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== cNMF ==
== cNMF ==
https://github.com/dylkot/cNMF/ (python)
* https://github.com/dylkot/cNMF/ (python)
 
* It takes a count matrix (N cells X G genes) as input and produces a (K x G) matrix of '''gene expression programs (GEPs)''' and a (N x K) matrix specifying the '''usage of each program''' for each cell in the data.
It takes a count matrix (N cells X G genes) as input and produces a (K x G) matrix of '''gene expression programs (GEPs)''' and a (N x K) matrix specifying the '''usage of each program''' for each cell in the data.
* [https://en.wikipedia.org/wiki/Gene_expression_programming Gene expression programming]
* [https://en.wikipedia.org/wiki/Non-negative_matrix_factorization NMF]


== CancerSubtypes ==
== CancerSubtypes ==

Revision as of 09:51, 21 November 2023

Resource

Workshops

Library preparation

  • Smart-Seq2
  • Drop-Seq
  • Fluidigm-C1

Guideline

Data analysis guidelines for single-cell RNA-seq in biomedical studies and clinical applications

Pipeline

GEO

Applications

Public data

Curated Cancer Cell Atlas

The Curated Cancer Cell Atlas – a comprehensive pan-cancer single-cell RNA-sequencing dataset, website

scRNAseq package

scRNAseq package from Bioconductor.

ExperimentHub package used by scRNAseq, DuoClustering2018 and other packages.

SRAToolkit

sinteractive  --gres=lscratch:800  --cpus-per-task=6 --mem=40g
  # the required memory can be checked by using the '''jobload''' command
  # the required local disk space can be obtained by 'ssh cnXXX' and run 'du -sh /lscratch/JobID'
  #    where the JobID is obtained through the jobload command

module load sratoolkit

fasterq-dump -t /lscratch/$SLURM_JOBID SRR2048331 # 1 min, 3.3G fastq file
fastq-dump --split-3 SRR2048331                   # 2 min 30 sec
pigz -p6 SRR12148211*.fastq  # vs gzip. 6 threads are used here 
                             # do not need to split threads,
                             # do not need to run separately

fastq-dump --split-3 --gzip SRR2048331   # 658M SRR2048331.fastq.gz (vs 626.9Mb in sra format)

fastq-dump --split-3 --gzip SRR12148213  # 862M + 2.5G for fastq.gz files (vs 2Gb in sra format)
                                         # 6.5G + 15G for fastq files
                                         # fastq/fastq.gz = 15/2.5 = 6
                                         # fastq.gz/sr = 3.4/2 = 1.7
# This is equivalent to 
# prefetch SRR12148213; cd SRR12148213
# fastq-dump --split-3 --gzip SRR12148213.sra # 45 min
# The 'prefetch' will create a new folder SRR12148213 and download SRR12148213.sra into it.
# Note the file sizes from _1.fastq and _2.fastq are quite different. 
# Another run. Check number of reads.
$ zcat SRR12148214_1.fastq.gz | wc -l
150635300
$ zcat SRR12148214_2.fastq.gz | wc -l
150635300

fasterq-dump -t /lscratch/$SLURM_JOB_ID \
  --split-files \
  -O /lscratch/$SLURM_JOBID SRR13526458; \
  pigz -p6 /lscratch/$SLURM_JOBID/SRR13526458*.fastq; \
  cp /lscratch/$SLURM_JOBID/SRR13526458*.fastq.gz ~/PDACPDX

sbatch --gres=lscratch:800 --mem=40g --cpus-per-task=6 --time=12:00:00 myscript

And the fasterq-dump's wiki and the help page 

$ fasterq-dump --help

Usage: fasterq-dump [ options ] [ accessions(s)... ]

Parameters:

  accessions(s)                    list of accessions to process


Options:

  -o|--outfile <path>              full path of outputfile (overrides usage
                                     of current directory and given accession)
  -O|--outdir <path>               path for outputfile (overrides usage of
                                     current directory, but uses given
                                     accession)
  -b|--bufsize <size>              size of file-buffer (dflt=1MB, takes
                                     number or number and unit)
  -c|--curcache <size>             size of cursor-cache (dflt=10MB, takes
                                     number or number and unit)
  -m|--mem <size>                  memory limit for sorting (dflt=100MB,
                                     takes number or number and unit)
  -t|--temp <path>                 path to directory for temp. files
                                     (dflt=current dir.)
  -e|--threads <count>             how many threads to use (dflt=6)
  -p|--progress                    show progress (not possible if stdout used)
  -x|--details                     print details of all options selected
  -s|--split-spot                  split spots into reads
  -S|--split-files                 write reads into different files
  -3|--split-3                     writes single reads into special file
     --concatenate-reads           writes whole spots into one file
  -Z|--stdout                      print output to stdout
  -f|--force                       force overwrite of existing file(s)
  -N|--rowid-as-name               use rowid as name (avoids using the name
                                     column)
     --skip-technical              skip technical reads
     --include-technical           explicitly include technical reads
  -P|--print-read-nr               include read-number in defline
  -M|--min-read-len <count>        filter by sequence-lenght
     --table <name>                which seq-table to use in case of pacbio
     --strict                      terminate on invalid read
  -B|--bases <bases>               filter output by matching against given
                                     bases
  -A|--append                      append to output-file, instead of
                                     overwriting it
     --ngc <path>                  <path> to ngc file
     --perm <path>                 <path> to permission file
     --location <location>         location in cloud
     --cart <path>                 <path> to cart file
     --disable-multithreading      disable multithreading
  -V|--version                     Display the version of the program
  -v|--verbose                     Increase the verbosity of the program
                                     status messages. Use multiple times for
                                     more verbosity.
  -L|--log-level <level>           Logging level as number or enum string.
                                     One of
                                     (fatal|sys|int|err|warn|info|debug) or
                                     (0-6) Current/default is warn
     --option-file file            Read more options and parameters from the
                                     file.
  -h|--help                        print this message

"fasterq-dump" version 2.10.9

Note that fastq.gz vs fastq file size is about 1:8 ratio. 

$ ls -lhogt | sed 's/^[^ ][^ ]*  *[^ ][^ ]* //'
 23G May  7 11:35 SRR10156301_2.fastq
 18G May  7 11:35 SRR10156301_1.fastq
3.1G May  7 11:02 SRR10156301_2.fastq.gz
1.4G May  7 11:02 SRR10156301_1.fastq.gz

Simulation

Splatter: Simulation Of Single-Cell RNA Sequencing Data

biorxiv. Genome Biol. 2017, splatter package. We can simulate data using 1) estimated parameters from an existing SingleCellExperiment object, or 2) user's specified parameters. 

suppressPackageStartupMessages({
  library(splatter)
  library(scater)
})

library(scRNAseq)
sce2 <- MacoskoRetinaData()
sce2

ncells <- 1000
params <- splatEstimate(as.matrix(assays(sce2)$counts[, 1:ncells]))
sim <- splatSimulate(params)

comparison <- compareSCEs(list(MacoskoRetina = sce2[, 1:ncells], Splat = sim))
ggplot(comparison$ColData, aes(x = sum, y = detected, colour = Dataset)) +
    geom_point()

splatPop

splatPop – simulating population scale single-cell RNA sequencing data

Preprocess

fastq file format, UMI

  • Single-cell RNA-seq data - raw data to count matrix
    • Reads with different UMI are biological duplicates - each read should be counted.
    • Reads with the same UMI are are technical duplicates - the UMIs should be collapsed to be counted as a single read.
  • Read1 contains barcode (represents cells) and UMI (represents reads), Read2 contains transcript sequence.
  • One case from scruff vignette.

Cell ranger

  • Chromium Single Cell Gene Expression
  • PDX/Multiple Species data. Creating a Reference Package with cellranger mkref
  • pbmc3k. The data is saved in filtered_gene_bc_matrices/hg19 directory with 3 files, barcodes.tsv (45K), genes.tsv (798K) and matrix.mtx (27M).
  • Getting started with Cell Ranger by Dave Tang
  • Workshop from UC Davis
  • *10x genomics single-cell RNAseq analysis from SRA data using Cell Ranger and Seurat, bioinformaticsworkbook.org (useful)
  • how to get/make/find fastq file from cellranger?
  • filtered_feature_bc_matrix vs raw_feature_bc_matrix. "Filtered feature-barcode matrix" contains only detected cellular barcodes. <raw_feature_bc_matrix> contains mainly empty barcodes. That is, the difference is the number of cells.
  • https://hpc.nih.gov/apps/cellranger.html
    • Reference genome 
      module load cellranger/5.0.0
echo $CELLRANGER_REF/refdata-gex-GRCh38-2020-A
# /fdb/cellranger/refdata-gex-GRCh38-2020-A
ls /fdb/cellranger/refdata-gex-GRCh38-2020-A
# fasta  genes  pickle  reference.json  star

ls /fdb/cellranger/
# refdata-cellranger-1.1.0         refdata-cellranger-GRCh38-1.2.0           refdata-cellranger-vdj-GRCh38-alts-ensembl-2.0.0
# refdata-cellranger-1.2.0         refdata-cellranger-GRCh38-3.0.0           refdata-cellranger-vdj-GRCh38-alts-ensembl-3.1.0
# refdata-cellranger-2.0.0         refdata-cellranger-GRCh38-and-mm10-3.1.0  refdata-cellranger-vdj-GRCh38-alts-ensembl-4.0.0
# refdata-cellranger-2020-A        refdata-cellranger-hg19-1.2.0             refdata-cellranger-vdj-GRCh38-alts-ensembl-5.0.0
# refdata-cellranger-2.1.0         refdata-cellranger-hg19-3.0.0             refdata-cellranger-vdj-GRCm38-alts-ensembl-2.2.0
# refdata-cellranger-2.2.0         refdata-cellranger-hg19_and_mm10-1.2.0    refdata-cellranger-vdj-GRCm38-alts-ensembl-3.1.0
# refdata-cellranger-3.0.0         refdata-cellranger-hg19_and_mm10-2.1.0    refdata-cellranger-vdj-GRCm38-alts-ensembl-4.0.0
# refdata-cellranger-3.1.0         refdata-cellranger-hg19-and-mm10-3.0.0    refdata-cellranger-vdj-GRCm38-alts-ensembl-5.0.0
# refdata-cellranger-4.0.0         refdata-cellranger-mm10-1.2.0             refdata-gex-GRCh38-2020-A
# refdata-cellranger-5.0.0         refdata-cellranger-mm10-2.1.0             refdata-gex-GRCh38_and_mm10-2020-A
# refdata-cellranger-ercc92-1.2.0  refdata-cellranger-mm10-3.0.0             refdata-gex-mm10-2020-A

ls /fdb/cellranger/refdata-gex-GRCh38-2020-A/genes
# genes.gtf 
wc -l /fdb/cellranger/refdata-gex-GRCh38-2020-A/genes/genes.gtf
# 2765974 /fdb/cellranger/refdata-gex-GRCh38-2020-A/genes/genes.gtf

# One column contains "gene_name"
head -n 6 /fdb/cellranger/refdata-gex-GRCh38-2020-A/genes/genes.gtf
##description: evidence-based annotation of the human genome (GRCh38), version 32 (Ensembl 98)
##provider: GENCODE
##contact: [email protected]
##format: gtf
##date: 2019-09-05
chr1	HAVANA	gene	29554	31109	.	+	.	gene_id "ENSG00000243485"; gene_version "5"; gene_type "lncRNA"; gene_name "MIR1302-2HG"; level 2; hgnc_id "HGNC:52482"; tag "ncRNA_host"; havana_gene "OTTHUMG00000000959.2";
    • cellranger mkfastq wraps Illumina's bcl2fastq to correctly demultiplex Chromium-prepared sequencing samples and to convert barcode and read data to FASTQ files.
    • cellranger count takes FASTQ files from cellranger mkfastq and performs alignment, filtering, and UMI counting. It uses the Chromium cellular barcodes to generate gene-cell matrices and perform clustering and gene expression analysis.
    • cellranger aggr aggregates results from cellranger count. How does cellranger aggr normalize for sequencing depth among multiple libraries?
    • cellranger reanalyze takes feature-barcode matrices produced by cellranger count or aggr and re-runs the dimensionality reduction, clustering, and gene expression algorithms.
    • The example code will generate an output directory with 
      $ tree s1/outs

├── analysis
│   ├── clustering
│   │   ├── graphclust
│   │   │   └── clusters.csv
│   │   ├── kmeans_10_clusters
│   │   │   └── clusters.csv
│   │   ├── kmeans_2_clusters
│   │   │   └── clusters.csv
│   │   ├── kmeans_3_clusters
│   │   │   └── clusters.csv
│   │   ├── kmeans_4_clusters
│   │   │   └── clusters.csv
│   │   ├── kmeans_5_clusters
│   │   │   └── clusters.csv
│   │   ├── kmeans_6_clusters
│   │   │   └── clusters.csv
│   │   ├── kmeans_7_clusters
│   │   │   └── clusters.csv
│   │   ├── kmeans_8_clusters
│   │   │   └── clusters.csv
│   │   └── kmeans_9_clusters
│   │       └── clusters.csv
│   ├── diffexp
│   │   ├── graphclust
│   │   │   └── differential_expression.csv
│   │   ├── kmeans_10_clusters
│   │   │   └── differential_expression.csv
│   │   ├── kmeans_2_clusters
│   │   │   └── differential_expression.csv
│   │   ├── kmeans_3_clusters
│   │   │   └── differential_expression.csv
│   │   ├── kmeans_4_clusters
│   │   │   └── differential_expression.csv
│   │   ├── kmeans_5_clusters
│   │   │   └── differential_expression.csv
│   │   ├── kmeans_6_clusters
│   │   │   └── differential_expression.csv
│   │   ├── kmeans_7_clusters
│   │   │   └── differential_expression.csv
│   │   ├── kmeans_8_clusters
│   │   │   └── differential_expression.csv
│   │   └── kmeans_9_clusters
│   │       └── differential_expression.csv
│   ├── pca
│   │   └── 10_components
│   │       ├── components.csv
│   │       ├── dispersion.csv
│   │       ├── features_selected.csv
│   │       ├── projection.csv
│   │       └── variance.csv
│   ├── tsne
│   │   └── 2_components
│   │       └── projection.csv
│   └── umap
│       └── 2_components
│           └── projection.csv
├── cloupe.cloupe
├── filtered_feature_bc_matrix
│   ├── barcodes.tsv.gz
│   ├── features.tsv.gz
│   └── matrix.mtx.gz   #  36601 x 25 after dim(Read10X(data.dir ="filtered_feature_bc_matrix"))
├── filtered_feature_bc_matrix.h5
├── metrics_summary.csv
├── molecule_info.h5
├── possorted_genome_bam.bam
├── possorted_genome_bam.bam.bai
├── raw_feature_bc_matrix
│   ├── barcodes.tsv.gz
│   ├── features.tsv.gz
│   └── matrix.mtx.gz  # 36601 x 737280
├── raw_feature_bc_matrix.h5
└── web_summary.html

31 directories, 41 files

      # The 2nd column contains gene symbol
$ zcat s1/outs/filtered_feature_bc_matrix/features.tsv.gz | head -5
ENSG00000243485	MIR1302-2HG	Gene Expression
ENSG00000237613	FAM138A	Gene Expression
ENSG00000186092	OR4F5	Gene Expression
ENSG00000238009	AL627309.1	Gene Expression
ENSG00000239945	AL627309.3	Gene Expression

# R
> x <- Read10X("s1/outs/filtered_feature_bc_matrix")
> dim(x)
[1] 36601    25
> x[1:5, 1:3]
5 x 3 sparse Matrix of class "dgCMatrix"
            AACGTTGGTTAAAGTG-1 AGTAGTCAGAGCTATA-1 ATCTGCCCATACTCTT-1
MIR1302-2HG                  .                  .                  .
FAM138A                      .                  .                  .
OR4F5                        .                  .                  .
AL627309.1                   .                  .                  .
AL627309.3                   .                  .                  .
> length(unique(rownames(x)))
[1] 36601
  • Why do we have Barcodes with Counts that are not Called Cells? Ambient RNA, A barcode is called a cell if Total UMI count > M/10 is called a cell, M = Cell with maximum total UMI counts (99th percentile of the top N)
  • Reads are considered duplicated, if they map to the same gene and have the same UMI. Instead of counting reads we will count number of unique UMIs per gene.

10x Genomics fragment schematic

10x Genomics fragment schematic

10x Molecule Info. This is referenced by the DropletUtils::read10xMolInfo() function.

plot

This is an example (GSM4088780) of part of "web_summary.html" output from running cellranger count 2.1.1.

Cellrangersum.png

We can create a similar plot using the DropletUtils::barcodeRanks() function. Here is the plot from running the example.

BarcodeRanks.png

  • 11,100 barcoded droplets
  • totals =colSums(mat). It has 877 unique values. Range is [0, 2344].
  • lower=100. Only cells satisfies the cutoff will be used in fitting the curve and find left.edge, right.edge, knee, inflection.
  • run.totals 877 values. Range [0, 2344]
  • run.rank 877 values. Range [1, 10900.5]
  • out = 11,100 x 3. Columns include rank, total, fitted
  • left.edge=68, rigth.edge=653. Associate with the x-axis. Relative to x, y.
  • knee=657, inflection=321 (inflection <- 10^(y[right.edge])). Used in the y-axis
  • In cellranger count, y-axis = UMI counts, x-axis = Barcodes.
  • In cellranger count, green points = cells, gray points = background.

emptyDrops() and DropletQC

emptyDrops() 

set.seed(0)
my.counts <- DropletUtils:::simCounts()

# Identify likely cell-containing droplets.
out <- emptyDrops(my.counts)
out
# DataFrame with 11100 rows and 5 columns
#           Total   LogProb    PValue   Limited        FDR
#       <integer> <numeric> <numeric> <logical>  <numeric>
# 1             2        NA        NA        NA         NA
# 2             9        NA        NA        NA         NA
# ...         ...       ...       ...       ...        ...
# 11099       191  -228.763 9.999e-05      TRUE 0.00013799
# 11100       198  -233.043 9.999e-05      TRUE 0.00013799

is.cell <- out$FDR <= 0.01
sum(is.cell, na.rm=TRUE)  # 943

DropletQC – improved identification of empty droplets and damaged cells in single-cell RNA-seq data

estimateAmbience()

estimateAmbience().

This function obtains an estimate of the composition of the ambient pool of RNA based on the barcodes with total UMI counts less than or equal to "lower". The default approach is to assume that all barcodes with total counts less than or equal to "lower" are empty.

It returns a numeric vector of length equal to nrow(m), containing the estimated the transcript proportions in the ambient solution. 

set.seed(0)
my.counts <- DropletUtils:::simCounts()
dim(my.counts) # [1]   100 11100

ambience <- estimateAmbience(my.counts)
head(ambience)
#        GENE1        GENE2        GENE3        GENE4        GENE5        GENE6 
# 0.0002213677 0.0003984648 0.0006331183 0.0008146427 0.0010935705 0.0012042561
length(ambience)  #  [1] 100

removeAmbience()

removeAmbience()

Estimate and remove the ambient profile from a count matrix, given pre-existing groupings of similar cells. This function is largely intended for plot beautification rather than real analysis.

It returns a numeric matrix-like object of the same dimensions as y, containing the counts after removing the ambient contamination. 

ngenes <- 1000
set.seed(1)
ambient <- runif(ngenes, 0, 0.1)  # length = 1000
cells <- c(runif(100) * 10, integer(900))  # length = 1000
y <- matrix(rpois(ngenes * 100, cells + ambient), nrow=ngenes)

# Pretending that all cells are in one group, in this example.
removed <- removeAmbience(y, ambient, groups=rep(1, ncol(y)))
dim(y)       # [1] 1000  100
dim(removed) # [1] 1000  100
summary(as.vector(y))
#    Min. 1st Qu.  Median    Mean 3rd Qu.    Max. 
#  0.0000  0.0000  0.0000  0.5423  0.0000 20.0000 
summary(as.vector(removed))
#    Min. 1st Qu.  Median    Mean 3rd Qu.    Max. 
#   0.000   0.000   0.000   0.515   0.000  20.000

Use tools other than cellranger

See Chapter 3 Processing Raw scRNA-seq Data and Chapter 4 Construction of expression matrix.

(non-UMI) Kallisto-Bustools or Alevin, featureCounts, RSEM. Note Kallisto can do both alignment and gene counting.

(UMI-based) UMI-tools & zUMIs, zUMIs: A fast and flexible pipeline to process RNA sequencing data with UMIs

Imputation

Evaluating imputation methods for single-cell RNA-seq data

PDX

WASP – a web-accessible single cell RNA-Seq processing

WASP – a versatile, web-accessible single cell RNA-Seq processing platform

Seurat*

  • Introduction vignette
    Purpose Code Comment
    Setup the Seurat Object pbmc.data <- Read10X(data.dir = "../data/pbmc3k/filtered_gene_bc_matrices/hg19/")
    pbmc <- CreateSeuratObject(counts = pbmc.data, project = "pbmc3k", min.cells = 3, min.features = 200)
    pbmc[["RNA"]]@counts     		Note the @ operator
    pbmc.data[c("CD3D", "TCL1A", "MS4A1"), 1:30]
    dense.size <- object.size(as.matrix(pbmc.data))
    QC pbmc[["percent.mt"]] <- PercentageFeatureSet(pbmc, pattern = "^MT-")
    VlnPlot(pbmc, features = c("nFeature_RNA", "nCount_RNA", "percent.mt"), ncol = 3)
    plot1 <- FeatureScatter(pbmc, feature1 = "nCount_RNA", feature2 = "percent.mt")
    plot2 <- FeatureScatter(pbmc, feature1 = "nCount_RNA", feature2 = "nFeature_RNA")
    plot1 + plot2
    pbmc <- subset(pbmc, subset = nFeature_RNA > 200 & nFeature_RNA < 2500 & percent.mt < 5)     		select cells with number of features in the range of (200,2500)
    Normalizing the data pbmc <- NormalizeData(pbmc, normalization.method = "LogNormalize", scale.factor = 10000)
    pbmc[["RNA"]]@data     		Note the @ operator
    feature selection pbmc <- FindVariableFeatures(pbmc, selection.method = "vst", nfeatures = 2000)
    top10 <- head(VariableFeatures(pbmc), 10)
    plot1 <- VariableFeaturePlot(pbmc)
    plot2 <- LabelPoints(plot = plot1, points = top10, repel = TRUE)
    plot1 + plot2
    Scaling the data all.genes <- rownames(pbmc)
    pbmc <- ScaleData(pbmc, features = all.genes)
    pbmc[["RNA"]]@scale.data     		Note the @ operator
    Perform linear dimensional reduction pbmc <- RunPCA(pbmc, features = VariableFeatures(object = pbmc))
    print(pbmc[["pca"]], dims = 1:5, nfeatures = 5)
    VizDimLoadings(pbmc, dims = 1:2, reduction = "pca")
    DimPlot(pbmc, reduction = "pca")
    DimHeatmap(pbmc, dims = 1, cells = 500, balanced = TRUE)
    DimHeatmap(pbmc, dims = 1:15, cells = 500, balanced = TRUE)
    Determine the ‘dimensionality’ of the dataset pbmc <- JackStraw(pbmc, num.replicate = 100)
    pbmc <- ScoreJackStraw(pbmc, dims = 1:20)
    Determine the ‘dimensionality’ of the dataset pbmc <- JackStraw(pbmc, num.replicate = 100)
    pbmc <- ScoreJackStraw(pbmc, dims = 1:20)
    JackStrawPlot(pbmc, dims = 1:15)
    ElbowPlot(pbmc)
    Cluster the cells pbmc <- FindNeighbors(pbmc, dims = 1:10)
    pbmc <- FindClusters(pbmc, resolution = 0.5)
    head(Idents(pbmc), 5)
    Non-linear transformation: UMAP/tSNE pbmc <- RunUMAP(pbmc, dims = 1:10)
    DimPlot(pbmc, reduction = "umap")
    Finding differentially expressed features cluster1.markers <- FindMarkers(pbmc, ident.1 = 2, min.pct = 0.25)
    head(cluster1.markers, n = 5)
    cluster5.markers <- FindMarkers(pbmc, ident.1 = 5, ident.2 = c(0, 3), min.pct = 0.25)
    head(cluster5.markers, n = 5)
    pbmc.markers <- FindAllMarkers(pbmc, only.pos = TRUE, min.pct = 0.25, logfc.threshold = 0.25)
    pbmc.markers %>% group_by(cluster) %>% top_n(n = 2, wt = avg_log2FC)
    cluster1.markers <- FindMarkers(pbmc, ident.1 = 0, logfc.threshold = 0.25, test.use = "roc", only.pos = TRUE)
    VlnPlot(pbmc, features = c("MS4A1", "CD79A"))
    VlnPlot(pbmc, features = c("NKG7", "PF4"), slot = "counts", log = TRUE)
    FeaturePlot(pbmc, features = c("MS4A1", "GNLY", "CD3E", "CD14", "FCER1A", "FCGR3A", "LYZ", "PPBP", "CD8A"))
    top10 <- pbmc.markers %>% group_by(cluster) %>% top_n(n = 10, wt = avg_log2FC)
    DoHeatmap(pbmc, features = top10$gene) + NoLegend()
    Assigning cell type identity to clusters new.cluster.ids <- c("Naive CD4 T", "CD14+ Mono", "Memory CD4 T", "B", "CD8 T", "FCGR3A+ Mono", "NK", "DC", "Platelet")
    names(new.cluster.ids) <- levels(pbmc)
    pbmc <- RenameIdents(pbmc, new.cluster.ids)
    DimPlot(pbmc, reduction = "umap", label = TRUE, pt.size = 0.5) + NoLegend()

seurat-wrappers

https://github.com/satijalab/seurat-wrappers

Seurat object

subset genes

<Method 1>: How to subset on genes and preserve idents? 

seurat.object # original seurat object
set.seed(1); genes.use <- sample(1:nrow(seurat.object), 1000)
subset.matrix <- seurat.object[['RNA']]@data[genes.use, ]
seurat.object_sub <- CreateSeuratObject(subset.matrix) 
seurat.object_sub <-  SetIdent(seurat.object_sub, 
                          value=as.factor([email protected]$manual_cluster_tree_addmodulescore))

<Method 2>: subset() function. See Seurat Command List.

<Method 3>: Seurat Weekly NO.2 || 我该如何取子集？(Downsampling)

Seurat methods

QC

QC part explanation: 

> pbmc <- CreateSeuratObject(counts = pbmc.data, project = "pbmc3k", min.cells = 3, min.features = 200)
> pbmc[["percent.mt"]] <- PercentageFeatureSet(pbmc, pattern = "^MT-")
> pbmc2 <- subset(pbmc, subset = nFeature_RNA > 200 & nFeature_RNA < 2500)
> pbmc
An object of class Seurat 
13714 features across 2700 samples within 1 assay 
Active assay: RNA (13714 features, 0 variable features)
> pbmc2
An object of class Seurat 
13714 features across 2695 samples within 1 assay 
Active assay: RNA (13714 features, 0 variable features)
> ind <- which(!colnames(pbmc[['RNA']]) %in% colnames(pbmc2[['RNA']]))
> ind
[1]  427 1060 1762 1878 2568
> tmp <- apply(pbmc[['RNA']]@data, 2, function(x) sum(x > 0)) # detected genes per cell
> range(tmp)
[1]  212 3400
> hist(tmp) # kind of normal
> which(tmp > 2500)
AGAGGTCTACAGCT-1 CCAGTCTGCGGAGA-1 GCGAAGGAGAGCTT-1 
             427             1060             1762 
GGCACGTGTGAGAA-1 TTACTCGAACGTTG-1 
            1878             2568 
> tmp2 <- colSums(pbmc[['RNA']]@data) # total counts per cell
> hist(tmp2)  # kind of normal

Normalization

  • NormalizeData(). log1p(value/colSums[cell-idx] *scale_factor). The scale_factor is 10,000 by default and this gives a good range on the normalized values (at least on the pbmc data). The cpp source code of LogNorm() in preprocessing.R. 
    R> range(log1p(pbmc[['RNA']]@data[,1]/sum(pbmc[['RNA']]@data[,1]) * 10000))
[1] 0.000000 5.753142

R> range(log1p(pbmc[['RNA']]@data[,1]/sum(pbmc[['RNA']]@data[,1]) * 1000))
[1] 0.000000 3.478712

R> range(log1p(pbmc[['RNA']]@data[,1]/sum(pbmc[['RNA']]@data[,1]) * 100000))
[1] 0.000000 8.052868

    And below is a comparison of the raw counts and normalized values: 

    R> pbmc2 <- NormalizeData(pbmc, normalization.method = "LogNormalize", 
                          scale.factor = 10000)
R> cbind(pbmc[['RNA']]@data[i, 1], pbmc2[['RNA']]@data[i, 1])
            [,1]     [,2]
MRPL20         1 1.635873
RPL22          1 1.635873
TNFRSF25       2 2.226555
EFHD2          1 1.635873
NECAP2         1 1.635873
AKR7A2         1 1.635873
CAPZB          1 1.635873
RPL11         41 5.138686
RP5-886K2.3    1 1.635873
PITHD1         1 1.635873

Integration datasets

DE analysis

Cluster-free DE analysis

https://twitter.com/satijalab/status/1635641897560559617 LEMUR and miloDE

Visualization

Azimuth

Azimuth - App for reference-based single-cell analysis

Public data examples

GUI/web applications

Asc-Seurat – Analytical single-cell Seurat-based web application 2021

tidyseurat

tidyseurat, paper in biorxiv & in Bioinformatics

Georges Seurat

Georges Seurat’s 162nd Birthday by Google

Bioconductor packages

Description Packages
Orchestrating Single-Cell Analysis with Bioconductor (OSCA)
simpleSingleCell is an earlier version 		See the list here
single cell RNA-Seq workflow (included in OSCA now) simpleSingleCell
visualization tools for single-cell and Bulk RNA Sequencing dittoSeq
An Introduction to Single-Cell RNA-sequencing Analysis in Bioconductor (中文, based on OSCA ebook） DropletUtils, scran, scater, SingleR, BiocFileCache
An Opinionated Computational Workflow for Single-Cell RNA-seq and ATAC-seq SingleCellExperiment, DropletUtils, scater, iSEE(shiny), scran, limma, edgeR
SplatPop: Simulating Population-Scale Single-Cell RNA-sequencing Data splatter
Importing alevin scRNA-seq counts into R/Bioconductor (check 'Get started') tximeta, SingleCellExperiment, fishpond, scran, Seurat

GUI

NIDAP

https://nidap.nih.gov/ which uses Code workbook (Code Workbook is an application that allows users to analyze and transform data using an intuitive graphical interface) for the interaction. The R/python analysis code can be accessed just like GEO2R from GEO. The background material is available on here.

SCTK2

Interactive analysis of single-cell data using flexible workflows with the Single-Cell Toolkit 2

Partek

Past Webinars

BingleSeq

BingleSeq: a user-friendly R package for bulk and single-cell RNA-Seq data analysis

Cellar

Interactive single-cell data analysis using Cellar 2022

ICARUS

ICARUS – an interactive web server for single cell RNA-seq analysis

How many cells are in the human body?

30-40 trillion (1012) cells

Power analysis

  • powsimR: power analysis for bulk and single cell RNA-seq experiments. It is time-consuming and error-prone to install lots of required packages. Better to have a Docker image. See my note here.
  • SCOPIT: sample size calculations for single-cell sequencing experiments
  • POWSC - Simulation, power evaluation and sample size recommendation for single-cell RNA-seq, Su 2020. 
    library(devtools)
install_github("haowulab/SC2P", build_vignettes=TRUE)
install_github("suke18/POWSC", build_vignettes = T, dependencies = T)

library(POWSC)
data("es_mef_sce")
sce = es_mef_sce[, colData(es_mef_sce)$cellTypes == "fibro"]
est_Paras = Est2Phase(sce)

sim_size = c(100, 400, 1000) # A numeric vector
pow_rslt = runPOWSC(sim_size = sim_size, 
                    est_Paras = est_Paras,
                    per_DE=0.05, 
                    DE_Method = "MAST", 
                    Cell_Type = "PW") 
# Note, using our previous developed tool SC2P is faster.

packageVersion("POWSC")  # [1] '0.1.0'
help(package="POWSC")
plot.POWSC(pow_rslt, Form="II", Cell_Type = "PW") 
summary.POWSC(pow_rslt, Form="II", Cell_Type = "PW")
#      (0,10] (10,20] (20,40] (40,80] (80,160] (160,Inf]
# 100  0.0171  0.1270  0.2877  0.2740   0.4909    0.6765
# 400  0.3200  0.5373  0.6486  0.7436   0.7959    0.8429
# 1000 0.5534  0.7424  0.8095  0.9067   0.9815    0.9077

Mitochondrion

Spike-in

  • RNA spike-in.
    • In the context of RNA-seq, “spike-in” data refers to the addition of a known quantity of synthetic RNA to a sample before sequencing. This synthetic RNA, known as an RNA spike-in, has a known sequence and is used to calibrate measurements and normalize the data. By comparing the observed signal from the spike-in RNA to its known quantity, researchers can correct for technical variability and improve the accuracy of their measurements.
    • An RNA spike-in is an RNA transcript of known sequence and quantity used to calibrate measurements in RNA hybridization assays, such as DNA microarray experiments, RT-qPCR, and RNA-Seq.
  • Spike-in genes are genes that are artificially added to a biological sample, typically at a known concentration, in order to serve as a reference or control for subsequent gene expression analysis.
    • Spike-in genes can be used to help normalize gene expression data and account for variations in sample preparation, handling, and sequencing. By adding known quantities of a spike-in gene to a sample, researchers can determine if the sample preparation and sequencing were performed correctly and if there were any systematic biases or variations in the data.
    • Spike-in genes are typically added to a sample at a constant concentration or a range of concentrations, depending on the specific experimental design. The spike-in genes can be used to monitor technical variability across samples and experiments, and to help ensure that any observed differences in gene expression are due to biological differences and not technical variations.
    • There are different types of spike-in genes that can be used depending on the experimental design, such as synthetic RNA molecules, bacterial genes, or exogenous control genes. Spike-in genes are commonly used in gene expression analysis techniques such as microarrays, RNA sequencing, and quantitative polymerase chain reaction (qPCR).
  • How genes can be added to a biological sample
    • Transfection
    • Microinjection
    • Electroporation
    • Viral delivery
  • Numerical example
    • Let's say a researcher wants to compare the gene expression levels of Sample A and Sample B. The researcher prepares both samples for RNA sequencing, and adds a known amount of a spike-in RNA molecule to both samples. The spike-in RNA molecule is designed to have a known sequence and a concentration of 50,000 copies per microliter.
    • The researcher then performs RNA sequencing on both samples and quantifies the expression levels of the spike-in RNA molecule and the endogenous genes. The following table shows the measured counts of the spike-in RNA molecule and a selected endogenous gene, GENE X, in both Sample A and Sample B:
    Sample Spike-in RNA GENE X
    A 100,000 500
    B 80,000 700
    • The formula for normalizing the counts of gene X to the counts of the spike-in RNA molecule is as follows:
    Normalized count = (Raw count of gene X / Raw count of spike-in RNA molecule) x 
                   (Average counts of spike-in RNA molecule across samples)
    • To account for this technical variation, the researcher can use the counts of the spike-in RNA molecule to normalize the counts of the endogenous genes. For example, the normalized counts of GENE X in Sample A and Sample B would be:
    Sample Normalized GENE X
    A 500/100,000*90,000
    B 700/80,000*90,000
  • scran vignette.
    • We perform some cursory quality control to remove cells with low total counts or high spike-in percentages.
    • An alternative approach is to normalize based on the spike-in counts. The idea is that the same quantity of spike-in RNA was added to each cell prior to library preparation. Size factors are computed to scale the counts such that the total coverage of the spike-in transcripts is equal across cells.
    • If we have spike-ins, we can use them to fit the trend (variance-mean relationship) instead.
  • Spike-in gene concentrations are known, normalization model existing technical variation by utilizing the difference between these known values and the values observed after processing. "Normalization Methods on Single-Cell RNA-seq Data: An Empirical Survey". Lytal 2020

CPM and glmpca package

Feature selection and dimension reduction for single-cell RNA-Seq based on a multinomial model, glmpca package. All codes are available in github. A talk by Hicks 2021.

Transformation

Comparison of transformations for single-cell RNA-seq data

AI

Quality control

Doublet

Downsampling

  • downsampleMatrix() from the scuttle package
    • downsampleMatrix: Downsample a count matrix to a desired proportion
    • Source code of downsampleMatrix
    R> library(scuttle)
R> sce <- mockSCE()
R> summary(colSums(counts(sce)))
   Min. 1st Qu.  Median    Mean 3rd Qu.    Max. 
 344769  366648  374586  373876  382028  396251 

R> downsampled2 <- downsampleMatrix(counts(sce), 
                                    prop = 10000/colSums(counts(sce)), bycol=TRUE)
R> colSums(downsampled2[, 1:5])
Cell_001 Cell_002 Cell_003 Cell_004 Cell_005 
   10000    10000    10000    10000    10000
  • downsampleReads() from the DropletUtils package which imports the scuttle package. 
    R> library(DropletUtils)
# Mocking up some 10X HDF5-formatted data.
R> out <- DropletUtils:::simBasicMolInfo(tempfile())
R> a <- read10xMolInfo(out)
R> names(a)
[1] "data"  "genes"
R> dim(a$data)
[1] 9508    5
R> str(a$genes)
 chr [1:20] "ENSG1" "ENSG2" "ENSG3" "ENSG4" "ENSG5" "ENSG6" "ENSG7" "ENSG8" ...
R> a$data[1:3, ]
DataFrame with 3 rows and 5 columns
         cell       umi gem_group      gene     reads
  <character> <integer> <integer> <integer> <integer>
1        GACT    781899         1         9        10
2        CAGC    747670         1         9         9
3        GTAC    614190         1         6        10
R> unique(a$data$gene)
 [1]  9  6 15  7 12 13  1 20  4  8  3 14 17 19 18 16  2  5 10 11
R> length(unique(a$data$gene))   # 20 genes
[1] 20
R> length(unique(a$data$cell))   # 256 cells
[1] 256

# Downsampling by the reads.
R> b<- downsampleReads(out, barcode.length=4, prop=0.5)
R> dim(b)
[1]  20 256
R> b[1:3, 1:5]
3 x 5 sparse Matrix of class "dgCMatrix"
      AAAA-1 AAAC-1 AAAG-1 AAAT-1 AACA-1
ENSG1      .      1      .      1      5
ENSG2      1      1      1      4      .
ENSG3      1      1      .      1      4
R> class(b)
[1] "dgCMatrix"
attr(,"package")
[1] "Matrix"
R> 4^4   # ATCG for each character of the barcode
[1] 256
R> 20*256  # So the combinations of (cell, gene) are not unique
[1] 5120
R> length(unique(a$data$umi)) # So UMIs are unique
[1] 9508
R> length(unique(a$data$gem_group))
[1] 1

    Change prop parameter. Notice the result of downsampling reads does not change much. 

    R> b1 <- downsampleReads(out, barcode.length=4, prop=.9999)
R> colSums(b1)[1:5]
AAAA-1 AAAC-1 AAAG-1 AAAT-1 AACA-1 
    41     37     28     36     35 
R> b2 <- downsampleReads(out, barcode.length=4, prop=0.2)
R> colSums(b2)[1:5]
AAAA-1 AAAC-1 AAAG-1 AAAT-1 AACA-1 
    37     31     23     31     28 
R> b5 <- downsampleReads(out, barcode.length=4, prop=0.5)
R> colSums(b5)[1:5]
AAAA-1 AAAC-1 AAAG-1 AAAT-1 AACA-1 
    41     37     28     36     35 
R> sum(b1)
[1] 9508
R> sum(b2)
[1] 8245
R> sum(b5)
[1] 9448
  • 7.5.2 Downsampling and log-transforming from OSCA
  • Down_Sample_Matrix()
  • zUMIs
  • Current best practices in single-cell RNA-seq analysis: a tutorial. While downsampling throws away data, it also increases technical dropout rates which CPM and other global scaling normalization methods do not. Thus, downsampling can deliver a more realistic representation of what cellular expression profiles would look like at similar count depths.
  • Huang 2020 Evaluation of Cell Type Annotation R Packages on Single-cell RNA-seq Data
    • downsample features (genes). down_sample_gene(). The idea is sample(gene_list, size = size, replace = FALSE, prob = prob_list) where gene_list is the list of detected genes and size is 5,000/10,000/15,000 and prob_list is the proportion of gene counts (defined by using rowSums()). We randomly downsampled the features from Fluidigm C1 data into 15,000, 10,000 and 5000 input genes, based on the original log count distribution.
    • downsample the reads into 25%, 50%, 75% of the original read depth (with two repetitions) on BAM files, and then realigned files following the method provided by the original manuscript.

Normalization

  • Introduction to single-cell RNA-seq. If using a 3’ or 5’ droplet-based method, the length of the gene will not affect the analysis because only the 5’ or 3’ end of the transcript is sequenced. However, if using full-length sequencing, the transcript length should be accounted for.
  • Bulk normalization methods
    • DESeq2: median of ratios (MoR) approach
    • edgeR: trimmed mean of M values
  • Count models for normalization of single-cell RNA-seq data. See Single Cell Genomics Day 2021

scone package: normalization

Dino

Normalization by distributional resampling of high throughput single-cell RNA-sequencing data Brown 2021

Batch correction

  • Question: scRNA-seq and the batch effects. The biggest problem I found is, since you have no idea what's the real data suppose to be.
  • https://www.biostars.org/p/316204/
  • Over-correction can happen.
    • Single-cell RNA-seq Analysis on NIDAP - Tutorial Part 2 40:00 (nih network is required). Groups differ by a single experimental modification (eg a knockoff gene or cells of similar lineage used to determine differentiation pattern). However batch correction is necessary in cases such as spike-in cells where we expect these cells should be clustered together.
  • Mutual nearest neighbors/MNN. Batch effects in single-cell RNA-sequencing data are corrected by matching mutual nearest neighbors Haghverdi 2018...
    • One assumption is the batch effect is almost orthogonal to the biological subspace. It also assumes the batch effect variation in much smaller than the biological effect variation between different cell types.
    • It is preferable to perform DE analyses using the uncorrected expression values with blocking on the batch, as discussed in Section 11.4... We suggest limiting the use of per-gene corrected values to visualization, e.g., when coloring points on a t-SNE plot by per-cell expression.
    • Sadly the R code of the paper does not have the file ('MAP2.csv') and the code does not work anymore (e.g. mnnCorrect() does not have $angles component). Check out the mnnpy module which still has an option to return angles.
  • SMNN can eliminate the overcorrection between different cell types & allow that batch effects are non-orthogonal to the biological differences. github source code. Note that the package is only available on github and it depends on the python mnnpy module. 
    library(reticulate)
use_python("/home/rstudio/.conda/envs/sc_env/bin/python")
py_config()  # confirm

library("SMNN")
data("data_SMNN")
dim(data_SMNN$batch1.mat)  # [1] 3000  400
data_SMNN$batch1.mat[1:5, 1:5]
dim(data_SMNN$batch2.mat)  # [1] 3000  500
# Maker genes
markers <- c("Col1a1", "Pdgfra", "Ptprc", "Pecam1")
# Corresponding cell type labels for each marker gene
cluster.info <- c(1, 1, 2, 3)
matched_clusters <- unifiedClusterLabelling(data_SMNN$batch1.mat, 
                                            data_SMNN$batch2.mat, 
                                            features.use = markers, 
                                            cluster.labels = cluster.info, 
                                            min.perc = 0.3)

corrected.results <- SMNNcorrect(batches = list(data_SMNN$batch1.mat, data_SMNN$batch2.mat), 
                                 batch.cluster.labels = matched_clusters, 
                                 k=20, 
                                 sigma=1, 
                                 cos.norm.in=TRUE, 
                                 cos.norm.out=TRUE, 
                                 subset.genes=rownames(data_SMNN$batch2.mat))
# NOTE: subset.genes parameter was added to fix a bug when I'm running the function
# [1] "Data preparation ..."
# Error: C stack usage  555609078676 is too close to the limit

> traceback()
3: py_call_impl(callable, dots$args, dots$keywords)
2: mnnpy$utils$transform_input_data(datas = batches.t, cos_norm_in = cos.norm.in, 
       cos_norm_out = cos.norm.out, var_index = as.character(c(0:ncol(batches.t1))), 
       var_subset = subset.index, n_jobs = n.jobs)
1: SMNNcorrect(batches = list(data_SMNN$batch1.mat, data_SMNN$batch2.mat), 
       batch.cluster.labels = matched_clusters, k = 20, sigma = 1, 
       cos.norm.in = TRUE, cos.norm.out = TRUE, subset.genes = rownames(data_SMNN$batch2.mat)[1:100])
  • https://broadinstitute.github.io/2019_scWorkshop/correcting-batch-effects.html
    • Seurat (CCA): RunMultiCCA() seems not available any more. Check out Integration and Label Transfer in Seurat 3 and Introduction to scRNA-seq integration FindIntegrationAnchors() & IntegrateData(). 
      # Create 2D UMAP DR plot for data before integration
# It can be seen there is a need to do batch effect correction
ifnb2 <- ifnb
ifnb2 <- NormalizeData(ifnb2)
ifnb2 <- FindVariableFeatures(ifnb2, selection.method = "vst", nfeatures = 2000)
ifnb2 <- ScaleData(ifnb2, verbose = FALSE)
ifnb2 <- RunPCA(ifnb2, npcs = 30, verbose = FALSE)
ifnb2 <- RunUMAP(ifnb2, reduction = "pca", dims = 1:30)
ifnb2 <- FindNeighbors(ifnb2, reduction = "pca", dims = 1:30)
ifnb2 <- FindClusters(ifnb2, resolution = 0.5)

# Compared to the 2D plot before and after integration
DimPlot(ifnb2, reduction = "umap", group.by = "stim") # not good, ideally two groups
                     # should be mixed together.
                     # It seems the blue color is on top of the salmon color.
DimPlot(immune.combined, reduction = "umap", group.by = "stim")
                     # Two groups are mixed. 

# Compare the cluster plots before and after integration
DimPlot(ifnb2, reduction = "umap", split.by = "stim") # not good, ideally the cluster
                     # pattern should be similar in both groups 
DimPlot(immune.combined, reduction = "umap", split.by = "stim") #good, the cluster
                     # patterns are similar in both groups

      FindConservedMarkers() and [email protected], ?FindConservedMarkers. ident.1 & ident.2 are used to define cells from the @active.ident component. For example, we can it to find genes that are conserved markers irrespective of stimulation/group condition in a certain cluster. Together with FeaturePlot() we can see the gene expression in 2D space (one plot per feature). The DotPlot() function with the split.by parameter can be useful for viewing conserved cell type markers across conditions 

      table([email protected]$stim, [email protected])

table(pbmc_small$groups, [email protected])
    • liger LIGER (NMF): liger:: optimizeALS(). Note that in order to use the R package, it is necessary to install hdf5 or libhdf5-dev library. Vignettes are available on github source but not on CRAN package.
    • Harmony: RunHarmony()

Clustering

# after I upgrade R from 4.0.x to 4.1.x
R> library(DuoClustering2018)
snapshotDate(): 2021-05-18
Warning message:
DEPRECATION: As of ExperimentHub (>1.17.2), default caching location has changed.
  Problematic cache: /Users/XXX/Library/Caches/ExperimentHub
  See https://bioconductor.org/packages/devel/bioc/vignettes/ExperimentHub/inst/doc/ExperimentHub.html#default-caching-location-update

NMF

cNMF

CancerSubtypes

library(CancerSubtypes)
data(GeneExp)
dim(GeneExp) # 1500 x 100
result=ExecuteCNMF(GeneExp,clusterNum=3,nrun=30)
dim(result$distanceMatrix)  # consensus/Similarity returned from nmf()
# [1] 100 100
round(unique(result$distanceMatrix), 3)
# [1] 1.000 0.000 0.167 0.067 0.933 0.100 0.433 0.900 0.833 0.567
#[11] 0.233 0.733 0.267 0.133 0.967 0.633 0.500 0.667 0.200 0.533
sil=silhouette_SimilarityMatrix(result$group, result$distanceMatrix)
plot(sil, col = c("red", "blue", "green"))
# Note that the average silhouette width is computed without considering groups.
# mean(c(.96, .98, 1)) # .98
# mean(sil[, 3]) #  0.9714631  match with the plot
# mean(tapply(sil[, 3], sil[, 1], mean)) # 0.9794998

pca <- prcomp(t(GeneExp))
plot(pca$x[,1], pca$x[, 2], col=result$group, pch=19) # black, red, green

seurat::BuildClusterTree

Significance Analysis for Clustering

Significance Analysis for Clustering with Single-Cell RNA-Sequencing Data 2022

Model

Feature selection

Feature selection revisited in the single-cell era Yang 2021

DE analysis

GSEA

AddModuleScore

library(Seurat)
data("pbmc_small")
cd_features <- list(c('CD79B', 'CD79A', 'CD19', 'CD180', 'CD200', 'CD3D',
  'CD2', 'CD3E', 'CD7', 'CD8A', 'CD14', 'CD1C', 'CD68', 'CD9', 'CD247'))
pbmc_small <- AddModuleScore(
  object = pbmc_small,
  features = cd_features,
  ctrl = 5,
  name = 'CD_Features'
)
head(pbmc_small, 3)

                  orig.ident nCount_RNA nFeature_RNA RNA_snn_res.0.8
ATGCCAGAACGACT SeuratProject         70           47               0
CATGGCCTGTGCAT SeuratProject         85           52               0
GAACCTGATGAACC SeuratProject         87           50               1
               letter.idents groups RNA_snn_res.1 CD_Features1
ATGCCAGAACGACT             A     g2             0    0.4420928
CATGGCCTGTGCAT             A     g1             0    0.4285719
GAACCTGATGAACC             B     g2             0    0.9217770

Tweedieverse

DE analysis with multiple samples

DE analysis with multiple samples. scuttle::aggregateAcrossCells()

Cell type deconvolution

Cell type annotation

GPT-4

Reference-free and cost-effective automated cell type annotation with GPT-4 in single-cell RNA-seq analysis

Cell-level metadata

Cell-level metadata are indispensable for documenting single-cell sequencing datasets Puntambekar 2021. Shiny app. Source code.

GSE137710 is a good example.

Cell subtypes

Single-cell RNA sequencing reveals specific cell subtypes that influence survival and determine molecular subtype classification of ovarian cancers

Some cell types

Cell-cell interactions

  • CellChat
    • Compute the communication probability 
      # load data
load("data_humanSkin_CellChat.rda")
data.input = data_humanSkin$data # normalized data matrix
meta = data_humanSkin$meta # a dataframe with rownames containing cell mata data
cell.use = rownames(meta)[meta$condition == "LS"] # extract the cell names from disease data
data.input = data.input[, cell.use]
meta = meta[cell.use, ]
unique(meta$labels) 

# Create a CellChat object
cellchat <- createCellChat(object = data.input, meta = meta, group.by = "labels")

# Add cell information
cellchat <- addMeta(cellchat, meta = meta)
cellchat <- setIdent(cellchat, ident.use = "labels") # set "labels" as default cell identity
levels(cellchat@idents) # show factor levels of the cell labels
#  [1] "APOE+ FIB"    "FBN1+ FIB"    "COL11A1+ FIB" "Inflam. FIB"  "cDC1"        
#  [6] "cDC2"         "LC"           "Inflam. DC"   "TC"           "Inflam. TC"  
# [11] "CD40LG+ TC"   "NKT" 
groupSize <- as.numeric(table(cellchat@idents))

# Set the ligand-receptor interaction database
CellChatDB <- CellChatDB.human 

# Preprocessing the expression data
CellChatDB.use <- subsetDB(CellChatDB, search = "Secreted Signaling") # use Secreted Signaling
cellchat@DB <- CellChatDB.use
cellchat <- subsetData(cellchat)
cellchat <- identifyOverExpressedGenes(cellchat)
cellchat <- identifyOverExpressedInteractions(cellchat)
cellchat <- projectData(cellchat, PPI.human)

# Compute the communication probability
cellchat <- computeCommunProb(cellchat)
cellchat <- computeCommunProbPathway(cellchat)
str(cellchat@net)
# List of 2
#  $ prob: num [1:12, 1:12, 1:656] 0 0 0 0 0 0 0 0 0 0 ...
#   ..- attr(*, "dimnames")=List of 3
#   .. ..$ : chr [1:12] "APOE+ FIB" "FBN1+ FIB" "COL11A1+ FIB" "Inflam. FIB" ...
#   .. ..$ : chr [1:12] "APOE+ FIB" "FBN1+ FIB" "COL11A1+ FIB" "Inflam. FIB" ...
#   .. ..$ : chr [1:656] "TGFB1_TGFBR1_TGFBR2" "TGFB2_TGFBR1_TGFBR2" "TGFB3_TGFBR1_TGFBR2"  "TGFB1_ACVR1B_TGFBR2" ...
#  $ pval: num [1:12, 1:12, 1:656] 1 1 1 1 1 1 1 1 1 1 ...
#   ..- attr(*, "dimnames")=List of 3
#   .. ..$ : chr [1:12] "APOE+ FIB" "FBN1+ FIB" "COL11A1+ FIB" "Inflam. FIB" ...
#   .. ..$ : chr [1:12] "APOE+ FIB" "FBN1+ FIB" "COL11A1+ FIB" "Inflam. FIB" ...
#   .. ..$ : chr [1:656] "TGFB1_TGFBR1_TGFBR2" "TGFB2_TGFBR1_TGFBR2" "TGFB3_TGFBR1_TGFBR2" "TGFB1_ACVR1B_TGFBR2" ...
str(cellchat@netP)
# List of 2
#  $ pathways: chr [1:13] "MIF" "GALECTIN" "CXCL" "COMPLEMENT" ...
#  $ prob    : num [1:12, 1:12, 1:13] 0 0 0 0 0 0 0 0 0 0 ...
#   ..- attr(*, "dimnames")=List of 3
#   .. ..$ : chr [1:12] "APOE+ FIB" "FBN1+ FIB" "COL11A1+ FIB" "Inflam. FIB" ...
#   .. ..$ : chr [1:12] "APOE+ FIB" "FBN1+ FIB" "COL11A1+ FIB" "Inflam. FIB" ...
#   .. ..$ : chr [1:13] "MIF" "GALECTIN" "CXCL" "COMPLEMENT" ...
sum(cellchat@net$prob)
# [1] 18.65038
    • The error I saw (R 4.1.1, CellChat 1.1.3) 
      cellchat <- netEmbedding(cellchat, type = "functional")
Manifold learning of the signaling networks for a single dataset 
Error in runUMAP(Similarity, min_dist = min_dist, n_neighbors = n_neighbors,  : 
  Cannot find UMAP, please install through pip (e.g. pip install umap-learn or reticulate::py_install(packages = 'umap-learn')).
    • cellchat netEmbedding 运行出错. It provides another solution (no need to install anything using pip) for calculating umap. 
      library(uwot)
cellchat <- netEmbedding(cellchat, umap.method = 'uwot',type = "functional")
    • Error in runUMAP(Similarity, min.dist = 0.3, n.neighbors = k)
    • function netEmbedding is unable to work #167
    • CellChat：细胞间相互作用分析利器 which contains two youtube links to the talks by the author

Integrating datasets

Benchmark

Meta-analysis of (single-cell method) benchmarks reveals the need for extensibility and interoperability

scanpy python package

Trajectory/pseudotime analysis

PAGA: graph abstraction reconciles clustering with trajectory inference through a topology preserving map of single cells

simulation

splatSimulatePaths() from splatter

monocle package for pseudotime analysis

slingshot

slingshot Tools for ordering single-cell sequencing

Spatial Transcriptomics Data Analysis

sincell package

SCell

SCell – integrated analysis of single-cell RNA-seq data

GEVM

Detection of high variability in gene expression from single-cell RNA-seq profiling. Two mouse scRNA-seq data sets were obtained from Gene Expression Omnibus (GSE65525 and GSE60361).

NMFEM

Detecting heterogeneity in single-cell RNA-Seq data by non-negative matrix factorization

Scater

Pre-processing, quality control, normalization and visualization of single-cell RNA-seq data in R

NDRindex

NDRindex: a method for the quality assessment of single-cell RNA-Seq preprocessing data

DEsingle

DEsingle – detecting three types of differential expression in single-cell RNA-seq data

totalVI

Joint probabilistic modeling of single-cell multi-omic data with totalVI

Copy number

Copy number by infercnv

  • https://github.com/broadinstitute/inferCNV/wiki
  • infercnv - Infer Copy Number Variation from Single-Cell RNA-Seq Data
  • We need to install jags software before we install the required package rjags. It seems OK to install this old version 4.3.0 of software on macOS Catalina 10.15.7. To install jags, right click the file 'JAGS-4.3.0.mpkg' and choose 'open'. Then follow the instruction of the installer to complete the installation.
  • https://www.jianshu.com/c/4c982303e19f?order_by=top
    • It seems the goal is to use the CNV data to cluster cells... All cells that clustered together with spiked in controls were labeled "nontumor" whereas the remaining clusters were labels as 'tumor'.
    • 使用inferCNV分析单细胞转录组中拷贝数变异. Or follow the help of plot_cnv(), infercnv::run()... inferCNV会输出一个 "map_metadata_from_infercnv.txt" (I don't see it) 文件用于记录每个细胞的元信息，所有信息都可以从该文件中进行提取。或者使用infercnv::add_to_seurat将信息直接增加到原来的seurat对象中。
    • CNS图表复现13—使用inferCNV来区分肿瘤细胞的恶性与否. 因为10X技术出来的单个细胞的reads数量太少，检测到的基因数量太少，但是inferCNV更新后有一个参数是cutoff ，就很清楚的指出来了：cutoff=1 works well for Smart-seq2, and cutoff=0.1 works well for 10x Genomics 说明它应用于10X单细胞转录组数据是没有问题的。
    • 是否是免疫细胞很容易区分那是否是肿瘤细胞呢？
  • Question: where to download the gene position file? The infercnv_genes_example object has 10338 genes already

Simpler plot 

data(infercnv_data_example) # data.frame 8252 x 20, genes x samples
data(infercnv_annots_example) # data.frame 20 x 1, 10 normal and 10 tumor
data(infercnv_genes_example) # data.frame 10338 x 3
infercnv_genes_example[1:3, ]
#             V2     V3     V4
# WASH7P    chr1  14363  29806
# LINC00115 chr1 761586 762902
# NOC2L     chr1 879584 894689
infercnv_object_example <- infercnv::CreateInfercnvObject(raw_counts_matrix=infercnv_data_example, 
                                                          gene_order_file=infercnv_genes_example,
                                                          annotations_file=infercnv_annots_example,
                                                          ref_group_names=c("normal"))

infercnv_object_example <- infercnv::run(infercnv_object_example,
                                         cutoff=1,
                                         out_dir=tempfile(), 
                                         cluster_by_groups=TRUE, 
                                         denoise=TRUE,
                                         HMM=FALSE,
                                         num_threads=2)
system("ls /var/folders/2q/slryb0rx4tj97t66v7l6pwvr_z6g3s/T//RtmpEuGgVy/file182d24b626381")
# 01_incoming_data.infercnv_obj                     infercnv.21_denoised.observations_dendrogram.txt
# 02_reduced_by_cutoff.infercnv_obj                 infercnv.21_denoised.png
# 03_normalized_by_depth.infercnv_obj               infercnv.21_denoised.references.txt
# 04_logtransformed.infercnv_obj                    infercnv.heatmap_thresholds.txt
# 08_remove_ref_avg_from_obs_logFC.infercnv_obj     infercnv.observation_groupings.txt
# 09_apply_max_centered_expr_threshold.infercnv_obj infercnv.observations.txt
# 10_smoothed_by_chr.infercnv_obj                   infercnv.observations_dendrogram.txt
# 11_recentered_cells_by_chr.infercnv_obj           infercnv.png
# 12_remove_ref_avg_from_obs_adjust.infercnv_obj    infercnv.preliminary.heatmap_thresholds.txt
# 14_invert_log_transform.infercnv_obj              infercnv.preliminary.observation_groupings.txt
# 15_no_subclustering.infercnv_obj                  infercnv.preliminary.observations.txt
# 21_denoise.NF_NA.SD_1.5.NL_FALSE.infercnv_obj     infercnv.preliminary.observations_dendrogram.txt
# expr.infercnv.21_denoised.dat                     infercnv.preliminary.png
# expr.infercnv.dat                                 infercnv.preliminary.references.txt
# expr.infercnv.preliminary.dat                     infercnv.references.txt
# infercnv.21_denoised.heatmap_thresholds.txt       preliminary.infercnv_obj
# infercnv.21_denoised.observation_groupings.txt    run.final.infercnv_obj
# infercnv.21_denoised.observations.txt

# Optional, what's the purpose of this plot function
#           when the last command already generated the heatmap?
plot_cnv(infercnv_object_example,
         out_dir=tempfile(),
         obs_title="Observations (Cells)",
         ref_title="References (Cells)",
         cluster_by_groups=TRUE,
         x.center=1,
         x.range="auto",
         hclust_method='ward.D',
         color_safe_pal=FALSE,
         output_filename="infercnv",
         output_format="png",
         png_res=300,
         dynamic_resize=0)
# ...
list.files("/var/folders/2q/slryb0rx4tj97t66v7l6pwvr_z6g3s/T//RtmpEuGgVy/file182d25df0c4ed")
# [1] "infercnv.heatmap_thresholds.txt"      "infercnv.observation_groupings.txt"  
# [3] "infercnv.observations_dendrogram.txt" "infercnv.observations.txt"           
# [5] "infercnv.png"                         "infercnv.references.txt"

More complicated plot 

infercnv_obj = CreateInfercnvObject(
  raw_counts_matrix=system.file("extdata", "oligodendroglioma_expression_downsampled.counts.matrix.gz", package = "infercnv"),
  annotations_file=system.file("extdata", "oligodendroglioma_annotations_downsampled.txt", package = "infercnv"),
  delim="\t", gene_order_file=system.file("extdata", "gencode_downsampled.EXAMPLE_ONLY_DONT_REUSE.txt", package = "infercnv"),
  ref_group_names=c("Microglia/Macrophage","Oligodendrocytes (non-malignant)")) 

infercnv_obj = infercnv::run(infercnv_obj,
                             cutoff=1, # cutoff=1 works well for Smart-seq2, and cutoff=0.1 works well for 10x Genomics
                             out_dir=tempfile(), 
                             cluster_by_groups=TRUE, 
                             denoise=TRUE,
                             HMM=TRUE) # HMM=TRUE will increase lots of computation
# 	STEP 1: incoming data
# 	STEP 02: Removing lowly expressed genes
# ...
# STEP 22: Denoising
# There were 50 or more warnings (use warnings() to see the first 50)

x <- read.delim(system.file("extdata", "oligodendroglioma_annotations_downsampled.txt", package = "infercnv"), header=F)
dim(x)  #  [1] 184   2;   184 cells only
unique(x[,2])  # 6 groups. Two of them represent references cells and the other 4 are the obs cells.
# [1] "Microglia/Macrophage"             "Oligodendrocytes (non-malignant)"
# [3] "malignant_MGH36"                  "malignant_MGH53"
# [5] "malignant_97"                     "malignant_93"
x[1:3, ]
#             V1                   V2
# 1 MGH54_P2_C12 Microglia/Macrophage
# 2 MGH36_P6_F03 Microglia/Macrophage
# 3 MGH53_P4_H08 Microglia/Macrophage

xx <- read.delim(system.file("extdata", "gencode_downsampled.EXAMPLE_ONLY_DONT_REUSE.txt", package = "infercnv"), header=F)
dim(xx)  #  [1] 10338     4
xx[1:2, ]
#          V1   V2     V3     V4
# 1    WASH7P chr1  14363  29806
# 2 LINC00115 chr1 761586 762902

Infercnv.png

Cell-cell interactions and communication

Deciphering cell–cell interactions and communication from gene expression Armingol 2021

Chromatin/scATAC-seq data

Signac

Signac is an extension of Seurat for the analysis, interpretation, and exploration of single-cell chromatin datasets.

GPU

NVIDIA Researchers Use AI to Spot Active Areas in Cell DNA

Single Cells and Single Nuclei

Webinars

Brain cells

PDAC

Single-cell analysis support the use of PDAC organoids as a clinically relevant model for in vitro studies of tumor heterogeneity 2021

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