--- title: "Workflow steps overview" author: "Author: Daniela Cassol (danielac@ucr.edu) and Thomas Girke (thomas.girke@ucr.edu)" date: "Last update: `r format(Sys.time(), '%d %B, %Y')`" output: BiocStyle::html_document: toc_float: true code_folding: show BiocStyle::pdf_document: default package: systemPipeR vignette: | %\VignetteEncoding{UTF-8} %\VignetteIndexEntry{systemPipeR: Workflow design and reporting generation environment} %\VignetteEngine{knitr::rmarkdown} fontsize: 14pt bibliography: bibtex.bib editor_options: chunk_output_type: console type: docs --- ```{r style, echo = FALSE, results = 'asis'} BiocStyle::markdown() options(width=80, max.print=1000) knitr::opts_chunk$set( eval=as.logical(Sys.getenv("KNITR_EVAL", "TRUE")), cache=as.logical(Sys.getenv("KNITR_CACHE", "TRUE")), tidy.opts=list(width.cutoff=80), tidy=TRUE) ``` ```{r setup, echo=FALSE, message=FALSE, warning=FALSE} suppressPackageStartupMessages({ library(systemPipeR) library(BiocParallel) library(Biostrings) library(Rsamtools) library(GenomicRanges) library(ggplot2) library(GenomicAlignments) library(ShortRead) library(ape) library(batchtools) library(magrittr) }) ``` ## Define environment settings and samples A typical workflow starts with generating the expected working environment containing the proper directory structure, input files, and parameter settings. To simplify this task, one can load one of the existing NGS workflows templates provided by _`systemPipeRdata`_ into the current working directory. The following does this for the _`rnaseq`_ template. The name of the resulting workflow directory can be specified under the _`mydirname`_ argument. The default _`NULL`_ uses the name of the chosen workflow. An error is issued if a directory of the same name and path exists already. On Linux and OS X systems one can also create new workflow instances from the command-line of a terminal as shown [here](http://bioconductor.org/packages/devel/data/experiment/vignettes/systemPipeRdata/inst/doc/systemPipeRdata.html#generate-workflow-template). To apply workflows to custom data, the user needs to modify the _`targets`_ file and if necessary update the corresponding _`.cwl`_ and _`.yml`_ files. A collection of pre-generated _`.cwl`_ and _`.yml`_ files are provided in the _`param/cwl`_ subdirectory of each workflow template. They are also viewable in the GitHub repository of _`systemPipeRdata`_ ([see here](https://github.com/tgirke/systemPipeRdata/tree/master/inst/extdata/param)). ```{r load_package, eval=FALSE} library(systemPipeR) library(systemPipeRdata) genWorkenvir(workflow="rnaseq", mydirname=NULL) setwd("rnaseq") ``` ## Read Preprocessing ### Preprocessing with _`preprocessReads`_ function The function _`preprocessReads`_ allows to apply predefined or custom read preprocessing functions to all FASTQ files referenced in a _`SYSargs2`_ container, such as quality filtering or adaptor trimming routines. The paths to the resulting output FASTQ files are stored in the _`output`_ slot of the _`SYSargs2`_ object. Internally, _`preprocessReads`_ uses the _`FastqStreamer`_ function from the _`ShortRead`_ package to stream through large FASTQ files in a memory-efficient manner. The following example performs adaptor trimming with the _`trimLRPatterns`_ function from the _`Biostrings`_ package. After the trimming step a new targets file is generated (here _`targets_trimPE.txt`_) containing the paths to the trimmed FASTQ files. The new targets file can be used for the next workflow step with an updated _`SYSargs2`_ instance, _e.g._ running the NGS alignments with the trimmed FASTQ files. Construct _`SYSargs2`_ object from _`cwl`_ and _`yml`_ param and _`targets`_ files. ```{r construct_SYSargs2_trim-se, echo = FALSE, eval=FALSE} targets <- system.file("extdata", "targets.txt", package="systemPipeR") dir_path <- system.file("extdata/cwl/preprocessReads/trim-se", package="systemPipeR") trim <- loadWorkflow(targets=targets, wf_file="trim-se.cwl", input_file="trim-se.yml", dir_path=dir_path) trim <- renderWF(trim, inputvars=c(FileName="_FASTQ_PATH1_", SampleName="_SampleName_")) trim ``` ```{r construct_SYSargs2_trim-pe, eval=FALSE} targetsPE <- system.file("extdata", "targetsPE.txt", package="systemPipeR") dir_path <- system.file("extdata/cwl/preprocessReads/trim-pe", package="systemPipeR") trim <- loadWorkflow(targets=targetsPE, wf_file="trim-pe.cwl", input_file="trim-pe.yml", dir_path=dir_path) trim <- renderWF(trim, inputvars=c(FileName1="_FASTQ_PATH1_", FileName2="_FASTQ_PATH2_", SampleName="_SampleName_")) trim output(trim)[1:2] ``` ```{r preprocessing, eval=FALSE} preprocessReads(args=trim, Fct="trimLRPatterns(Rpattern='GCCCGGGTAA', subject=fq)", batchsize=100000, overwrite=TRUE, compress=TRUE) ``` The following example shows how one can design a custom read preprocessing function using utilities provided by the _`ShortRead`_ package, and then run it in batch mode with the _'preprocessReads'_ function (here on paired-end reads). ```{r custom_preprocessing, eval=FALSE} filterFct <- function(fq, cutoff=20, Nexceptions=0) { qcount <- rowSums(as(quality(fq), "matrix") <= cutoff, na.rm=TRUE) # Retains reads where Phred scores are >= cutoff with N exceptions fq[qcount <= Nexceptions] } preprocessReads(args=trim, Fct="filterFct(fq, cutoff=20, Nexceptions=0)", batchsize=100000) ``` ### Preprocessing with TrimGalore! [TrimGalore!](http://www.bioinformatics.babraham.ac.uk/projects/trim_galore/) is a wrapper tool to consistently apply quality and adapter trimming to fastq files, with some extra functionality for removing Reduced Representation Bisulfite-Seq (RRBS) libraries. ```{r trimGalore, eval=FALSE} targets <- system.file("extdata", "targets.txt", package="systemPipeR") dir_path <- system.file("extdata/cwl/trim_galore/trim_galore-se", package="systemPipeR") trimG <- loadWorkflow(targets=targets, wf_file="trim_galore-se.cwl", input_file="trim_galore-se.yml", dir_path=dir_path) trimG <- renderWF(trimG, inputvars=c(FileName="_FASTQ_PATH1_", SampleName="_SampleName_")) trimG cmdlist(trimG)[1:2] output(trimG)[1:2] ## Run Single Machine Option trimG <- runCommandline(trimG[1], make_bam = FALSE) ``` ### Preprocessing with Trimmomatic ```{r trimmomatic, eval=FALSE} targetsPE <- system.file("extdata", "targetsPE.txt", package="systemPipeR") dir_path <- system.file("extdata/cwl/trimmomatic/trimmomatic-pe", package="systemPipeR") trimM <- loadWorkflow(targets=targetsPE, wf_file="trimmomatic-pe.cwl", input_file="trimmomatic-pe.yml", dir_path=dir_path) trimM <- renderWF(trimM, inputvars=c(FileName1="_FASTQ_PATH1_", FileName2="_FASTQ_PATH2_", SampleName="_SampleName_")) trimM cmdlist(trimM)[1:2] output(trimM)[1:2] ## Run Single Machine Option trimM <- runCommandline(trimM[1], make_bam = FALSE) ``` ## FASTQ quality report The following _`seeFastq`_ and _`seeFastqPlot`_ functions generate and plot a series of useful quality statistics for a set of FASTQ files including per cycle quality box plots, base proportions, base-level quality trends, relative k-mer diversity, length and occurrence distribution of reads, number of reads above quality cutoffs and mean quality distribution. The function _`seeFastq`_ computes the quality statistics and stores the results in a relatively small list object that can be saved to disk with _`save()`_ and reloaded with _`load()`_ for later plotting. The argument _`klength`_ specifies the k-mer length and _`batchsize`_ the number of reads to a random sample from each FASTQ file. ```{r fastq_quality, eval=FALSE} fqlist <- seeFastq(fastq=infile1(trim), batchsize=10000, klength=8) pdf("./results/fastqReport.pdf", height=18, width=4*length(fqlist)) seeFastqPlot(fqlist) dev.off() ```
**Figure 1:** FASTQ quality report

Parallelization of FASTQ quality report on a single machine with multiple cores. ```{r fastq_quality_parallel_single, eval=FALSE} f <- function(x) seeFastq(fastq=infile1(trim)[x], batchsize=100000, klength=8) fqlist <- bplapply(seq(along=trim), f, BPPARAM = MulticoreParam(workers=4)) seeFastqPlot(unlist(fqlist, recursive=FALSE)) ``` Parallelization of FASTQ quality report via scheduler (_e.g._ Slurm) across several compute nodes. ```{r fastq_quality_parallel_cluster, eval=FALSE} library(BiocParallel); library(batchtools) f <- function(x) { library(systemPipeR) targetsPE <- system.file("extdata", "targetsPE.txt", package="systemPipeR") dir_path <- system.file("extdata/cwl/preprocessReads/trim-pe", package="systemPipeR") trim <- loadWorkflow(targets=targetsPE, wf_file="trim-pe.cwl", input_file="trim-pe.yml", dir_path=dir_path) trim <- renderWF(trim, inputvars=c(FileName1="_FASTQ_PATH1_", FileName2="_FASTQ_PATH2_", SampleName="_SampleName_")) seeFastq(fastq=infile1(trim)[x], batchsize=100000, klength=8) } resources <- list(walltime=120, ntasks=1, ncpus=4, memory=1024) param <- BatchtoolsParam(workers = 4, cluster = "slurm", template = "batchtools.slurm.tmpl", resources = resources) fqlist <- bplapply(seq(along=trim), f, BPPARAM = param) seeFastqPlot(unlist(fqlist, recursive=FALSE)) ``` ## NGS Alignment software After quality control, the sequence reads can be aligned to a reference genome or transcriptome database. The following sessions present some NGS sequence alignment software. Select the most accurate aligner and determining the optimal parameter for your custom data set project. For all the following examples, it is necessary to install the respective software and export the `PATH` accordingly. If it is available [Environment Module](http://modules.sourceforge.net/) in the system, you can load all the request software with _`moduleload(args)`_ function. ### Alignment with `HISAT2` using _`SYSargs2`_ The following steps will demonstrate how to use the short read aligner `Hisat2` [@Kim2015-ve] in both interactive job submissions and batch submissions to queuing systems of clusters using the _`systemPipeR's`_ new CWL command-line interface. The parameter settings of the aligner are defined in the `hisat2-mapping-se.cwl` and `hisat2-mapping-se.yml` files. The following shows how to construct the corresponding *SYSargs2* object, here *args*. ```{r hisat_SYSargs2_object, eval=TRUE} targets <- system.file("extdata", "targets.txt", package="systemPipeR") dir_path <- system.file("extdata/cwl/hisat2/hisat2-se", package="systemPipeR") args <- loadWorkflow(targets=targets, wf_file="hisat2-mapping-se.cwl", input_file="hisat2-mapping-se.yml", dir_path=dir_path) args <- renderWF(args, inputvars=c(FileName="_FASTQ_PATH1_", SampleName="_SampleName_")) args cmdlist(args)[1:2] output(args)[1:2] ``` Subsetting _`SYSargs2`_ class slots for each workflow step. ```{r subset, eval=TRUE} subsetWF(args, slot="input", subset='FileName')[1:2] ## Subsetting the input files for this particular workflow subsetWF(args, slot="output", subset=1, index=1)[1:2] ## Subsetting the output files for one particular step in the workflow subsetWF(args, slot="step", subset=1)[1] ## Subsetting the command-lines for one particular step in the workflow subsetWF(args, slot="output", subset=1, index=1, delete=TRUE)[1] ## DELETING specific output files ``` Build `Hisat2` index. ```{r hisat_index, eval=FALSE} dir_path <- system.file("extdata/cwl/hisat2/hisat2-idx", package="systemPipeR") idx <- loadWorkflow(targets=NULL, wf_file="hisat2-index.cwl", input_file="hisat2-index.yml", dir_path=dir_path) idx <- renderWF(idx) idx cmdlist(idx) ## Run runCommandline(idx, make_bam = FALSE) ``` #### Interactive job submissions in a single machine To simplify the short read alignment execution for the user, the command-line can be run with the *`runCommandline`* function. The execution will be on a single machine without submitting to a queuing system of a computer cluster. This way, the input FASTQ files will be processed sequentially. By default *`runCommandline`* auto detects SAM file outputs and converts them to sorted and indexed BAM files, using internally the `Rsamtools` package [@Rsamtools]. Besides, *`runCommandline`* allows the user to create a dedicated results folder for each workflow and a sub-folder for each sample defined in the *targets* file. This includes all the output and log files for each step. When these options are used, the output location will be updated by default and can be assigned to the same object. ```{r runCommandline_align, eval=FALSE} runCommandline(args, make_bam = FALSE) ## generates alignments and writes *.sam files to ./results folder args <- runCommandline(args, make_bam = TRUE) ## same as above but writes files and converts *.sam files to sorted and indexed BAM files. Assigning the new extention of the output files to the object args. ``` If available, multiple CPU cores can be used for processing each file. The number of CPU cores (here 4) to use for each process is defined in the _`*.yml`_ file. With _`yamlinput(args)['thread']`_ one can return this value from the _`SYSargs2`_ object. #### Parallelization on clusters Alternatively, the computation can be greatly accelerated by processing many files in parallel using several compute nodes of a cluster, where a scheduling/queuing system is used for load balancing. For this the *`clusterRun`* function submits the computing requests to the scheduler using the run specifications defined by *`runCommandline`*. To avoid over-subscription of CPU cores on the compute nodes, the value from _`yamlinput(args)['thread']`_ is passed on to the submission command, here _`ncpus`_ in the _`resources`_ list object. The number of independent parallel cluster processes is defined under the _`Njobs`_ argument. The following example will run 18 processes in parallel using for each 4 CPU cores. If the resources available on a cluster allow running all 18 processes at the same time then the shown sample submission will utilize in total 72 CPU cores. Note, *`clusterRun`* can be used with most queueing systems as it is based on utilities from the _`batchtools`_ package which supports the use of template files (_`*.tmpl`_) for defining the run parameters of different schedulers. To run the following code, one needs to have both a conf file (see _`.batchtools.conf.R`_ samples [here](https://mllg.github.io/batchtools/)) and a template file (see _`*.tmpl`_ samples [here](https://github.com/mllg/batchtools/tree/master/inst/templates)) for the queueing available on a system. The following example uses the sample conf and template files for the Slurm scheduler provided by this package. ```{r clusterRun_args, eval=FALSE} library(batchtools) resources <- list(walltime=120, ntasks=1, ncpus=4, memory=1024) reg <- clusterRun(args, FUN = runCommandline, more.args = list(args=args, make_bam=TRUE, dir=FALSE), conffile = ".batchtools.conf.R", template = "batchtools.slurm.tmpl", Njobs=18, runid="01", resourceList=resources) getStatus(reg=reg) waitForJobs(reg=reg) ``` Check and update the output location if necessary. ```{r output, eval=FALSE} args <- output_update(args, dir=FALSE, replace=TRUE, extension=c(".sam", ".bam")) ## Updates the output(args) to the right location in the subfolders output(args) ``` #### Create new targets file To establish the connectivity to the next workflow step, one can write a new *targets* file with the *`writeTargetsout`* function. The new *targets* file serves as input to the next *`loadWorkflow`* and *`renderWF`* call. ```{r writeTargetsout, eval=FALSE} names(clt(args)) writeTargetsout(x=args, file="default", step = 1, new_col = "FileName", new_col_output_index = 1, overwrite = TRUE) ``` #### Alignment with `HISAT2` and `SAMtools` Alternatively, it possible to build an workflow with `HISAT2` and `SAMtools`. ```{r hisat_alignment, eval=FALSE} targets <- system.file("extdata", "targets.txt", package="systemPipeR") dir_path <- system.file("extdata/cwl/workflow-hisat2/workflow-hisat2-se", package="systemPipeR") WF <- loadWorkflow(targets=targets, wf_file="workflow_hisat2-se.cwl", input_file="workflow_hisat2-se.yml", dir_path=dir_path) WF <- renderWF(WF, inputvars=c(FileName="_FASTQ_PATH1_", SampleName="_SampleName_")) WF cmdlist(WF)[1:2] output(WF)[1:2] ``` ### Alignment with _`Tophat2`_ The NGS reads of this project can also be aligned against the reference genome sequence using `Bowtie2/TopHat2` [@Kim2013-vg; @Langmead2012-bs]. Build _`Bowtie2`_ index. ```{r bowtie_index, eval=FALSE} dir_path <- system.file("extdata/cwl/bowtie2/bowtie2-idx", package="systemPipeR") idx <- loadWorkflow(targets=NULL, wf_file="bowtie2-index.cwl", input_file="bowtie2-index.yml", dir_path=dir_path) idx <- renderWF(idx) idx cmdlist(idx) ## Run in single machine runCommandline(idx, make_bam = FALSE) ``` The parameter settings of the aligner are defined in the `tophat2-mapping-pe.cwl` and `tophat2-mapping-pe.yml` files. The following shows how to construct the corresponding *SYSargs2* object, here *tophat2PE*. ```{r tophat2-pe, eval=FALSE} targetsPE <- system.file("extdata", "targetsPE.txt", package = "systemPipeR") dir_path <- system.file("extdata/cwl/tophat2/tophat2-pe", package="systemPipeR") tophat2PE <- loadWorkflow(targets = targetsPE, wf_file = "tophat2-mapping-pe.cwl", input_file = "tophat2-mapping-pe.yml", dir_path = dir_path) tophat2PE <- renderWF(tophat2PE, inputvars = c(FileName1 = "_FASTQ_PATH1_", FileName2 = "_FASTQ_PATH2_", SampleName = "_SampleName_")) tophat2PE cmdlist(tophat2PE)[1:2] output(tophat2PE)[1:2] ## Run in single machine tophat2PE <- runCommandline(tophat2PE[1], make_bam = TRUE) ``` Parallelization on clusters. ```{r tophat2-pe_parallel, eval=FALSE} resources <- list(walltime=120, ntasks=1, ncpus=4, memory=1024) reg <- clusterRun(tophat2PE, FUN = runCommandline, more.args = list(args=tophat2PE, make_bam=TRUE, dir=FALSE), conffile = ".batchtools.conf.R", template = "batchtools.slurm.tmpl", Njobs=18, runid="01", resourceList=resources) waitForJobs(reg=reg) ``` Create new targets file ```{r writeTargetsout_tophat2PE, eval=FALSE} names(clt(tophat2PE)) writeTargetsout(x=tophat2PE, file="default", step = 1, new_col = "tophat2PE", new_col_output_index = 1, overwrite = TRUE) ``` ### Alignment with _`Bowtie2`_ (_e.g._ for miRNA profiling) The following example runs _`Bowtie2`_ as a single process without submitting it to a cluster. Building the index: ```{r bowtie2_index, eval=FALSE} dir_path <- system.file("extdata/cwl/bowtie2/bowtie2-idx", package="systemPipeR") idx <- loadWorkflow(targets=NULL, wf_file="bowtie2-index.cwl", input_file="bowtie2-index.yml", dir_path=dir_path) idx <- renderWF(idx) idx cmdlist(idx) ## Run in single machine runCommandline(idx, make_bam = FALSE) ``` Building all the command-line: ```{r bowtie2_SYSargs2, eval=FALSE} targetsPE <- system.file("extdata", "targetsPE.txt", package = "systemPipeR") dir_path <- system.file("extdata/cwl/bowtie2/bowtie2-pe", package="systemPipeR") bowtiePE <- loadWorkflow(targets = targetsPE, wf_file = "bowtie2-mapping-pe.cwl", input_file = "bowtie2-mapping-pe.yml", dir_path = dir_path) bowtiePE <- renderWF(bowtiePE, inputvars = c(FileName1 = "_FASTQ_PATH1_", FileName2 = "_FASTQ_PATH2_", SampleName = "_SampleName_")) bowtiePE cmdlist(bowtiePE)[1:2] output(bowtiePE)[1:2] ``` Running all the jobs to computing nodes. ```{r bowtie2_cluster, eval=FALSE} resources <- list(walltime=120, ntasks=1, ncpus=4, memory=1024) reg <- clusterRun(bowtiePE, FUN = runCommandline, more.args = list(args=bowtiePE, dir = FALSE), conffile = ".batchtools.conf.R", template = "batchtools.slurm.tmpl", Njobs = 18, runid = "01", resourceList = resources) getStatus(reg = reg) ``` Alternatively, it possible to run all the jobs in a single machine. ```{r bowtie2_sm, eval=FALSE} bowtiePE <- runCommandline(bowtiePE) ``` Create new targets file. ```{r writeTargetsout_bowtiePE, eval=FALSE} names(clt(bowtiePE)) writeTargetsout(x=bowtiePE, file="default", step = 1, new_col = "bowtiePE", new_col_output_index = 1, overwrite = TRUE) ``` ### Alignment with _`BWA-MEM`_ (_e.g._ for VAR-Seq) The following example runs BWA-MEM as a single process without submitting it to a cluster. ##TODO: add reference Build the index: ```{r bwa_index, eval=FALSE} dir_path <- system.file("extdata/cwl/bwa/bwa-idx", package="systemPipeR") idx <- loadWorkflow(targets=NULL, wf_file="bwa-index.cwl", input_file="bwa-index.yml", dir_path=dir_path) idx <- renderWF(idx) idx cmdlist(idx) # Indexes reference genome ## Run runCommandline(idx, make_bam = FALSE) ``` Running the alignment: ```{r bwa-pe_alignment, eval=FALSE} targetsPE <- system.file("extdata", "targetsPE.txt", package = "systemPipeR") dir_path <- system.file("extdata/cwl/bwa/bwa-pe", package="systemPipeR") bwaPE <- loadWorkflow(targets = targetsPE, wf_file = "bwa-pe.cwl", input_file = "bwa-pe.yml", dir_path = dir_path) bwaPE <- renderWF(bwaPE, inputvars = c(FileName1 = "_FASTQ_PATH1_", FileName2 = "_FASTQ_PATH2_", SampleName = "_SampleName_")) bwaPE cmdlist(bwaPE)[1:2] output(bwaPE)[1:2] ## Single Machine bwaPE <- runCommandline(args= bwaPE, make_bam=FALSE) ## Cluster library(batchtools) resources <- list(walltime = 120, ntasks = 1, ncpus = 4, memory = 1024) reg <- clusterRun(bwaPE, FUN = runCommandline, more.args = list(args=bwaPE, dir = FALSE), conffile = ".batchtools.conf.R", template = "batchtools.slurm.tmpl", Njobs = 18, runid = "01", resourceList = resources) getStatus(reg = reg) ``` Create new targets file. ```{r writeTargetsout_bwaPE, eval=FALSE} names(clt(bwaPE)) writeTargetsout(x=bwaPE, file="default", step = 1, new_col = "bwaPE", new_col_output_index = 1, overwrite = TRUE) ``` ### Alignment with _`Rsubread`_ (_e.g._ for RNA-Seq) The following example shows how one can use within the \Rpackage{systemPipeR} environment the R-based aligner \Rpackage{Rsubread}, allowing running from R or command-line. ```{r rsubread, eval=FALSE} ## Build the index: dir_path <- system.file("extdata/cwl/rsubread/rsubread-idx", package="systemPipeR") idx <- loadWorkflow(targets = NULL, wf_file = "rsubread-index.cwl", input_file = "rsubread-index.yml", dir_path = dir_path) idx <- renderWF(idx) idx cmdlist(idx) runCommandline(args= idx, make_bam = FALSE) ## Running the alignment: targets <- system.file("extdata", "targets.txt", package = "systemPipeR") dir_path <- system.file("extdata/cwl/rsubread/rsubread-se", package="systemPipeR") rsubread <- loadWorkflow(targets = targets, wf_file = "rsubread-mapping-se.cwl", input_file = "rsubread-mapping-se.yml", dir_path = dir_path) rsubread <- renderWF(rsubread, inputvars = c(FileName = "_FASTQ_PATH1_", SampleName = "_SampleName_")) rsubread cmdlist(rsubread)[1] ## Single Machine rsubread <- runCommandline(args=rsubread[1]) ``` Create new targets file. ```{r writeTargetsout_rsubread, eval=FALSE} names(clt(rsubread)) writeTargetsout(x=rsubread, file="default", step = 1, new_col = "rsubread", new_col_output_index = 1, overwrite = TRUE) ``` ### Alignment with _`gsnap`_ (_e.g._ for VAR-Seq and RNA-Seq) Another R-based short read aligner is _`gsnap`_ from the _`gmapR`_ package [@Wu2010-iq]. The code sample below introduces how to run this aligner on multiple nodes of a compute cluster. ```{r gsnap, eval=FALSE} ## Build the index: dir_path <- system.file("extdata/cwl/gsnap/gsnap-idx", package="systemPipeR") idx <- loadWorkflow(targets = NULL, wf_file = "gsnap-index.cwl", input_file = "gsnap-index.yml", dir_path = dir_path) idx <- renderWF(idx) idx cmdlist(idx) runCommandline(args= idx, make_bam = FALSE) ## Running the alignment: targetsPE <- system.file("extdata", "targetsPE.txt", package = "systemPipeR") dir_path <- system.file("extdata/cwl/gsnap/gsnap-pe", package="systemPipeR") gsnap <- loadWorkflow(targets = targetsPE, wf_file = "gsnap-mapping-pe.cwl", input_file = "gsnap-mapping-pe.yml", dir_path = dir_path) gsnap <- renderWF(gsnap, inputvars = c(FileName1 = "_FASTQ_PATH1_", FileName2 = "_FASTQ_PATH2_", SampleName = "_SampleName_")) gsnap cmdlist(gsnap)[1] output(gsnap)[1] ## Cluster library(batchtools) resources <- list(walltime = 120, ntasks = 1, ncpus = 4, memory = 1024) reg <- clusterRun(gsnap, FUN = runCommandline, more.args = list(args=gsnap, make_bam=FALSE), conffile = ".batchtools.conf.R", template = "batchtools.slurm.tmpl", Njobs = 18, runid = "01", resourceList = resources) getStatus(reg = reg) gsnap <- output_update(gsnap, dir=FALSE, replace=TRUE, extension=c(".sam", ".bam")) ``` Create new targets file. ```{r writeTargetsout_gsnap, eval=FALSE} names(clt(gsnap)) writeTargetsout(x=gsnap, file="default", step = 1, new_col = "gsnap", new_col_output_index = 1, overwrite = TRUE) ``` ## Create symbolic links for viewing BAM files in IGV The genome browser IGV supports reading of indexed/sorted BAM files via web URLs. This way it can be avoided to create unnecessary copies of these large files. To enable this approach, an HTML directory with Http access needs to be available in the user account (_e.g._ _`home/publichtml`_) of a system. If this is not the case then the BAM files need to be moved or copied to the system where IGV runs. In the following, _`htmldir`_ defines the path to the HTML directory with http access where the symbolic links to the BAM files will be stored. The corresponding URLs will be written to a text file specified under the `_urlfile`_ argument. ```{r igv, eval=FALSE} symLink2bam(sysargs=args, htmldir=c("~/.html/", "somedir/"), urlbase="http://myserver.edu/~username/", urlfile="IGVurl.txt") ``` ## Read counting for mRNA profiling experiments Create _`txdb`_ (needs to be done only once). ```{r create_txdb, eval=FALSE} library(GenomicFeatures) txdb <- makeTxDbFromGFF(file="data/tair10.gff", format="gff", dataSource="TAIR", organism="Arabidopsis thaliana") saveDb(txdb, file="./data/tair10.sqlite") ``` The following performs read counting with _`summarizeOverlaps`_ in parallel mode with multiple cores. ```{r read_counting_multicore, eval=FALSE} library(BiocParallel) txdb <- loadDb("./data/tair10.sqlite") eByg <- exonsBy(txdb, by="gene") outpaths <- subsetWF(args, slot="output", subset=1, index=1) bfl <- BamFileList(outpaths, yieldSize=50000, index=character()) multicoreParam <- MulticoreParam(workers=4); register(multicoreParam); registered() counteByg <- bplapply(bfl, function(x) summarizeOverlaps(eByg, x, mode="Union", ignore.strand=TRUE, inter.feature=TRUE, singleEnd=TRUE)) # Note: for strand-specific RNA-Seq set 'ignore.strand=FALSE' and for PE data set 'singleEnd=FALSE' countDFeByg <- sapply(seq(along=counteByg), function(x) assays(counteByg[[x]])$counts) rownames(countDFeByg) <- names(rowRanges(counteByg[[1]])); colnames(countDFeByg) <- names(bfl) rpkmDFeByg <- apply(countDFeByg, 2, function(x) returnRPKM(counts=x, ranges=eByg)) write.table(countDFeByg, "results/countDFeByg.xls", col.names=NA, quote=FALSE, sep="\t") write.table(rpkmDFeByg, "results/rpkmDFeByg.xls", col.names=NA, quote=FALSE, sep="\t") ``` Please note, in addition to read counts this step generates RPKM normalized expression values. For most statistical differential expression or abundance analysis methods, such as _`edgeR`_ or _`DESeq2`_, the raw count values should be used as input. The usage of RPKM values should be restricted to specialty applications required by some users, _e.g._ manually comparing the expression levels of different genes or features. Read counting with _`summarizeOverlaps`_ using multiple nodes of a cluster. ```{r read_counting_multinode, eval=FALSE} library(BiocParallel) f <- function(x) { library(systemPipeR); library(BiocParallel); library(GenomicFeatures) txdb <- loadDb("./data/tair10.sqlite") eByg <- exonsBy(txdb, by="gene") args <- systemArgs(sysma="param/tophat.param", mytargets="targets.txt") outpaths <- subsetWF(args, slot="output", subset=1, index=1) bfl <- BamFileList(outpaths, yieldSize=50000, index=character()) summarizeOverlaps(eByg, bfl[x], mode="Union", ignore.strand=TRUE, inter.feature=TRUE, singleEnd=TRUE) } resources <- list(walltime=120, ntasks=1, ncpus=4, memory=1024) param <- BatchtoolsParam(workers = 4, cluster = "slurm", template = "batchtools.slurm.tmpl", resources = resources) counteByg <- bplapply(seq(along=args), f, BPPARAM = param) countDFeByg <- sapply(seq(along=counteByg), function(x) assays(counteByg[[x]])$counts) rownames(countDFeByg) <- names(rowRanges(counteByg[[1]])); colnames(countDFeByg) <- names(outpaths) ``` Useful commands for monitoring the progress of submitted jobs ```{r process_monitoring, eval=FALSE} getStatus(reg=reg) outpaths <- subsetWF(args, slot="output", subset=1, index=1) file.exists(outpaths) sapply(1:length(outpaths), function(x) loadResult(reg, id=x)) # Works after job completion ``` #### Read and alignment count stats Generate a table of read and alignment counts for all samples. ```{r align_stats1, eval=FALSE} read_statsDF <- alignStats(args) write.table(read_statsDF, "results/alignStats.xls", row.names=FALSE, quote=FALSE, sep="\t") ``` The following shows the first four lines of the sample alignment stats file provided by the _`systemPipeR`_ package. For simplicity the number of PE reads is multiplied here by 2 to approximate proper alignment frequencies where each read in a pair is counted. ```{r align_stats2, eval=TRUE} read.table(system.file("extdata", "alignStats.xls", package="systemPipeR"), header=TRUE)[1:4,] ``` Parallelization of read/alignment stats on single machine with multiple cores. ```{r align_stats_parallel, eval=FALSE} f <- function(x) alignStats(args[x]) read_statsList <- bplapply(seq(along=args), f, BPPARAM = MulticoreParam(workers=8)) read_statsDF <- do.call("rbind", read_statsList) ``` Parallelization of read/alignment stats via scheduler (_e.g._ Slurm) across several compute nodes. ```{r align_stats_parallel_cluster, eval=FALSE} library(BiocParallel); library(batchtools) f <- function(x) { library(systemPipeR) targets <- system.file("extdata", "targets.txt", package="systemPipeR") dir_path <- "param/cwl/hisat2/hisat2-se" ## TODO: replace path to system.file args <- loadWorkflow(targets=targets, wf_file="hisat2-mapping-se.cwl", input_file="hisat2-mapping-se.yml", dir_path=dir_path) args <- renderWF(args, inputvars=c(FileName="_FASTQ_PATH1_", SampleName="_SampleName_")) args <- output_update(args, dir=FALSE, replace=TRUE, extension=c(".sam", ".bam")) alignStats(args[x]) } resources <- list(walltime=120, ntasks=1, ncpus=4, memory=1024) param <- BatchtoolsParam(workers = 4, cluster = "slurm", template = "batchtools.slurm.tmpl", resources = resources) read_statsList <- bplapply(seq(along=args), f, BPPARAM = param) read_statsDF <- do.call("rbind", read_statsList) ``` ## Read counting for miRNA profiling experiments Download miRNA genes from miRBase. ```{r read_counting_mirna, eval=FALSE} system("wget ftp://mirbase.org/pub/mirbase/19/genomes/My_species.gff3 -P ./data/") gff <- import.gff("./data/My_species.gff3") gff <- split(gff, elementMetadata(gff)$ID) bams <- names(bampaths); names(bams) <- targets$SampleName bfl <- BamFileList(bams, yieldSize=50000, index=character()) countDFmiR <- summarizeOverlaps(gff, bfl, mode="Union", ignore.strand=FALSE, inter.feature=FALSE) # Note: inter.feature=FALSE important since pre and mature miRNA ranges overlap rpkmDFmiR <- apply(countDFmiR, 2, function(x) returnRPKM(counts=x, gffsub=gff)) write.table(assays(countDFmiR)$counts, "results/countDFmiR.xls", col.names=NA, quote=FALSE, sep="\t") write.table(rpkmDFmiR, "results/rpkmDFmiR.xls", col.names=NA, quote=FALSE, sep="\t") ``` ## Correlation analysis of samples The following computes the sample-wise Spearman correlation coefficients from the _`rlog`_ (regularized-logarithm) transformed expression values generated with the _`DESeq2`_ package. After transformation to a distance matrix, hierarchical clustering is performed with the _`hclust`_ function and the result is plotted as a dendrogram ([sample\_tree.pdf](./results/sample_tree.pdf)). ```{r sample_tree_rlog, eval=TRUE} library(DESeq2, warn.conflicts=FALSE, quietly=TRUE); library(ape, warn.conflicts=FALSE) countDFpath <- system.file("extdata", "countDFeByg.xls", package="systemPipeR") countDF <- as.matrix(read.table(countDFpath)) colData <- data.frame(row.names=targets.as.df(targets(args))$SampleName, condition=targets.as.df(targets(args))$Factor) dds <- DESeqDataSetFromMatrix(countData = countDF, colData = colData, design = ~ condition) d <- cor(assay(rlog(dds)), method="spearman") hc <- hclust(dist(1-d)) plot.phylo(as.phylo(hc), type="p", edge.col=4, edge.width=3, show.node.label=TRUE, no.margin=TRUE) ```
**Figure 2:** Correlation dendrogram of samples for _`rlog`_ values.

Alternatively, the clustering can be performed with _`RPKM`_ normalized expression values. In combination with Spearman correlation the results of the two clustering methods are often relatively similar. ```{r sample_tree_rpkm, eval=FALSE} rpkmDFeBygpath <- system.file("extdata", "rpkmDFeByg.xls", package="systemPipeR") rpkmDFeByg <- read.table(rpkmDFeBygpath, check.names=FALSE) rpkmDFeByg <- rpkmDFeByg[rowMeans(rpkmDFeByg) > 50,] d <- cor(rpkmDFeByg, method="spearman") hc <- hclust(as.dist(1-d)) plot.phylo(as.phylo(hc), type="p", edge.col="blue", edge.width=2, show.node.label=TRUE, no.margin=TRUE) ``` ## DEG analysis with _`edgeR`_ The following _`run_edgeR`_ function is a convenience wrapper for identifying differentially expressed genes (DEGs) in batch mode with _`edgeR`_'s GML method [@Robinson2010-uk] for any number of pairwise sample comparisons specified under the _`cmp`_ argument. Users are strongly encouraged to consult the [_`edgeR`_](\href{http://www.bioconductor.org/packages/devel/bioc/vignettes/edgeR/inst/doc/edgeRUsersGuide.pdf) vignette for more detailed information on this topic and how to properly run _`edgeR`_ on data sets with more complex experimental designs. ```{r edger_wrapper, eval=TRUE} targetspath <- system.file("extdata", "targets.txt", package="systemPipeR") targets <- read.delim(targetspath, comment="#") cmp <- readComp(file=targetspath, format="matrix", delim="-") cmp[[1]] countDFeBygpath <- system.file("extdata", "countDFeByg.xls", package="systemPipeR") countDFeByg <- read.delim(countDFeBygpath, row.names=1) edgeDF <- run_edgeR(countDF=countDFeByg, targets=targets, cmp=cmp[[1]], independent=FALSE, mdsplot="") ``` Filter and plot DEG results for up and down-regulated genes. Because of the small size of the toy data set used by this vignette, the _FDR_ value has been set to a relatively high threshold (here 10%). More commonly used _FDR_ cutoffs are 1% or 5%. The definition of '_up_' and '_down_' is given in the corresponding help file. To open it, type _`?filterDEGs`_ in the R console. ```{r edger_deg_counts, eval=TRUE} DEG_list <- filterDEGs(degDF=edgeDF, filter=c(Fold=2, FDR=10)) ```
**Figure 3:** Up and down regulated DEGs identified by _`edgeR`_.

```{r edger_deg_stats, eval=TRUE} names(DEG_list) DEG_list$Summary[1:4,] ``` ## DEG analysis with _`DESeq2`_ The following _`run_DESeq2`_ function is a convenience wrapper for identifying DEGs in batch mode with _`DESeq2`_ [@Love2014-sh] for any number of pairwise sample comparisons specified under the _`cmp`_ argument. Users are strongly encouraged to consult the [_`DESeq2`_](http://www.bioconductor.org/packages/devel/bioc/vignettes/DESeq2/inst/doc/DESeq2.pdf) vignette for more detailed information on this topic and how to properly run _`DESeq2`_ on data sets with more complex experimental designs. ```{r deseq2_wrapper, eval=TRUE} degseqDF <- run_DESeq2(countDF=countDFeByg, targets=targets, cmp=cmp[[1]], independent=FALSE) ``` Filter and plot DEG results for up and down-regulated genes. ```{r deseq2_deg_counts, eval=TRUE} DEG_list2 <- filterDEGs(degDF=degseqDF, filter=c(Fold=2, FDR=10)) ```
**Figure 4:** Up and down regulated DEGs identified by _`DESeq2`_.

## Venn Diagrams The function _`overLapper`_ can compute Venn intersects for large numbers of sample sets (up to 20 or more) and _`vennPlot`_ can plot 2-5 way Venn diagrams. A useful feature is the possibility to combine the counts from several Venn comparisons with the same number of sample sets in a single Venn diagram (here for 4 up and down DEG sets). ```{r vennplot, eval=TRUE} vennsetup <- overLapper(DEG_list$Up[6:9], type="vennsets") vennsetdown <- overLapper(DEG_list$Down[6:9], type="vennsets") vennPlot(list(vennsetup, vennsetdown), mymain="", mysub="", colmode=2, ccol=c("blue", "red")) ```
**Figure 5:** Venn Diagram for 4 Up and Down DEG Sets.

## GO term enrichment analysis of DEGs ### Obtain gene-to-GO mappings The following shows how to obtain gene-to-GO mappings from _`biomaRt`_ (here for _A. thaliana_) and how to organize them for the downstream GO term enrichment analysis. Alternatively, the gene-to-GO mappings can be obtained for many organisms from Bioconductor's _`*.db`_ genome annotation packages or GO annotation files provided by various genome databases. For each annotation, this relatively slow preprocessing step needs to be performed only once. Subsequently, the preprocessed data can be loaded with the _`load`_ function as shown in the next subsection. ```{r get_go_biomart, eval=FALSE} library("biomaRt") listMarts() # To choose BioMart database listMarts(host="plants.ensembl.org") m <- useMart("plants_mart", host="plants.ensembl.org") listDatasets(m) m <- useMart("plants_mart", dataset="athaliana_eg_gene", host="plants.ensembl.org") listAttributes(m) # Choose data types you want to download go <- getBM(attributes=c("go_id", "tair_locus", "namespace_1003"), mart=m) go <- go[go[,3]!="",]; go[,3] <- as.character(go[,3]) go[go[,3]=="molecular_function", 3] <- "F"; go[go[,3]=="biological_process", 3] <- "P"; go[go[,3]=="cellular_component", 3] <- "C" go[1:4,] dir.create("./data/GO") write.table(go, "data/GO/GOannotationsBiomart_mod.txt", quote=FALSE, row.names=FALSE, col.names=FALSE, sep="\t") catdb <- makeCATdb(myfile="data/GO/GOannotationsBiomart_mod.txt", lib=NULL, org="", colno=c(1,2,3), idconv=NULL) save(catdb, file="data/GO/catdb.RData") ``` ### Batch GO term enrichment analysis Apply the enrichment analysis to the DEG sets obtained in the above differential expression analysis. Note, in the following example the _FDR_ filter is set here to an unreasonably high value, simply because of the small size of the toy data set used in this vignette. Batch enrichment analysis of many gene sets is performed with the _`GOCluster_Report`_ function. When _`method="all"`_, it returns all GO terms passing the p-value cutoff specified under the _`cutoff`_ arguments. When _`method="slim"`_, it returns only the GO terms specified under the _`myslimv`_ argument. The given example shows how one can obtain such a GO slim vector from BioMart for a specific organism. ```{r go_enrichment, eval=FALSE} load("data/GO/catdb.RData") DEG_list <- filterDEGs(degDF=edgeDF, filter=c(Fold=2, FDR=50), plot=FALSE) up_down <- DEG_list$UporDown; names(up_down) <- paste(names(up_down), "_up_down", sep="") up <- DEG_list$Up; names(up) <- paste(names(up), "_up", sep="") down <- DEG_list$Down; names(down) <- paste(names(down), "_down", sep="") DEGlist <- c(up_down, up, down) DEGlist <- DEGlist[sapply(DEGlist, length) > 0] BatchResult <- GOCluster_Report(catdb=catdb, setlist=DEGlist, method="all", id_type="gene", CLSZ=2, cutoff=0.9, gocats=c("MF", "BP", "CC"), recordSpecGO=NULL) library("biomaRt") m <- useMart("plants_mart", dataset="athaliana_eg_gene", host="plants.ensembl.org") goslimvec <- as.character(getBM(attributes=c("goslim_goa_accession"), mart=m)[,1]) BatchResultslim <- GOCluster_Report(catdb=catdb, setlist=DEGlist, method="slim", id_type="gene", myslimv=goslimvec, CLSZ=10, cutoff=0.01, gocats=c("MF", "BP", "CC"), recordSpecGO=NULL) ``` ### Plot batch GO term results The _`data.frame`_ generated by _`GOCluster_Report`_ can be plotted with the _`goBarplot`_ function. Because of the variable size of the sample sets, it may not always be desirable to show the results from different DEG sets in the same bar plot. Plotting single sample sets is achieved by subsetting the input data frame as shown in the first line of the following example. ```{r plot_go_enrichment, eval=FALSE} gos <- BatchResultslim[grep("M6-V6_up_down", BatchResultslim$CLID), ] gos <- BatchResultslim pdf("GOslimbarplotMF.pdf", height=8, width=10); goBarplot(gos, gocat="MF"); dev.off() goBarplot(gos, gocat="BP") goBarplot(gos, gocat="CC") ``` ![](GOslimbarplotMF.png)
**Figure 6:** GO Slim Barplot for MF Ontology.

## Clustering and heat maps The following example performs hierarchical clustering on the _`rlog`_ transformed expression matrix subsetted by the DEGs identified in the above differential expression analysis. It uses a Pearson correlation-based distance measure and complete linkage for cluster join. ```{r hierarchical_clustering, eval=FALSE} library(pheatmap) geneids <- unique(as.character(unlist(DEG_list[[1]]))) y <- assay(rlog(dds))[geneids, ] pdf("heatmap1.pdf") pheatmap(y, scale="row", clustering_distance_rows="correlation", clustering_distance_cols="correlation") dev.off() ```
**Figure 7:** Heat map with hierarchical clustering dendrograms of DEGs.

# References