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\author{Vanja Haberle \footnote{vanja.haberle@gmail.com}}

\title{\textsf{CAGEr: an R package for CAGE (Cap Analysis of Gene Expression) data analysis and promoterome mining}}

\date{April 25, 2016}

\begin{document}
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\tableofcontents

%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Introduction}
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
This document briefly describes how to use the package \Rpackage{CAGEr}. 
\Rpackage{CAGEr} is a Bioconductor-compliant R package designed to manipulate, analyse and visualise 
Cap Analysis of Gene Expression (CAGE) sequencing data. CAGE (\cite{Kodzius:2006}) is a high-throughput method for transcriptome
analysis that utilizes "cap-trapping" (\cite{Carninci:1996}), a technique based on the biotinylation of the 7-methylguanosine cap of Pol II transcripts, 
to pulldown the 5'-complete cDNAs reversely transcribed from the captured transcripts. A linker sequence is ligated to the 5' end of the cDNA and 
a specific restriction enzyme is used to cleave off a short fragment from the 5' end. Resulting fragments are then amplified and sequenced using 
massive parallel high-throughput sequencing technology, which results in a large number of short sequenced tags that can be mapped back to 
the referent genome to infer the exact position of the transcription start sites (TSSs) used for transcription of captured RNAs (Figure~\ref{fig:CAGEprotocol}). 
Number of CAGE tags supporting each TSS gives the information on relative frequency of its usage and can be used as a measure of expression 
from that specific TSS. Thus, CAGE provides information on two aspects of capped transcriptome: genome-wide 1bp-resolution map of transcription 
start sites and transcript expression levels. This information can be used for various analyses, from 5' centered expression profiling (\cite{Takahashi:2012}) to 
studying promoter architecture (\cite{Carninci:2006}).

\begin{figure}
\centering
\includegraphics[width = 150mm]{images/CAGEprotocol.pdf}
\caption{Overview of CAGE experiment}
\label{fig:CAGEprotocol}
\end{figure}

CAGE samples derived from various organisms (genomes) can be analysed by \Rpackage{CAGEr} and the only limitation is the availability of the referent genome as a \Rpackage{BSgenome} package in case when raw mapped CAGE tags are processed. \Rpackage{CAGEr} provides a comprehensive workflow that starts from mapped CAGE tags and includes reconstruction of TSSs and promoters and their visualisation, as well as more specialized downstream analyses like promoter width, expression profiling and
differential TSS usage. It can use both Binary Sequence Alignment Map (BAM) files of aligned CAGE tags or 
files with genomic locations of TSSs and number of supporting CAGE tags as input. If BAM files are provided \Rpackage{CAGEr} constructs TSSs from aligned 
CAGE tags and counts the number of tags supporting each TSS, while allowing filtering out low-quality tags and removing technology-specific bias. 
It further performs normalization of raw CAGE tag count, clustering of TSSs into tag clusters (TC) and their aggregation across multiple CAGE experiments into promoters 
to construct the promoterome. Various methods for normalization and clustering of TSSs are supported. Exporting data into different types of track files allows various visualisations 
of TSSs and clusters (promoters) in the UCSC Genome Browser, which facilitate generation of hypotheses. \Rpackage{CAGEr} manipulates multiple CAGE experiments 
at once and performs analyses across datasets, including expression profiling and detection of differential TSS usage (promoter shifting). Multicore option for parallel 
processing is supported on Unix-like platforms, which significantly reduces computing time.
\newline
\newline
Here are some of the functionalities provided in this package:

\begin{itemize}
\item
Reading in multiple CAGE datasets from various sources; user provided BAM or TSS input files, public CAGE datasets from accompanying data package.
\item
Correcting systematic G nucleotide addition bias at the 5' end of CAGE tags.
\item
Plotting pairwise scatter plots, calculating correlation between datasets and merging datasets.
\item
Normalizing raw CAGE tag count: simple tag per million (tpm) or power-law based normalization (\cite{Balwierz:2009}).
\item
Clustering individual TSSs into tag clusters (TCs) and aggregating clusters across multiple CAGE datasets to create a set of consensus promoters.
\item
Making bedGraph or BED files of individual TSSs or clusters for visualisation in the genome browser.
\item
Expression clustering of individual TSSs or consensus promoters into distinct expression profiles using common clustering algorithms.
\item
Calculating promoter width based on the cumulative distribution of CAGE signal along the promoter.
\item
Scoring and statistically testing differential TSS usage (promoter shifting) and detecting promoters that shift between two samples.
\end{itemize}

Several data packages are accompanying \Rpackage{CAGEr} package. They contain majority of the up-to-date publicly available CAGE data produced by major consortia including FANTOM and ENCODE. These include \Rpackage{FANTOM3and4CAGE} package available from Bioconductor, as well as \Rpackage{ENCODEprojectCAGE} and \Rpackage{ZebrafishDevelopmentalCAGE} packages available from \href{http://promshift.genereg.net/CAGEr/}{http://promshift.genereg.net/CAGEr/}. In addition, direct fetching of TSS data from FANTOM5 web resource (the largest collection of TSS data for human and mouse) from within \Rpackage{CAGEr} is also available. These are all valuable resources of genome-wide TSSs in various tissue/cell types for various model organisms that can be used directly in R. 
Section 5 in this vignette describes how these public datasets can be included into a workflow provided by \Rpackage{CAGEr}. For further information on the content of the data packages and the list of available CAGE datasets please refer to the vignette of the corresponding data package.
\newline
For further details on the implemented methods and for citing the \Rpackage{CAGEr} package in your work please refer to \cite{Haberle:2015}.


%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Input data for \Rpackage{CAGEr}}
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\Rpackage{CAGEr} package supports three types of CAGE data input: 

\begin{enumerate}
\item
\textbf{Sequenced CAGE tags mapped to the genome}: either BAM (Binary Sequence Alignment Map) files of sequenced CAGE tags aligned to the referent genome (including the paired-end data such as CAGEscan) or BED files of CAGE tags (fragments).
\item
\textbf{CAGE detected TSSs (CTSSs)}: tab separated files with genomic coordinates of TSSs and number of tags supporting each TSS. The file should not contain 
a header and the data must be organized in four columns: 
\begin{itemize}
\item
name of the chromosome: names must match the names of chromosomes in the corresponding \Rpackage{BSgenome} package
\item
1-based coordinate of the TSS on the chromosome
\item
genomic strand: should be either + or -
\item
number of CAGE tags supporting that TSS
\end{itemize}
\item
\textbf{Publicly available CAGE datasets from R data package}: Several data packages containing CAGE data for various organisms produced by major consortia are accompanying this package. Selected subset of these data can be used as input for \Rpackage{CAGEr}.
\end{enumerate}

The type and the format of the input files is specified at the beginning of the workflow, when the \Rcode{CAGEset} object is created (section 4.2). This is done by setting the \Rcode{inputFilesType} argument, which accepts the following self-explanatory options referring to formats mentioned above: \Rcode{"bam", "bamPairedEnd", "bed", "ctss", "CTSStable"}.
\newline
In addition, the package provides a method for coercing a \Rcode{data.frame} object containing single base-pair TSS information into a \Rcode{CAGEset} object (as described in section 4.2.1), which can be further used in the workflow described below. 

%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Getting started}
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
To load the \Rpackage{CAGEr} package into your R envirnoment type:
<<>>=
library(CAGEr)
@

In this tutorial we will be using data from zebrafish (\emph{Danio rerio}) that was mapped to the danRer7 assembly of the genome. 
Therefore, the corresponding genome package \Rpackage{BSgenome.Drerio.UCSC.danRer7} has to be installed and available to
load by typing:
<<>>=
library(BSgenome.Drerio.UCSC.danRer7)
@

In case the data is mapped to a genome that is not readily available through \Rpackage{BSgenome} package (not in the list returned by \Rfunction{available.genomes()} function), 
a custom \Rpackage{BSgenome} package has to be build and installed first. (See the vignette within the \Rpackage{BSgenome} package for instructions on how to build 
a custom genome package). The built genome should then be loaded by calling \Rcode{library()} and the \Rcode{genomeName} argument should be set to the name of the build genome 
package when creating a \Robject{CAGEset} object (see the section \emph{Creating \Robject{CAGEset} object} below).


%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{\Rpackage{CAGEr} workflow}
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%

\subsection{Specifying input files}
The subset of zebrafish (\emph{Danio rerio}) developmental time-series CAGE data generated by (\cite{Nepal:2013})
will be used for the demonstration of the workflow and the provided functionalities. 
Files with genomic coordinates of TSSs detected by CAGE in 4 zebrafish developmental stages are included in this package 
in the \Rcode{extdata} subdirectory. The files contain TSSs from a part of chromosome 17 (26,000,000-46,000,000), and there 
are two files for one of the developmental stages (two independent replicas). The data in files is organized in four tab separated 
columns as described above in the section about  input data formats.
First we have to define paths to the input files:
<<>>=
inputDir <- system.file("extdata", package = "CAGEr")
pathsToInputFiles <- list.files(inputDir, full.names = TRUE)
basename(pathsToInputFiles)
@

\subsection{Creating a \Robject{CAGEset} object}
We start the workflow by creating a \Robject{CAGEset} object, which is a container 
for storing CAGE datasets and all the results that will be generated by applying specific functions. 
\Rcode{CAGEset} is created by providing name of the referent genome, paths to input files, type of 
input files and labels of individual CAGE datasets (samples):
<<>>=
myCAGEset <- new("CAGEset", genomeName = "BSgenome.Drerio.UCSC.danRer7",
		inputFiles = pathsToInputFiles, inputFilesType = "ctss",
		sampleLabels = c("zf_30p_dome", "zf_high", 
		"zf_prim6_rep1", "zf_prim6_rep2", "zf_unfertilized_egg"))
@

To display the created object type:
<<>>=
myCAGEset
@

The supplied information can be seen in the \Rcode{Input data information} section, whereas all other slots are still empty, since no data has been read yet and no analysis conducted.

\subsubsection{Coercing a \Rcode{data.frame} into a \Rcode{CAGEset} object}
In addition to creating an empty \Robject{CAGEset} object and reading in the TSS data from specified input files (as described in the following section), a \Rcode{CAGEset} object can be created directly by coercing a \Rcode{data.frame} containing single base-pair TSS information. To be able to do the coercion into a \Robject{CAGEset} the \Rcode{data.frame} must conform with the following:
\begin{itemize}
\item
the \Rcode{data.frame} must have at least 4 columns
\item
the first three columns must be named \Rcode{chr}, \Rcode{pos} and \Rcode{strand}, and should contain chromosome name, 1-based genomic coordinate of the TSS (positive integer) and TSS strand information (\Rcode{+} or \Rcode{-}), respectively
\item
these first three columns must be of the class \Rcode{character}, \Rcode{integer} and \Rcode{character}, respectively
\item
all additional columns must be of the class \Rcode{integer} and should contain raw CAGE tag counts (non-negative integer) supporting each TSS in different samples (columns). At least one such column with tag counts must be present.
\item
the names of the columns containing tag counts must begin with a letter, and these column names are used as sample labels in the resulting \Robject{CAGEset} object.
\end{itemize}

An example of such \Rcode{data.frame} is shown below:
<<>>=
TSS.df <- read.table(system.file("extdata/Zf.unfertilized.egg.chr17.ctss", 
                                package = "CAGEr"))
# make sure the column names are as required
colnames(TSS.df) <- c("chr", "pos", "strand", "zf_unfertilized_egg")
# make sure the column classes are as required
TSS.df$chr <- as.character(TSS.df$chr)
TSS.df$pos <- as.integer(TSS.df$pos)
TSS.df$strand <- as.character(TSS.df$strand)
TSS.df$zf_unfertilized_egg <- as.integer(TSS.df$zf_unfertilized_egg)
head(TSS.df)
@

This \Rcode{data.frame} can now be coerced to a \Robject{CAGEset} object, which will fill the corresponding slots of the object with provided TSS information:
<<>>=
myCAGEset.coerced <- as(TSS.df, "CAGEset")
myCAGEset.coerced
@

\subsection{Reading in the data}
In case when the CAGE / TSS data is to be read from input files, an empty \Robject{CAGEset} object with information about the files is first created as described above in the Section 4.2. To actually read in the data into the object we use the following function:
<<>>=
getCTSS(myCAGEset)
@
This function reads in the data from the provided files in the order they were specified in the \Rcode{inputFiles} argument. 
It creates a single set of all TSSs detected across all input datasets (union of TSSs) and a table with counts of CAGE tags supporting each TSS in 
every dataset. (Note that in case when a \Robject{CAGEset} object is created by coercion from a \Rcode{data.frame} there is no need to call the above function to read in the data, as the data will be loaded into \Rcode{CAGEset} during coercion). Genomic coordinates of all TSSs and numbers of supporting CAGE tags in every input sample can be retrieved by typing:
<<>>=
ctss <- CTSStagCount(myCAGEset)
head(ctss)
@
Note that the samples are ordered in the way they were supplied when creating the \Robject{CAGEset} object and will be presented in that 
order in all the results and plots. To check sample labels and their ordering type:
<<>>=
sampleLabels(myCAGEset)
@
In addition, a colour is assigned to each sample, which is consistently used to depict that sample in all the plots. By default a rainbow palette of colours is used 
and the hexadecimal format of the assigned colours can be seen as names attribute of sample labels shown above. The colours can be changed to taste at any point 
in the workflow using the \Rfunction{setColors} function.

\subsection{Correlation between samples}
After the data has been read we can start exploring the data by looking at the correlation between the samples. The  \Rfunction{plotCorrelation} 
function will plot pairwise scatter plots of CAGE tag count per TSS and calculate correlation between all possible pairs of samples. A tag count 
threshold can be set, so that only TSSs with tag count above the threshold (either in one or both samples) are considered when calculating 
correlation. Three different correlation measures are supported: Pearson's, Spearman's and Kendall's correlation coefficients.
<<eval=FALSE>>=
corr.m <- plotCorrelation(myCAGEset, samples = "all", method = "pearson")
@
The command above will create a PNG file with pairwise scatter plots and calculated correlation coefficients (Figure~\ref{fig:CorrelationScatterPlots}), 
and will return a matrix with correlation coefficients. Note that the correlation is calculated using raw tag counts on a linear scale, however, the 
scatter plots are plotted on a logarithmic scale for the convenience. Figure~\ref{fig:CorrelationScatterPlots} shows that early developmental stages 
correlate very well with each other, whereas their correlation with later prim6 stage is lower. There is also a very good correlation between the two 
replicas for prim6 developmental stage.

\begin{figure}[h!]
\centering
\includegraphics[width=110mm,height=109mm]{images/Pairwise_tag_count_correlation.jpg}
\caption{Correlation of raw CAGE tag counts per TSS}
\label{fig:CorrelationScatterPlots}
\end{figure}


Based on calculated correlation we might want to merge and/or rearrange some of the datasets. To rearrange the samples in the
temporal order of the zebrafish development (unfertilized egg -> high -> 30 percent dome -> prim6) and to merge the two replicas for the 
prim6 developmental stage we use the \Rfunction{mergeSamples} function:
<<>>=
mergeSamples(myCAGEset, mergeIndex = c(3,2,4,4,1), 
			mergedSampleLabels = c("zf_unfertilized_egg", 
			"zf_high", "zf_30p_dome", "zf_prim6"))
@
The \Rcode{mergeIndex} argument controls which samples will be merged and how the final dataset will be ordered. Samples labeled by the 
same number (in our case samples three and four) will be merged together by summing 
number of CAGE tags per TSS. The final set of samples will be ordered in the ascending order of values provided in \Rcode{mergeIndex} and 
will be labeled by the labels provided in the \Rcode{mergedSampleLabels} argument.
Note that \Rfunction{mergeSamples} function resets all slots with results of downstream analyses, so in case there were any results in the 
\Robject{CAGEset} object prior to merging, they will be removed.

\subsection{Normalization}
Library sizes (number of total sequenced tags) of individual experiments differ, thus normalization is required to make
them comparable. The \Rfunction{librarySizes} function returns the total number of CAGE tags in each sample:
<<>>=
librarySizes(myCAGEset)
@
\Rpackage{CAGEr} package supports both simple tags per million normalization and power-law based normalization. 
It has been shown that many CAGE datasets follow a power-law distribution (\cite{Balwierz:2009}). Plotting the number of CAGE tags (X-axis) 
against the number of TSSs that are supported by <= of that number of tags (Y-axis) results in a distribution that can be approximated by a power-law. 
On a log-log scale this reverse cumulative distribution will manifest as a monotonically decreasing linear function, which can be defined as
\newline 
\Rcode{y = -1 * alpha * x + beta}  
\newline
and is fully determined by the slope \Rcode{alpha} and total number of tags T (which together with \Rcode{alpha} determines the value of \Rcode{beta}).
\newline
To check whether our CAGE datasets follow power-law distribution and in which range of values, we can use the \Rfunction{plotReverseCumulatives} function:
<<eval=FALSE>>=
plotReverseCumulatives(myCAGEset, fitInRange = c(5, 1000), onePlot = TRUE)
@
This will create a PDF file with reverse cumulative plots (Figure~\ref{fig:ReverseCumulatives}) in your working directory. In addition, a power-law distribution will be fitted to each reverse cumulative 
using values in the specified range (denoted with dashed lines in Figure~\ref{fig:ReverseCumulatives}) and the value of alpha will be reported for each sample (shown in the brackets in 
the Figure~\ref{fig:ReverseCumulatives} legend). The plots can help in choosing the optimal parameters for power-law based normalization. We can see that the reverse cumulative 
distributions look similar and follow the power-law in the central part of the CAGE tag counts values with a slope between -1.1 and -1.3. Thus, we choose a range from 5 to 1000 tags 
to fit a power-law, and we normalize all samples to a referent power-law distribution with a total of 50,000 tags and slope of -1.2 (\Rcode{alpha = 1.2}). (Note that since this example dataset 
contains only data from one part of chromosome 17 and the total number of tags is very small, we normalize to a referent distribution with a similarly small number of tags. When analyzing 
full datasets it is reasonable to set total number of tags for referent distribution to one million to get normalized tags per million values.)

\begin{figure}[h!]
\centering
\includegraphics[width=90mm,height=90mm]{images/CTSS_reverse_cumulatives_all_samples.pdf}
\caption{Reverse cumulative distribution of CAGE tags}
\label{fig:ReverseCumulatives}
\end{figure}

To perform normalization we pass these parameters to the \Rfunction{normalizeTagCount} function:
<<>>=
normalizeTagCount(myCAGEset, method = "powerLaw", 
		fitInRange = c(5, 1000), alpha = 1.2, T = 5*10^4)
@
The normalization is performed as described in (\cite{Balwierz:2009}): 
\begin{enumerate}
\item
Power-law is fitted to the reverse cumulative distribution in the specified range of CAGE tags values to each sample separately.
\item
A referent power-law distribution is defined based on the provided \Rcode{alpha} (slope in the log-log representation) and \Rcode{T} (total number of tags) parameters. Setting \Rcode{T} to 1 million results in normalized tags per million (tpm) values.
\item
Every sample is normalized to the defined referent distribution, \emph{i.e.} given the parameters that approximate its own power-law distribution it is calculated how many tags would each TSS have in the referent power-law distribution.
\end{enumerate}

In addition to the two provided normalization methods, a pass-through option "none" can be set as \Rcode{method} parameter to keep using raw tag counts in all downstream steps. Note that \Rfunction{normalizeTagCount} function has to be applied to 
\Robject{CAGEset} object before moving to next steps. Thus, in order to keep using raw tag counts run the function with \Rcode{method="none"}. In that case, all results and parameters in the further steps that would normally refer to 
normalized CAGE signal (denoted as tpm), will actually be raw tag counts.

\subsection{Exporting CAGE signal to bedGraph}
CAGE data can be visualized in the genomic context by exporting raw or normalized CAGE tag counts to a bedGraph (or BigWig) file and uploading (or linking) the file 
to a genome browser. Positions of TSSs and tag counts supporting them are exported using \Rfunction{exportCTSStoBedGraph} function:
<<eval=FALSE>>=
exportCTSStoBedGraph(myCAGEset, values = "normalized", 
				format = "bedGraph", oneFile = TRUE)
@
This will produce a single bedGraph file with multiple annotated tracks that can be directly visualized as custom tracks in the genome browser (Figure~\ref{fig:CTSSbedGraph}).
There are two tracks per sample; one for TSSs on the plus strand and the other for the minus strand. Values for TSSs on minus strand are shown as negative and are pointing downwards in the browser.
\newline
Alternatively, the tracks can be exported to a binary BigWig format:
<<eval=FALSE>>=
exportCTSStoBedGraph(myCAGEset, values = "normalized", format = "BigWig")
@
which will produce two BigWig files per sample (one for TSSs on the plus strand and the other for the minus strand) and an accompanying text file with track headers.

\begin{figure}[h!]
\centering
\includegraphics[width=160mm]{images/CTSSbedGraph.pdf}
\caption{CAGE data bedGraph track visualized in the UCSC Genome Browser}
\label{fig:CTSSbedGraph}
\end{figure}

\subsection{CTSS clustering}
Transcription start sites are found in the promoter region of a gene and reflect the transcriptional activity of that promoter (Figure~\ref{fig:CTSSbedGraph}). 
TSSs in the close proximity of each other give rise to a functionally equivalent set of transcripts and are likely regulated by the same promoter elements. Thus, TSSs can 
be spatially clustered into larger transcriptional units, called tag clusters (TCs) that correspond to individual promoters. \Rpackage{CAGEr} supports three methods for spatial clustering of TSSs 
along the genome, two "ab initio" methods driven by the data itself, as well as assigning TSSs to predefined genomic regions:
\begin{enumerate}
\item
simple distance-based clustering in which two neighbouring TSSs are joined together if they are closer than some specified distance (greedy algorithm)
\item
parametric clustering of data attached to sequences based on the density of the signal (\cite{Frith:2007}, \href{http://www.cbrc.jp/paraclu/}{http://www.cbrc.jp/paraclu/})
\item
counting TSSs and their signal in a set of user supplied genomic regions (\emph{e.g.} annotation derived promoter regions or other regions of interest)
\end{enumerate}
These functionalities are provided in the \Rfunction{clusterCTSS} function, which accepts additional arguments for controlling which CTSSs will be included in the 
clustering as well as for refining the final set of tag clusters.
\newline
We will perform a simple distance-based clustering using 20bp as a maximal allowed distance between two neighbouring TSSs. Prior to 
clustering we will filter out low-fidelity TSSs - the ones supported by less than 2 normalized tag counts in all of the samples.
<<>>=
clusterCTSS(object = myCAGEset, threshold = 1, thresholdIsTpm = TRUE, 
		nrPassThreshold = 1, method = "distclu", maxDist = 20, 
		removeSingletons = TRUE, keepSingletonsAbove = 5)
@
Our final set of tag clusters will not include singletons (clusters with only one TSS), unless the normalized signal is above 5, \emph{i.e.} it is a reasonably supported TSS.
The \Rfunction{clusterCTSS} function creates a set of clusters for each sample separately; for each cluster it returns the genomic coordinates, counts the number of TSSs within 
the cluster, determines the position of the most frequently used (dominant) TSS, calculates the total CAGE signal within the cluster and CAGE signal supporting the dominant TSS only.
We can extract tag clusters for a desired sample from the \Robject{CAGEset} object by calling \Rfunction{tagClusters} function:
<<>>=
tc <- tagClusters(myCAGEset, sample = "zf_unfertilized_egg")
head(tc)
@

\subsection{Promoter width}
Genome-wide mapping of TSSs using CAGE has initially revealed two major classes of promoters in mammals (\cite{Carninci:2006}), with respect to the number and distribution of TSSs within the promoter. 
They have been further correlated with differences in the underlying sequence and the functional classes of the genes they regulate, as well as the organization of the chromatin around them.
These are:
 \begin{itemize}
 \item
 "broad" promoters with multiple TSSs characterized by a high GC content and overlap with a CpG island, which are associated with widely expressed or developmentally regulated genes
 \item
 "sharp" promoters with one dominant TSS often associated with a TATA-box at a fixed upstream distance, which often regulate tissue-specific transcription
 \end{itemize}
Thus, the width of the promoter is an important characteristic that distinguishes different functional classes of promoters. \Rpackage{CAGEr} package analyzes promoter width across all samples present 
in the \Robject{CAGEset} object. It defines promoter width by taking into account both the positions and the CAGE signal at TSSs along the tag cluster, thus making it more robust with respect 
to total expression and local level of noise at the promoter. Width of every tag cluster is calculated as following:
\begin{enumerate}
\item
Cumulative distribution of CAGE signal along the cluster is calculated.
\item
Positions of two selected quantiles are determined. At the 5' end the position of the "lower" quantile \Rcode{qLow} is determined, which is defined as the point that divides the cluster into two parts, 
such that the 5' part contains \Rcode{< qLow * 100\%} of the CAGE signal of that cluster. Accordingly, position of the "upper" quantile \Rcode{qUp} is determined near the 3' end, which is defined as the point that 
divides the cluster into two parts such that the 5' part contains \Rcode{>= qUp * 100\%} of the CAGE signal of that cluster.
\item
Promoter width is defined as the distance (in base pairs) between the two quantiles. This \textbf{interquantile width} marks the central part of the cluster that contains \Rcode{>= (qUp - qLow) * 100\%} of the CAGE signal.
\end{enumerate}
The procedure is schematically shown in Figure~\ref{fig:CumulativeDistribution}. 

\begin{figure}[h!]
\centering
\includegraphics[width=100mm]{images/CumulativeDistributionAndQuantiles.pdf}
\caption{Cumulative distribution of CAGE signal and definition of interquantile width}
\label{fig:CumulativeDistribution}
\end{figure}

Required computations are done using \Rfunction{cumulativeCTSSdistribution} and \Rfunction{quantilePositions} functions, which calculate cumulative distribution for 
every tag cluster in each of the samples and determine the positions of selected quantiles, respectively:
<<>>=
cumulativeCTSSdistribution(myCAGEset, clusters = "tagClusters")
quantilePositions(myCAGEset, clusters = "tagClusters", qLow = 0.1, qUp = 0.9)
@

Tag clusters and their interquantile width can be retrieved by calling \Rfunction{tagClusters} function:
<<>>=
tc <- tagClusters(myCAGEset, sample = "zf_unfertilized_egg",
		returnInterquantileWidth = TRUE,  qLow = 0.1, qUp = 0.9)
head(tc)
@

Interquantile width can also be visualized in a gene-like representation in the UCSC genome browser by exporting the data into a BED file:
<<eval=FALSE>>=
exportToBed(object = myCAGEset, what = "tagClusters", 
		qLow = 0.1, qUp = 0.9, oneFile = TRUE)
@
In this gene-like representation (Figure~\ref{fig:tagClustersBed}), the oriented line shows the full span of the cluster, filled block marks the interquantile width and a single base-pair thick 
block denotes the position of the dominant TSS.

\begin{figure}[h!]
\centering
\includegraphics[width=160mm]{images/TagClustersBed.pdf}
\caption{Tag clusters visualization in the genome browser}
\label{fig:tagClustersBed}
\end{figure}

Once the cumulative distributions and the positions of quantiles have been calculated, the histograms of interquantile width can be plotted to 
globally compare the promoter width across different samples (Figure~\ref{fig:TagClustersInterquantileWidth}):
<<eval=FALSE>>=
plotInterquantileWidth(myCAGEset, clusters = "tagClusters", 
		tpmThreshold = 3, qLow = 0.1, qUp = 0.9)
@
Significant difference in the promoter width might indicate global differences in the modes of gene regulation between the two samples.
The histograms can also help in choosing an appropriate width threshold for separating sharp and broad promoters.

\begin{figure}[h!]
\centering
\includegraphics[width=140mm]{images/TagClustersInterquantileWidth.pdf}
\caption{Distribution of promoter interquantile width}
\label{fig:TagClustersInterquantileWidth}
\end{figure}


\subsection{Creating consensus promoters across samples}
Tag clusters are created for each sample individually and they are often sample-specific, thus can be present in one sample but absent in another. In addition, 
in many cases tag clusters do not coincide perfectly within the same promoter region, or there might be two clusters in one sample and only one larger in the other. 
To be able to compare genome-wide transcriptional activity across samples and to perform expression profiling, a single set of consensus clusters needs to be created. 
This is done using \Rfunction{aggregateTagClusters} function, which aggregates tag clusters from all samples into a single set of non-overlapping consensus clusters:
<<>>=
aggregateTagClusters(myCAGEset, tpmThreshold = 5, 
		qLow = 0.1, qUp = 0.9, maxDist = 100)
@
Tag clusters can be aggregated using their full span (from start to end) or using positions of previously calculated quantiles as their boundaries. Only tag clusters above 
given tag count threshold will be considered and two clusters will be aggregated together if their boundaries (i.e. either starts and ends or positions of quantiles) are \Rcode{<= maxDist} apart.
Final set of consensus clusters can be retrieved by:
<<>>=
consensusCl <- consensusClusters(myCAGEset)
head(consensusCl)
@
which will return genomic coordinates and sum of CAGE signal across all samples for each consensus cluster (the \Rcode{tpm} column).
\newline
Analogously to tag clusters, analysis of promoter width can be performed for consensus clusters as well, using the same \Rfunction{cumulativeCTSSdistribution}, \Rfunction{quantilePositions} 
and \Rfunction{plotInterquantileWidth} functions as described above, but by setting 
the \Rcode{clusters} parameter to \Rcode{"consensusClusters"}.
\newline
Although consensus clusters are created to represent consensus across all samples, they obviously have different CAGE signal and can have different width or position of the dominant TSS in the different samples. Sample-specific information on consensus clusters can be retrieved with the \Rfunction{consensusClusters} function, by specifying desired sample name (analogous to retrieving tag clusters):
<<eval=FALSE>>=
consensusCl <- consensusClusters(myCAGEset, sample = "zf_unfertilized_egg",
		    returnInterquantileWidth = TRUE,  qLow = 0.1, qUp = 0.9)
@
This will, in addition to genomic coordinates of the consensus clusters, which are constant across all samples, also return the position of the dominant TSS, the CAGE signal (tpm) and the interquantile width specific for a given sample. Note that when specifying individual sample, only the consensus clusters that have some CAGE signal in that sample will be returned (which will be a subset of all consensus clusters).


\subsection{Expression profiling}
Since CAGE signal reflects level of transcription from a given TSS or promoter it can be used for 5' centered expression profiling.
Expression clustering can be done at level of individual CTSSs or at level of entire promoters (consensus clusteres). In the former case, 
feature vector containing log transformed and scaled normalized CAGE signal at individual TSS across multiple samples is used as input 
for clustering algorithm, whereas in the latter case CAGE signal within the entire consensus cluster is used. \Rpackage{CAGEr} package 
supports two unsupervised clustering algorithms: kmeans and self-organizing maps (SOM). Both algorithms require to specify number of clusters in advance.
\newline
We will perform expression clustering at the level of entire promoter using SOM algorithm and applying it only to promoters with normalized 
CAGE signal \emph{>= 15} in at least one sample.
<<>>=
getExpressionProfiles(myCAGEset, what = "consensusClusters", tpmThreshold = 10, 
		nrPassThreshold = 1, method = "som", xDim = 4, yDim = 2)
@
Distribution of expression across samples for 8 clusters returned by SOM (4 x 2 map) can be visualized using \Rfunction{plotExpressionProfiles} function 
as shown in Figure~\ref{fig:ConsensusClustersExpressionProfiles}:
<<eval=FALSE>>=
plotExpressionProfiles(myCAGEset, what = "consensusClusters")
@

\begin{figure}[h!]
\centering
\includegraphics[width=160mm]{images/ConsensusClustersExpressionProfiles.pdf}
\caption{Expression clusters}
\label{fig:ConsensusClustersExpressionProfiles}
\end{figure}

Each cluster is shown in different color and is marked by its label and the number of elements (promoters) in the cluster.
We can extract promoters belonging to a specific cluster by typing:
<<>>=
class3_1 <- extractExpressionClass(myCAGEset, 
		what = "consensusClusters", which = "3_1")
head(class3_1)
@

Consensus clusters and information on their expression profile can be exported to a BED file, which allows visualization of the promoters in the 
genome browser colored in the color of the expression cluster they belong to (Figure~\ref{fig:ConsensusClustersBed}):
<<eval=FALSE>>=
exportToBed(myCAGEset, what = "consensusClusters", 
		colorByExpressionProfile = TRUE)
@

\begin{figure}[h!]
\centering
\includegraphics[width=160mm]{images/ConsensusClustersBed.pdf}
\caption{Consensus clusters colored by expression profile in the genome browser}
\label{fig:ConsensusClustersBed}
\end{figure}

Expression profiling of individual TSSs is done using the same procedure as described above for consensus clusters, only by setting \Rcode{what} parameter 
to \Rcode{"CTSS"} in all of the functions.


\subsection{Shifting promoters}
As shown in Figure~\ref{fig:tagClustersBed}, TSSs within the same promoter region can be used differently in different samples. Thus, although the overall transcription level from a promoter 
does not change between the samples, the differential usage of TSSs or "promoter shifting" may indicate changes in the regulation of transcription from that promoter, which cannot be detected 
by expression profiling. To detect this promoter shifting, a method described in \cite{Haberle:2014} has been implemented in \Rpackage{CAGEr}. Shifting can be detected between two 
individual samples or between two groups of samples. In the latter case, samples are first merged into groups and then compared in the same way as two individual samples. For all promoters a 
shifting score is calculated based on the difference in the cumulative distribution of CAGE signal along that promoter in the two samples. In addition, a more general assessment of differential TSS 
usage is obtained by performing Kolmogorov-Smirnov test on the cumulative distributions of CAGE signal, as described below. Thus, prior to shifting score calculation and statistical testing, we have to 
calculate cumulative distribution along all consensus clusters:
<<>>=
cumulativeCTSSdistribution(myCAGEset, clusters = "consensusClusters")
@
Next, we calculate a shifting score and P-value of Kolmogorov-Smirnov test for all promoters comparing two specified samples:
<<>>=
scoreShift(myCAGEset, groupX = "zf_unfertilized_egg", groupY = "zf_prim6",
		testKS = TRUE, useTpmKS = FALSE)
@
This function will calculate shifting score as illustrated in Figure~\ref{fig:ShiftingScore}. Values of shifting score are in range between -Inf and 1. Positive values can be interpreted as the proportion of transcription initiation 
in the sample with lower expression that is happening "outside" (either upstream or downstream) of the region used for transcription initiation in the other sample. In contrast, negative values 
indicate no physical separation, \emph{i.e.} the region used for transcription initiation in the sample with lower expression is completely contained within the region used for transcription 
initiation in the other sample. Thus, shifting score detects only the degree of upstream or downstream shifting, but does not detect more general changes in TSS rearrangement in the region, 
\emph{e.g.} narrowing or broadening of the region used for transcription.
\newline
To assess any general change in the TSS usage within the promoter region, a two-sample Kolmogorov-Smirnov (K-S) test on cumulative 
sums of CAGE signal along the consensus cluster is performed. Cumulative sums in both samples are scaled to range between 0 and 1 and are considered to be empirical cumulative distribution functions (ECDF) 
reflecting sampling of TSS positions during transcription initiation. K-S test is performed to assess whether the two underlying probability distributions differ. To obtain a P-value 
(\emph{i.e.} the level at which the null-hypothesis can be rejected), sample sizes that generated the ECDFs are required, in addition to actual K-S statistics calculated from ECDFs. 
These are derived either from raw tag counts, \emph{i.e.} exact number of times each TSS in the cluster was sampled during sequencing (when \Rcode{useTpmKS = FALSE}), or from normalized 
tpm values (when \Rcode{useTpmKS = TRUE}). P-values obtained from K-S tests are further corrected for multiple testing using Benjamini and Hochenberg (BH) method and for each P-value a corresponding false-discovery rate (FDR) is also reported.

\begin{figure}[h!]
\centering
\includegraphics[width=130mm]{images/ShiftingScore.pdf}
\caption{Calculation of shifting score}
\label{fig:ShiftingScore}
\end{figure}


We can select a subset of promoters with shifting score and/or FDR above specified threshold:
<<>>=
shifting.promoters <- getShiftingPromoters(myCAGEset, 
		tpmThreshold = 5, scoreThreshold = 0.6,
		fdrThreshold = 0.01)
head(shifting.promoters)
@
The \Rfunction{getShiftingPromoters} function returns genomic coordinates, shifting score and P-value (FDR) of the promoters, as well as the value of CAGE signal and position of the dominant TSS in the two 
compared (groups of) samples. Figure~\ref{fig:ShiftingPromoter} shows the difference in the CAGE signal between the two compared samples for one of the selected high-scoring shifting promoters.

\begin{figure}[h!]
\centering
\includegraphics[width=160mm]{images/ShiftingPromoter.pdf}
\caption{Example of shifting promoter}
\label{fig:ShiftingPromoter}
\end{figure}

\newpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Accessing public CAGE datasets}
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\subsection{Available CAGE data resources}
There are several large collections of CAGE data available that provide single base-pair resolution TSSs for numerous human and mouse primary cells, cell lines and tissues. Together with several minor datasets for other model organisms (\emph{Drosophila melanogaster}, \emph{Danio rerio}) they are a valuable resource that provides cell/tissue type and developmental stage specific TSSs essential for any type of promoter centred analysis. By enabling direct and user-friendly import of TSSs for selected samples into R, \emph{CAGEr} facilitates the integration of these precise TSS data with other genomic data types.
Each of the available CAGE data resources accessible from within \emph{CAGEr} is explained in more detail further below.
\subsubsection{FANTOM5}
FANTOM consortium provides single base-pair resolution TSS data for numerous human and mouse primary cells, cell lines and tissues. The main FANTOM5 publication (\cite{Consortium:2014hz}) released \textasciitilde1000 human and \textasciitilde400 mouse CAGE samples that cover the vast majority of human primary cell types, mouse developmental tissues and a large number of commonly used cell lines. These data is available from FANTOM web resource at \href{http://fantom.gsc.riken.jp/5/data/}{http://fantom.gsc.riken.jp/5/data/} in the form of TSS files with genomic coordinates and number of tags mapping to each TSS detected by CAGE. The list of all available samples for both human and mouse (as presented in the Supplementary Table 1 of the publication) has been included in \emph{CAGEr} to facilitate browsing, searching and selecting samples of interest. TSSs for selected samples are then fetched directly form the web resource and imported into a \Rcode{CAGEset} object enabling their further manipulation with \emph{CAGEr}.
\subsubsection{FANTOM3 and 4}
Previous FANTOM projects (3 and 4) (\cite{Consortium:2005kp,Faulkner:2009fw,Suzuki:2009gy}) produced CAGE datasets for multiple human and mouse tissues as well as several timecourses, including differentiation of a THP-1 human myeloid leukemia cell line. All this TSS data has been grouped into datasets by the organism and tissue of origin and has been collected into an R data package named \emph{FANTOM3and4CAGE}, which is available from Bioconductor (\href{http://www.bioconductor.org/packages/release/data/experiment/html/FANTOM3and4CAGE.html}{http://www.bioconductor.org/packages/release/data/experiment/html/FANTOM3and4CAGE.html}). The vignette accompanying the package provides information on available datasets and lists of samples. When the data package is installed, \emph{CAGEr} can import the TSSs for selected samples directly into a \Rcode{CAGEset} object for further manipulation.
\subsubsection{ENCODE cell lines}
ENCODE consortium produced CAGE data for common human cell lines (\cite{Djebali:2012hc}), which were used by ENCODE for various other types of genome-wide analyses. The advantage of this dataset is that it enables the integration of precise TSSs from a specific cell line with many other genome-wide data types provided by ENCODE for the same cell line. However, the format of CAGE data provided by ENCODE at UCSC (\href{http://genome.ucsc.edu/ENCODE/dataMatrix/encodeDataMatrixHuman.html}{http://genome.ucsc.edu/ENCODE/dataMatrix/encodeDataMatrixHuman.html}) includes only raw mapped CAGE tags and their coverage along the genome, and coordinates of enriched genomic regions (peaks), which do not take advantage of the single base-pair resolution provided by CAGE. To address this, we have used the raw CAGE tags to derive single base-pair resolution TSSs and collected them into an R data package named \emph{ENCODEprojectCAGE}. This data package is available for download from CAGEr web site at \href{http://promshift.genereg.net/CAGEr}{http://promshift.genereg.net/CAGEr} and includes TSSs for 36 different cell lines fractionated by cellular compartment. The vignette accompanying the package provides information on available datasets and lists of individual samples. Once the package has been downloaded and installed, \emph{CAGEr} can access it to import TSS data for selected subset of samples for further manipulation and integration.
\subsubsection{Zebrafish developmental timecourse}
Precise TSSs are also available for zebrafish (\emph{Danio Rerio}) from CAGE data published by Nepal \emph{et al.} (\cite{Nepal:2013}). The timecourse covering early embryonic development of zebrafish includes 12 developmental stages. The TSS data has been collected into an R data package named \emph{ZebrafishDevelopmentalCAGE}, which is available for download from CAGEr web site at \href{http://promshift.genereg.net/CAGEr}{http://promshift.genereg.net/CAGEr}. As with other data packages mentioned above, once the package is installed \emph{CAGEr} can use it to import stage-specific single base pair TSSs into a \Rcode{CAGEset} object.

\subsection{Importing public TSS data for manipulation in \emph{CAGEr}}
The data from above mentioned resources can be imported into a \Rcode{CAGEset} object using the \Rcode{importPublicData()} function.
The \Rcode{importPublicData()} function has four arguments: \Rcode{source}, \Rcode{dataset}, \Rcode{group} and \Rcode{sample}. Argument \Rcode{source} accepts one of the following values: \Rcode{"FANTOM5"}, \Rcode{"FANTOM3and4"}, \Rcode{"ENCODE"}, or \Rcode{"ZebrafishDevelopment"}, which refer to one of the four resources listed above.
The following sections explain how to utilize this function for each of the four currently supported resources.

\subsubsection{FANTOM5 human and mouse samples}
Lists of all human and mouse CAGE samples produced within FANTOM5 project are available in \emph{CAGEr}. To load the information on human samples type:
<<>>=
data(FANTOM5humanSamples)
head(FANTOM5humanSamples)
nrow(FANTOM5humanSamples)
@
There are 988 human samples in total and for each the following information is provided:

\begin{itemize}
\item
\Rcode{sample}: a unique name/label of the sample which should be provided to \Rcode{importPublicData()} function to retrieve given sample
\item
\Rcode{type}: type of sample, which can be "cell line", "primary cell" or "tissue"
\item
\Rcode{description}: short description of the sample as provided in FANTOM5 main publication (\cite{Consortium:2014hz})
\item
\Rcode{library\_id}: unique ID of the CAGE library within FANTOM5
\item
\Rcode{data\_url}: URL to corresponding CTSS file at FANTOM5 web resource from which the data is fetched
\end{itemize}

Provided information facilitates searching for samples of interest, \emph{e.g.} we can search for astrocyte samples:
<<>>=
astrocyteSamples <- FANTOM5humanSamples[grep("Astrocyte", 
				FANTOM5humanSamples[,"description"]),]
astrocyteSamples
@
Analogous information is provided for mouse samples, which can be loaded by typing:
<<>>=
data(FANTOM5mouseSamples)
head(FANTOM5mouseSamples)
nrow(FANTOM5mouseSamples)
@

To import TSS data for samples of interest from FANTOM5 we use the \Rcode{importPublicData()} function and set the argument \Rcode{source = "FANTOM5"}. The \Rcode{dataset} argument can be set to either \Rcode{"human"} or \Rcode{"mouse"}, and the \Rcode{sample} argument is provided by a vector of sample lables/names. For example, names of astrocyte samples from above are:
<<>>=
astrocyteSamples[,"sample"]
@
and to import first three samples type:
<<>>=
astrocyteCAGEset <- importPublicData(source = "FANTOM5", dataset = "human", 
					sample = astrocyteSamples[1:3,"sample"])
astrocyteCAGEset
@

The resulting \Rcode{astrocyteCAGEset} is a \Rcode{CAGEset} object that can be included in the \emph{CAGEr} workflow described above to perform normalisation, clustering, visualisation, etc.

\subsubsection{\emph{FANTOM3and4CAGE} data package}

To use TSS data from FANTOM3 and FANTOM4 projects, a data package \emph{FANTOM3and4CAGE} has to be installed and loaded.  This package is available from Bioconductor and can be installed by calling:
<<eval=FALSE>>=
source("http://bioconductor.org/biocLite.R")
biocLite("FANTOM3and4CAGE")
@
For the list of available datasets with group and sample labels for specific human or mouse samples load the data package and get list of samples:
<<>>=
library(FANTOM3and4CAGE)

data(FANTOMhumanSamples)
head(FANTOMhumanSamples)
data(FANTOMmouseSamples)
head(FANTOMmouseSamples)
@

In the above data frames, the columns \Rcode{dataset}, \Rcode{group} and \Rcode{sample} provide values that should be passed to corresponding arguments in \Rcode{importPublicData()} function. For example to import human kidney normal and malignancy samples call:
<<>>=
kidneyCAGEset <- importPublicData(source = "FANTOM3and4",
			dataset = "FANTOMtissueCAGEhuman", 
			group = "kidney", sample = c("kidney", "malignancy"))
kidneyCAGEset
@
When the samples belong to different groups or different datasets, it is necessary to provide the dataset and group assignment for each sample separately:
<<>>=
mixedCAGEset <- importPublicData(source = "FANTOM3and4",
		dataset = c("FANTOMtissueCAGEmouse", "FANTOMtissueCAGEmouse", 
		"FANTOMtimecourseCAGEmouse"), group = c("liver", "liver", 
		"liver_under_constant_darkness"), 
		sample = c("cloned_mouse", "control_mouse", "4_hr"))
mixedCAGEset
@

For more details about datasets available in the \emph{FANTOM3and4CAGE} data package please refer to the vignette accompanying the package.

\subsubsection{\emph{ENCODEprojectCAGE} data package}

TSS data derived from ENCODE CAGE datasets has been collected into \emph{ENCODEprojectCAGE} data package, which is available for download from the \emph{CAGEr} web site (\href{http://promshift.genereg.net/CAGEr/}{http://promshift.genereg.net/CAGEr/}). Downloaded package can be installed from local using \Rcode{install.packages()} function from within R and used with \emph{CAGEr} as described below.
List of datasets available in this data package can be obtained like this:
<<eval=FALSE>>=
library(ENCODEprojectCAGE)

data(ENCODEhumanCellLinesSamples)
@

The information provided in this data frame is analogous to the one in previously discussed data package and provides values to be used with \Rcode{importPublicData()} function. The command to import whole cell CAGE samples for three different cell lines would look like this:
<<eval=FALSE>>=
ENCODEset <- importPublicData(source = "ENCODE", 
	dataset = c("A549", "H1-hESC", "IMR90"), 
	group = c("cell", "cell", "cell"), sample = c("A549_cell_rep1", 
	"H1-hESC_cell_rep1", "IMR90_cell_rep1"))
@

For more details about datasets available in the \emph{ENCODEprojectCAGE} data package please refer to the vignette accompanying the package.

\subsubsection{\emph{ZebrafishDevelopmentalCAGE} data package}

The zebrafish TSS data for 12 developmental stages is collected in \emph{ZebrafishDevelopmentalCAGE} data package, which is also available for download from the \emph{CAGEr} web site (\href{http://promshift.genereg.net/CAGEr/}{http://promshift.genereg.net/CAGEr/}). It can be installed from local using \Rcode{install.packages()} function.
To get a list of samples within the package type:
<<eval=FALSE>>=
library(ZebrafishDevelopmentalCAGE)

data(ZebrafishSamples)
@
In this package there is only one dataset called \Rcode{ZebrafishCAGE} and all samples belong to the same group 
called \Rcode{development}. To import selected samples from this dataset type:
<<eval=FALSE>>=
zebrafishCAGEset <- importPublicData(source = "ZebrafishDevelopment", 
			dataset = "ZebrafishCAGE", group = "development", 
			sample = c("zf_64cells", "zf_prim6"))
@

For more details please refer to the vignette accompanying the data package.
\newline
\newline
Importing TSS data from any of the four above explained resources results in the \Rcode{CAGEset} object that can be directly included into the workflow provided by \emph{CAGEr} to perform normalisation, clustering, promoter width analysis, visualisation, \emph{etc.} This high-resolution TSS data can then easily be integrated with other genomic data types to perform various promoter-centred analyses, which does not rely on annotation but uses precise and matched cell/tissue type TSSs.


%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Session Info}
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
<<>>=
sessionInfo()
@

%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%\newpage
\bibliographystyle{apalike}
\bibliography{CAGEr}
\end{document}
