + annotate = "in_tissue_factor", + # specify a custom palette + pal = c("Yes" = "yellow", "No" = "red") +) + + # use ggplot2::guides() to override legend titles in ggspavis + ggplot2::guides( + fill = ggplot2::guide_legend( + title = "Spot overlaps tissue", + override.aes = list(size = 2) + ) + ) +``` + +- The purple tissue overhang on the left isn't colored at all - indeed, there aren't spots at those coordinates outside the slide +- Red points are those to filter out - they are uninformative since they don't overlap tissue. + +```{r filter_in_tissue, live = TRUE} +# keep only spots that are in the tissue and save to filtered_spe +filtered_spe <- spe[, spe$in_tissue == 1] + +# print resulting filtered_spe +filtered_spe +``` + + +Now, we're down to 4120 spots from the original 4992, but that's still a pretty good amount. +But not all of these spots are necessarily good quality, so we'll want to do some additional QC filtering next. + +## Filtering low-quality spots + +### Filtering with global QC thresholds + +As a first step towards filtering, we can borrow some approaches from scRNA-seq and calculate some quality-control measures. +Let's do that and have a look - we'll get our mitochondrial genes for QC calculations and we'll use `scran::addPerCellQC()` like we might for single-cell. + +```{r get mitochondrial genes} +# read in a table of mitochondrial genes and extract ids +mito_genes <- readr::read_tsv(mito_file) |> + # filter to only the genes that are found in our dataset + dplyr::filter(gene_id %in% rownames(filtered_spe)) |> + # create a vector from the gene_id column + dplyr::pull(gene_id) +``` + +```{r calculate qc, live = TRUE} +filtered_spe <- scuttle::addPerCellQC( + filtered_spe, + subsets = list(mito = mito_genes) +) + +# print resulting colData to see QC stats +colData(filtered_spe) |> head() +``` + +`ggspavis` comes with a helpful QC plotter (makes histograms by default but has a couple more options!). +These are `ggplot2` objects, so we can use `ggplot2` code with them like add a title to each. + +```{r global qc distributions} +#| fig.width: 12 +#| fig.height: 4 + +# Extract the colData into a data frame for plotting +coldata_df <- colData(filtered_spe) |> + as.data.frame() |> + # relocate barcodes to an actual column instead of rownames + tibble::rownames_to_column("barcode") + +# Define density plots for each statistic +qc_plot_sum <- ggplot2::ggplot(coldata_df) + + ggplot2::aes(x = sum) + + ggplot2::geom_density(fill = "lightblue") + + ggplot2::labs(title = "Total unique UMIs") + +qc_plot_detected <- ggplot2::ggplot(coldata_df) + + ggplot2::aes(x = detected) + + ggplot2::geom_density(fill = "steelblue") + + ggplot2::labs(title = "Total detected genes") + +qc_plot_mito <- ggplot2::ggplot(coldata_df) + + ggplot2::aes(x = subsets_mito_percent) + + ggplot2::geom_density(fill = "slateblue") + + ggplot2::labs(title = "Mitochondrial %") + + +# Plot together with patchwork +qc_plot_sum + qc_plot_detected + qc_plot_mito +``` + +These distributions all look unimodal without too much skew, which is different from how distributions from single-cell data might look. + +Placeholder for discussion about differences/similarities in technical artifacts between data types: + +- One of the reasons we filter on these metrics in single cell is due to potential differences in cell capture across droplets +- This isn't a factor in spatial where QC stats are calculated per spot aka for aggregates of cells, extremes end up getting smoothed out + - Differences in capture aren't due to heterogeneity from droplet capture but because of tissue biology/prep: + - We have distinct regions of tissue or groups of cells that have less genes detected than other ones (biology) or because the tissue didn't lay on the slide completely flat or wasn't completely permeabilized in all spots equally + +For example, let's consider the mitochondria percent distribution: + +- We do see a long right-tail for mitochondrial percentages, but the values are all really low which doesn't suggest any major quality issues. +- The lower values here vs in single-cell data make sense because cells weren't stressed in the same way they would be for single-cell library prep; more likely to tell you about the tissue quality itself (biology) and not spot quality (technical) + +Let's plot the same stats on the slide. +We'll include the H&E plot here as well for an immediate comparison. + +```{r global qc coords} +#| message: FALSE +#| fig.width: 10 + +he_plot <- ggspavis::plotVisium(filtered_spe, spots = FALSE, zoom = TRUE) + + ggplot2::ggtitle("H&E") + + # add styling to match other panels + ggplot2::theme( + plot.title = ggplot2::element_text( + hjust = 0.5, + margin = ggplot2::margin(0, 0, 0.1, 0) + ), + panel.border = ggplot2::element_rect(color = "black", linewidth = 0.25) + ) + +qc_spots_sum <- ggspavis::plotCoords(filtered_spe, annotate="sum", point_size = 1) + + # use distinct palette + ggplot2::scale_color_distiller(palette = "Blues", direction = 1) + + ggplot2::ggtitle("Total unique UMIs") + +qc_spots_detected <- ggspavis::plotCoords(filtered_spe, annotate="detected", point_size = 1) + + # use distinct palette + ggplot2::scale_color_distiller(palette = "YlOrRd", direction = 1) + + ggplot2::ggtitle("Total detected genes") + +qc_spots_mito <- ggspavis::plotCoords(filtered_spe, annotate="subsets_mito_percent", point_size = 1) + + # use distinct palette + ggplot2::scale_color_distiller(palette = "Greens", direction = 1) + + ggplot2::ggtitle("Mitochondrial %") + +# wrap plots with patchwork into 2x2 grid +patchwork::wrap_plots( + he_plot, + qc_spots_sum, + qc_spots_detected, + qc_spots_mito, + nrow = 2 +) +``` + +Now, we start to see there's more to the story. +There is spatial heterogeneity in these QC stats: + +- the likely stromal regions have distinctly higher mitochondrial percentages (although again, these values are pretty low all around!) and fewer detected UMIs/genes +- the likely blastema regions tend to have more detected UMIs/genes + +This tells us that there is local structure in the data that we might like our QC approach to take into consideration. +Using these global thresholds, we can see that QC is confounded by biology. +If we filter these spots, we'll be removing spots associated with a particular tissue region. +This shows that global thresholds may not be suitable for data with this kind of heterogeneity, so a different approach could be warranted. +Worth noting, this is not strictly a spatial transcriptomics issue; there can be quite a bit of heterogeneity in scRNA-seq data too depending on what was sequenced! +Always plot your data! + +### Filtering with local QC thresholds + + +## Normalization + +## Exploring marker gene expression + + +## Session info + +To conclude, we'll run the `sessionInfo()` command to print out exactly what system setting and R package versions were used to run this code. + +```{r} +sessionInfo() +``` + diff --git a/spatial/spatial.Rproj b/spatial/spatial.Rproj new file mode 100644 index 00000000..98df0873 --- /dev/null +++ b/spatial/spatial.Rproj @@ -0,0 +1,16 @@ +Version: 1.0 + +RestoreWorkspace: No +SaveWorkspace: No +AlwaysSaveHistory: No + +EnableCodeIndexing: Yes +UseSpacesForTab: Yes +NumSpacesForTab: 2 +Encoding: UTF-8 + +RnwWeave: Sweave +LaTeX: pdfLaTeX + +AutoAppendNewline: Yes +StripTrailingWhitespace: Yes diff --git a/workshop-releases.md b/workshop-releases.md index 043037d3..48695a35 100644 --- a/workshop-releases.md +++ b/workshop-releases.md @@ -76,7 +76,7 @@ You can also test the Docker image locally, which may be more convenient for tes To test the Docker image locally, you can use the following commands to pull the Docker image and run launch the container with an RStudio server session: ```bash -docker pull --platform linux/amd64 ccdl/training_rstudio:edge +docker pull ccdl/training_rstudio:edge docker run \ -e PASSWORD={PASSWORD} \ -p 8787:8787 \