In jianhong/ChIPpeakAnno: Batch annotation of the peaks identified from either ChIP-seq, ChIP-chip experiments, or any experiments that result in large number of genomic interval data

knitr::opts_chunk$set(message = FALSE, 
                      warning = FALSE,
                              tidy = FALSE, 
                              fig.width = 6, 
                              fig.height = 6,
                              fig.align = "center")
# Handle the biofilecache error
library(BiocFileCache)
bfc <- BiocFileCache()
res <- bfcquery(bfc, 
                "annotationhub.index.rds", 
                field = "rname", 
                exact = TRUE) 
bfcremove(bfc, rids = res$rid)

Introduction

Several experimental techniques including ChIP-ChIP [@blat1999], ChIP-seq [@barski2007; @johnson2007], and CUT&RUN [@skene2017] have gained widespread use in molecular biology. These techniques enable the study of protein-DNA interactions by identifying the genomic regions that are associated with specific proteins, such as transcription factor binding sites, or epigenetic markers like histone modification. Through these approaches, researchers gain valuable insights into the regulation of genes and the structure of chromatin.

The analysis involves several key procedures as outlined below, you can also integrate other omics data to gain a more comprehensive view of the gene regulation mechanisms. The steps where r Biocpkg("ChIPpeakAnno") package can be utilized are displayed in pink boxes.

png <- system.file("extdata", "dataFlow.png", package = "ChIPpeakAnno")

knitr::include_graphics(png)

Explanations of selected terms:

• Peak calling: identify the genomic regions of significant read enrichment (peaks), which represent the binding sites of the protein or epigenetic marker of interest
• Peak annotation: associate peaks with nearby genomic features, usually gene transcripts, to understand their functional implications
• Enrichment analysis: identify the gene ontology (GO) terms or biological pathways that are associated with the annotated genes
• Motif analysis: identify the enriched DNA patterns within peaks to understand the binding preference of the target protein

Various algorithms have been developed to identify peaks from experimental data [@thomas2017]. Once the peak files are obtained, they are commonly converted into formats like BED and its variants. These formats allow the data to be easily loaded into genome browsers, such as the UCSC Genome Browser, as custom tracks. This enables investigators to examine the proximity of the peaks to different genomic features like promoters, enhancers, or other regulatory elements. However manually navigating through the genome browser can be a daunting task due to the typically large number of peaks that are spread across the genome.

The Bioconductor package r Biocpkg("ChIPpeakAnno") is a pioneering tool that facilitates the batch annotation of peaks obtained from various technologies. One notable feature of r Biocpkg("ChIPpeakAnno") is its ability to dynamically retrieve up-to-date annotations by leveraging the r Biocpkg("biomaRt") package. This enables users to access the most current annotation files. Additionally, users can supply their own annotation data or utilize existing annotation packages. Furthermore, r Biocpkg("ChIPpeakAnno") provides functions to retrieve sequences surrounding the peaks, which can be utilized for peak validation via PCR and motif discovery [@Durinck2005]. The package also facilitates the identification of enriched GO or pathway terms. In addition, r Biocpkg("ChIPpeakAnno") includes several helper functions that aid in visualizing binding patterns and comparing peak profiles across multiple samples or experimental conditions.

r Biocpkg("ChIPpeakAnno") has been continuously enhanced since its initial release in 2010, as evident from its active development^[https://github.com/jianhong/ChIPpeakAnno/blob/devel/NEWS]. New features are being regularly added based on user requests. If you have any usage related questions, please search for solutions or post new questions on the Bioconductor Support Site, which is actively monitored by many seasoned users and developers. For feature request, bug report, or any other concerns, raise an issue on the ChIPpeakAnno GitHub repository. Your input and contributions will be greatly appreciated and carefully considered.

Quick demo

This section provides a quick example on utilizing r Biocpkg("ChIPpeakAnno") to annotate peaks identified with MACS or MACS2 [@zhang2008]. Several steps are typically involved and are discussed in more details in the following sections:

• Convert peaks to GRanges (Section \@ref(readinpeak))
• Handle replicates (Section \@ref(handlereplicates))
• Prepare annotation file (Section \@ref(prepanno))
• Annotate peaks (Section \@ref(annopeaks))
• Add other feature IDs (Section \@ref(addids))

From Section \@ref(readinpeak) to \@ref(profilecompare), we delve into detailed use cases, showcasing the versatility of r Biocpkg("ChIPpeakAnno") in various scenarios. Section \@ref(workflow1) to \@ref(workflow2) present several commonly employed analytical pipelines, offering step-by-step illustrations for different settings.

Convert peaks to GRanges {#readinpeakdemo}

The r Biocpkg("ChIPpeakAnno") package provides an example peak file in narrowPeak format, which is generated by MACS. To work with this example file, we need to locate it with system.file and convert it into a GRanges object using toGRanges function.

library(ChIPpeakAnno)

macs_peak <- system.file("extdata", "MACS_peaks.xls", 
                         package = "ChIPpeakAnno")
macs_peak_gr <- toGRanges(macs_peak, format = "MACS")
head(macs_peak_gr, n = 2)

Prepare annotations {#prepannos}

This section demonstrates how to prepare annotation file from Ensembl package. The Ensembl package is generally favored, as it offers more comprehensive and superior annotation. For more information, please refer to Section \@ref(prepanno).

# Use Ensembl annotation package:
library(EnsDb.Hsapiens.v86)

ensembl.hs86.transcript <- transcripts(EnsDb.Hsapiens.v86)

Annotate peaks

# Use Ensembl annotation package:
macs_peak_ensembl <- annotatePeakInBatch(macs_peak_gr, 
                                         AnnotationData = ensembl.hs86.transcript)
head(macs_peak_ensembl, n = 2)

For more regarding the metadata columns, please refer to the Section \@ref(findnearest). Of note, the metadata does not come with "gene symbol" column. We can add it using the addGeneIDs function as below.

Add gene symbols

To add gene symbols, we can leverage either the organism annotation package, namely r Biocpkg("org.Hs.eg.db"), or the r Biocpkg("bioRmart") package. The latter offers the most current information but may have a slower performance as it requires querying the BioMart databases in real time. Once the required package is loaded, you can use the addGeneIDs function to add gene symbols. The following example utilizes the r Biocpkg("bioRmart") method.

library(biomaRt)
mart <- useMart(biomart = "ENSEMBL_MART_ENSEMBL",
                dataset = "hsapiens_gene_ensembl")

# Use Ensembl annotation package:
macs_peak_ensembl <- addGeneIDs(annotatedPeak = macs_peak_ensembl, 
                                mart = mart,
                                feature_id_type = "ensembl_transcript_id",
                                IDs2Add = "hgnc_symbol")
head(macs_peak_ensembl, n = 2)

Now, a hgnc_symbol metadata column has been successfully inserted to the resulting GRanges object. It is crucial to specify the correct feature_id_type and IDs2Add for the function to work properly. By default, feature_id_type = "ensembl_gene_id".

The following sections demonstration each topic in more details.

Convert peaks to GRanges {#readinpeak}

Peak files are represented as GRanges objects in r Biocpkg("ChIPpeakAnno"). To facilitate the conversion of commonly used peak formats including BED (and its variants), GFF, and CSV, we have implemented a helper function called toGRanges.

# Convert GFF to GRanges:
macs_peak_gff <- system.file("extdata", "GFF_peaks_hg38.gff", 
                             package = "ChIPpeakAnno")
macs_peak_gr1 <- toGRanges(macs_peak_gff, format = "GFF", header = FALSE)
head(macs_peak_gr1, n = 2)

# Convert BED to GRanges:
macs_peak_bed <- system.file("extdata", "MACS_output_hg38.bed", 
                             package = "ChIPpeakAnno")
macs_peak_gr2 <- toGRanges(macs_peak_bed, format = "BED", header = FALSE)
head(macs_peak_gr2, n = 2)

Handle replicates {#handlereplicates}

Obtain final peak set from replicates

For experiments with two or more biological replicates, there are several strategies^[https://ro-che.info/articles/2018-07-11-chip-seq-consensus] to retrieve a final peak set:

• Take the intersection of peaks:
• Most conservative, reduces noise by considering the most robust and consistently enriched peaks
• May result in small number of peaks with narrower widths
• Take the union of all peaks: may lead to large number of peaks
• Least conservative, provides a more comprehensive view of the binding landscape
• Useful for identifying condition-specific or rare binding events
• Merge the overlapping peaks: may lead to peaks with broader widths
• Can be achieved with ChIPpeakAnno::findOverlapsOfPeaks
• Use peak coverage voting to determine a final set:
• Can be achieved with r Biocpkg("consensusSeekeR")
• Statistical model based approaches that leverage peak coverage and/or read depth information

Investigators may choose one of these strategies based on their research questions and the quality of their data.

Assess the concordance of peak replicates

Concordant peaks refer to the peaks that come from different biological replicates and exhibit a high degree of overlap in their genomic coordinates and signal intensity. The presence of concordant peaks indicates that the observed signal is likely to be true positive rather than arising from random noise or technical artifacts. You can evaluate the concordance of replicates by visualizing the peak overlappings.

Find overlapping peaks

Here are examples demonstrating how to identify overlapping peaks using the sample peaks provided in the r Biocpkg("ChIPpeapAnno") package. By default, two peaks are considered "overlapping" if "one range has its start or end strictly inside the other (maxgap = -1L)". If you set maxgap = 1000, two peaks will be classified as "overlapping" if the number of positions that separate them is less than 1000. Additionally, specifying minoverlap = 100 ensures that only peaks with a minimum of 100 overlapping positions are treated as "overlapping". When 0 < minoverlap < 1, the function will identify overlaps based on the percentage covered of peak interval.

# For two samples:
data(peaks1)
data(peaks2)
head(peaks1, n = 2)
head(peaks2, n = 2)
ol <- findOverlapsOfPeaks(peaks1, peaks2)
names(ol)

# For three samples:
data(peaks3)
ol2 <- findOverlapsOfPeaks(peaks1, peaks2, peaks3,
                           connectedPeaks = "min")

# Find peaks with at least 90% overlap:
ol3 <- findOverlapsOfPeaks(peaks1, peaks2, minoverlap = 0.9)

The results are an object of overlappingPeaks. This object provides comprehensive information about the overlappings, allowing for further visualization and interpretation:

• venn_cnt: an object of VennCounts, which can be used to draw a Venn diagram
• peaklist: a list of GRanges objects that contains all merged overlapping peaks or unique peaks
• uniquePeaks: an object of GRanges that consists of all unique peaks
• mergedPeaks: an object of GRanges that consists of all merged overlapping peaks
• peaksInMergedPeaks: an object of GRanges that consists of all peaks in each samples involved in the overlapping peaks
• overlappingPeaks: a list of data frames that consists of the annotation of all overlapping peaks
• all.peaks: a list of GRanges objects that consists the input peaks with formatted rownames

To determine the number of peaks that are unique to each peak set (i.e., not overlapping with any peaks in other set), you can use the following code:

# For peaks1:
length(ol$peaklist[["peaks1"]])

# For peaks2:
length(ol$peaklist[["peaks2"]])

To obtain the merged overlapping peaks, you have two options:

• You can use ol$mergedPeaks to obtain all the merged overlapping peaks from all peak sets in ol. This means that if you have multiple peak sets, this command will output all the peaks that overlap with peaks from any other peak sets
• Alternatively, you can use ol$peaklist[["peaks1///peaks2"]], where "peaks1///peaks2" represents a GRanges object that contains all the merged overlapping peaks from peaks1 and peaks2. When you only have two peak sets, ol$mergedPeaks is equivalent to ol$peaklist[["peaks1///peaks2"]]. However, if more peak sets are involved, e.g. three peak sets, ol$peaklist[["peaks1///peaks2///peaks3"]] would represent merged overlapping peaks that consist of peaks from all three sets. In other words, ol$mergedPeaks is equivalent to ol$peaklist[["peaks1///peaks2"]] plus ol$peaklist[["peaks2///peaks3"]] and ol$peaklist[["peaks1///peaks2///peaks3"]]

Visualize the overlaps using Venn diagram {#venn}

The output from findOverlapsOfPeaks can be directly fed to makeVennDiagram to draw a Venn diagram and evaluate the degree of overlap between peak sets. Additionally, the makeVennDiagram function also calculates a P-value, which indicates whether the observed overlap is statistically significant or not.

# For two samples:
venn <- makeVennDiagram(ol, totalTest = 100,
                        fill = c("#009E73", "#F0E442"),
                        col = c("#D55E00", "#0072B2"),
                        cat.col = c("#D55E00", "#0072B2"))

# For three samples:
venn2 <- makeVennDiagram(ol2, totalTest = 100,
                         fill = c("#CC79A7", "#56B4E9", "#F0E442"),
                         col = c("#D55E00", "#0072B2", "#E69F00"),
                         cat.col = c("#D55E00", "#0072B2", "#E69F00"))

# For peaks overlap at least 90%:
venn3 <- makeVennDiagram(ol3, totalTest = 100,
                         fill = c("#009E73", "#F0E442"),
                         col = c("#D55E00", "#0072B2"),
                         cat.col = c("#D55E00", "#0072B2"))

The parameter totalTest refers to the total number of potential peaks that are considered in the hypergeometric test. It should be set to a value larger than the largest number of peaks in the replicates. Refer to Section \@ref(hypertest) on how to choose totalTest. Since P-value is sensitive to the choice of totalTest, we recommend using permutation test, implemented as peakPermTest in Biocpkg("ChIPpeakAnno"). For details, go to Section \@ref(permtest).

Use other tools to plot Venn diagram

Users can leverage the ol$venn_cnt object and choose other tools to draw Venn diagram. Below illustrates how to use r CRANpkg("Vennerable") library for plotting.

if (!library(Vennerable)) {
  install.packages("Vennerable", repos="http://R-Forge.R-project.org")
  library(Vennerable)
}

n <- which(colnames(ol$vennCounts) == "Counts") - 1
set_names <- colnames(ol$vennCounts)[1:n]
weight <- ol$vennCounts[, "Counts"]
names(weight) <- apply(ol$vennCounts[, 1:n], 1, base::paste, collapse = "")
v <- Venn(SetNames = set_names, Weight = weight)
plot(v)

Visualize the distribution of relative positions for overlapping peaks

As mentioned earlier, two peaks are considered as "overlapping" if their distances are within maxgap. To visualize the distribution of the relative positions of peaks1 to peaks2, we can create a pie chart using the ol$overlappingPeaks object. This object contains detailed information about the relative positions of peaks for each pair of peak sets. For instance, ol$overlappingPeaks[["peaks1\\\peaks2"]] represents the relative positions of overlapping peaks between peaks1 and peaks2.

names(ol$overlappingPeaks)
dim(ol$overlappingPeaks[["peaks1///peaks2"]])
ol$overlappingPeaks[["peaks1///peaks2"]][1:2, ]
unique(ol$overlappingPeaks[["peaks1///peaks2"]][["overlapFeature"]])
pie1(table(ol$overlappingPeaks[["peaks1///peaks2"]]$overlapFeature))

The column overlapFeature describes the relative positions of peaks between peaks1 and peaks2:

• upstream: the peak from peaks1 is located upstream of the peak from peaks2 and is within a distance of maxgap
• downstream: the peak from peaks1 is located downstream of the peak from peaks2 and is within a distance of maxgap
• overlapStart: the peak from peaks1 overlaps with the start of peak from peaks2
• overlapEnd: the peak from peaks1 overlaps with the end of peak from peaks2
• inside: the peak from peaks1 is completely contained within the peak from peaks2
• includeFeature: the peak from peaks1 contains the entire range of the peak from peaks2. Note that the term Feature here refers to the peak in peaks2 rather than a genomic feature as used in peak annotation context

The utilization of a Venn diagram, in conjunction with a pie chart, enables a more comprehensive evaluation of peak concordance among biological replicates.

Prepare annotation file {#prepanno}

An annotation file contains genomic coordinates and other relevant information for various genomic features, such as genes, transcripts, promoters, enhancers, and more. By comparing the peaks with these coordinates, researchers can determine which features are most enriched or associated with the peaks, which can help them understand the functional relevance of the peaks and provide insights into the potential regulatory elements or genes involved.

Commonly used annotations

Popular annotation files come from two sources and are typically stored in tab-delimited formats, such as GTF or BED.

library(knitr)

Resource <- c("Ensembl", "NCBI RefSeq")
Generated_by <- c("EMBL-EBI", "NCBI")
Annotation_criteria <- c("Comprehensive (most transcripts)", "Conservative (fewer transcripts)")
Gene_id_name <- c("Ensembl gene ID", "NCBI Gene ID, or entrezGene ID, or entrez ID")
URL <- c("https://ftp.Ensembl.org/pub/", "https://ftp.ncbi.nlm.nih.gov/refseq/")
df <- data.frame(Resource, Generated_by, Annotation_criteria, URL)

kable(df, caption = 'Commonly used annotation resources')

In addition, the UCSC Genome Browser team provides processed annotations based on the above resources that can be visualized easily with the browser. For human and mouse, there are four GTF files provided respectively.

library(knitr)

Human <- c("[hg38.refGene.gtf.gz](https://hgdownload.soe.ucsc.edu/goldenPath/hg38/bigZips/genes/hg38.refGene.gtf.gz)",
           "[hg38.ncbiRefSeq.gtf.gz](https://hgdownload.soe.ucsc.edu/goldenPath/hg38/bigZips/genes/hg38.ncbiRefSeq.gtf.gz)",
           "[hg38.knownGene.gtf.gz](https://hgdownload.soe.ucsc.edu/goldenPath/hg38/bigZips/genes/hg38.knownGene.gtf.gz)")
Mouse <- c("[mm10.refGene.gtf.gz](https://hgdownload.soe.ucsc.edu/goldenPath/mm10/bigZips/genes/mm10.refGene.gtf.gz)",
           "[mm10.ncbiRefSeq.gtf.gz](https://hgdownload.soe.ucsc.edu/goldenPath/mm10/bigZips/genes/mm10.ncbiRefSeq.gtf.gz)",
           "[mm10.knownGene.gtf.gz](https://hgdownload.soe.ucsc.edu/goldenPath/mm10/bigZips/genes/mm10.knownGene.gtf.gz)")
Remark <- c("Based on RefSeq transcripts aligned by UCSC followed by manual curation",
            "Based on RefSeq transcripts as aligned by NCBI",
            "Based on Ensembl gene models")
df <- data.frame(Human, Mouse, Remark)

kable(df, caption = "UCSC Genome Browser hosted annotation files")

We would recommend using EnsDb annotations because they are usually more comprehensive and consistent.

Bioconductor supported TxDb and EnsDb {#txdbensdb}

In Bioconductor, the tab-delimited files are often converted into TxDb or EnsDb object. Both can be created with the tab delimited files mentioned above. While EnsDb is tailored to Ensembl annotations and contains additional information such as gene_name, symbol, and gene_biotype, the TxDb counterpart is typically created from RefSeq or UCSC Genome Browser hosted annotations.

There is a comprehensive list of pre-built Bioconductor maintained TxDb and EnsDb packages such as r Biocpkg("TxDb.Hsapiens.UCSC.hg38.knownGene") and r Biocpkg("EnsDb.Hsapiens.v86"). The list is regularly updated and can be found here.

Obtain EnsDb and TxDb from AnnotationHub

Users can also retrieve annotations using the r Biocpkg("AnnotationHub") package. By default, r Biocpkg("AnnotationHub") uses a snapshot that matches the version of Bioconductor being used, so it may be slightly more up-to-date compared to the pre-built packages. Additionally, users have the option to switch to an earlier version^[https://bioconductor.org/packages/release/bioc/vignettes/AnnotationHub/inst/doc/AnnotationHub.html#configuring-annotationhub-objects]. Below are examples of how to obtain annotations using r Biocpkg("AnnotationHub").

library(AnnotationHub)

ah <- AnnotationHub()

# Obtain EnsDb:
EnsDb_Mmusculus_all <- query(ah, pattern = c("Mus musculus", "EnsDb"))
head(EnsDb_Mmusculus_all, n = 2)
EnsDb_Mmusculus <- EnsDb_Mmusculus_all[["AH53222"]]
class(EnsDb_Mmusculus)

# Obtain TxDb:
TxDb_Mmusculus_all <- query(ah, pattern = c("Mus musculus", "TxDb"))
head(TxDb_Mmusculus_all, n = 2)
TxDb_Mmusculus <- TxDb_Mmusculus_all[["AH52264"]]
class(TxDb_Mmusculus)

Build custom EnsDb and TxDb

To create the most up-to-date or custom TxDb and EnsDb objects, you can utilize functions such as makeTxDbFromUCSC, makeTxDbFromEnsembl, and ensDbFromGRanges from the r Biocpkg("GenomicFeatures")^[https://bioconductor.org/packages/devel/bioc/vignettes/GenomicFeatures/inst/doc/GenomicFeatures.html] package and the r Biocpkg("Ensembldb")^[https://www.bioconductor.org/packages/devel/bioc/vignettes/Ensembldb/inst/doc/Ensembldb.html#102_Building_annotation_packages] package.

Use biomaRt {#usebiomart}

The r Biocpkg("biomaRt") package offers a convenient interface to the BioMart databases that are prominently maintained by Ensembl. By querying r Biocpkg("biomaRt"), you can access the latest available annotations from Ensembl. Check this vignette for details.

r Biocpkg("ChIPpeakAnno") provides a helper function called getAnnotation, which simplifies the retrieval of desired annotations by leveraging r Biocpkg("biomaRt"). Here is an example:

library(biomaRt)

listMarts()
head(listDatasets(useMart("ENSEMBL_MART_ENSEMBL")), n = 2)
mart <- useMart(biomart = "ENSEMBL_MART_ENSEMBL",
                dataset = "mmusculus_gene_ensembl")
anno_from_biomart <- getAnnotation(mart, 
                                   featureType = "transcript")
head(anno_from_biomart, n = 2)

Convert annotations into GRanges

To annotate peaks, the chosen annotation file needs to be converted into GRanges class first. This can be achieved with toGRanges function, accessor functions, or getAnnotation function.

Use toGRanges function

r Biocpkg("ChIPpeakAnno") offers a helper function called toGRanges, which can convert annotations from various formats including GFF, BED, CSV, TxDb, and EnsDb into GRanges. Below is an example.

library(EnsDb.Hsapiens.v86)

anno_ensdb_transcript <- toGRanges(EnsDb.Hsapiens.v86, 
                                   feature = "transcript")
head(anno_ensdb_transcript, n = 2)

Similarly, set feature = "gene" to retrieve gene annotations in GRanges format.

Use accessor functions

If you are working with TxDb or EnsDb objects, you can obtain annotations in GRanges format with various accessor functions such as genes and transcripts, which are designed for easy fetching of the desired information.

anno_ensdb_transcript <- transcripts(EnsDb.Hsapiens.v86)

head(anno_ensdb_transcript, n = 2)

Use getAnnotation function

The output from getAnnotation function is in GRanges format, refer to Section \@ref(usebiomart) for details.

Visualize peak distributions

Plotting peak distributions is a valuable quality control measure as it provides an overview of the localization of peaks across the genome. Unexpected distributions can indicate potential issues with the data. In addition, you can select appropriate annotation file depending on the distribution of your peaks. For instance, if peaks are enriched near promoters, you may focus on annotating them with nearby transcripts.

Plot peak distributions relative to genomic features

The binOverFeature function can be used to plot the distribution of peak counts relative to a specific genomic feature. The following example shows the distribution of peaks in the macs_peak_gr2 dataset relative to the gene feature (transcription start site (TSS)).

library(TxDb.Hsapiens.UCSC.hg38.knownGene)

annotation_data <- transcripts(TxDb.Hsapiens.UCSC.hg38.knownGene)
binOverFeature(macs_peak_gr2, 
               featureSite = "FeatureStart",
               nbins = 20,
               annotationData = annotation_data,
               xlab = "peak distance from TSS (bp)", 
               ylab = "peak count", 
               main = "Distribution of aggregated peak numbers around TSS")

By default, featureSite = "FeatureStart" meaning that distance is calculated as peak to feature start (i.e. TSS for transcript). The plot above, describing the distribution of peaks around TSS, exhibits a signal summit around TSS, a characteristic pattern observed in peaks obtained from transcription factor binding experiments.

You can also plot peak distribution over multiple genomic features including exon, intron, enhancer, proximal promoter, 5' UTR and 3' UTR in a single bar graph using assignChromosomeRegion.

chromosome_region <- assignChromosomeRegion(macs_peak_gr2,
                                            TxDb = TxDb.Hsapiens.UCSC.hg38.knownGene,
                                            nucleotideLevel = FALSE,
                                            precedence=c("Promoters",
                                                         "immediateDownstream", 
                                                         "fiveUTRs", 
                                                         "threeUTRs",
                                                         "Exons", 
                                                         "Introns"))

# optional helper function to rotate x-axis labels for barplot(): 
# ref: https://stackoverflow.com/questions/10286473/rotating-x-axis-labels-in-r-for-barplot
rotate_x <- function(data, rot_angle) {
  plt <- barplot(data, xaxt = "n")
  text(plt, par("usr")[3], 
       labels = names(data), 
       srt = rot_angle, adj = c(1.1,1.1), 
       xpd = TRUE, cex = 0.6)
}

rotate_x(chromosome_region[["percentage"]], 45)

By default, nucleotideLevel = FALSE meaning that peaks are treated as ranges when determining overlaps with genomic features. If a peak intersects with multiple features, the feature assignment is determined by the order specified in the precedence argument. If precedence is not set, counts for each overlapping feature will be incremented. Otherwise, if nucleotideLevel = TRUE, the summit of the peak (a single position, not suitable for broad peaks) will be used when determining overlaps.

Plot peak distributions over different feature levels

In addition to inspecting the peak enrichment pattern by plotting the distribution against genomic features, users can plot distributions over different feature levels, each containing multiple categories, using the genomicElementDistribution function.

• Transcript Level: "geneLevel"

  • "Promoter"
  • "Gene body"
  • "Downstream"
  • "Distal Intergenic"

• Exon Level: "Exons"

  • "5' UTR"
  • "CDS"
  • "3' URT"
  • "Other exon"

• Exon/Intron/Intergenic: "ExonIntron"

  • "Exon"
  • "Intron"
  • "Intergenic"

Please note that peaks can be classified into multiple categories from different levels, leading to the total percentage of annotated features being greater than 100%. At each level, since a peak spans a genomic range, it may overlap with multiple categories of features. In such cases, by default nucleotideLevel = FALSE, which means that the precedence is determined by the order listed in the labels argument.

The genomicElementDistribution function accepts either a single peak object or a list of peak objects as input. If a single peak object if provided, a pie chart will be created; if a list of peak objects is provided, a bar graph will be created.

Pie graph with one peak set

genomicElementDistribution(macs_peak_gr1, 
                           TxDb = TxDb.Hsapiens.UCSC.hg38.knownGene)

As can be seen, a significant number of peaks originate from promoter regions.

Bar graph with a list of peak sets

macs_peaks <- GRangesList(rep1 = macs_peak_gr1,
                          rep2 = macs_peak_gr2)
genomicElementDistribution(macs_peaks, 
                           TxDb = TxDb.Hsapiens.UCSC.hg38.knownGene)

The consistent patterns from "rep1" and "rep2" indicate a high correlation between them.

Plot peak overlaps for multiple features {#upset}

We can create an UpSet plot to view peak overlaps across multiple genomic features.

library(UpSetR)

res <- genomicElementUpSetR(macs_peak_gr1,
                            TxDb.Hsapiens.UCSC.hg38.knownGene)
upset(res[["plotData"]], 
      nsets = length(colnames(res$plotData)), 
      nintersects = NA)

UpSet plot can be considered as a "high dimensional Venn diagram" that allows for the visualization of overlaps for multiple sets. For example, in the above plot, it is evident that the feature set "gene body" (from the "Transcript Level") and the feature set "intron" (from the "Exon/Intron/Intergenic") share the highest number of common peaks. Type ?genomicElementUpSetR for the definition of distal_promoter.

Annotate peaks {#annopeaks}

With the annotation data, you can assign the peaks identified in your experiments to nearby features of your choice such as genes and transcripts. This process is known as peak annotation. The annotatePeakInBatch function offers a highly flexible approach to perform peak annotation with various output option.

For example, you can annotate peaks based on their nearest (output = "nearestLocation") or overlapping (output = "overlapping") features using the peak-centric method. Alternatively, you can annotate peaks based on their relative locations to features using the feature-centric method. For example, if a peak is located upstream of a gene within a certain distance (e.g. promoter region), you can assign that gene to the peak (output = "upstream"). The bindingRegion option allows for even more flexibility in specifying teh relative locations. Detailed explanations see below.

The choice between the peak-centric and feature-centric methods depends on your research question, although most users initially opt for the peak-centric approach as it is easier to interpret.

Peak-centric method {#peakcentric}

You can assign the nearest or overlapping features to your peaks by setting the output option to the following values:

• output: criteria for assigning features
  • "nearestLocation": (default) output the features that are nearest to the peaks based on the absolute value of the difference between the peak location and the feature location, as calculated by abs(PeakLocForDistance - FeatureLocForDistance)
  • "overlapping": output features that overlap with the peaks within the maximum distance specified by the maxgap parameter
  • "both": output both the nearest feature and overlapping features that are not the nearest
  • "shortestDistance": output the features that have the shortest distance to the peaks; the "shortest distance" is determined from either end of the feature to either end of the peak

Other relevant parameters:

• PeakLocForDistance: location of the peak for distance calculation

  • "middle": (recommended) use the center of the peak
  • "start": (default) use the peak edge that is smaller in coordinate
  • "end": use the peak edge that is larger in coordinate
  • "endMinusStart": use "end" defined above if feature is on the plus strand, and "start" if minus strand

• FeatureLocForDistance: location of the feature for distance calculation

  • "middle": use the center of the feature
  • "start": use the feature edge that is smaller in coordinate
  • "end": use the feature edge that is large in coordinate
  • "TSS": (default) use "start" defined above if feature is on the plus strand and "end" if on the minus strand
  • "geneEnd": use "end" defined above if feature is on the plus strand and "start" if on the minus strand

• maxgap: the maximum gap that is allowed between the peak and the feature ranges for them to be considered as overlapping; it is defined as the number of bases that separates the two ranges, the gap between two adjacent ranges is 0

The following diagram illustrates how to annotate peaks using the peak-centric method. When output = "nearestLocation", the distance between the peak and the feature is calculated as abs(PeakLocForDistance - FeatureLocForDistance); for demonstration, PeakLocForDistance = "start" and FeatureLocForDistance = "TSS".

png <- system.file("extdata", "annoPeakCentric.png", package = "ChIPpeakAnno")

knitr::include_graphics(png)

Feature-centric method {#featurecentric}

Peaks can also be annotated based on their relative locations to nearby features. For example, by setting output = "upstream", peaks will be annotated to features that they are located upstream of.

Relative peak-to-feature location

You can use the following options to specify the desired relative locations of the peaks to the features.

• output: criteria for assigning features
  • "upstream": output the feature if the peak resides upstream of it
  • "upstrem&inside": output the feature if the peak resides upstream of it or contained within it
  • "inside": output the feature if the peak is completely contained within it
  • "inside&downstream": output the feature if the peak resides within it or downstream of it
  • "downstream": output the feature if the peak resides downstream of it
  • "upstreamORdownstream": output the feature if the peak resides upstream or downstream of it

Other relevant parameter:

• maxgap: allowed gap when determining overlap between two ranges; will be ignored if output = "inside"

The following diagram illustrates how peaks are annotated using the feature-centric method.

png <- system.file("extdata", "annoFeatureCentric1.png", package = "ChIPpeakAnno")

knitr::include_graphics(png)

When using the feature-centric method, the annotatePeakInBatch function will output the feature as long as the peak range overlaps with the target region, except when output = "inside". In this case, the entire peak range must be completely contained within the feature range to output it.

More customization with bindingRegion

The bindingRegion parameter, which refers to the regions that the target TF binds to, adds further flexibility to the feature-centric method when defining the target region. For example, if you would like to annotate peaks to features where the peaks are located within specific regions relative to them, such as from 2000bp upstream to 500bp downstream of TSS (where most promoters are found), you can set output = "overlapping", FeatureLocForDistance = "TSS", and bindingRegion = c(-2000, 500). Once bindingRegion is specified, the maxgap will be ignored.

The following diagram demonstrates several examples of how to use bindingRegion to define target regions.

png <- system.file("extdata", "annoFeatureCentric2.png", package = "ChIPpeakAnno")

knitr::include_graphics(png)

Bi-directional promoters are genomic regions that are located upstream of the TSS of two adjacent genes that are transcribed in opposite directions [@adachi2002]. Those promoters typically regulate the expression of both genes. To annotate peaks located in such regions, you can use output = "overlapping", FeatureLocForDistance = "TSS", and bindingRegion = c(-5000, 3000). By default, select = "all", meaning that all overlapping features will be outputted. If you want to annotate peaks to features with the nearest bi-directional promoters, you can use output = "nearestBiDirectionalPromoters" along with bindingRegion. In this setting, at most one feature will be reported from each direction. When using output = "nearestBiDirectionalPromoters", both maxgap and FeatureLocForDistance will be ignored.

To illustrate, we can use the myPeakList provided by r Biocpkg("ChIPpeakAnno"). As different major genome releases (e.g. hg19 vs hg38) may have variations in feature coordinates, it is highly recommended to use an annotation file that matches the genome version used when generating your peak file. In the case of myPeakList, the peaks were originally called against hg18, so it is necessary to use a matching annotation file created with hg18. Alternatively, you can use the rtracklayer::liftOver() function to convert myPeakList to hg38 coordinates. For step-by-step instructions, refer to "Step3" in Section \@ref(custompool).

Example 1: find the nearest features {#findnearest}

We can annotate the peaks by assigning the nearby features to them. This can be achieved by setting output = "nearestLocation". When this option is used, the results may include "overlapping" features as long as they are the nearest ones to the peaks.

library(TxDb.Hsapiens.UCSC.hg18.knownGene)

data(myPeakList)
peak_to_anno <- myPeakList[1:100]
anno_data <- transcripts(TxDb.Hsapiens.UCSC.hg18.knownGene)

annotated_peak <- annotatePeakInBatch(peak_to_anno, 
                                      output = "nearestLocation",
                                      PeakLocForDistance = "start",
                                      FeatureLocForDistance = "TSS",
                                      AnnotationData = anno_data)
head(annotated_peak, n = 2)

Here is a breakdown of the options:

• output = "nearestLocation": (default)
  • output the nearest features calculated as abs(PeakLocForDistance - FeatureLocForDistance). The output can consist of both "strictly nearest features (non-overlapping)" and "overlapping features" as long as it is the nearest
• PeakLocForDistance = "start" (default):
  • "start" means using the 5' end (relative to plus strand) of the peak to calculate the distance to features
• FeatureLocForDistance = "TSS" (default):
  • "TSS" means using the 5' end (relative to plus strand) for features on the plus strand and use the 3' end (also relative to plus strand) for features on the minus strand to calculate the distance to features

Here is a breakdown of the resulting metadata columns:

• peak: id of the peak
• feature: id of the feature such as Ensembl gene ID
• start_position, end_position, feature_strand: feature coordinates
• insideFeature: relative location of the peak to the feature
  • "upstream": peak resides upstream of the feature
  • "downstream": peak resides downstream of the feature
  • "inside": peak resides inside the feature
  • "overlapStart": peak overlaps with the start of the feature
  • "overlapEnd": peak overlaps with the end of the feature
  • "includeFeature": peak includes the feature entirely
• distancetoFeature: peak-to-feature distance as calculated with abs(PeakLocForDistance - FeatureLocForDistance)
• shortestDistance: the shortest distance from either end of the peak to either end the feature
• fromOverlappingOrNearest: relevant only when output = "both"; "nearestLocation" indicates that the feature is the closest to the peak, can overlap with the peak; "Overlapping" means that the feature overlaps with the peak and it is not the nearest; when output = "nearestLocation", this column should consist of only "NearestLocation"

Example 2: find the nearest and overlapping features {#findboth}

In addition, annotatePeakInBatch can also report both the nearest features and overlapping features by setting output = "both" and the maxgap parameter. For example, the following command outputs the nearest features plus all overlapping features that are within 100bp away.

annotated_peak <- annotatePeakInBatch(peak_to_anno, 
                                      AnnotationData = anno_data,
                                      output = "both",
                                      maxgap = 100)
head(annotated_peak, n = 4)

Now, the fromOverlappingOrNearest column consists of both "NearestLocation" and "Overlapping" categories.

Visualize peak-to-feature distances

The relative location distribution of peak-to-feature can be visualized using the information stored in the insideFeature metadata column.

pie1(table(annotated_peak$insideFeature))

Use custom annotation data

You also have the option to create and pass user-defined feature coordinates in GRanges format as annotationData. For example, if you have a list of transcript factor binding sites obtained by literature mining and would like to use it to annotate your peaks. you can convert the TF binding site coordinates into a GRanges object and then pass that object into annotatePeakInBatch.

TF_binding_sites <- GRanges(seqnames = c("1", "2", "3", "4", "5", "6", "1", 
                                         "2", "3", "4", "5", "6", "6", "6", 
                                         "6", "6", "5"),
                            ranges = IRanges(start = c(967659, 2010898, 2496700, 
                                                       3075866, 3123260, 3857500,
                                                       96765, 201089, 249670, 
                                                       307586, 312326, 385750, 
                                                       1549800, 1554400, 1565000, 
                                                       1569400, 167888600),
                                             end = c(967869, 2011108, 2496920, 
                                                     3076166,3123470, 3857780,
                                                     96985, 201299, 249890, 307796, 
                                                     312586, 385960, 1550599, 
                                                     1560799, 1565399, 1571199, 
                                                     167888999),
                                             names = paste("t", 1:17, sep = "")),
                            strand = c("*", "*", "*", "*", "*", "*", "*", "*", "*", 
                                       "*", "*", "*", "*", "*", "*", "*", "*"))

annotated_peak2 <- annotatePeakInBatch(peaks1, AnnotationData = TF_binding_sites)
head(annotated_peak2, n = 2)

Another example of using user-defined AnnotationData is to annotate peaks by promoters. A promoter is typically defined as the DNA sequence located immediately upstream of the TSS of a gene. The specific size of a promoter can vary depending on the gene, its regulatory complexity, and the species being studied. In practice, the promoter region can be defined as 2000bp upstream and 500bp downstream from the TSS. To prepare a custom annotation file containing only promoters, users can leverage the accessor function promoters. Similar results can be obtained using the feature-centric approach mentioned in Section \@ref(featurecentric).

promoter_regions <- promoters(TxDb.Hsapiens.UCSC.hg18.knownGene, 
                              upstream = 2000, downstream = 500)
head(promoter_regions, n = 2)

annotated_peak3 <- annotatePeakInBatch(peak_to_anno, 
                                       AnnotationData = promoter_regions)
head(annotated_peak3, n = 2)

Add other feature IDs {#addids}

Depending on the annotation file you use, the feature assigned to your peaks may have different feature IDs. For example, if you annotate your peaks with genes using the TxDb.Hsapiens.UCSC.hg38.knownGene annotation file, the feature id provided in the annotation file will be "entrez ID". On the other hand, if you annotate your peaks using the EnsDb.Hsapiens.v86 annotation file, the feature id in the annotation will be "Ensembl gene ID".

anno_txdb <- genes(TxDb.Hsapiens.UCSC.hg38.knownGene)
head(anno_txdb$gene_id, n = 5)

anno_ensdb <- genes(EnsDb.Hsapiens.v86)
head(anno_ensdb$gene_id, n = 5)

The feature id in the annotation file will be listed under the "feature" metadata column in your annotated peak GRanges object. To link these feature IDs to other IDs such as "symbol", you can use the addGeneIDs function.

The addGeneIDs function can accept either a vector of feature IDs or an annotated peak GRanges object as input. It works by creating a mapping between the input feature IDs and the IDs to be linked, using either an organism annotation dataset (OrgDb object, such as org.Hs.eg.db) or a BioMart dataset (Mart object, such as useMart(biomart = "Ensembl", dataset = "hsapiens_gene_Ensembl")).

To use the function correctly, you need to provide the input feature ID type using the feature_id_type argument, and specify the feature ID types that need to be linked via the IDs2Add argument. The supported feature_id_type and IDs2Add differ between OrgDb and Mart. Below summarizes the commonly used ID types.

• Use OrgDb

  • feature_id_type: input feature ID type
    • "ensembl_gene_id"
    • "entrez_id"
    • "gene_symbol"
    • "gene_alias"
    • "refseq_id"
  • IDs2Add: features ID types to be linked
    • "ensembl"
    • "ensembltrans"
    • "ensemblprot"
    • "entrez_id"
    • "refseq"
    • "symbol"

• Use Mart

  • feature_id_type: input feature ID type, use listFilters(mart) to see the full list
    • "ensembl_gene_id"
    • "ensembl_transcript_id"
    • "ensembl_exon_id"
    • "external_gene_name"
    • "entrezgene_id"
  • IDs2Add: feature ID types to be linked, use listAttributes(mart) to see the full list
    • "ensembl_gene_id"
    • "ensembl_transcript_id"
    • "ensembl_exon_id"
    • "gene_biotype"
    • "transcript_biotype"
    • "entrezgene_id"
    • "hgnc_symbol"
    • "entrezgene_trans_name"

Example1: find gene symbols for a vector of entrez IDs

The following example demonstrates how to use the addGeneIDs function to find gene symbols for a vector of entrez IDs using OrgDb. Note that the "org.Hs.eg.db" package name must be quoted.

library(org.Hs.eg.db)
library(TxDb.Hsapiens.UCSC.hg38.knownGene)

ucsc.hg38.knownGene <- genes(TxDb.Hsapiens.UCSC.hg38.knownGene)
entrez_ids <- head(ucsc.hg38.knownGene$gene_id, n = 10)
print(entrez_ids)

res <- addGeneIDs(entrez_ids, 
                  orgAnn = "org.Hs.eg.db", 
                  feature_id_type = "entrez_id",
                  IDs2Add = "symbol")
head(res, n = 3)

Example2: add gene symbols to annotated peaks

For this example, we annotate the macs_peak_gr2 (obtained in Section \@ref(readinpeak)) using transcript information from TxDb, and add gene symbols to them.

txdb.hg38.transcript <- transcripts(TxDb.Hsapiens.UCSC.hg38.knownGene)
head(txdb.hg38.transcript, n = 4)
head(names(txdb.hg38.transcript), n = 4)

It appears that the txdb.hg38.transcript contains a metadata column called tx_name, which holds the Ensembl transcript IDs. However, the annotatePeakInBatch function requires this information to be stored as the names of the txdb.hg38.trancsript object (names(txdb.hg38.transcript)). This is necessary to generate annotated peaks that are compatible with the addGeneIDs function. To achieve this, we need to extract the transcript IDs and assign them as names. Below is how.

tr_id <- txdb.hg38.transcript$tx_name
tr_id <- sub("\\..*$", "", tr_id) # get rid of the trailing version number
names(txdb.hg38.transcript) <- tr_id
head(txdb.hg38.transcript, n = 4)

Now, we can annotate macs_peak_gr2 with annotatePeakInBatch.

res <- annotatePeakInBatch(macs_peak_gr2, 
                           AnnotationData = txdb.hg38.transcript)
head(res, n = 2)

As anticipated, the resulting feature column contains Ensembl transcript IDs. Next, we utilize the mart option to add gene symbols.

library(biomaRt)

mart <- useMart(biomart = "ENSEMBL_MART_ENSEMBL",
                dataset = "hsapiens_gene_ensembl")
res <- addGeneIDs(res, 
                  mart = mart,
                  feature_id_type = "ensembl_transcript_id",
                  IDs2Add = "hgnc_symbol")
head(res, n = 2)

Use listMarts to show available biomart and use listDatasets to show available dataset. Be aware that, unlike using OrgDb option, we must supply hgnc_symbol instead of symbol for the IDs2Add argument.

Enrichment analysis

Enrichment analysis is a crucial step in determining whether specific biological processes, pathways, or functional categories are over-represented among the genes associated with the peaks. This analysis provides functional implications for the annotated peaks. Two commonly used method for enrichment analysis are gene ontology (GO) and pathway enrichment.

The GO is a structured vocabulary that categorizes genes and their products into three main categories: Molecular Function (what it does), Biological Process (how it does it), and Cellular Component (where it does it). A pathway, on the other hand, refers to a set of predefined genes involved in a coordinated sequence of molecular events or cellular processes that collectively perform a specific biological function. Examples of biological pathways include the "Glycolysis pathway", which breaks down glucose into pyruvate; and the "MAPK signaling pathway", which is involved in cell proliferation, differentiation, and response to external stimuli. While GO enrichment analysis provides more general insights, pathway analysis focuses specifically on predefined pathways. Both methods are commonly practiced and can be achieved with the getEnrichedGO and getEnrichedPATH function in r Biocpkg("ChIPpeakAnno").

In the following demonstration, we first annotate macs_peak_gr2 (obtained in Section \@ref(readinpeakdemo)) using genes from EnsDb, and perform GO and pathway enrichment analysis.

anno_data <- genes(EnsDb.Hsapiens.v86)
annotated_peak4 <- annotatePeakInBatch(macs_peak_gr2,
                                       AnnotationData = anno_data,
                                       output = "both")

enriched_go <- getEnrichedGO(annotated_peak4, 
                             orgAnn = "org.Hs.eg.db", 
                             feature_id_type = "ensembl_gene_id")
enrichmentPlot(enriched_go, label_wrap = 60)

Please ensure to enclose the "org.Hs.eg.db" in quotes, and that the feature_id_type matches the ID type stored in the feature metadata column of your annotated peak object. If you are using genes from TxDb for peak annotation, the feature is likely entrez_id. If you are using genes from EnsDb for annotation, the feature should be ensembl_gene_id. The code snippet below shows the number of enriched GO terms in each category, "bp" for "biological process", "cc" for "cellular component", and "mf" for "molecular function". By default, multiAdjMethod = NULL meaning that no multiple testing correction is performed.

length(enriched_go[["bp"]][["go.id"]])
length(enriched_go[["cc"]][["go.id"]])
length(enriched_go[["mf"]][["go.id"]])

It turns out that if using the subset of annotated macs_peak_gr2 leads to zero enriched GO terms under the "bp" and "cc" categories, and 6 hits under the "mf" category. It suggests taht there are no significantly enriched "bp" or "cc" terms associated with the subset of peaks. However, there are 6 significantly enriched "mf" terms.

head(enriched_go[["mf"]], n = 2)

Pathway enrichment analysis can be performed using popular databases such as Reactome and KEGG (Kyoto Encyclopedia of Genes and Genomes). Reactome is renowned for its detailed and expert-curated information, while KEGG offers a broader scope of information, including pathways, diseases, drugs, and more organisms. The getEnrichedPATH function can use either database by specifying the pathAnn parameter.

For demonstration, we will use the built-in dataset annotatedPeaks with Reactome database. Be aware that the feature_id_type for this dataset is ensembl_gene_id. To switch to the KEGG database, simply set pathAnn = "KEGGREST".

library(reactome.db)

data(annotatedPeak)
enriched_path <- getEnrichedPATH(annotatedPeak,
                                 orgAnn = "org.Hs.eg.db",
                                 feature_id_type = "ensembl_gene_id",
                                 pathAnn = "reactome.db")

To visualize the enriched pathways, we can use the enrichmentPlot function.

enrichmentPlot(enriched_path)

To use getEnrichedGO and getEnrichedPATH, an OrgDb annotation package is necessary. For species that are less common and do not have a valid OrgDb available, users can find alternative methods in this post.

Motif analysis

Sequence motif refer to a recurring pattern in DNA that is believed to have a biological function. These motifs often indicate binding sites for proteins like TFs, some other motifs play a role at the RNA level such as ribosome binding and transcription termination. For peaks obtained through experiments like TF ChIP-seq, motif analysis aids in validating expected binding factors or discovering the TFBSs, while unanticipated motifs suggest the presence of co-binding factors.

r Biocpkg("ChIPpeakAnno") provides several functions that are related to motif analysis, details see below. + getAllPeakSequence: obtain genomic sequences around peaks + Obtained sequences can be used for motif discovery or PCR validation + oligoSummary: find consensus sequences (motifs) in peak sequences + summarizePatternInPeaks: check if given motifs appear in peak sequences

Obtain sequences surrounding the peaks

Here is an example of how to retrieve the peak sequences, including 20bp upstream and 20bp downstream, for the macs_peak_gr2 peaks obtained in Section \@ref(readinpeak).

library(BSgenome.Hsapiens.UCSC.hg38)

sequence_around_peak <- getAllPeakSequence(macs_peak_gr2, 
                                           upstream = 20,
                                           downstream = 20, 
                                           genome = BSgenome.Hsapiens.UCSC.hg38)
head(sequence_around_peak, n = 2)

The genome argument can accept either a BSgenome object or a Mart object. It is important to ensure that the genome version used matches the one used for creating the peak file. You can find a full list of available BSgenome objects on this site.

The following example demonstrates on how to use the mart option. It is slower compared to using a BSgenome because it queries the BioMart for annotations on the fly if AnnotationData is not set. Refer to Section \@ref(usebiomart) for more on Mart.

library(biomaRt)

mart <- useMart(biomart="ENSEMBL_MART_ENSEMBL",
                dataset="hsapiens_gene_ensembl")

sequence_around_peak <- getAllPeakSequence(macs_peak_gr2[1], 
                                           upstream = 20,
                                           downstream = 20, 
                                           genome = mart)

To save the sequences into fasta, use the write2FASTA function.

write2FASTA(sequence_around_peak, file = "macs_peak_gr2.fa")

Discover consensus sequences (motifs) in the peaks

The oligoSummary function utilizes Markov models to ascertain if a motif is enriched in a set of sequences relative to the background. As a prerequisite, we must first calculate the frequencies of all combinations of short oligonucleotides of a specified length in the background [@leung1996over]. This can be accomplished using the oligoFrequency function. In the example below, we aim to identify consensus sequences of length 6.

freqs <- oligoFrequency(BSgenome.Hsapiens.UCSC.hg38$chr1)
motif_summary <- oligoSummary(sequence_around_peak, 
                              oligoLength = 6,
                              MarkovOrder = 3,
                              freqs = freqs,
                              quickMotif = TRUE)

Here is a breakdown of the arguments:

• oligoLength: the length of the motifs to search for
• MarkovOrder: the order of a Markov chain, which determines how much the current state in the chain depends on previous states; simply put, a zero-order Markov chain does not consider any context or dependencies between adjacent elements, while higher-order Markov chains capture longer-range dependencies in sequences, making them more suitable for modeling complex sequence patterns
• freqs: the background motif frequencies used for the Markov model
• quickMotif: whether or not to generate the motif matrices

The resulting motif_summary is a list containing three elements:

• zscore: the Z-score of each oligonucleotide, which quantifies how many standard deviations away the observed count is from the expected count; a high positive score suggests that the motif is significantly over-represented in the peaks
• counts: the count numbe of each oligonucleotide
• motifs: a list of motif matrices when quickMotif = TRUE

We can use histogram (hist) to visualize the resulting Z-scores and labels top hits with the text function. Below, we label the name of the motif that has the highest Z-score.

zscore <- sort(motif_summary$zscore)
h <- hist(zscore, breaks = 100, main = "Histogram of Z-score")
text(x = zscore[length(zscore)], 
     y = h$counts[length(h$counts)] + 1, 
     labels = names(zscore[length(zscore)]), 
     adj = 0, 
     srt = 90)

To illustrate how the Z-score can be influenced by the enrichment level of various motifs, the following code utilizes simulated data. We first generate 5000 sequences with lengths ranging between 100 and 2000 nucleotides. Subsequently,we randomly introduce motif 1 into 10% of the sequences and motif 2 into 15% of the sequences.

set.seed(1)

# motifs to simulate
simulate_motif_1 <- "AATTTA"
simulate_motif_2 <- "TGCATG"

# sample 5000 sequences with lengths ranging from 100 to 2000 nucleotides
# randomly insert motif_1 to 10% of the sequences, and motif_2 to 15% of the sequences
simulation_seqs <- sapply(sample(c(1, 2, 0), 
                                 5000,
                                 prob = c(0.1, 0.15, 0.75),
                                 replace = TRUE), 
                           function(x) {
                             seq <- sample(c("A", "T", "C", "G"),
                                           sample.int(1900, 1) + 100, 
                                           replace = TRUE)
                             insert_pos <- sample.int(length(seq) - 6, 1)
                             if (x == 1) {
                               seq[insert_pos:(insert_pos + 5)] <- strsplit(simulate_motif_1, "")[[1]]
                             } else if (x == 2) {
                               seq[insert_pos:(insert_pos + 5)] <- strsplit(simulate_motif_2, "")[[1]]
                             }
                             paste(seq, collapse = "")
                           }
)

motif_summary_simu <- oligoSummary(simulation_seqs, 
                                   oligoLength = 6, 
                                   MarkovOrder = 3, 
                                   quickMotif = TRUE)
zscore_simu <- sort(motif_summary_simu$zscore, 
                    decreasing = TRUE)
h_simu <- hist(x = zscore_simu, 
               breaks = 100, 
               main = "Histogram of Z-score for simulation data")
text(x = zscore_simu[1:2],  
     y = rep(5, 2), 
     labels = names(zscore_simu[1:2]), 
     adj = 0, 
     srt = 90)

As evident from the simulation results, the higher the probability of the motif, the larger the resulting Z-score. In addition, you can visualize the motif using the r Biocpkg("motifStack") package.

library(motifStack)

pfm <- new("pfm", mat = motif_summary_simu$motifs[[1]],
           name = "sample motif 1")
motifStack(pfm)

To loop through each element in motif_summary$motifs, we can use the mapply function.

pfms <- mapply(function(motif, id) { new("pfm", mat = motif, name = as.character(id)) },
               motif_summary$motifs,
               seq_along(motif_summary$motifs))

motifStack(pfms[[1]])

Scan pre-defined sequence patterns in the peaks

If you have a list of motifs (sequence patterns), the summarizePatternInPeaks function can be utilized to determine whether they are present in the peaks.

example_pattern_file <- system.file("extdata/examplePattern.fa",
                                    package = "ChIPpeakAnno")
readLines(example_pattern_file)
pattern_in_peak <- summarizePatternInPeaks(patternFilePath = example_pattern_file,
                                           BSgenomeName = BSgenome.Hsapiens.UCSC.hg38,
                                           peaks = macs_peak_gr2[1:200],
                                           bgdForPerm = "chromosome",
                                           nperm = 100, 
                                           method = "permutation.test")

pattern_in_peak$motif_enrichment
head(pattern_in_peak$motif_occurrence, n = 2)

The summarziePatternInPeaks function provides two distinct methods (method = c("binom.test", "permutation.test")) for evaluating the significance of given motif enrichment within target peaks. When utilizing the "binom.test" method, the expected frequencies for each motif must be determined first, which can be achieved via expectFrequencyMethod = c("Markov", "Naive"). When employing the "permutation.test" method, users need to provide parameters such as the number of permutation ("nperm"), the significance level ("alpha"), and methods how background sequences are determined (bgdForPerm = c("shuffle", "chromosome"), chromosome = c("asPeak", "random")).

The outcome from summarizePatternInPeaks comprises two tables. The "motif_enrichment" table succinctly summarizes the enrichment statistics for each motif. Specifically, when method = "binom.test", the last column "pValueBinomTest" represents the P-values resulting from the binomial test. Conversely, when method = "permutation.test", the last column "cutOffPermutationTest" denotes the threshold value derived from the permutation test, determined by the specified signifiance level. The "motif_occurrence" table contains detailed information regarding the occurrences of each motif within every peak.

Alternative tools

Besides using r Biocpkg("ChIPpeakAnno"), users can extract the sequences with the getAllPeakSequence function and employ other tools such as MEME Suite for motif-related analysis.

Peak profile comparison {#profilecompare}

When analyzing two or more peak sets from different experiments (e.g. two TFs), it can be insightful to examine whether the peak profiles correlate, and if they do, how do they compare to each other. r Biocpkg("ChIPpeakAnno") not only offers functions to assess if there is a significant overlap among multiple peak sets, but it also provides visualization functions for easy comparison of peak patterns side-by-side.

The significance of overlap can be determined with hypergeometric test or permutation test, both of which are available in r Biocpkg("ChIPpeakAnno"). Please be advised that, due to the inherent challenges in estimating the totalTest parameter for the hypergeometric test, we strongly discourage the use of this approach.

Use hypergeometric test to determine overlap among peak sets {#hypertest}

The hypergeometric test is grounded in the principle of the hypergeometric distribution, which is a probability distribution that describes the likelihood of drawing a specific number of successes (k) from a sample size of n, given a finite population (N) that contains K successes when sampling is done without replacement. The null hypothesis posits that the sample is drawn randomly from the population, implying that the there is no overlap between the two sets of peaks. A small P-value indicates that the null should be rejected, indicating a significant overlap between the two sets of peaks.

In r Biocpkg("ChIPpeakAnno"), the hypergeometric test is incorporated into the makeVennDiagram function, as detailed in Section \@ref(venn). The following example illustrates how to compute the hypergeometric test P-values for peak sets from three TF ChIP-seq experiments.

tf1 <- toGRanges(system.file("extdata/TAF.broadPeak", package = "ChIPpeakAnno"),
                 format = "broadPeak")
tf2 <- toGRanges(system.file("extdata/Tead4.broadPeak", package = "ChIPpeakAnno"),
                 format = "broadPeak")
tf3 <- toGRanges(system.file("extdata/YY1.broadPeak", package = "ChIPpeakAnno"),
                 format = "broadPeak")

To effectively apply the hypergeometric test, we first need to estimate the total number of binding sites, i.e. totalTest. The selection of totalTest affects the stringency of the test, with smaller values resulting in a more conservative outcome (larger P-values). For practical guidance on how to choose an appropriate value for totalTest, you can refer to this post.

In our example, we assume that potential binding regions (which include coding regions and promoter regions) constitute 3% (fairly rough estimation) of the entire genome. Since the example data is derived from chromosome 2, we can estimate the number of total binding sites as (length of chr2) * 3% / (mean peak length).

overlapping_peaks <- findOverlapsOfPeaks(tf1, 
                                         tf2, 
                                         tf3, 
                                         connectedPeaks = "keepAll")
mean_peak_width <- mean(width(unlist(GRangesList(overlapping_peaks[["all.peaks"]]))))

total_binding_sites <- length(BSgenome.Hsapiens.UCSC.hg38[["chr2"]]) * 0.03 / mean_peak_width
venn1 <- makeVennDiagram(overlapping_peaks, 
                         totalTest = total_binding_sites, 
                         connectedPeaks = "keepAll", 
                         fill = c("#CC79A7", "#56B4E9", "#F0E442"),
                         col = c("#D55E00", "#0072B2", "#E69F00"),
                         cat.col = c("#D55E00", "#0072B2", "#E69F00"))

For the P-values of each peak pair:

venn1[["p.value"]]

For the overlapping peak counts:

venn1[["vennCounts"]]

The parameter connectedPeaks denotes how to calculate peak counts when multiple peaks from different groups overlap. If connectedPeaks = "keepFirstListConsistent", the counts from the first group will be consistently displayed in the plot. If connectedPeaks = "keepAll", you will see multiple numbers: all original peak counts for each group will be displayed in parentheses, while the count of the minimally involved peaks will be displayed outside the parentheses.

venn2 <- makeVennDiagram(overlapping_peaks, 
                         totalTest = total_binding_sites, 
                         connectedPeaks = "keepFirstListConsistent", 
                         fill = c("#CC79A7", "#56B4E9", "#F0E442"),
                         col = c("#D55E00", "#0072B2", "#E69F00"),
                         cat.col = c("#D55E00", "#0072B2", "#E69F00"))

Given that all the P-values are extremely small, we must reject the null hypothesis. This indicates that each pair of peak sets significantly overlaps with each other. A major drawback of this approach is the necessity to estimate a totalTest, which could dramatically affect the test results. For instance, if we choose "2%" instead of "3%" in the above example, the P-value for tf1 vs. tf2 increases to 0.49, meaning we can no longer reject the null. To circumvent the requirement of totalTest, we have integrated a permutation test in the peakPermTest function.

Use permutation test to determine overlap among peak sets {#permtest}

The permutation test is a non-parametric test, which means it does not require the data to adhere to any specific distribution. The test statistic is determined according to the observed data, and the null distribution of the test statistic is estimated using a permutation (re-sampling) procedure.

In our scenario, the number of overlapping peaks is treated as the test statistic. Its null distribution is estimated by first re-sampling peaks from a random peak list (the peak pool, which represents all potential binding sites) followed by counting the number of overlapping peaks. The random peak list is generated using the distributions found from the input peaks, ensuring that the peak widths and relative binding positions to the features (such as TSS and geneEnd) follow the same distributions as the input peaks. When the null hypothesis is valid, the number of overlapping peaks is not significantly different from what would be expected by chance. Here are some sample codes using peakPermTest.

Example1 demonstrates a permutation test for non-relevant peak sets.

library(TxDb.Hsapiens.UCSC.hg38.knownGene)

txdb <- TxDb.Hsapiens.UCSC.hg38.knownGene
set.seed(123)

# Example1: non-relevant peak sets
utr5 <- unique(unlist(fiveUTRsByTranscript(txdb)))
utr3 <- unique(unlist(threeUTRsByTranscript(txdb)))

utr5 <- utr5[sample.int(length(utr5), 1000)]
utr3 <- utr3[sample.int(length(utr3), 1000)]

pt1 <- peakPermTest(peaks1 = utr3, 
                    peaks2 = utr5,
                    TxDb = txdb, 
                    maxgap = 500,
                    seed = 1)
plot(pt1)

png <- system.file("extdata", "permTest1.png", package = "ChIPpeakAnno")

knitr::include_graphics(png)

Example2 demonstrates a permutation test for highly relevant peak sets.

# Example2: highly relevant peak sets
cds <- unique(unlist(cdsBy(txdb)))
ol <- findOverlaps(cds, utr3, maxgap = 1)
peaks2 <- c(cds[sample.int(length(cds), 500)],
            cds[queryHits(ol)][sample.int(length(ol), 500)])
pt2 <- peakPermTest(peaks1 = utr3,
                    peaks2 = peaks2,
                    TxDb = txdb,
                    maxgap = 500,
                    seed = 1)
plot(pt2)

png <- system.file("extdata", "permTest2.png", package = "ChIPpeakAnno")

knitr::include_graphics(png)

As observed, for highly relevant peak sets, the P-value is very small.

Use custom peak pool {#custompool}

The peakPermTest function can automatically generate a peak pool given the peak binding type (bindingType = c("TSS", "geneEnd")), annotation type (featureType = c("transcript", "exon")), and annotation data (TxDb). Additionally, users have the option to construct the peak pool from actual binding sites derived from experimental data, with hot spots removed. Hot spots refer to genomic regions that have a high likelihood of being bound by many TFs in ChIP-seq experiments [@yip2012]. We recommend removing hot spots prior to the permutation test to prevent overestimation of the association between two input peak sets. Users are also advised to remove ENCODE blacklist regions. The blacklists were constructed by identifying consistently problematic regions across independent cell lines and types of experiments for each species in the ENCODE and modENCODE datasets [@encode2012].

Below is an example of creating a peak pool for the human genome using the TF binding site clusters downloaded from ENCODE. The following steps are involved:

• Step1: download TF binding sites
• Step2: download hot spots
• Step3: liftover hot spots to hg38 (if necessary)
• Step4: select peak sets to test
• Step5: remove hot spots from binding pool
• Step6: perform permutation test

# Step1: download TF binding sites
temp <- tempfile()
download.file(file.path("https://hgdownload.cse.ucsc.edu/",
                        "goldenpath/",
                        "hg38/",
                        "encRegTfbsClustered/",
                        "encRegTfbsClusteredWithCells.hg38.bed.gz"), 
              temp)
df_tfbs <- read.delim(gzfile(temp, "r"), header = FALSE)
unlink(temp)

colnames(df_tfbs)[1:4] <- c("seqnames", "start", "end", "TF")
tfbs_hg38 <- GRanges(as.character(df_tfbs$seqnames),
                     IRanges(df_tfbs$start, df_tfbs$end),
                     TF = df_tfbs$TF)

# Step2: download hot spots
base_url <- "http://metatracks.encodenets.gersteinlab.org/metatracks/"

temp1 <- tempfile()
temp2 <- tempfile()
download.file(file.path(base_url, 
                        "HOT_All_merged.tar.gz"), 
              temp1)
download.file(file.path(base_url, 
                        "HOT_intergenic_All_merged.tar.gz"),
              temp2)
untar(temp1, exdir = dirname(temp1))
untar(temp2, exdir = dirname(temp1))
bedfiles <- dir(dirname(temp1), "bed$")
hot_spots_hg19 <- sapply(file.path(dirname(temp1), bedfiles), toGRanges, format = "BED")
unlink(temp1)
unlink(temp2)

names(hot_spots_hg19) <- gsub("_merged.bed", "", bedfiles)
hot_spots_hg19 <- sapply(hot_spots_hg19, unname)
hot_spots_hg19 <- GRangesList(hot_spots_hg19)

# Step3: liftover hot spots to hg38
library(R.utils)

temp_chain_gz <- tempfile()
temp_chain <- tempfile()
base_url_chain <- "http://hgdownload.cse.ucsc.edu/goldenpath/"
download.file(file.path(base_url_chain, 
                        "hg19/liftOver/",
                        "hg19ToHg38.over.chain.gz"),
              temp_chain_gz)
gunzip(filename = temp_chain_gz, destname = temp_chain)
chain_file <- import.chain(hg19_to_hg38)
unlink(temp_chain_gz)
unlink(temp_chain)

hot_spots_hg38 <- liftOver(hot_spots_hg19, chain_file)

# Step4: select peak sets to test
tfbs_hg38_by_TF <- split(tfbs_hg38, tfbs_hg38$TF)
TAF1 <- tfbs_hg38_by_TF[["TAF1"]]
TEAD4 <- tfbs_hg38_by_TF[["TEAD4"]]

# Step5: remove hot spots from binding pool
tfbs_hg38 <- subsetByOverlaps(tfbs_hg38, hot_spots_hg38, invert=TRUE)
tfbs_hg38 <- reduce(tfbs_hg38)

# Step6: perform permutation test
pool <- new("permPool", 
            grs = GRangesList(tfbs_hg38), 
            N = length(TAF1))
pt3 <- peakPermTest(TAF1, TEAD4, pool = pool, ntimes = 500)
plot(pt3)

png <- system.file("extdata", "permTest3.png", package = "ChIPpeakAnno")

knitr::include_graphics(png)

Visualize TSS enrichment signal of multiple experiments {#multiexp}

If you have peak files obtained from multiple TF ChIP-seq experiments and would like to compare their enriched signals around TSS using raw signals such as read coverage. r Biocpkg("ChIPpeakAnno") provides two functions, featureAlignedHeatmap and featureAlignedDistribution, to visualize their binding patterns side-by-side.

To illustrate this, we first need to prepare both the peak data and coverage information (bigWig). This involves four steps:

• Step1: read in example peak files
• Step2: find peaks that are shared by all
• Step3: read in example coverage files
• Step4: visualize binding patterns

# Step1: read in example peak files
extdata_path <- system.file("extdata", package = "ChIPpeakAnno")
broadPeaks <- dir(extdata_path, "broadPeak")
gr_TFs <- sapply(file.path(extdata_path, broadPeaks), toGRanges, format = "broadPeak")
names(gr_TFs) <- gsub(".broadPeak", "", broadPeaks)
names(gr_TFs)

Now, we have imported peak files for three TFs ("TAF", "Tead", and "YY1") into the gr_TFs object. The next step is to identify overlapping peaks to ensure that we are comparing binding patterns over the same genomic regions.

# Step2: find peaks that are shared by all
ol <- findOverlapsOfPeaks(gr_TFs)
gr_TFs_ol <- ol$peaklist$`TAF///Tead4///YY1`

# Step3: read in example coverage files
# here we read in coverage data from -2000bp to -2000bp of each shared peak center
gr_TFs_ol_center <- reCenterPeaks(gr_TFs_ol, width = 4000) # use the center of the peaks and extend 2000bp upstream and downstream to obtain peaks with uniform length of 4000bp

bigWigs <- dir(extdata_path, "bigWig")
coverage_list <- sapply(file.path(extdata_path, bigWigs), 
                        import, # rtracklayer::import
                        format = "BigWig",
                        which = gr_TFs_ol_center,
                        as = "RleList")

names(coverage_list) <- gsub(".bigWig", "", bigWigs)
names(coverage_list)

Heatmap

# Step4: visualize binding patterns
sig <- featureAlignedSignal(coverage_list, gr_TFs_ol_center)

# since the bigWig files are only a subset of the original files,
# filter to keep peaks that are with coverage data for all peak sets
keep <- rowSums(sig[[1]]) > 0 & rowSums(sig[[2]]) > 0 & rowSums(sig[[3]]) > 0
sig <- sapply(sig, function(x) x[keep, ], simplify = FALSE)
gr_TFs_ol_center <- gr_TFs_ol_center[keep]

featureAlignedHeatmap(sig, gr_TFs_ol_center,
                      upper.extreme=c(3, 0.5, 4))

By default, the rows in the heatmap are ordered by the total coverage per row from the first sample ("TAF" in this example). We can reorder the rows by tuning the sortBy option. For example, setting sortBy = "YY1" will order the rows by the "YY1" sample in the dataset. You can also sort the rows based on hierarchical clustering results. Here is a demonstration:

# perform hierarchical clustering on rows
sig_rowsums <- sapply(sig, rowSums, na.rm = TRUE)
row_distance <- dist(sig_rowsums)
hc <- hclust(row_distance)

# use hierarchical clustering order to sort
gr_TFs_ol_center$sort_by <- hc$order
featureAlignedHeatmap(sig, gr_TFs_ol_center, 
                      upper.extreme = c(3, 0.5, 4),
                      sortBy = "sort_by")

Density plot

Additionally, we can create a density plot using the featureAlignedDistribution function.

featureAlignedDistribution(sig, gr_TFs_ol_center,
                           type = "l")

Common workflow 1: single TF with replicates {#workflow1}

For experiments targeting a single TF with replicates, a common analytic strategy is outlined below.

• Step1: import data
• Convert BED/GFF files into GRanges
• Find overlapping peaks among replicates
• Add metadata (optional)
• Visualize replicate concordance using a Venn diagram
• Step2: prepare annotation file
• Step3: visualize peak distribution
• Step4: annotate peaks
• Find possible enhancers
• Find potential bi-directional promoters
• Step5: perform enrichment analysis
• Step6: conduct motif analysis

Step1: import data

Common peak formats such as BED, GFF, and MACS can be converted into GRanges format using the toGRanges function. After importing the data, concordance across peak replicates will be evaluated with findOverlapsOfPeaks and makeVennDiagram. Be aware that the metadata columns will be dropped for the merged overlapping peaks. To add them back, we can use the addMetadata function. For example, addMetadata(ol, colNames = "score", FUN = mean) will add a "score" column to each merged overlapping peak by taking the mean score of individual peaks involved.

library(ChIPpeakAnno)

# Convert BED/GFF into GRanges
bed1 <- system.file("extdata", "MACS_output_hg38.bed", 
                    package = "ChIPpeakAnno")
gr1 <- toGRanges(bed1, format = "BED", header = FALSE)
gff1 <- system.file("extdata", "GFF_peaks_hg38.gff", 
                    package = "ChIPpeakAnno")
gr2 <- toGRanges(gff1, format = "GFF", header = FALSE)

# Find overlapping peaks
ol <- findOverlapsOfPeaks(gr1, gr2)

# Add "score" metadata column to overlapping peaks
ol <- addMetadata(ol, colNames = "score", FUN = mean) 
head(ol$mergedPeaks, n = 2)

venn <- makeVennDiagram(ol,
                        fill = c("#009E73", "#F0E442"),
                        col = c("#D55E00", "#0072B2"),
                        cat.col = c("#D55E00", "#0072B2"))

For the P-value of hypergeometric test:

venn[["p.value"]]

For overlapping peak counts:

venn[["vennCounts"]]

As observed, the extremely small P-value suggests a high relevance between the two peak sets, reflecting good consistency among experimental replicates.

Step2: prepare annotation file

Similar to the peak files, the annotation file must also be converted into a GRanges object. Annotation GRanges can be constructed from not only BED, GFF, and user-defined text files, but also from EnsDb and TxDb objects using the toGRanges function. For EnsDb and TxDb objects, annotation can also be prepared with accessor functions, as detailed in Section \@ref(txdbensdb). Note that the version of genome used to create the annotation file must match the genome used for peak calling, as feature positions may vary across different genome releases. As an example, if you are using Mus_musculus.v103 for read mapping, it's best to use EnsDb.Mmusculus.v103 for annotation.

library(EnsDb.Hsapiens.v86)

ensembl.hs86.gene <- toGRanges(EnsDb.Hsapiens.v86, feature = "gene")
head(ensembl.hs86.gene, n = 2)

Step3: visualize peak distribution

Peak distribution relative to features

Now, given the merged overlapping peaks and annotation data, we can visualize the distribution of the distance from the merged overlapping peaks to the nearest features, such as genes (TSSs), using the binOverFeature function.

binOverFeature(ol$mergedPeaks, 
               nbins = 20,
               annotationData = ensembl.hs86.gene,
               xlab = "peak distance from TSSs (bp)", 
               ylab = "peak count", 
               main = "Distribution of aggregated peak numbers around TSS")

Peak distribution over multiple feature levels

We can use the genomicElementDistribution to summarize the distribution of peaks over different types of genomic features such exon, intron, enhancer, UTR. When inputting a single peak file, a pie graph will be generated.

library(TxDb.Hsapiens.UCSC.hg38.knownGene)

genomicElementDistribution(ol$mergedPeaks, 
                           TxDb = TxDb.Hsapiens.UCSC.hg38.knownGene)

As can be seen, a significant number of peaks originate from promoter regions, which is consistent with the signature of peaks obtained from Tf binding experiments. When inputting a list of peak sets (e.g. replicates), a bar graph will be generated.

macs_peaks <- GRangesList(rep1 = gr1,
                          rep2 = gr2)
genomicElementDistribution(macs_peaks, 
                           TxDb = TxDb.Hsapiens.UCSC.hg38.knownGene)

According to the bar graph, the two peak replicates are consistent.

Peak overlappings for multiple features

To visualize peak overlap for multiple feature sets, we can utilize the UpSet plot (for details, see Section \@ref(upset)).

library(UpSetR)

res <- genomicElementUpSetR(ol$mergedPeak,
                            TxDb.Hsapiens.UCSC.hg38.knownGene)
upset(res[["plotData"]], 
      nsets = length(colnames(res$plotData)), 
      nintersects = NA)

Step4: annotate peaks

Based on the above distribution of aggregated peak numbers around the TSS and the distribution of peaks in different chromosomal regions, most of the peaks locate near the TSS. Therefore, it is reasonable to adopt the feature-centric method to annotate the peaks residing in the promoter regions. Promoters can be specified with the bindingRegion parameter. In the following example, the promoter region is defined as 2000bp upstream and 500bp downstream of the TSS (bindingRegion = c(-2000, 500)).

ol_anno <- annotatePeakInBatch(ol$mergedPeak, 
                               AnnotationData = ensembl.hs86.gene, 
                               output = "nearestBiDirectionalPromoters",
                               bindingRegion = c(-2000, 500))
head(ol_anno, n = 2)

You can export the annotation to a CSV file:

ol_anno <- unname(ol_anno) # remove names to avoid duplicate row.names error
ol_anno$peakNames <- NULL # remove peakNames to avoid unimplemented type 'list' error
write.csv(as.data.frame(ol_anno), "ol_anno.csv")

To visualize the distribution of peaks around features:

pie1(table(ol_anno$insideFeature))

Find peaks located in bi-directional promoters

In addition to using the output = "nearestBiDirectionalPromoters" option, r Biocpkg("ChIPpeakAnno") provides another helper function called peaksNearBDP to fetch statistics for peaks situated in bi-directional promoters.

peaks_near_BDP <- peaksNearBDP(ol$mergedPeaks, 
                    AnnotationData = ensembl.hs86.gene, 
                    MaxDistance = 5000) 
# MaxDistance will be translated into "bindingRegion = 
# c(-MaxDistance, MaxDistance)" internally

peaks_near_BDP$n.peaks
peaks_near_BDP$n.peaksWithBDP
peaks_near_BDP$percentPeaksWithBDP

head(peaks_near_BDP$peaksWithBDP, n = 2)

Find peaks located in enhancers

Enhancers are DNA sequences that can amplify gene expression and are frequently used as biomarkers for cancer diagnosis and treatment. They can be identified using a variety of experimental methods such as 3C, 5C, and HiC [@lieberman2009]. With enhancers obtained through these experimental techniques, we can locate peaks that locate to potential enhancer regions. The following example uses 5C data derived with the hg19 genome assembly, hence, it's necessary to use a matching annotation file.

library(EnsDb.Hsapiens.v75)

ensembl.hs75.gene <- toGRanges(EnsDb.Hsapiens.v75, feature = "gene")

DNA5C <- system.file("extdata", 
                     "wgEncodeUmassDekker5CGm12878PkV2.bed.gz",
                     package="ChIPpeakAnno")
DNAinteractiveData <- toGRanges(gzfile(DNA5C))
# the example bed.gz file can also be downloaded from:
# https://hgdownload.cse.ucsc.edu/goldenpath/hg19/encodeDCC/wgEncodeUmassDekker5C/wgEncodeUmassDekker5CGm12878PkV2.bed.gz

peaks_near_enhancer <-  findEnhancers(peaks = ol$mergedPeaks,
                                      annoData = ensembl.hs75.gene, 
                                      DNAinteractiveData = DNAinteractiveData)
head(peaks_near_enhancer, n = 2)

Step5: perform enrichment analysis

With annotated peaks, we can use the getEnrichedGO and getEnrichedPATH functions to respectively retrieve a list of enriched GO or pathway terms.

library(org.Hs.eg.db)

enriched_go <- getEnrichedGO(annotatedPeak = ol_anno, 
                             orgAnn = "org.Hs.eg.db", 
                             feature_id_type = "ensembl_gene_id",
                             condense = TRUE)
enrichmentPlot(enriched_go)

Likewise, the following pertains to pathway enrichment analysis.

library(reactome.db)

enriched_pathway <- getEnrichedPATH(annotatedPeak = ol_anno,
                                    orgAnn = "org.Hs.eg.db", 
                                    pathAnn = "KEGGREST",
                                    maxP = 0.05)
enrichmentPlot(enriched_pathway)

# To remove the common suffix " - Homo sapiens (human)":
enrichmentPlot(enriched_pathway, label_substring_to_remove = " - Homo sapiens \\(human\\)")

Step6: conduct motif analysis

To perform motif analysis, we first need to extract sequences surrounding the peaks, then acquire the consensus sequences. We can also visualize the top motifs discovered.

library(BSgenome.Hsapiens.UCSC.hg38)

seq_around_peak <- getAllPeakSequence(ol$mergedPeaks, 
                                      upstream = 20,
                                      downstream = 20, 
                                      genome = BSgenome.Hsapiens.UCSC.hg38)
head(seq_around_peak, n = 2)

We can use the write2FASTA function to store the result in a FASTA file. The following code snippet generates Z-scores for short oligos of length 6.

freqs <- oligoFrequency(BSgenome.Hsapiens.UCSC.hg38$chr1)
motif_summary <- oligoSummary(seq_around_peak, 
                              oligoLength = 6,
                              MarkovOrder = 3,
                              freqs = freqs,
                              quickMotif = TRUE)
zscore <- sort(motif_summary$zscore)
h <- hist(zscore, 
          breaks = 100, 
          main = "Histogram of Z-score")
text(x = zscore[length(zscore)], 
     y = h$counts[length(h$counts)] + 1, 
     labels = names(zscore[length(zscore)]), 
     adj = 0, 
     srt = 90)

We can use motifStack to visualize the top discovered motifs.

library(motifStack)

pfm <- new("pfm", mat = motif_summary$motifs[[1]],
           name = "sample motif 1")
motifStack(pfm)

Common workflow 2: comparing binding profiles for multiple TFs {#workflow2}

Given two or more peak sets from different TFs, it might be intriguing to examine if the peak profiles are correlated, and if so, how does the peak patterns compare to each other. The workflow presented here demonstrates how to compare the binding profiles of three TFs. The steps are outlined below.

• Step1: import data
• Step2: determine if there is significant overlap among peak sets
• Step3: visualize and compare the binding patterns
• Heatmap
• Density plot

Step1: import data

tf1 <- toGRanges(system.file("extdata/TAF.broadPeak", package = "ChIPpeakAnno"),
                 format = "broadPeak")
tf2 <- toGRanges(system.file("extdata/Tead4.broadPeak", package = "ChIPpeakAnno"),
                 format = "broadPeak")
tf3 <- toGRanges(system.file("extdata/YY1.broadPeak", package = "ChIPpeakAnno"),
                 format = "broadPeak")

Step2: determine if there is significant overlap among peak sets

To examine the associations across different peak sets, r Biocpkg("ChIPpeakAnno") implements both hypergeometric test (makeVennDiagram, for details see Section \@ref(hypertest)) and permutation test (peakPermTest, for details see Section \@ref(permtest)). For demonstration, here we use hypergeometric test.

library(BSgenome.Hsapiens.UCSC.hg38)

overlapping_peaks <- findOverlapsOfPeaks(tf1, 
                                         tf2, 
                                         tf3, 
                                         connectedPeaks = "keepAll")
mean_peak_width <- mean(width(unlist(GRangesList(overlapping_peaks[["all.peaks"]]))))

total_binding_sites <- length(BSgenome.Hsapiens.UCSC.hg38[["chr2"]]) * 0.03 / mean_peak_width
venn1 <- makeVennDiagram(overlapping_peaks, 
                         totalTest = total_binding_sites, 
                         connectedPeaks = "keepAll", 
                         fill = c("#CC79A7", "#56B4E9", "#F0E442"),
                         col = c("#D55E00", "#0072B2", "#E69F00"),
                         cat.col = c("#D55E00", "#0072B2", "#E69F00"))

For the P-values of each peak pair:

venn1[["p.value"]]

Given that the P-values are all extremely low, there is a significant overlap between each pair of peak sets.

Step3: visualize and compare the binding patterns

We can leverage heatmap and density plot to effectively visualize and compare the binding patterns of multiple TFs. For detailed instructions, refer to \@ref(multiexp).

ol_tfs <- findOverlapsOfPeaks(tf1, tf2, tf3, 
                              connectedPeaks = "keepAll")
gr_ol_tfs <- ol_tfs$peaklist$`tf1///tf2///tf3`

TF_width <- width(gr_ol_tfs)
gr_ol_tfs_center <- reCenterPeaks(gr_ol_tfs, width = 4000)

extdata_path <- system.file("extdata", package = "ChIPpeakAnno")
bigWigs <- dir(extdata_path, "bigWig")
coverage_list <- sapply(file.path(extdata_path, bigWigs), 
                        import, # rtracklayer::import
                        format = "BigWig",
                        which = gr_ol_tfs_center,
                        as = "RleList")

names(coverage_list) <- gsub(".bigWig", "", bigWigs)

sig <- featureAlignedSignal(coverage_list, gr_ol_tfs_center)
# since the bigWig files are only a subset of the original files,
# filter to keep peaks that are with coverage data for all peak sets
keep <- rowSums(sig[[1]]) > 0 & rowSums(sig[[2]]) > 0 & rowSums(sig[[3]]) > 0
sig <- sapply(sig, function(x) x[keep, ], simplify = FALSE)
gr_ol_tfs_center <- gr_ol_tfs_center[keep]

featureAlignedHeatmap(sig, gr_ol_tfs_center,
                      upper.extreme=c(3, 0.5, 4))

By default, the rows are arranged according to the total coverage per row from the first sample. Below is an example of how to sort by the third sample ("YY1") in the dataset through tuning the sortBy option.

featureAlignedHeatmap(sig, gr_ol_tfs_center,
                      upper.extreme=c(3, 0.5, 4),
                      sortBy = "YY1")

To generate a density plot, use the featureAlignedDistribution function.

featureAlignedDistribution(sig, gr_ol_tfs_center, 
                           type="l")

As observed, the binding of "YY1" is significantly stronger, while the binding of "Tead" is considerably weaker.

Have questions?

For questions related to usage, please post your queries on the Bioconductor Support Site. If you wish to report a bug or request a new feature, kindly raise an issue on the ChIPpeakAnno GitHub repository.

Selected Q & A

How to import peaks?

r Biocpkg("ChIPpeakAnno") provides a helper function, toGRanges, specifically designed for importing files. Users also have the option to use other tools, such as rtracklayer::import.

How to properly annotate peaks from ChIP-seq data?

r Biocpkg("ChIPpeakAnno") offers a variety of options for annotating peaks, which can be specified with the output argument in the annotatePeakInBatch function.

• If you are interested in the nearest features surrounding the peaks, set output = "nearestLocation", output = "shortestDistance", or output = "overlapping". This is a peak-centric method, for details see Section \@ref(peakcentric).
• If you are interested in the peaks binding to the promoter regions, you can set output = "upstream". For more flexibility, you can adjust the bindingRegion option. This is a feature-centric method, for details see Section \@ref(featurecentric)

Should I use annotatePeakInBatch or annoPeaks?

You should use annotatePeakInBatch as it serves as the primary wrapper function that internally calls annoPeaks. Historically, annotatePeakInBatch was primarily composed of peak-centric methods. Over time, feature-centric methods, initially implemented in the annoPeaks function, were integrated into annotatePeakInBatch.

How to prepare annotation data for annotatePeakInBatch?

Ensure that the annotation data is in GRanges format for annotatePeakInBatch to work. You can convert TxDb, EnsDb, or even GFF and BED, etc. into GRanges using the toGRanges function. Additionally, you can leverage the getAnnotation function to fetch annotation data from BioMart. If needed, you can even create a personalized annotation file. For detailed instructions, see Section \@ref(prepanno).

Can I output either the nearest or overlapping features?

This question pertains to this post.

When you set output = "both", both the "nearest" and "overlapping" features will be output. If your preference is to assign only one type of feature to each peak (either "nearest" or "overlapping", with a preference for "overlapping" in cases where the "nearest" feature is not "overlapping"), you can use the following strategy: first, annotate peaks with overlapping features; second, annotate peaks lacking overlapping features with the nearest features; last, concatenate the two sets of results. The example codes are provided below:

library(ensembldb)
library(EnsDb.Hsapiens.v75)
data(myPeakList)
annoData <- annoGR(EnsDb.Hsapiens.v75)

# Step1: annotate peaks to the overlapping features, if "select = 'all'", multiple features can be assigned to a single peak.
anno_overlapping <- annotatePeakInBatch(myPeakList, 
                                        AnnotationData = annoData, 
                                        output = "overlapping", 
                                        select = "first")
anno_overlapping_non_na <- anno_overlapping[!is.na(anno_overlapping$feature)]

# Step2: annotate peaks that are without overlapping features to nearest features
myPeakList_non_overlapping <- myPeakList[!(names(myPeakList) %in% anno_overlapping_non_na$peak)]  
anno_nearest <- annotatePeakInBatch(myPeakList_non_overlapping,
                                    AnnotationData = annoData, 
                                    output = "nearestLocation", 
                                    select = "first")

# Step3: concatenate the two
anno_final <- c(anno_overlapping_non_na, anno_nearest)

The provided code allocates either the "overlapping" or "nearest" feature to each peak. If the "overlapping" feature is not the "nearest", only the "overlapping" one will be reported.

Why is the sum of peak numbers in the Venn diagram NOT equal to the sum of the peaks in the original peak lists?

This question is a common scenario in calculating the intersection of peaks. Peaks, being a range of continuous points rather than single points, present a challenge in determining the overlapping regions. Consider the intersection of two lists of range objects: list A (1~3, 4~5, 7~9) and list B (2~8), the number of overlapping ranges depends on the reference chosen, if we use list B as the reference, the output would be one range; however, if we use list A as the reference, the output would be three ranges.

How does findOverlapsOfPeaks count the number of overlapping peaks?

This question is in response to this post.

When counting the number of overlapping peaks using findOverlapsOfPeaks, the connectedPeaks option comes into play. If multiple peaks involve in any group of connected or overlapping peaks in any input peak list, setting connectedPeaks = "merge" will increment the overlapping counts by one. On the other hand, setting connectedPeaks = "min" will add the minimal number of involved peaks in each group of connected or overlapped peaks to the overlapping counts. If connectedPeaks = "keepAll", it will add the number of involved peaks for each peak list to the corresponding overlapping counts. In addition, it will output counts as if connectedPeaks was set to "min".

For examples, if 5 peaks in group1 overlap with 2 peaks in group 2:

• Setting connectedPeaks = "merge" will add 1 to the overlapping counts
• Setting connectedPeaks = "min" will add 2 to the overlapping counts
• Setting connectedPeaks = "keepAll" will add 5 peaks to "count.group1", 2 to "count.group2", and 2 to the overlapping counts

Is there a way to show the number of peaks in original peak lists?

You have the option to configure connectedPeaks = "keepAll" in both the findOverlapsOfPeaks and makeVennDiagram functions.

How to extract the original peak IDs of the overlapping peaks?

In the output of findOverlapsOfPeaks, each element in the peak list contains a metadata column names peakNames, which is a CharacterList. This CharacterList is a list of the contributing peak IDs with prefixes, e.g. "peaks1_peakname1", "peaksi_peaknamej", where "i" denotes the peak group i and "j" denotes the specific peak j within that group. Users can retrieve the original peak name by splitting these strings. Here are the sample codes:

library(ChIPpeakAnno)
library(reshape2)

# Step1: read in two peak files
bed <- system.file("extdata", 
                   "MACS_output.bed",
                   package = "ChIPpeakAnno")
gff <- system.file("extdata", 
                   "GFF_peaks.gff", 
                   package = "ChIPpeakAnno")
gr1 <- toGRanges(bed, format = "BED", 
                 header = FALSE)
gr2 <- toGRanges(gff, format = "GFF", 
                 header = FALSE, skip = 3)
names(gr2) <- seq(length(gr2)) # add names to gr2 for Step4

# Step2: find overlapping peaks
ol <- findOverlapsOfPeaks(gr1, gr2)
peakNames <- ol$peaklist[['gr1///gr2']]$peakNames

# Step3: extract original peak names
peakNames <- melt(peakNames, 
                  value.name = "merged_peak_id") # reshape df
head(peakNames, n = 2)

peakNames <- cbind(peakNames[, 1], 
                   do.call(rbind, 
                           strsplit(as.character(peakNames[, 3]), "__")))

colnames(peakNames) <- c("merged_peak_id", "group", "peakName")
head(peakNames, n = 2)

# Step4: split by peak group
gr1_subset <- gr1[peakNames[peakNames[, "group"] %in% "gr1", "peakName"]]
gr2_subset <- gr2[peakNames[peakNames[, "group"] %in% "gr2", "peakName"]]
head(gr1_subset, n = 2)
head(gr2_subset, n = 2)

Alternatively, the following snippet should also work.

gr1_renamed <- ol$all.peaks$gr1
gr2_renamed <- ol$all.peaks$gr2
head(gr1_renamed, n = 2)
head(gr2_renamed, n = 2)

peakNames <- melt(ol$peaklist[['gr1///gr2']]$peakNames, 
                  value.name = "merged_peak_id")
gr1_subset <- gr1_renamed[peakNames[grepl("^gr1", peakNames[, 3]), 3]]
gr2_subset <- gr2_renamed[peakNames[grepl("^gr2", peakNames[, 3]), 3]]
head(gr1_subset, n = 2)
head(gr2_subset, n = 2)

How to select the proper number for totalTest when using makeVennDiagram?

During the evaluation of the association between two sets of peak data using the hypergeometric distribution, it is essential to determine the total number of potential binding sites, the totalTest parameter in the makeVennDiagram. This parameter should have a value larger than the largest number of peaks in the peak list. The choice of totalTest influences teh stringency of the test, with smaller values yielding more stringent tests and larger P-values. The computation time for calculating P-values remains independent of the totalTest value. For practical guidance on selecting an appropriate totalTest value, please refer to this post.

How to cite ChIPpeakAnno

If you use r Biocpkg("ChIPpeakAnno") in your work, please cite it as follows:

citation(package = "ChIPpeakAnno")

Session info

Here is the output of sessionInfo() on the system on which this document was compiled running pandoc r rmarkdown::pandoc_version():

sessionInfo()

jianhong/ChIPpeakAnno documentation built on Jan. 4, 2025, 5:27 p.m.