Last updated on 2025-11-25 | Edit this page

Overview

Questions

What are the different choices to consider when planning an RNA-seq experiment?
How does one process the raw fastq files to generate a table with read counts per gene and sample?
Where does one find information about annotated genes for a given organism?
What are the typical steps in an RNA-seq analysis?

Objectives

Explain what RNA-seq is.
Describe some of the most common design choices that have to be made before running an RNA-seq experiment.
Provide an overview of the procedure to go from the raw data to the read count matrix that will be used for downstream analysis.
Show some common types of results and visualizations generated in RNA-seq analyses.

What are we measuring in an RNA-seq experiment?

Illustration of part of the central dogma of molecular biology, where DNA is transcribed to RNA, and intronic sequences are spliced out

In order to produce an RNA molecule, a stretch of DNA is first transcribed into mRNA. Subsequently, intronic regions are spliced out, and exonic regions are combined into different isoforms of a gene.

Illustration of the major experimental steps of an RNA-seq experiment

(figure adapted from Martin & Wang (2011)).

In a typical RNA-seq experiment, RNA molecules are first collected from a sample of interest. After a potential enrichment for molecules with polyA tails (predominantly mRNA), or depletion of otherwise highly abundant ribosomal RNA, the remaining molecules are fragmented into smaller pieces (there are also long-read protocols where entire molecules are considered, but those are not the focus of this lesson). It is important to keep in mind that because of the splicing excluding intronic sequences, an RNA molecule (and hence a generated fragment) may not correspond to an uninterrupted region of the genome. The RNA fragments are then reverse transcribed into cDNA, whereafter sequencing adapters are added to each end. These adapters allow the fragments to attach to the flow cell. Once attached, each fragment will be heavily amplified to generate a cluster of identical sequences on the flow cell. The sequencer then determines the sequence of the first 50-200 nucleotides of the cDNA fragments in each such cluster, starting from one end, which corresponds to a read. Many data sets are generated with so called paired-end protocols, in which the fragments are read from both ends. Millions of such reads (or pairs of reads) will be generated in an experiment, and these will be represented in a (pair of) FASTQ files. Each read is represented by four consecutive lines in such a file: first a line with a unique read identifier, next the inferred sequence of the read, then another identifier line, and finally a line containing the base quality for each inferred nucleotide, representing the probability that the nucleotide in the corresponding position has been correctly identified.

Discussion

Challenge: Discuss the following points with your neighbor

What are potential advantages and disadvantages of paired-end protocols compared to only sequencing one end of a fragment?
What quality assessment can you think of that would be useful to perform on the FASTQ files with read sequences?

Experimental design considerations

Before starting to collect data, it is essential to take some time to think about the design of the experiment. Experimental design concerns the organization of experiments with the purpose of making sure that the right type of data, and enough of it, is available to answer the questions of interest as efficiently as possible. Aspects such as which conditions or sample groups to consider, how many replicates to collect, and how to plan the data collection in practice are important questions to consider. Many high-throughput biological experiments (including RNA-seq) are sensitive to ambient conditions, and it is often difficult to directly compare measurements that have been done on different days, by different analysts, in different centers, or using different batches of reagents. For this reason, it is very important to design experiments properly, to make it possible to disentangle different types of (primary and secondary) effects.

A classification of many different factors affecting measurements obtained from an experiment into treatment, biological, technical and error effects

(figure from Lazic (2017)).

Discussion

Challenge: Discuss with your neighbor

Why is it essential to have replicates?

Importantly, not all replicates are equally useful, from a statistical point of view. One common way to classify the different types of replicates is as ‘biological’ and ‘technical’, where the latter are typically used to test the reproducibility of the measurement device, while biological replicates inform about the variability between different samples from a population of interest. Another scheme classifies replicates (or units) into ‘biological’, ‘experimental’ and ‘observational’. Here, biological units are entities we want to make inferences about (e.g., animals, persons). Replication of biological units is required to make a general statement of the effect of a treatment - we can not draw a conclusion about the effect of drug on a population of mice by studying a single mouse only. Experimental units are the smallest entities that can be independently assigned to a treatment (e.g., animal, litter, cage, well). Only replication of experimental units constitute true replication. Observational units, finally, are entities at which measurements are made.

To explore the impact of experimental design on the ability to answer questions of interest, we are going to use an interactive application, provided in the ConfoundingExplorer package.

Discussion

Challenge

Launch the ConfoundingExplorer application and familiarize yourself with the interface.

Discussion

Challenge

For a balanced design (equal distribution of replicates between the two groups in each batch), what is the effect of increasing the strength of the batch effect? Does it matter whether one adjusts for the batch effect or not?
For an increasingly unbalanced design (most or all replicates of one group coming from one batch), what is the effect of increasing the strength of the batch effect? Does it matter whether one adjusts for the batch effect or not?

RNA-seq quantification: from reads to count matrix

Illustration of a set of reads generated by a sequencer, and genomic and transcriptomic reference sequences

The read sequences contained in the FASTQ files from the sequencer are typically not directly useful as they are, since we do not have the information about which gene or transcript they originate from. Thus, the first processing step is to attempt to identify the location of origin for each read, and use this to obtain an estimate of the number of reads originating from a gene (or another features, such as an individual transcript). This can then be used as a proxy for the abundance, or expression level, of the gene. There is a plethora of RNA quantification pipelines, and the most common approaches can be categorized into three main types:

Align reads to the genome, and count the number of reads that map within the exons of each gene. This is the one of simplest methods. For species for which the transcriptome is poorly annotated, this would be the preferred approach. Example: STAR alignment to GRCm39 + Rsubread featureCounts
Align reads to the transcriptome, quantify transcript expression, and summarize transcript expression into gene expression. This approach can produce accurate quantification results (independent benchmarking), particularly for high-quality samples without DNA contamination. Example: RSEM quantification using rsem-calculate-expression --star on the GENCODE GRCh38 transcriptome + tximport
Pseudoalign reads against the transcriptome, using the corresponding genome as a decoy, quantifying transcript expression in the process, and summarize the transcript-level expression into gene-level expression. The advantages of this approach include: computational efficiency, mitigation of the effect of DNA contamination, and GC bias correction. Example: salmon quant --gcBias + tximport

At typical sequencing read depth, gene expression quantification is often more accurate than transcript expression quantification. However, differential gene expression analyses can be improved by having access also to transcript-level quantifications.

Other tools used in RNA-seq quantification include: TopHat2, bowtie2, kallisto, HTseq, among many others.

The choice of an appropriate RNA-seq quantification depends on the quality of the transcriptome annotation, the quality of the RNA-seq library preparation, the presence of contaminating sequences, among many factors. Often, it can be informative to compare the quantification results of multiple approaches.

The quantification method is species- and experiment-dependent, and often requires large amounts of computing resources, so we will give you a overview of how to perform quantification, and scripts to run the quantification. We will use pre-computed quantifications for the exercises in this workshop.

Discussion

Challenge: Discuss the following points with your neighbor

Which of the mentioned RNA-Seq quantification tools have you heard about? Do you know other pros and cons of the methods?
Have you done your own RNA-Seq experiment? If so, what quantification tool did you use and why did you choose it?
Do you have access to specific tools / local bioinformatics expert / computational resources for quantification? If you don’t, how might you gain access?

Finding the reference sequences

In order to quantify abundances of known genes or transcripts from RNA-seq data, we need a reference database informing us of the sequence of these features, to which we can then compare our reads. This information can be obtained from different online repositories. It is highly recommended to choose one of these for a particular project, and not mix information from different sources. Depending on the quantification tool you have chosen, you will need different types of reference information. If you are aligning your reads to the genome and investigating the overlap with known annotated features, you will need the full genome sequence (provided in a fasta file) and a file that tells you the genomic location of each annotated feature (typically provided in a gtf file). If you are mapping your reads to the transcriptome, you will instead need a file with the sequence of each transcript (again, a fasta file).

If you are working with mouse or human samples, the GENCODE project provides well-curated reference files.
Ensembl provides reference files for a large set of organisms, including plants and fungi.
UCSC also provides reference files for many organisms.

Discussion

Challenge

Download the latest mouse transcriptome fasta file from GENCODE. What do the entries look like? Tip: to read the file into R, consider the readDNAStringSet() function from the Biostrings package.

Where are we heading towards in this workshop?

In this workshop, we will discuss and practice: downloading publicly available RNA-seq data, performing quality assessment of the raw data, quantifying gene expression, and performing differential expression analysis to identify genes that are expressed at different levels between two or more conditions. We will also cover some aspects of data visualization and interpretation of the results.

The outcome of a differential expression analysis is often represented using graphical representations, such as MA plots and heatmaps (see below for examples).

An example MA plot An example heatmap

In the following episodes we will learn, among other things, how to generate and interpret these plots. It is also common to perform follow-up analyses to investigate whether there is a functional relationship among the top-ranked genes, so called gene set (enrichment) analysis, which will also be covered in a later episode.

Key Points

RNA-seq is a technique of measuring the amount of RNA expressed within a cell/tissue and state at a given time.
Many choices have to be made when planning an RNA-seq experiment, such as whether to perform poly-A selection or ribosomal depletion, whether to apply a stranded or an unstranded protocol, and whether to sequence the reads in a single-end or paired-end fashion. Each of the choices have consequences for the processing and interpretation of the data.
Many approaches exist for quantification of RNA-seq data. Some methods align reads to the genome and count the number of reads overlapping gene loci. Other methods map reads to the transcriptome and use a probabilistic approach to estimate the abundance of each gene or transcript.
Information about annotated genes can be accessed via several sources, including Ensembl, UCSC and GENCODE.

Content from Downloading and organizing files

Last updated on 2025-11-25 | Edit this page

Overview

Questions

What files are required to process raw RNA seq reads?
Where do we obtain raw reads, reference genomes, annotations, and transcript sequences?
How should project directories be organized to support a smooth workflow?
What practical considerations matter when downloading and preparing RNA seq data?

Objectives

Identify the essential inputs for RNA seq analysis: FASTQ files, reference genome, annotation, and transcript sequences.
Learn where and how to download RNA seq datasets and reference materials.
Create a clean project directory structure suitable for quality control, alignment, and quantification.
Understand practical issues such as compressed FASTQ files and adapter trimming.

Introduction

Before we can perform quality control, mapping, or quantification, we must first gather the files required for RNA seq analysis and organize them in a reproducible way. This episode introduces the essential inputs of an RNA seq workflow and demonstrates how to obtain and structure them on a computing system.

We begin by discussing raw FASTQ reads, reference genomes, annotation files, and transcript sequences. We then download the dataset used throughout the workshop and set up the directory structure that supports downstream steps.

Most FASTQ files arrive compressed as .fastq.gz files. Modern RNA seq tools accept compressed files directly, so uncompressing them is usually unnecessary.

Prerequisite

Should I uncompress FASTQ files?

In almost all RNA seq workflows, you should keep FASTQ files compressed. Tools such as FastQC, HISAT2, STAR, salmon, and fastp can read .gz files directly. Keeping files compressed saves space and reduces input and output overhead.

Many library preparation protocols introduce adapter sequences at the ends of reads. For most alignment based analyses, explicit trimming is not required because aligners soft clip adapter sequences automatically.

Prerequisite

Should I trim adapters for RNA seq analysis?

In most cases, no. Aligners such as HISAT2 and STAR soft clip adapter sequences. Overly aggressive trimming can reduce mapping rates or distort read length distributions. Trimming is needed only for specific applications such as transcript assembly where uniform read lengths are important.
More information: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4766705/

Downloading the data and project organization

For this workshop, we use a publicly available dataset that investigates the effects of cardiomyocyte specific Nat10 knockout on heart development in mice (BioProject PRJNA1254208, GEO GSE295332). The study includes three biological replicates per group, sequenced on an Illumina NovaSeq X Plus in paired end mode.

Before downloading files, we create a reproducible directory layout for raw data, scripts, mapping outputs, and count files.

Challenge

Create the directories needed for this episode

Create a working directory for the workshop (e.g., rnaseq-workshop). Inside it, create four subdirectories:

data : raw FASTQ files, reference genome, annotation
scripts : custom scripts, SLURM job files
results : alignment, counts and various other outputs

Show me the solution

BASH

cd $RCAC_SCRATCH
mkdir -p rnaseq-workshop/{data,scripts,results}

The resulting directory structure should look like this:

BASH

aseethar@scholar-fe03:[aseethar] $ pwd
/scratch/scholar/aseethar
aseethar@scholar-fe03:[aseethar] $ tree rnaseq-workshop/
rnaseq-workshop/
├── results
├── data
└── scripts

What files do I need for RNA-seq analysis?

Prerequisite

You will need:

Raw reads (FASTQ)
- Usually .fastq.gz files containing nucleotide sequences and per-base quality scores.

For alignment based workflows

Reference genome (FASTA)
- Contains chromosome or contig sequences.
Annotation file (GTF/GFF)
- Describes gene models: exons, transcripts, coordinates.

For transcript-level quantification workflows

Transcript sequences (FASTA)
- Required for transcript based quantification with tools such as salmon, kallisto, and RSEM.

Reference files must be consistent with each other (same genome version). To ensure a smooth analysis workflow, keep these files logically organized. Good directory structure minimizes mistakes, simplifies scripting, and supports reproducibility.

Downloading the data

We now download:

the GRCm39 reference genome
the corresponding GENCODE annotation
transcript sequences (if using transcript based quantification)
raw FASTQ files from SRA

Reference genome and annotation

We will use GENCODE GRCm39 (M38) as the reference.

Challenge

Annotation

Visit the GENCODE mouse page: https://www.gencodegenes.org/mouse/.
Which annotation file (GTF/GFF3 files section) should you download, and why?

Show me the solution

Use the basic gene annotation in GTF format (e.g., gencode.vM38.primary_assembly.basic.annotation.gtf.gz).
It contains essential gene and transcript models without pseudogenes or other biotypes not typically used for RNA-seq quantification.

Challenge

Reference genome

Select the corresponding Genome sequence (GRCm39) FASTA (e.g., GRCm39.primary_assembly.genome.fa.gz).

Show me the solution

Transcript sequences

Transcript level quantification requires a transcript FASTA file consistent with the annotation. GENCODE provides transcript sequences for all transcripts in GRCm39 (gencode.vM38.transcripts.fa.gz).

Challenge

Transcript sequence file

Where on the GENCODE page can you find the transcript FASTA file?

Show me the solution

The transcript FASTA file is available in the same release directory as the genome and annotation files (under FASTA files). It is named gencode.vM38.transcripts.fa.gz.

Transcript sequences file in FASTA format

Downloading all reference files

BASH

cd ${SCRATCH}/rnaseq-workshop

GTFlink="https://ftp.ebi.ac.uk/pub/databases/gencode/Gencode_mouse/release_M38/gencode.vM38.primary_assembly.basic.annotation.gtf.gz"
GENOMElink="https://ftp.ebi.ac.uk/pub/databases/gencode/Gencode_mouse/release_M38/GRCm39.primary_assembly.genome.fa.gz"
CDSLink="https://ftp.ebi.ac.uk/pub/databases/gencode/Gencode_mouse/release_M38/gencode.vM38.transcripts.fa.gz"

# Download to 'data'
wget -P data ${GENOMElink}
wget -P data ${GTFlink}
wget -P data ${CDSLink}

# Uncompress
cd data
gunzip GRCm39.primary_assembly.genome.fa.gz
gunzip gencode.vM38.primary_assembly.basic.annotation.gtf.gz
gunzip gencode.vM38.transcripts.fa.gz

The transcript file header has multiple pieces of information which often interfere with downstream analysis. We create a simplified version containing only the transcript IDs.

BASH

grep ">" gencode.vM38.transcripts.fa |head -1
cut -f 1 -d "|" gencode.vM38.transcripts.fa > gencode.vM38.transcripts-clean.fa
grep ">" gencode.vM38.transcripts-clean.fa |head -1

Raw reads

The experiment we use investigates how NAT10 influences mouse heart development.

Study: The effects of NAT10 on heart development in mice
Organism: Mus musculus
Design: Nat10^flox/flox vs Nat10-CKO (n=3 per group), P10 hearts
Sequencing: Illumina NovaSeq X Plus, paired-end
BioProject: PRJNA1254208
GEO: GSE295332

Challenge

Where do you find FASTQ files?

GEO provides metadata and experimental details, but FASTQ files are stored in the SRA. How do you obtain the complete list of sequencing runs?

Show me the solution

Navigate to the SRA page for the BioProject and use Run Selector to list runs.
On the Run Selector page, click Accession List to download all SRR accession IDs.

Download FASTQ files with SRA Toolkit

BASH

sinteractive -A workshop -p cpu -N 1 -n 4 --time=1:00:00
cd ${SCRATCH}/rnaseq-workshop/data
module load biocontainers
module load sra-tools
while read SRR; do 
   fasterq-dump --threads 4 --progress $SRR;
   gzip ${SRR}_1.fastq
   gzip ${SRR}_2.fastq
done<SRR_Acc_List.txt

Since downloading may take a long time, we provide a pre-downloaded copy of all FASTQ files. The directory, after downloading, should look like this:

BASH

data
├── annot.tsv
├── gencode.vM38.primary_assembly.basic.annotation.gtf
├── gencode.vM38.transcripts.fa
├── GRCm39.primary_assembly.genome.fa
├── mart.tsv
├── SRR33253285_1.fastq.gz
├── SRR33253285_2.fastq.gz
├── SRR33253286_1.fastq.gz
├── SRR33253286_2.fastq.gz
├── SRR33253287_1.fastq.gz
├── SRR33253287_2.fastq.gz
├── SRR33253288_1.fastq.gz
├── SRR33253288_2.fastq.gz
├── SRR33253289_1.fastq.gz
├── SRR33253289_2.fastq.gz
├── SRR33253290_1.fastq.gz
├── SRR33253290_2.fastq.gz
└── SRR_Acc_List.txt

Key Points

RNA-seq requires three main inputs: FASTQ reads, a reference genome (FASTA), and gene annotation (GTF/GFF).
In lieu of a reference genome and annotation, transcript sequences (FASTA) can be used for transcript-level quantification.
Keeping files compressed and well-organized supports reproducible analysis.
Reference files must match in genome build and version.
Public data repositories such as GEO and SRA provide raw reads and metadata.
Tools like wget and fasterq-dump enable programmatic, reproducible data retrieval.

Content from Quality control of RNA-seq reads

Last updated on 2025-11-25 | Edit this page

Overview

Questions

How do we check the quality of raw RNA-seq reads?
What information do FastQC and MultiQC provide?
How do we decide if trimming is required?
What QC issues are common in Illumina RNA-seq data?

Objectives

Run FastQC and MultiQC on RNA-seq FASTQ files.
Understand how to interpret the important FastQC modules.
Learn which FastQC warnings are expected for RNA-seq.
Understand why trimming is usually unnecessary for standard RNA-seq.
Identify samples that may require closer inspection.

Introduction

Before mapping reads to a genome, it is important to evaluate the quality of the raw RNA-seq data.
Quality control helps reveal problems such as adapter contamination, low quality cycles, and unexpected technical issues.

In this episode, we will run FastQC and MultiQC on the mouse heart dataset and interpret the results.
We will also explain when trimming is useful and when it can be harmful for alignment based RNA-seq workflows.

FastQC evaluates individual FASTQ files. MultiQC summarizes all FastQC reports into one view, which is helpful for comparing replicates.

Callout

Why do we perform QC on FASTQ files?

Quality control answers several questions:

Are sequencing qualities stable across cycles?
Are adapters present at high levels?
Do different replicates show similar patterns?
Are there outliers that indicate failed libraries?

QC provides confidence that downstream steps will be reliable.

Running FastQC

FastQC generates an HTML report for each FASTQ file. For paired-end datasets, you can run it on both R1 and R2 together or separately.

We will create a directory for QC output and then run FastQC on all FASTQ files.

Run FastQC on your FASTQ files

Use the commands below to run FastQC on all reads in the data directory.

BASH

cd $SCRATCH/rnaseq-workshop
mkdir -p results/qc_fastq/

sinteractive -A workshop -p cpu -N 1 -n 4 --time=1:00:00
module load biocontainers
module load fastqc

fastqc data/*.fastq.gz --outdir results/qc_fastq/ --threads 4

BASH

qc
├── SRR33253285_1_fastqc.html
├── SRR33253285_1_fastqc.zip
├── SRR33253285_2_fastqc.html
├── SRR33253285_2_fastqc.zip
├── SRR33253286_1_fastqc.html
├── SRR33253286_1_fastqc.zip
├── SRR33253286_2_fastqc.html
├── SRR33253286_2_fastqc.zip
├── SRR33253287_1_fastqc.html
├── SRR33253287_1_fastqc.zip
├── SRR33253287_2_fastqc.html
├── SRR33253287_2_fastqc.zip
├── SRR33253288_1_fastqc.html
├── SRR33253288_1_fastqc.zip
├── SRR33253288_2_fastqc.html
├── SRR33253288_2_fastqc.zip
├── SRR33253289_1_fastqc.html
├── SRR33253289_1_fastqc.zip
├── SRR33253289_2_fastqc.html
├── SRR33253289_2_fastqc.zip
├── SRR33253290_1_fastqc.html
├── SRR33253290_1_fastqc.zip
├── SRR33253290_2_fastqc.html
└── SRR33253290_2_fastqc.zip

Understanding the FastQC report

FastQC produces several diagnostic modules. RNA-seq users often misinterpret certain modules, so we focus on the ones that matter most.

Callout

Which FastQC modules are important for RNA-seq?

Most informative modules:

Per base sequence quality
Per sequence quality scores
Per base sequence content
Per sequence GC content
Adapter content
Overrepresented sequences

Modules that commonly show warnings for RNA-seq, even when libraries are fine:

Per sequence GC content (RNA-seq reflects transcript GC bias, not whole genome)
K-mer content (poly A tails and priming artifacts often trigger warnings)

Warnings do not always indicate problems. Context and comparison across samples matter more than individual icons.

Key modules explained

Below are guidelines for interpreting the most useful modules.

Per base sequence quality

Shows the distribution of quality scores at each position across all reads. You want high and stable quality across the read. A small quality drop at the end of R2 is common and usually not a problem.

Per tile sequence quality

Reports whether specific regions of the flowcell produced lower quality reads. Uneven tiles can indicate instrument issues or bubble formation, but this module rarely shows problems in modern NovaSeq data.

Per sequence quality scores

Shows how many reads have high or low overall quality. A good dataset has most reads with high scores and very few reads with low quality.

Per base sequence content

Shows the proportion of A, C, G, and T at each cycle. For genomic resequencing, all four lines should be close to 25 percent. RNA-seq usually shows imbalanced composition in early cycles due to biased priming and transcript composition. Mild imbalance at the start of the read is normal and does not require trimming.

Per sequence GC content

Shows the GC distribution across all reads. RNA-seq often fails this test because expressed transcripts are not GC neutral. Only pronounced multi-modal or extremely shifted distributions are concerning, for example when contamination from another organism is suspected.

Per base N content

Reports the percentage of unidentified bases (N) at each position. This should be near zero for modern Illumina runs. High N content indicates failed cycles or poor sequencing chemistry.

Sequence length distribution

Shows the distribution of read lengths. Standard RNA-seq libraries should have a narrow peak at the expected read length. Variable lengths usually appear after trimming or for specialized protocols such as small RNA.

Sequence duplication levels

Reports how many reads are exact duplicates. Some duplication is normal in RNA-seq because highly expressed genes produce many identical fragments. Very high duplication, especially combined with low library size, can indicate low library complexity or overamplification.

Overrepresented sequences

Lists specific sequences that occur more often than expected. Common sources include ribosomal RNA fragments, poly A tails, adapter sequence, or reads from highly expressed transcripts. Some overrepresented sequences are expected, but very high levels may indicate contamination or poor rRNA depletion.

Adapter content

Shows whether any read cycles contain adapter sequences. Small amounts at the extreme ends are common and usually harmless. Consistently high adapter content across many cycles suggests short inserts or poorly prepared libraries.

Aggregating reports with MultiQC

FastQC generates a separate HTML file per sample. MultiQC consolidates all reports and allows easy comparison of replicates.

Run MultiQC on the FastQC outputs

Generate a single combined report using MultiQC.

BASH

cd rnaseq-workshop
module load biocontainers
module load multiqc

cd $SCRATCH/rnaseq-workshop
multiqc results/qc_fastq/ -o results/qc_fastq/

Example MultiQC summary across all samples

Trimming adapter sequences

Before deciding whether to trim reads, it is essential to understand what adapters are and why they appear in sequencing data.

Adapters are short synthetic DNA sequences ligated to both ends of each fragment during library preparation. They provide primer binding sites needed for cluster amplification and sequencing. Ideally, the sequenced fragment is long enough that the machine reads only biological DNA. However, when the insert is shorter than the read length, the sequencer runs into the adapter sequence, which produces adapter contamination at the ends of reads.

Many users assume that adapters must always be trimmed, but this is not true for typical RNA-seq workflows.

Discussion

Do we need to trim reads for standard RNA-seq workflows?

For alignment based RNA-seq, adapter trimming is usually not required. Modern aligners such as HISAT2 and STAR detect and soft clip adapter sequence automatically. Soft-clipped bases do not contribute to mismatches or mapping penalties, so adapters generally do not harm alignment quality.

In contrast, unnecessary trimming can introduce new problems:

shorter reads and lower mapping power
uneven trimming across samples and artificial differences
reduction in read complexity
broken read pairing for paired-end sequencing
more parameters to track for reproducibility

Trimming is recommended only in specific situations:

very short inserts, for example degraded RNA or ancient DNA
adapter sequence dominating read tails, for example more than 10-15 percent adapter content
transcript assembly workflows such as StringTie or Scallop, which benefit from uniform read lengths
small RNA or miRNA libraries, where inserts are intentionally very short and adapter presence is guaranteed

Exercise: evaluate the QC results

Challenge

Inspect your MultiQC report

Using the MultiQC HTML report, answer:

Do all replicates show similar per base sequence quality profiles?
Is adapter content high enough to impact analysis?
Is any sample behaving differently from the rest, for example in number of reads, quality, duplication, or GC percentage?

Show me the solution

Typical interpretation for this dataset (students should compare with their own):

All replicates show high, stable sequence quality.
Adapter signal appears only at the extreme ends of reads for selected samples and is less than 4 percent.
No samples show concerning patterns or biases.
Trimming is therefore unnecessary for this dataset.

What if trimming is required?

Optional reference: how to trim adapters with fastp (not part of the hands-on)

Sometimes you may encounter datasets where trimming is appropriate, for example short inserts, small RNA, or visibly high adapter content in QC. Although we will not trim reads in this workshop, here is a minimal example you can use in your own projects if trimming becomes necessary.

fastp is a fast, widely used tool that can automatically detect adapters, trim them, filter low quality reads, and generate QC reports.

Example: trimming paired-end reads with `fastp`

BASH

module load biocontainers
module load fastp

fastp \
  -i SRRXXXXXXX_1.fastq.gz \
  -I SRRXXXXXXX_2.fastq.gz \
  -o SRRXXXXXXX_1.trimmed.fastq.gz \
  -O SRRXXXXXXX_2.trimmed.fastq.gz \
  --detect_adapter_for_pe \
  --thread 8 \
  --html fastp_report.html \
  --json fastp_report.json

What this command does

automatically detects adapters (--detect_adapter_for_pe)
trims low quality bases (default behaviour)
keeps paired-end reads synchronized
writes trimmed output (*.trimmed.fastq.gz)
generates QC reports (HTML and JSON)

When to use this

Use trimming only if:

QC shows strong adapter presence
your library prep is known to produce short inserts
you are working with small RNA or degraded RNA
you are doing transcript assembly where uniform read length matters

For standard RNA-seq, trimming is usually unnecessary.

Summary

Key Points

FastQC and MultiQC provide essential diagnostics for RNA-seq data.
Some FastQC warnings are normal for RNA-seq and do not indicate problems.
Aligners soft clip adapters, so trimming is usually unnecessary for alignment based RNA-seq.
QC helps detect outliers before alignment and quantification.

Content from A. Genome-based quantification (STAR + featureCounts)

Last updated on 2025-11-25 | Edit this page

Overview

Questions

How do we map RNA-seq reads to a reference genome?
How do we determine library strandness before alignment?
How do we quantify reads per gene using featureCounts?
How do we submit mapping jobs on RCAC clusters with SLURM?

Objectives

Check RNA-seq library strandness using Salmon.
Index a reference genome for STAR.
Map reads to the genome using STAR.
Quantify gene level counts with featureCounts.
Interpret basic alignment and counting statistics.

Introduction

In this episode we move from raw FASTQ files to genome based gene counts.
This workflow has three major steps:

Determine library strandness: STAR does not infer strandness automatically. featureCounts requires the correct setting.
Map reads to the genome using STAR: STAR is a splice aware aligner designed for RNA-seq data.
Count mapped reads per gene using featureCounts

This represents one of the two standard quantification strategies in RNA-seq.
The alternative method (B. transcript based quantification with Salmon or Kallisto) will be covered in Episode 4B.

Step 1: Checking RNA-seq library strandness

Callout

Why strandness matters

RNA-seq libraries can be:

unstranded
stranded forward
stranded reverse

Stranded protocols are common in modern kits, but old GEO and SRA datasets are often unstranded.
featureCounts must be given the correct strand setting. If the wrong setting is used, most reads will be counted against incorrect genes.

STAR does not infer strandness automatically, so we must determine it before alignment.

Detecting strandness with Salmon

Salmon can infer strandness directly from raw FASTQ files without any alignment.
We run Salmon with -l A, which tells it to explore all library types.

BASH

sinteractive -A workshop -q standby -p cpu -N 1 -n 4 --time=1:00:00

cd $SCRATCH/rnaseq-workshop
mkdir -p results/strand_check

module load biocontainers
module load salmon

salmon index --transcripts data/gencode.vM38.transcripts-clean.fa \
             --index data/salmon_index \
             --threads 4

salmon quant --index data/salmon_index \
             --libType A \
             --mates1 data/SRR33253286_1.fastq.gz \
             --mates2 data/SRR33253286_2.fastq.gz \
             --output results/strand_check \
             --threads 4

The important output is in results/strand_check/lib_format_counts.json:

What does this file look like?

JSON

{
    "read_files": "[ data/SRR33253286_1.fastq.gz, data/SRR33253286_2.fastq.gz]",
    "expected_format": "IU",
    "compatible_fragment_ratio": 1.0,
    "num_compatible_fragments": 21088696,
    "num_assigned_fragments": 21088696,
    "num_frags_with_concordant_consistent_mappings": 19116702,
    "num_frags_with_inconsistent_or_orphan_mappings": 2510542,
    "strand_mapping_bias": 0.5117980078362889,
    "MSF": 0,
    "OSF": 0,
    "ISF": 9783890,
    "MSR": 0,
    "OSR": 0,
    "ISR": 9332812,
    "SF": 1234245,
    "SR": 1276297,
    "MU": 0,
    "OU": 0,
    "IU": 0,
    "U": 0
}

Interpreting lib_format_counts.json

This JSON file reports Salmon’s automatic library type detection.

"expected_format": "IU": This is Salmon’s conclusion. “I” means Inward (correct for paired-end reads) and “U” means Unstranded.

"strand_mapping_bias": 0.512: This is the key evidence. A value near 0.5 (50%) indicates that reads mapped equally to both the sense and antisense strands, the definitive sign of an unstranded library.

"ISF" and "ISR": These are the counts for Inward-Sense-Forward (9.78M) and Inward-Sense-Reverse (9.33M) fragments. Because these values are almost equal, they confirm the ~50/50 split seen in the strand bias.

Common results:

Code	Meaning
IU	unstranded
ISR	stranded reverse (paired end)
ISF	stranded forward (paired end)
SR	stranded reverse (single end)
SF	stranded forward (single end)

Conclusion: The data is unstranded. We will record this and supply it to STAR/featureCounts.

Step 2: Preparing the reference genome for alignment

STAR requires a genome index. Indexing builds a searchable data structure that speeds up alignment.

Required input:

genome FASTA file
annotation file (GTF or GFF3)
output directory for the index

Optionally, annotation improves splice junction detection and yields better alignments.

Callout

Why STAR?

Splice-aware
Extremely fast
Produces high-quality alignments
Well supported and widely used
Compatible with downstream tools such as featureCounts, StringTie, and RSEM

Optional alternatives include HISAT2 and Bowtie2, but STAR remains the standard for high-depth Illumina RNA-seq.

Challenge

Exercise: Read the STAR manual

Check the “Generating genome indexes” section of the
STAR manual

Which options are required for indexing?

Show me the solution

Key options:

--runMode genomeGenerate
--genomeDir directory to store the index
--genomeFastaFiles the FASTA file
--sjdbGTFfile annotation (recommended)
--runThreadN number of threads
--sjdbOverhang read length minus one (100 is a safe default)

Indexing the Genome with STAR

Below is a minimal SLURM job script for building the STAR genome index. Create a file named index_genome.sh in $SCRATCH/rcac_rnaseq/scripts with this content:

BASH

#!/bin/bash
#SBATCH --nodes=1
#SBATCH --ntasks=1
#SBATCH --cpus-per-task=20
#SBATCH --account=workshop
#SBATCH --partition=cpu
#SBATCH --time=1:00:00
#SBATCH --job-name=star_index
#SBATCH --output=cluster-%x.%j.out
#SBATCH --error=cluster-%x.%j.err

module load biocontainers
module load star

data_dir="${SCRATCH}/rnaseq-workshop/data"
genome="${data_dir}/GRCm39.primary_assembly.genome.fa"
gtf="${data_dir}/gencode.vM38.primary_assembly.basic.annotation.gtf"
indexdir="${data_dir}/star_index"

STAR \
    --runMode genomeGenerate \
    --runThreadN ${SLURM_CPUS_ON_NODE} \
    --genomeDir ${indexdir} \
    --genomeFastaFiles ${genome} \
    --sjdbGTFfile ${gtf}

Submit the job:

BASH

sbatch index_genome.sh

Indexing requires about 30 to 45 minutes.

What will the output look like?

The index directory is now ready for use in mapping and should have contents like this: Location: $SCRATCH/rcac_rnaseq/data Contents of star_index/:

BASH

star_index/
├── chrLength.txt
├── chrNameLength.txt
├── chrName.txt
├── chrStart.txt
├── exonGeTrInfo.tab
├── exonInfo.tab
├── geneInfo.tab
├── Genome
├── genomeParameters.txt
├── Log.out
├── SA
├── SAindex
├── sjdbInfo.txt
├── sjdbList.fromGTF.out.tab
├── sjdbList.out.tab
└── transcriptInfo.tab

Callout

Do I need the GTF for indexing?

No, but it is strongly recommended.
Including it improves splice junction detection and increases mapping accuracy.

Step 3: Mapping reads with STAR

Once the genome index is ready, we can align reads.

For aligning reads, important STAR options are:

--runThreadN
--genomeDir
--readFilesIn
--readFilesCommand zcat for gzipped FASTQ files
--outSAMtype BAM SortedByCoordinate
--outSAMunmapped Within

STAR’s defaults are tuned for mammalian genomes. If working with plants, fungi, or repetitive genomes, additional tuning may be required.

Challenge

Exercise: Write a STAR alignment command

Write a basic STAR alignment command using your genome index and paired FASTQ files.

Show me the solution

BASH

STAR \
    --runThreadN 12 \
    --genomeDir data/star_index \
    --readFilesIn data/SRR1234567_1.fastq.gz data/SRR1234567_2.fastq.gz \
    --readFilesCommand zcat \
    --outFileNamePrefix results/mapping/SRR1234567 \
    --outSAMtype BAM SortedByCoordinate \
    --outSAMunmapped Within

Mapping many FASTQ files with a SLURM array

When having multiple samples, array jobs simplify submission.

First, we will create samples.txt file with just the sample names

BASH

cd $SCRATCH/rnaseq-workshop/data
ls *_1.fastq.gz | sed 's/_1.fastq.gz//' > ${SCRATCH}/rnaseq-workshop/scripts/samples.txt

BASH

#!/bin/bash
#SBATCH --nodes=1
#SBATCH --ntasks=1
#SBATCH --cpus-per-task=20
#SBATCH --account=workshop
#SBATCH --partition=cpu
#SBATCH --time=8:00:00
#SBATCH --job-name=read_mapping
#SBATCH --array=1-6
#SBATCH --output=cluster-%x.%j.out
#SBATCH --error=cluster-%x.%j.err

module load biocontainers
module load star

FASTQ_DIR="$SCRATCH/rnaseq-workshop/data"
GENOME_INDEX="$SCRATCH/rnaseq-workshop/data/star_index"
OUTDIR="$SCRATCH/rnaseq-workshop/results/mapping"

mkdir -p $OUTDIR

SAMPLE=$(sed -n "${SLURM_ARRAY_TASK_ID}p" samples.txt)

R1=${FASTQ_DIR}/${SAMPLE}_1.fastq.gz
R2=${FASTQ_DIR}/${SAMPLE}_2.fastq.gz

STAR \
    --runThreadN ${SLURM_CPUS_ON_NODE} \
    --genomeDir $GENOME_INDEX \
    --readFilesIn $R1 $R2 \
    --readFilesCommand zcat \
    --outFileNamePrefix $OUTDIR/$SAMPLE \
    --outSAMunmapped Within \
    --outSAMtype BAM SortedByCoordinate

Submit:

BASH

sbatch map_reads.sh

Discussion

Why use an array job?

Think about this:

We have six samples, each with two FASTQ files
Having six separate slurm jobs means:
- Copy-pasting the same command six times
- Higher chance of typos or mistakes
if mapping with single slurm file, sequential execution = slower
Samples can run independently

Why is a job array a good choice?

Arrays allow us to run identical commands across many inputs:

All samples are processed the same way
Runs in parallel
Simplifies submission
Reduces copy-paste errors
Minimizes runtime cost on the cluster

Step 4: Assessing alignment quality

STAR produces several useful files, particularly:

Log.final.out
Contains mapping statistics, including % uniquely mapped.
Aligned.sortedByCoord.out.bam
Ready for downstream quantification.

To summarize alignment statistics across samples, we use MultiQC:

BASH

cd $SCRATCH/rnaseq-workshop
module load biocontainers
module load multiqc

multiqc results/mapping -o results/qc_alignment

Key metrics to review

Uniquely mapped reads — typically 60–90% is good
Multi-mapped reads — depends on genome, but >20% is concerning
Unmapped reads — check reasons (mismatches? too short?)
Mismatch rate — high values indicate quality or contamination issues

Challenge

Exercise: Examine your alignment metrics

Using your MultiQC alignment report:

Which sample has the highest uniquely mapped percentage?
Are any samples clear outliers?
Does mapped/unmapped reads appear consistent across samples?

Show me the solution

Example interpretation for this dataset:

All samples have ~80% uniquely mapped reads.
No sample is an outlier.
mapped/unmapped reads consistent across samples

Step 5: Quantifying gene counts with featureCounts

Once the reads have been aligned and sorted, we can quantify how many read pairs map to each gene. This gives us gene-level counts that we will later use for differential expression. For alignment-based RNA-seq workflows, featureCounts is the recommended tool because it is fast, robust, and compatible with most gene annotation formats.

featureCounts uses two inputs:

A set of aligned BAM files (sorted by coordinate)
A GTF annotation file describing gene models

It assigns each read (or read pair) to a gene based on the exon boundaries.

Callout

Important: stranded or unstranded?

featureCounts needs to know whether your dataset is stranded.

-s 0 unstranded
-s 1 stranded forward
-s 2 stranded reverse

From our Salmon strandness check, this dataset behaves as unstranded, so we will use -s 0.

Running featureCounts

We will create a new directory for count output:

BASH

cd $SCRATCH/rnaseq-workshop/scripts
mkdir -p $SCRATCH/rnaseq-workshop/results/counts

Below is an example SLURM script to count all BAM files at once.

BASH

#!/bin/bash
#SBATCH --nodes=1
#SBATCH --ntasks=1
#SBATCH --cpus-per-task=16
#SBATCH --account=workshop
#SBATCH --partition=cpu
#SBATCH --time=1:00:00
#SBATCH --job-name=featurecounts
#SBATCH --output=cluster-%x.%j.out
#SBATCH --error=cluster-%x.%j.err

module load biocontainers
module load subread

GTF="$SCRATCH/rnaseq-workshop/data/gencode.vM38.annotation.gtf"
BAMS="$SCRATCH/rnaseq-workshop/results/mapping/*.bam"
output_dir="$SCRATCH/rnaseq-workshop/results/counts"

featureCounts \
  -T ${SLURM_CPUS_PER_TASK} \
  -p \
  -s 0 \
  -t exon \
  -g gene_id \
  -a ${GTF} \
  -o ${output_dir}/gene_counts.txt \
  ${BAMS}

Submit the job:

BASH

sbatch count_features.sh

gene_counts.txt will contain:

one row per gene
one column per sample

Callout

Generate a clean count matrix

BASH

cd $SCRATCH/rnaseq-workshop/results/counts
cut -f 1,7- gene_counts.txt |\
  sed 's+/scratch/negishi/'$USER'/rnaseq-workshop/results/mapping/++g' |\
  grep -v "^#" |\
  sed 's/Aligned.sortedByCoord.out.bam//g' \
  > gene_counts_clean.txt

Step 6: QC on counts

Before proceeding to differential expression, it is good practice to perform some QC on the count data. We can again use MultiQC to summarize featureCounts statistics:

BASH

cd $SCRATCH/rnaseq-workshop
mkdir -p results/qc_counts
module load biocontainers
module load multiqc
multiqc results/counts -o results/qc_counts

Challenge

Exercise: Examine your counting metrics

Using your MultiQC featureCounts report:

Which sample has the lowest percent assigned, and what are common reasons for lower assignment?
Which unassigned category is the largest, and what does it indicate?
Does the sample with the most assigned reads also have the highest percent assigned?
What does “Unassigned: No Features” mean, and why might it occur?
Are multi-mapped reads a concern for RNA-seq differential expression?

Show me the solution

Example interpretation for this dataset:

SRR33253289 has the lowest assignment rate. Causes include low complexity, incomplete annotation, or more intronic reads.
Unmapped or No Features dominate. These reflect reads that do not align or do not overlap annotated exons.
No. Total assigned reads depend on depth, while percent assigned reflects library quality.
Reads mapped outside annotated exons, often from intronic regions, unannotated transcripts, or incomplete annotation.
Multi-mapping is expected for paralogs and repeats. It is usually fine as long as the rate is consistent across samples.

Summary

Key Points

STAR is a fast splice-aware aligner widely used for RNA-seq.
Genome indexing requires FASTA and optionally GTF for improved splice detection.
Mapping is efficiently performed using SLURM array jobs.
Alignment statistics (from Log.final.out + MultiQC) must be reviewed.
Strandness should be inferred using aligned BAM files.
Final BAM files are ready for downstream counting.
featureCounts is used to obtain gene-level counts for differential expression.
Exons are counted and summed per gene.
The output is a simple matrix for downstream statistical analysis.

Content from B. Transcript-based quantification (Salmon)

Last updated on 2025-11-25 | Edit this page

Overview

Questions

How do we quantify expression without genome alignment?
What inputs does Salmon require?
How do we run Salmon for paired-end RNA-seq data?
How do we interpret transcript-level outputs (TPM, NumReads)?
How do we summarize transcripts to gene-level counts using tximport?

Objectives

Build a transcriptome index for Salmon.
Quantify transcript abundance directly from FASTQ files.
Understand key Salmon output files.
Summarize transcript-level estimates to gene-level counts.
Prepare counts for downstream differential expression.

Introduction

In the previous episode, we mapped reads to the genome using STAR and generated gene-level counts with featureCounts.

In this episode, we use an alternative quantification strategy:

Build a transcriptome index
Quantify expression directly from FASTQ files using Salmon (without alignment)
Summarize transcript estimates back to gene-level counts

This workflow is faster, uses less storage, and models transcript-level uncertainty.

Callout

When should I use transcript-based quantification?

Salmon is ideal when:

You do not need alignment files.
You want transcript-level TPMs.
You want fast quantification with bias correction.
Storage is limited (no BAM files generated).

It is not ideal if you need splice junctions, variant calling, or visualization in IGV (all those depends on alignment files).

Step 1: Preparing the transcriptome reference

Salmon requires a transcriptome FASTA file (all annotated transcripts). We previously downloaded transcripts file from GENCODE for this purpose (gencode.vM38.transcripts.fa).

Callout

Why transcriptome choice matters

Transcript-level quantification inherits all assumptions of the annotation.
Missing or incorrect transcripts → biased TPM estimates.

Building the Salmon index

A Salmon index is lightweight and quick to build. You may have already built this in Episode 4A.

BASH

sinteractive -A workshop -p cpu -N 1 -n 4 --time=1:00:00

cd $SCRATCH/rnaseq-workshop
mkdir -p data/salmon_index

module load biocontainers
module load salmon

salmon index \
    --transcripts data/gencode.vM38.transcripts-clean.fa \
    --index data/salmon_index \
    --threads 4

This creates a directory containing Salmon’s k-mer index of all transcripts.

What will the index contain?

salmon_index/
├── complete_ref_lens.bin        # IMPORTANT: full transcript lengths used in TPM/effLength calculations
├── ctable.bin                   # internal k-mer lookup table (not user-relevant)
├── ctg_offsets.bin              # internal offsets for reference sequences
├── duplicate_clusters.tsv       # internal duplicate k-mer cluster info
├── info.json                    # IMPORTANT: index metadata (k-mer size, transcripts, build parameters)
├── mphf.bin                     # minimal perfect hash function for fast lookup
├── pos.bin                      # internal k-mer position table
├── pre_indexing.log             # IMPORTANT: log of preprocessing and FASTA parsing steps
├── rank.bin                     # internal rank/select bitvector structure
├── refAccumLengths.bin          # cumulative reference lengths (internal)
├── ref_indexing.log             # IMPORTANT: main indexing log; good for debugging build problems
├── reflengths.bin               # IMPORTANT: transcript lengths used in quantification math
├── refseq.bin                   # IMPORTANT: transcript sequences stored in binary format (core index content)
├── seq.bin                      # internal sequence encoding structures
└── versionInfo.json             # IMPORTANT: Salmon version + environment info for reproducibility

Step 2: Quantifying transcript abundances

The core Salmon command is salmon quant. We use the transcript index and paired-end FASTQ files.

BASH

mkdir -p $SCRATCH/rnaseq-workshop/results/salmon_quant

Example command:

BASH

salmon quant \
    --index salmon_index \
    --libType IU \
    --mates1 data/SRR33253286_1.fastq.gz \
    --mates2 data/SRR33253286_2.fastq.gz \
    --output salmon_quant/SRR33253286 \
    --threads 16 \
    --validateMappings \
    --seqBias \
    --gcBias

Important flags:

--libType IU → unstranded, inward-oriented paired-end (Replace with ISR, ISF, etc. based on your strandness.)
--validateMappings → improves mapping accuracy
--seqBias, --gcBias → corrects sequence- and GC-related biases

Callout

Correct library type is essential

A wrong --libType silently distorts transcript abundances. Use Salmon’s strandness detection from Episode 4A if unsure.

Step 3: Running Salmon for many samples

Again, we use a SLURM array.

First, create a sample list (or reuse the one from Episode 4A):

BASH

cd $SCRATCH/rnaseq-workshop/data
ls *_1.fastq.gz | sed 's/_1.fastq.gz//' > $SCRATCH/rnaseq-workshop/scripts/samples.txt

Array job:

BASH

#!/bin/bash
#SBATCH --nodes=1
#SBATCH --ntasks=1
#SBATCH --cpus-per-task=16
#SBATCH --account=workshop
#SBATCH --partition=cpu
#SBATCH --time=4:00:00
#SBATCH --job-name=salmon_quant
#SBATCH --array=1-6
#SBATCH --output=cluster-%x.%j.out
#SBATCH --error=cluster-%x.%j.err

module load biocontainers
module load salmon

DATA="$SCRATCH/rnaseq-workshop/data"
INDEX="$SCRATCH/rnaseq-workshop/data/salmon_index"
OUT="$SCRATCH/rnaseq-workshop/results/salmon_quant"

mkdir -p $OUT

SAMPLE=$(sed -n "${SLURM_ARRAY_TASK_ID}p" samples.txt)

R1=${DATA}/${SAMPLE}_1.fastq.gz
R2=${DATA}/${SAMPLE}_2.fastq.gz

salmon quant \
  --index $INDEX \
  --libType IU \
  --mates1 $R1 \
  --mates2 $R2 \
  --output $OUT/$SAMPLE \
  --threads ${SLURM_CPUS_ON_NODE} \
  --validateMappings \
  --seqBias \
  --gcBias

Submit:

BASH

sbatch quant_salmon.sh

The typical run time is ~5 minutes per sample.

Discussion

Why use an array job here?

Unlike the STAR workflow in 4A, Salmon runs quickly and produces small output. The advantage of array jobs here is not speed, but consistency, all samples are quantified with identical parameters and metadata. This ensures that transcript-level TPM and count estimates are directly comparable across samples.

Inspect results

After Salmon finishes, each sample directory contains a small set of output files.
Only a few of these are important for routine analysis:

SRR33253285/
├── quant.sf                       # IMPORTANT: transcript-level TPM, counts, effective lengths
├── lib_format_counts.json         # IMPORTANT: inferred library type (IU, ISR, ISF)
├── aux_info
│   ├── meta_info.json             # IMPORTANT: mapping rate, fragment counts, bias flags
│   ├── fld.gz                     # fragment length distribution (binary format)
│   ├── expected_bias.gz           # bias model used for correction
│   ├── observed_bias.gz           # observed sequence and GC bias
│   └── ambig_info.tsv             # ambiguous equivalence class summaries (advanced QC only)
├── libParams
│   └── flenDist.txt               # fragment length distribution in a human-readable format
├── cmd_info.json                  # command and parameters used to run Salmon
└── logs
└── salmon_quant.log               # main log file for troubleshooting

What is inside quant.sf?

This file contains the transcript-level quantification results:

Name: transcript ID
Length: transcript length
EffectiveLength: length adjusted for fragment distribution and bias
TPM: within-sample normalized abundance
NumReads: estimated number of reads assigned to that transcript

Example:

Name                       Length  EffectiveLength TPM             NumReads
ENSMUST00000193812.2       1070    775.000         0.000000        0.000
ENSMUST00000082908.3       110     110.000         0.000000        0.000
ENSMUST00000162897.2       4153    3483.630        0.013994        1.000
ENSMUST00000159265.2       2989    2694.000        0.000000        0.000
ENSMUST00000070533.5       3634    3339.000        0.000000        0.000
ENSMUST00000192857.2       480     185.000         0.000000        0.000
ENSMUST00000195335.2       2819    2524.000        0.000000        0.000
ENSMUST00000192336.2       2233    1938.000        0.000000        0.000
ENSMUST00000194099.2       2309    2014.000        0.000000        0.000

Key QC metrics to review

Open aux_info/meta_info.json and check:

percent_mapped: proportion of fragments assigned to transcripts
library_types: Salmon inferred library orientation, which should match your --libType
fragment length statistics: mean and standard deviation of inferred fragment lengths

These metrics provide a first-pass sanity check before summarizing the results to gene-level counts.

Callout

What does Salmon count?

NumReads is a model-based estimate, not raw counts of reads. TPM values are within-sample normalized and should not be directly compared across samples.

Step 4: Summarizing to gene-level counts (`tximport`)

Most differential expression tools (DESeq2, edgeR) require gene-level counts. We convert transcript estimates → gene-level matrix.

Prepare tx2gene mapping

We will need a mapping file that links the transcript IDs to gene IDs. The gencode.vM38.primary_assembly.basic.annotation.gtf file contains this information.

BASH

cd $SCRATCH/rnaseq-workshop/data
awk -F'\t' '$3=="transcript" {
    match($9, /transcript_id "([^"]+)"/, tx);
    match($9, /gene_id "([^"]+)"/, gene);
    print tx[1] "\t" gene[1]
}' gencode.vM38.primary_assembly.basic.annotation.gtf > tx2gene.tsv

Next, we need to process this mapping in R. Load the modules and start the R session, first:

BASH

module load biocontainers
module load r-rnaseq
R

In the R session, read in the tx2gene mapping:

R

setwd(paste0(Sys.getenv("SCRATCH"),"/rnaseq-workshop"))
library(readr)
library(tximport)

tx2gene <- read_tsv("data/tx2gene.tsv", col_types = "cc", col_names = FALSE)
samples <- read_tsv("scripts/samples.txt", col_names = FALSE)
files <- file.path("results/salmon_quant", samples$X1, "quant.sf")
names(files) <- samples$X1

txi <- tximport(files,
                type = "salmon",
                tx2gene = tx2gene,
                countsFromAbundance = "lengthScaledTPM")
saveRDS(txi, file = "results/salmon_quant/txi.rds")

txi$counts now contains gene-level counts suitable for DESeq2.

R

> head(txi$counts)
                      SRR33253285 SRR33253286 SRR33253287 SRR33253288
ENSMUSG00000000001.5     473.4426  553.784929  826.996853  676.910126
ENSMUSG00000000003.16      0.0000    0.000000    0.000000    0.000000
ENSMUSG00000000028.16     40.5783   33.966518   55.305501   97.845071
ENSMUSG00000000031.20      0.0000    0.000000    0.000000    0.000000
ENSMUSG00000000037.18     19.2220    9.599673   16.951493   20.226549
ENSMUSG00000000049.12      0.0000    5.022073    1.984551    1.016411
                      SRR33253289 SRR33253290
ENSMUSG00000000001.5   724.117586  813.848375
ENSMUSG00000000003.16    0.000000    0.000000
ENSMUSG00000000028.16   41.691461   63.812279
ENSMUSG00000000031.20    0.000000    0.000000
ENSMUSG00000000037.18   19.680740   18.945058
ENSMUSG00000000049.12    0.993724    9.186882

Callout

Why use lengthScaledTPM?

This method leverages Salmon’s bias correction while generating stable counts for statistical modeling.

Step 5: QC of quantification metrics

Use MultiQC to verify:

BASH

cd $SCRATCH/rnaseq-workshop
module load biocontainers
module load multiqc

multiqc results/salmon_quant -o results/qc_salmon

Inspect:

mapping rates
fragment length distributions
library type consistency

Challenge

Exercise: examine your Salmon outputs

Using your MultiQC summary and Salmon fragment length plots:

What is the percent aligned for each sample, and are any samples outliers?
Is the inferred library type consistent across samples?
How similar are the fragment length distributions across samples?
Does the fragment length curve show unusual peaks or multimodality?
Do CFR and M bias values suggest any strand or positional bias?

Show me the solution

Example interpretation for this dataset:

All samples show about 89 to 91 percent aligned. No sample is an outlier and the range is narrow.
The inferred library type is consistent across samples and matches an unstranded protocol.
Fragment length distributions are nearly identical. All samples peak around the same length and have similar spread.
The fragment length curves are smooth and unimodal. There are no secondary peaks or irregular shoulders.
CFR is 100 percent for all samples and M bias is about 0.5, indicating balanced, strand-neutral coverage without detectable end bias.

Summary

Key Points

Salmon performs fast, alignment-free transcript quantification.
A transcriptome FASTA is required for building the index.
salmon quant uses FASTQ files directly with bias correction.
Transcript-level outputs include TPM and estimated counts.
tximport converts transcript estimates to gene-level counts.
Gene-level counts from Salmon are suitable for DESeq2.

Content from Gene-level QC and differential expression (DESeq2)

Last updated on 2025-11-25 | Edit this page

Overview

Questions

How do we explore RNA seq count data before running DESeq2.
How do we restrict analysis to protein coding genes.
How do we perform differential expression with DESeq2.
How do we visualize sample relationships and DE genes.
How do we save a useful DE results table with annotation and expression values.

Objectives

Import count and sample metadata into R.
Attach biotype and gene symbol annotation to Ensembl gene IDs.
Explore library sizes, variance stabilizing transforms, distances, and PCA.
Run DESeq2 for a two group comparison.
Create a volcano plot with gene symbols.
Export joined DE results including normalized counts and annotation.

Introduction

In this episode we move from count matrices to differential expression analysis and visualization.

The workflow has four parts:

Import counts and sample metadata.
Exploratory analysis on protein coding genes (quality checks).
Differential expression analysis with DESeq2.
Visualization and export of annotated DE results.

Prerequisite

What you need for this episode

gene_counts_clean.txt generated from featureCounts
a samples.csv file describing the experimental groups
RStudio session on Scholar using Open OnDemand

While we created the count matrix in the previous episode, we still need to create a sample metadata file. This file should contain at least two columns: sample names matching the count matrix column names, and the experimental condition (e.g., control vs treatment). Simply copy/paste the following into a text file and save it in your scripts directory as samples.csv:

sample,condition
SRR33253285,Nat10-CKO
SRR33253286,Nat10-CKO
SRR33253287,Nat10-CKO
SRR33253288,Nat10floxflox
SRR33253289,Nat10flox/flox
SRR33253290,Nat10flox/flox

Also, create a direcotry for DESeq2 results:

BASH

mkdir -p results/deseq2

All analyses are performed in R through Open OnDemand

Step 1: Start Open OnDemand R session and prepare data

We will be using the OOD to start an interactive session on Scholar: https://gateway.negishi.rcac.purdue.edu

Login using your Purdue credentials after clicking the above link
Click on “Interactive Apps” in the top menu, and select “RStudio (Bioconductor)”
Fill the job submission form as follows:
- queue: workshop
- Walltime: 4
- Number of cores: 4
Click “Launch” and wait for the RStudio session to start
Once the session starts, you’ll will be able to click on “Connect to RStudio server” which will open the RStudio interface.

Once in RStudio, load the necessary libraries:

Setup: load packages and data

R

library(RColorBrewer)
library(EnsDb.Mmusculus.v79)
library(ensembldb)
library(tidyverse)
library(DESeq2)
library(ggplot2)
library(pheatmap)
library(readr)
library(dplyr)
library(ComplexHeatmap)
library(ggrepel)
library(vsn)
# Modify the path as needed
setwd("/scratch/negishi/aseethar/rnaseq-workshop")

countsFile <- "results/counts/gene_counts_clean.txt"
# prepare this file first!
groupFile  <- "scripts/samples.csv"

coldata <-
  read.csv(
    groupFile,
    row.names = 1,
    header = TRUE,
    stringsAsFactors = TRUE
  )
coldata$condition <- as.factor(coldata$condition)

cts <- as.matrix(read.delim(countsFile, row.names = 1, header = TRUE))

We also load gene level annotation tables prepared earlier.

R

mart <-
  read.csv(
    "data/mart.tsv",
    sep = "\t",
    header = TRUE
  )

annot <-
  read.csv(
    "data/annot.tsv",
    sep = "\t",
    header = TRUE
  )

How were these data prepared?

Since the gene IDs in the count matrix are Ensembl IDs, it will be hard to interpret the results without annotation. We will need to use the org.Mm.eg.db package for attaching gene symbols, so we can interpret the results better.0

R

library(biomaRt)
library(tidyverse)
# select Mart
ensembl = useMart("ENSEMBL_MART_ENSEMBL")
# we need to use the correct dataset for mouse
# we will query the available datasets and filter for mouse (GRCm39)
listDatasets(ensembl) %>%
  filter(str_detect(description, "GRCm39"))
# we can use the dataset name to set the dataset
ensembl = useDataset("mmusculus_gene_ensembl", mart = ensembl)
# our counts have Ensembl gene IDs with version numbers
# lets find how it is referred in biomaRt
listFilters(ensembl) %>%
  filter(str_detect(name, "ensembl"))
# so we will now set the filter type accordingly
filterType <- "ensembl_gene_id_version"
# from our counts data, get the list of gene IDs
filterValues <- rownames(counts)
# take a look at the available attributes (first 20)
listAttributes(ensembl) %>%
     head(20)
# we will retrieve ensembl gene id, ensembl gene id with version and external gene name
attributeNames <- c('ensembl_gene_id',
                    'ensembl_gene_id_version',
                    'external_gene_name',
                    'gene_biotype',
                    'description')
# get the annotation
annot <- getBM(
  attributes = attributeNames,
  filters = filterType,
  values = filterValues,
  mart = ensembl
)
saveRDS(annot, file = "results/counts/gene_annotation.rds")
# also save as tsv for future use
write.table(
  annot,
  file = "data/mart.tsv",
  sep = "\t",
  quote = FALSE,
  row.names = FALSE
)
# save a simplified annotation file with only essential columns
mart %>% 
select(ensembl_gene_id, ensembl_gene_id_version, external_gene_name) %>%
  write.table(
    file = "data/annot.tsv",
    sep = "\t",
    quote = FALSE,
    row.names = FALSE

The objects loaded here form the foundation for the entire workflow. cts contains raw gene counts from featureCounts. coldata contains the metadata that describes each sample and defines the comparison groups. mart and annot contain functional information for each gene so we can attach gene symbols and biotypes later in the analysis.

Callout

Sample metadata and design

coldata must have row names that match the count matrix column names. The condition column defines the groups for DESeq2 (Nat10-CKO vs Nat10flox/flox).

Exploratory analysis

Attach annotation and filter to protein coding genes

We first join counts with biotype information from mart and filter to protein coding genes.

R

cts_tbl <- cts %>%
  as.data.frame() %>%
  rownames_to_column("ensembl_gene_id_version")

cts_annot <- cts_tbl %>%
  left_join(mart, by = "ensembl_gene_id_version")

sum(is.na(cts_annot$gene_biotype))

Joining annotation allows us to move beyond raw Ensembl IDs. This step attaches gene symbols, biotypes, and descriptions so that later plots and tables are interpretable. At this point the dataset is still large and includes many noncoding genes, pseudogenes, and biotypes we won’t analyze further.

Summarize gene biotypes:

R

biotype_df <- cts_annot %>%
  dplyr::count(gene_biotype, name = "n") %>%
  arrange(desc(n))

ggplot(biotype_df, aes(x = reorder(gene_biotype, n), y = n)) +
  geom_col(fill = "steelblue") +
  coord_flip() +
  theme_minimal() +
  labs(
    x = "Gene biotype",
    y = "Number of genes",
    title = "Distribution of gene biotypes in annotation"
  )

Distribution of gene biotypes in annotation

Visualizing biotype composition gives a sense of how many genes belong to each functional class. RNA-seq quantifies all transcribed loci, but many biotypes (snRNA, TEC, pseudogenes) are not suitable for differential expression with DESeq2 because they tend to be lowly expressed or unstable. This motivates filtering to protein-coding genes for QC and visualization.

Filter to protein coding genes and lowly expressed genes:

R


cts_coding <- cts_annot %>%
    filter(gene_biotype == "protein_coding") %>%
    filter(rowSums(across(all_of(rownames(coldata)))) > 5) %>%
    select(ensembl_gene_id_version, all_of(rownames(coldata))) %>%
    column_to_rownames("ensembl_gene_id_version")

dim(cts_coding)

[1] 15839     6

Callout

Why filter protein coding genes and low expression

Noncoding biotypes (lncRNA, pseudogene, etc.) can be biologically important. Filtering lowly expressed genes reduces noise in exploratory analyses. Genes with near-zero counts across all samples inflate variance estimates and obscure real biological structure. The threshold of 5 total counts is a conservative filter that keeps genes with minimal but genuine expression while removing background noise.

Library size differences

We summarize total counts per sample for protein coding genes.

R

libSize <- colSums(cts_coding) %>%
  as.data.frame() %>%
  rownames_to_column("sample") %>%
  rename(total_counts = 2) %>%
  mutate(total_millions = total_counts / 1e6)

ggplot(libSize, aes(x = sample, y = total_counts)) +
  geom_col(fill = "steelblue") +
  theme_minimal() +
  labs(
    x = "Sample",
    y = "Total assigned reads",
    title = "Total reads per sample"
  ) +
  theme(
    axis.text.x = element_text(angle = 45, hjust = 1)
  )

Library size differences affect normalization. Although DESeq2 corrects for this through size factors, inspecting raw library sizes ensures no sample has abnormally low sequencing depth, which could compromise downstream analyses.

Create DESeq2 object for exploratory plots

R

dds <- DESeqDataSetFromMatrix(
  countData = cts_coding,
  colData   = coldata,
  design    = ~ condition
)

dds <- estimateSizeFactors(dds)

Inspect size factors vs library size:

R

ggplot(
  data.frame(
    libSize    = colSums(assay(dds)),
    sizeFactor = sizeFactors(dds),
    Group      = dds$condition
  ),
  aes(x = libSize, y = sizeFactor, col = Group)
) +
  geom_point(size = 5) +
  theme_bw() +
  labs(x = "Library size", y = "Size factor")

Variance stabilization and mean SD plots

Raw count data have a strong mean–variance dependency: low count genes have high variance, and high count genes show lower relative variability. This makes Euclidean distance and PCA misleading on raw counts. VST removes this dependency so that downstream clustering is biologically meaningful rather than driven by count scale.

Inspect mean vs standard deviation plot before transformation:

R

meanSdPlot(assay(dds), ranks = FALSE)

Apply variance stabilizing transform (VST):

R

vsd <- DESeq2::vst(dds, blind = TRUE)
meanSdPlot(assay(vsd), ranks = FALSE)

average counts vs variance after transformation

Callout

Why use VST

Counts have mean dependent variance. Variance stabilizing transform (VST) makes the variance roughly constant across the range, which improves distance based methods and PCA.

Sample to sample distances

The distance heatmap helps answer a key question: Do samples group by biological condition or by an unwanted variable (batch, sequencing run, contamination)? This is one of the fastest ways to detect outliers.

R

sampleDists       <- dist(t(assay(vsd)))
sampleDistMatrix  <- as.matrix(sampleDists)
rownames(sampleDistMatrix) <- colnames(vsd)
colnames(sampleDistMatrix) <- NULL
colors <- colorRampPalette(rev(brewer.pal(9, "Blues")))(255)

pheatmap(
  sampleDistMatrix,
  clustering_distance_rows = sampleDists,
  clustering_distance_cols = sampleDists,
  col                      = colors
)

PCA on top variable genes

PCA projects samples into low-dimensional space and reveals dominant sources of variation. Ideally, samples separate by biological condition along one of the top PCs. If not, the signal from the condition may be weak or confounded by other factors.

R

rv <- rowVars(assay(vsd))
select <- order(rv, decreasing = TRUE)[seq_len(min(500, length(rv)))]

pca <- prcomp(t(assay(vsd)[select, ]))
percentVar <- pca$sdev ^ 2 / sum(pca$sdev ^ 2)

intgroup    <- "condition"
intgroup.df <- as.data.frame(colData(vsd)[, intgroup, drop = FALSE])

group <- if (length(intgroup) == 1) {
  factor(apply(intgroup.df, 1, paste, collapse = " : "))
}

d <- data.frame(
  PC1 = pca$x[, 1],
  PC2 = pca$x[, 2],
  intgroup.df,
  name = colnames(vsd)
)

ggplot(d, aes(PC1, PC2, color = condition)) +
  scale_shape_manual(values = 1:12) +
  scale_color_manual(values = c(
    "Nat10-CKO"       = "#00b462",
    "Nat10flox/flox"  = "#980c80"
  )) +
  theme_bw() +
  theme(legend.title = element_blank()) +
  geom_point(size = 2, stroke = 2) +
  geom_text_repel(aes(label = name)) +
  xlab(paste("PC1", round(percentVar[1] * 100, 2), "% variance")) +
  ylab(paste("PC2", round(percentVar[2] * 100, 2), "% variance"))

Challenge

Exercise: interpret exploratory plots

Using the library size, distance heatmap, and PCA:

Are there any obvious outlier samples.
Do replicates from the same condition cluster together.
Does any sample look inconsistent with its metadata label.

Show me the solution

Example interpretation:

Library sizes are comparable across samples and fall within a narrow range.
Samples cluster by condition in both distance heatmaps and PCA.
No sample appears as a clear outlier relative to its group.

Differential gene expression analysis

For DE testing we use the full count matrix (cts) without filtering by biotype. This preserves all features counted by featureCounts.

R

dds <- DESeqDataSetFromMatrix(
  countData = cts_coding,
  colData   = coldata,
  design    = ~ condition
)

You can either run the individual DESeq2 steps:

R

dds <- estimateSizeFactors(dds)
dds <- estimateDispersions(dds)
dds <- nbinomWaldTest(dds)

Single command to run the DESeq2 pipeline

Above three steps can be run together using the DESeq function:

R

dds <- DESeq(dds)

estimating size factors
  Note: levels of factors in the design contain characters other than
  letters, numbers, '_' and '.'. It is recommended (but not required) to use
  only letters, numbers, and delimiters '_' or '.', as these are safe characters
  for column names in R. [This is a message, not a warning or an error]
estimating dispersions
gene-wise dispersion estimates
mean-dispersion relationship
  Note: levels of factors in the design contain characters other than
  letters, numbers, '_' and '.'. It is recommended (but not required) to use
  only letters, numbers, and delimiters '_' or '.', as these are safe characters
  for column names in R. [This is a message, not a warning or an error]
final dispersion estimates
fitting model and testing

Inspect dispersion estimates:

R

plotDispEsts(dds)

Obtain results for the contrast of interest (Nat10flox/flox vs Nat10-CKO):

R

res <-
  results(
    dds,
    contrast = c(
      "condition",
      "Nat10flox/flox",
      "Nat10-CKO"
    )
  )

summary(res)

TXT

out of 15839 with nonzero total read count
adjusted p-value < 0.1
LFC > 0 (up)       : 3195, 20%
LFC < 0 (down)     : 3398, 21%
outliers [1]       : 362, 2.3%
low counts [2]     : 308, 1.9%
(mean count < 1)
[1] see 'cooksCutoff' argument of ?results
[2] see 'independentFiltering' argument of ?results

Order the results by p value to see the top DE genes:

R

head(res[order(res$pvalue), ])

log2 fold change (MLE): condition Nat10flox/flox vs Nat10-CKO
Wald test p-value: condition Nat10flox.flox vs Nat10.CKO
DataFrame with 6 rows and 6 columns
       baseMean log2FoldChange     lfcSE      stat       pvalue         padj
      <numeric>      <numeric> <numeric> <numeric>    <numeric>    <numeric>
18512   4645.33       -5.17453 0.1335387  -38.7493  0.00000e+00  0.00000e+00
940     5041.68       -2.88015 0.1039939  -27.6953 7.94364e-169 6.02485e-165
13789  14362.27       -2.40229 0.0951377  -25.2506 1.11514e-140 5.63852e-137
16183  42766.77       -4.68585 0.1923780  -24.3575 4.82857e-131 1.83111e-127
18278   7055.06       -3.48409 0.1535305  -22.6932 5.23170e-114 1.58719e-110
270     1469.83       -2.87951 0.1298316  -22.1788 5.50606e-109 1.39202e-105

Each row represents one gene. Key columns: baseMean: average normalized count across all samples log2FoldChange: direction and magnitude of change between groups pvalue / padj: statistical significance after multiple testing correction Genes with NA values often have too few reads to estimate dispersion and should not be interpreted.

Create a summary table:

R

log2fc_cut <- log2(1.5)

res_df <- as.data.frame(res)

summary_table <- tibble(
    total_genes = nrow(res_df),
    sig = sum(res_df$padj < 0.05, na.rm = TRUE),
    up  = sum(res_df$padj < 0.05 & res_df$log2FoldChange >  log2fc_cut, na.rm = TRUE),
    down= sum(res_df$padj < 0.05 & res_df$log2FoldChange < -log2fc_cut, na.rm = TRUE)
)
print(summary_table)

# A tibble: 1 × 4
  total_genes   sig    up  down
        <int> <int> <int> <int>
1       15839  5658  2180  2363

Keep gene IDs:

R

res_df <- as.data.frame(res)
res_df$ensembl_gene_id_version <- rownames(res_df)

Joining results with expression and annotation

Get normalized counts:

R

norm_counts    <- counts(dds, normalized = TRUE)
norm_counts_df <- as.data.frame(norm_counts)
norm_counts_df$ensembl_gene_id_version <- rownames(norm_counts_df)

Join DE results, normalized counts, and annotation:

R

res_annot <- res_df %>%
  dplyr::left_join(norm_counts_df,
                   by = "ensembl_gene_id_version") %>%
  dplyr::left_join(mart,
                   by = "ensembl_gene_id_version")

Joining normalized counts and annotation makes the output biologically interpretable. The final table becomes a comprehensive results object containing: - gene identifiers - gene symbols - functional descriptions - differential expression statistics - normalized expression values This is the format most researchers expect when examining results or importing them into downstream tools.

Define significance labels for plotting:

R

log2fc_cut <- log2(1.5)

res_annot <- res_annot %>%
  dplyr::mutate(
    label = dplyr::coalesce(external_gene_name, ensembl_gene_id_version),
    sig = dplyr::case_when(
      padj <= 0.05 & log2FoldChange >=  log2fc_cut ~ "up",
      padj <= 0.05 & log2FoldChange <= -log2fc_cut ~ "down",
      TRUE ~ "ns"
    )
  )

Visualization: volcano plot

The volcano plot summarizes statistical significance (padj) and effect size (log2FoldChange) simultaneously. Genes far left or right show strong expression differences. Genes high on the plot pass stringent significance thresholds. Labels help identify the most biologically meaningful genes.

R

ggplot(
  res_annot,
  aes(
    x     = log2FoldChange,
    y     = -log10(padj),
    col   = sig,
    label = label
  )
) +
  geom_point(alpha = 0.6) +
  scale_color_manual(values = c(
    "up"   = "firebrick",
    "down" = "dodgerblue3",
    "ns"   = "grey70"
  )) +
  geom_text_repel(
    data = dplyr::filter(res_annot, sig != "ns"),
    max.overlaps       = 12,
    min.segment.length = Inf,
    box.padding        = 0.3,
    seed               = 42,
    show.legend        = FALSE
  ) +
  theme_classic() +
  xlab("log2 fold change") +
  ylab("-log10 adjusted p value") +
  ggtitle("Volcano plot: Nat10-CKO vs Nat10flox/flox")

Volcano plot of differential expression results

Callout

Interpreting the volcano

Points to the right are higher in Nat10flox/flox, to the left are higher in Nat10-CKO. Points near the top have stronger statistical support (small adjusted p values). Colored points mark genes that pass both fold change and padj thresholds.

Saving results

Saving both the full table and the significant subset allows flexibility: - The full table is needed for pathway analyses and reproducibility. - The subset table focuses on interpretable and biologically meaningful DE genes.

R

readr::write_tsv(
  res_annot,
  "results/deseq2/DESeq2_results_joined.tsv"
)

Filter to significant genes (padj ≤ 0.05 and fold change ≥ 1.5):

R

sig_res <- res_annot %>%
  dplyr::filter(
    padj <= 0.05,
    abs(log2FoldChange) >= log2fc_cut
  )

readr::write_tsv(
  sig_res,
  "results/deseq2/DESeq2_results_sig.tsv"
)

Challenge

Exercise: explore top DE genes

Using sig_res:

Which genes have the largest positive log2 fold change.
Which genes have the largest negative log2 fold change.
Are any of the top genes known to be related to Nat10 function or the studied phenotype.

Show me the solution

Example interpretation:

Several strongly up regulated genes are involved in cell cycle and RNA processing.
Strongly down regulated genes may include transcription factors and signaling components.
Cross referencing top hits with prior literature or databases can reveal mechanistic hypotheses.

Summary

Key Points

Counts and sample metadata must be aligned before building DESeq2 objects.
Biotype annotation allows restriction to protein coding genes for exploratory QC.
Variance stabilizing transform, distance heatmaps, and PCA help detect outliers and batch effects.
DESeq2 provides a complete pipeline from raw counts to differential expression.
Joining DE results with normalized counts and gene annotation yields a useful final table.
Volcano plots summarize differential expression using fold change and adjusted p values.

Content from Gene set enrichment analysis

Last updated on 2025-11-25 | Edit this page

Overview

Questions

How do we identify functional pathways and biological themes from DE genes.
How do we run over representation analysis for GO, KEGG, and MSigDB sets.
How do we create simple visualizations of enriched terms.
How do we interpret enrichment results in a biologically meaningful way.

Objectives

Prepare a gene list from DESeq2 results.
Run GO enrichment using clusterProfiler.
Run KEGG pathway enrichment with organism specific data.
Perform MSigDB Hallmark enrichment.
Produce barplots and dotplots for enriched terms.

Introduction

Differential expression gives a list of up regulated and down regulated genes.
To understand the underlying biology, we examine which functional gene sets are enriched.
This process is called over representation analysis (ORA). ORA identifies pathways or gene sets that contain more DE genes than expected by chance.

Here we will use clusterProfiler, org.Mm.eg.db, and msigdbr to run GO, KEGG, and Hallmark analyses on mouse data.

Callout

What ORA requires

A background universe of all tested genes (usually all genes in the DESeq2 dataset).
A subset of genes considered significant (for example padj < 0.05).
A gene identifier type that matches the annotation database (Ensembl or Entrez).

Step 1: Load DE results and prepare gene lists

Start your R session:

BASH

sinteractive -A rcac -p cpu -N 1 -n 4 --time=02:00:00
module load biocontainers
module load r-rnaseq
R

Load data:

R

library(DESeq2)
library(clusterProfiler)
library(org.Mm.eg.db)
library(msigdbr)
library(dplyr)
library(ggplot2)
library(readr)

Load DESeq2 results from Episode C:

R

res <- read_csv("DESeq2_results_shrunken.csv", show_col_types = FALSE)

Define:

universe = all genes tested
significant set = padj < 0.05

R

universe <- res$X1
sig <- res %>% filter(padj < 0.05) %>% pull(X1)

Convert Ensembl IDs to Entrez (required for GO and KEGG):

R

id_map <- bitr(universe,
               fromType = "ENSEMBL",
               toType = "ENTREZID",
               OrgDb = org.Mm.eg.db)

Merge IDs into results:

R

res2 <- left_join(res, id_map, by = c("X1" = "ENSEMBL"))
sig_entrez <- res2 %>% filter(padj < 0.05) %>% pull(ENTREZID)
universe_entrez <- res2$ENTREZID

Step 2: GO enrichment

Run GO Biological Process ORA:

R

ego <- enrichGO(gene = sig_entrez,
                universe = universe_entrez,
                OrgDb = org.Mm.eg.db,
                ont = "BP",
                readable = TRUE)

View top terms:

R

head(ego)

Plot:

R

dotplot(ego, showCategory = 20) +
    ggtitle("GO Biological Process enrichment")

Discussion

Why GO is useful

GO BP terms provide high level summaries of biological activity.
Strongly biased terms (for example mitochondrial or translation) often correspond to global shifts in cell state or stress.

Step 3: KEGG enrichment

KEGG requires Entrez IDs and organism code "mmu" for mouse.

R

ekegg <- enrichKEGG(gene = sig_entrez,
                    universe = universe_entrez,
                    organism = "mmu")

Plot KEGG enrichment:

R

barplot(ekegg, showCategory = 15)

Callout

KEGG caveat

KEGG Entrez annotations may not perfectly match recent Ensembl builds.
If a gene is missing, it will be silently dropped.

Step 4: MSigDB Hallmark enrichment

Load Hallmark gene sets for mouse:

R

hallmark <- msigdbr(species = "Mus musculus", category = "H")
hallmark_list <- split(hallmark$entrez_gene, hallmark$gs_name)

Run ORA:

R

ehall <- enricher(gene = sig_entrez,
                  universe = universe_entrez,
                  TERM2GENE = hallmark_list)

Plot:

R

dotplot(ehall, showCategory = 20) +
    ggtitle("MSigDB Hallmark enrichment")

Inspect enriched Hallmark sets

R

head(ehall)

Callout

Why Hallmark sets are recommended

Hallmark pathways are curated to reduce redundancy.
They are easier to interpret compared to long lists of GO terms.

Step 5: Saving results

R

write_csv(as.data.frame(ego), "GO_enrichment.csv")
write_csv(as.data.frame(ekegg), "KEGG_enrichment.csv")
write_csv(as.data.frame(ehall), "Hallmark_enrichment.csv")

Step 6: Interpretation guidelines

Meaningful enrichment patterns should:

involve pathways relevant to your experimental design
contain multiple DE genes in the same direction
show consistent themes across GO, KEGG, and Hallmark sets

Pathways enriched only by one DE gene or very small sets should be interpreted cautiously.

Challenge

Exercise: interpret enrichment results

Using your GO, KEGG, and Hallmark outputs:

Identify the pathways with the smallest adjusted p values.
Are the enriched pathways consistent with the expected phenotype.
Which pathway collections (GO, KEGG, Hallmark) give the clearest signal.

Show me the solution

Example interpretation:

Several GO terms related to RNA processing and cell cycle appear enriched.
Hallmark sets provide clearer summaries of biological themes compared to GO.
KEGG pathways show fewer hits but corroborate the GO findings.

Summary

Key Points

ORA requires a universe and a significant gene set.
Ensembl IDs must be converted to Entrez for GO and KEGG.
clusterProfiler provides GO BP and KEGG ORA.
Hallmark sets offer high quality curated pathways for interpretation.
Dotplots and barplots are effective for visualizing enrichment results.

Overview

Questions

Objectives

What are we measuring in an RNA-seq experiment?

Challenge: Discuss the following points with your neighbor

Experimental design considerations

Challenge: Discuss with your neighbor

Challenge

Challenge

RNA-seq quantification: from reads to count matrix

Challenge: Discuss the following points with your neighbor

Finding the reference sequences

Challenge

Where are we heading towards in this workshop?

Overview

Questions

Objectives

Introduction

Should I uncompress FASTQ files?

Should I trim adapters for RNA seq analysis?

Downloading the data and project organization

Create the directories needed for this episode

Show me the solution

BASH

BASH

What files do I need for RNA-seq analysis?

Downloading the data

Reference genome and annotation

Annotation

Show me the solution

Reference genome

Show me the solution

Transcript sequences

Transcript sequence file

Show me the solution

Downloading all reference files

BASH

BASH

Raw reads

Where do you find FASTQ files?

Show me the solution

Download FASTQ files with SRA Toolkit

BASH

BASH

Overview

Questions

Objectives

Introduction

Why do we perform QC on FASTQ files?

Running FastQC

Run FastQC on your FASTQ files

BASH

BASH

Understanding the FastQC report

Which FastQC modules are important for RNA-seq?

Key modules explained

Per base sequence quality

Per tile sequence quality

Per sequence quality scores

Per base sequence content

Per sequence GC content

Per base N content

Sequence length distribution

Sequence duplication levels

Overrepresented sequences

Adapter content

Aggregating reports with MultiQC

Run MultiQC on the FastQC outputs

BASH

Trimming adapter sequences

Do we need to trim reads for standard RNA-seq workflows?

Exercise: evaluate the QC results

Inspect your MultiQC report

Show me the solution

What if trimming is required?

Example: trimming paired-end reads with fastp

BASH

What this command does

When to use this

Summary

Example: trimming paired-end reads with `fastp`

Interpreting `lib_format_counts.json`