%\VignetteEngine{knitr::knitr}
%\VignetteIndexEntry{Base Functions and Classes for MS-based Proteomics}
%\VignetteKeywords{Mass Spectrometry, MS, MSMS, Proteomics, Infrastructure, Bioinformatics, quantitative }
%\VignettePackage{MSnbase-demo}

\documentclass[12pt, oneside]{article}

<<style, eval=TRUE, echo=FALSE, results="asis">>=
BiocStyle::latex()
@

\bioctitle[\Biocpkg{MSnbase} introduction]{
  \Biocpkg{MSnbase}: labelled and label-free MS2 data pre-processing,
  visualisation and quantification.
}

\author{
  Laurent Gatto\footnote{\email{lg390@cam.ac.uk}} and Sebastian Gibb
}


\begin{document}

\maketitle

%% Abstract and keywords %%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\vskip 0.3in minus 0.1in
\hrule
\begin{abstract}
  This vignette describes the functionality implemented in the
  \Biocpkg{MSnbase} package.  \Biocpkg{MSnbase} aims at (1)
  facilitating the import, processing, visualisation and
  quantification of mass spectrometry data into the \R environment
  \cite{Rstat} by providing specific data classes and methods and (2)
  enabling the utilisation of throughput-high data analysis pipelines
  provided by the Bioconductor \cite{Gentleman2004} project.
\end{abstract}
\textit{Keywords}: Mass Spectrometry (MS), proteomics, infrastructure, quantitative.
\vskip 0.1in minus 0.05in
\hrule
\vskip 0.2in minus 0.1in
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%

\newpage

\tableofcontents

<<environment, cache=FALSE, echo=FALSE>>=
suppressPackageStartupMessages(library("ggplot2"))
suppressPackageStartupMessages(library("MSnbase"))
suppressPackageStartupMessages(library("zoo"))
suppressPackageStartupMessages(require("Rdisop"))
suppressPackageStartupMessages(require("pRolocdata"))
suppressPackageStartupMessages(require("pRoloc"))
library("grid")
library("reshape2")
@

\newpage

%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\input{Foreword.tex}

\input{Bugs.tex}

\newpage

%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Introduction}\label{sec:intro}

\Biocpkg{MSnbase} \cite{Gatto2012} aims are providing a reproducible
research framework to proteomics data analysis.  It should allow
researcher to easily mine mass spectrometry data, explore the data and
its statistical properties and visually display these.

\Biocpkg{MSnbase} also aims at being compatible with the
infrastructure implemented in Bioconductor, in particular
\Biocpkg{Biobase}. As such, classes developed specifically for
proteomics mass spectrometry data are based on the \Rclass{eSet} and
\Rclass{ExpressionSet} classes. The main goal is to assure seamless
compatibility with existing meta data structure, accessor methods and
normalisation techniques.

This vignette illustrates \Biocpkg{MSnbase} utility using a dummy
data sets provided with the package without describing the underlying
data structures. More details can be found in the package, classes,
method and function documentations. A description of the classes is
provided in the \texttt{MSnbase-development} vignette.

\input{NoteAboutSpeedAndMemory.tex}

%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Data structure and content}\label{sec:data}

\subsection{Importing experiments}\label{sec:io}

\Biocpkg{MSnbase} is able to import raw MS data stored in one of the
\texttt{XML}-based formats as well as peak lists in the \texttt{mfg}
format\footnote{%%
  Mascot Generic Format --
  \url{http://www.matrixscience.com/help/data\_file\_help.html\#GEN}}

\paragraph{Raw data} The \texttt{XML}-based formats, \texttt{mzXML}
\cite{Pedrioli2004}, \texttt{mzData} \cite{Orchard2007} and
\texttt{mzML} \cite{Martens2010} can be imported with the
\Rfunction{readMSData} function, as illustrated below (see
\Rfunction{?readMSData} for more details).


<<readdata, echo=TRUE, cache=FALSE, tidy=FALSE>>=
file <- dir(system.file(package = "MSnbase", dir = "extdata"),
            full.names = TRUE, pattern = "mzXML$")
rawdata <- readMSData(file, msLevel = 2, verbose = FALSE)
@
%% $

Only spectra of a give MS level can be loaded at a time by setting the
\texttt{msLevel} parameter accordingly.  In this document, we will use
the \Robject{itraqdata} data set, provided with \Biocpkg{MSnbase}. It
includes feature metadata, accessible with the \Rfunction{fData}
accessor.  The metadata includes identification data for the
\Sexpr{length(itraqdata)} MS2 spectra.

\paragraph{MSnbase 2.0 } The new major version of \Biocpkg{MSnbase}
uses a new \textit{on-disk} data storage model (see the
\textit{benchmarking} vignette for more details). The new data backend
is compatible with the orignal \textit{in-memory} model. To make use
of the new infrastructure, read your raw data using
\Rfunction{readMSData2}, rather than \Rfunction{readMSData}. All
existing operations work irrespective of the backend.

\paragraph{Peak lists} Peak lists can often be exported after spectrum
processing from vendor-specific software and are also used as input to
search engines.  Peak lists in \texttt{mgf} format can be imported
with the function \Rfunction{readMgfData} (see
\Rfunction{?readMgfData} for details) to create experiment objects.
Experiments or individual spectra can be exported to an \texttt{mgf}
file with the \Rfunction{writeMgfData} methods (see
\Rfunction{?writeMgfData} for details and examples).

\bigskip

\paragraph{Experiments with multiple runs} Although it is possible to
load and process multiple files serially and later merge the resulting
quantitation data as show in section \ref{sec:combine} (page
\pageref{sec:combine}), it is also feasible to load several raw data
files at once.  Here, we report the analysis of an LC-MSMS experiment
were 14 liquid chromatography (LC) fractions were loaded using
\Rfunction{readMSData} on a 32-cores servers with 128 Gb of RAM.  It
took about 90 minutes to read the 14 uncentroided \texttt{mzXML} raw
files (4.9 Gb on disk in total) and create a 3.3 Gb raw data object
(an \Rclass{MSnExp} instance, see next section).  Quantitation of 9
reporter ions (\Rclass{iTRAQ9} object, see \ref{sec:reporterions})
for 88690 features was performed in parallel on 16 processors and took
76 minutes.  The resulting quantitation data was only 22.1 Mb and
could easily be further processed and analysed on a standard laptop
computer.

Since verions 1.13.5, parallel support is provided by the
\Biocpkg{BiocParallel} and various backends including multicore
(forking), simple networf network of workstations (SNOW) using
sockets, forking or MPI among others.

\bigskip

See also section \ref{sec:io2} to import quantitative data stored in
spreadsheets into \R for further processing using
\Biocpkg{MSnbase}. The \texttt{MSnbase-io} vignette gives a general
overview of \Biocpkg{MSnbase}'s input/ouput capabilites.

\subsection{MS experiments}\label{sec:msnexp}

Raw data is contained in \Rclass{MSnExp} objects, that stores all the
spectra of an experiment, as defined by one or multiple raw data
files.

<<MSnExp, cache=FALSE, echo=TRUE>>=
library("MSnbase")
itraqdata
head(fData(itraqdata))
@

<<experimentsize, echo=FALSE, cache=FALSE>>=
sz <- sum(sapply(assayData(itraqdata), object.size)) +
  object.size(itraqdata)
sz <- as.numeric(sz)
sz <- round(sz/(1024^2), 2)
@

As illustrated above, showing the experiment textually displays it's
content:
\begin{itemize}
\item Information about the raw data, i.e. the spectra.
\item Specific information about the experiment processing\footnote{%%
    this part will be automatically updated when the object is
    modified with it's \textit{ad hoc} methods, as illustrated later}
  and package version.  This slot can be accessed with the
  \Rfunction{processingData} method.
\item Other meta data, including experimental phenotype, file name(s)
  used to import the data, protocol data, information about features
  (individual spectra here) and experiment data. Most of these are
  implemented as in the \Rclass{eSet} class and are described in more
  details in their respective manual pages. See \Rfunction{?MSnExp}
  and references therein for additional background information.

  The experiment meta data associated with an \Rclass{MSnExp}
  experiment is of class \Rclass{MIAPE}. It stores general
  information about the experiment as well as MIAPE (Minimum
  Information About a Proteomics Experiment) information
  \cite{Taylor2007, Taylor2008}.  This meta-data can be accessed with
  the \Rfunction{experimentData} method.  When available, a summary of
  MIAPE-MS data can be printed with the \Rfunction{msInfo} method.
  See \Rfunction{?MIAPE} for more details.
\end{itemize}

\subsection{Spectra objects}\label{sec:spectra}

The raw data is composed of the \Sexpr{length(itraqdata)} MS spectra.
The spectra are named individually
(\Sexpr{paste(paste(head(featureNames(itraqdata)),collapse=", "),",
  ...",sep="")}) and stored in a \Robject{environment}. They can be
accessed individually with \texttt{itraqdata[["X1"]]} or
\texttt{itraqdata[[1]]}, or as a list with
\texttt{spectra(itraqdata)}.  As we have loaded our experiment
specifying \texttt{msLevel=2}, the spectra will all be of level 2 (or
higher, if available).

<<Spectrum, cache=FALSE, echo=TRUE>>=
sp <- itraqdata[["X1"]]
sp
@

Attributes of individual spectra or of all spectra of an experiment
can be accessed with their respective methods:
\Rfunction{precursorCharge} for the precursor charge,
\Rfunction{rtime} for the retention time, \Rfunction{mz} for the MZ
values, \Rfunction{intensity} for the intensities, ... see the
\Rclass{Spectrum}, \Rclass{Spectrum1} and \Rclass{Spectrum2}
manuals for more details.

<<accessors, cache=FALSE, echo=TRUE>>=
peaksCount(sp)
head(peaksCount(itraqdata))
rtime(sp)
head(rtime(itraqdata))
@

\subsection{Reporter ions}\label{sec:reporterions}

Reporter ions are defined with the \Rclass{ReporterIons} class.
Specific peaks of interest are defined by a MZ value, a with around
the expected MZ and a name (and optionally a colour for plotting, see
section \ref{sec:plotting}). \Rclass{ReporterIons} instances are
required to quantify reporter peaks in \Rclass{MSnExp}
experiments. Instances for the most commonly used isobaric tags like
iTRAQ 4-plex and 8-plex and TMT 6- and 10-plex tags are already
defined in \Biocpkg{MSnbase}. See \Rfunction{?ReporterIons} for
details about how to generate new \Rclass{ReporterIons} objects.

<<ReporterIons>>=
iTRAQ4
TMT10
@

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Plotting raw data}\label{sec:plotting}

\subsection{MS data space}\label{sec:msmaps}

The \Rclass{MSmap} class can be used to isolate specific slices of
interest from a complete MS acquisition by specifying $m/z$ and
retention time ranges. One needs a raw data file in a format supported
by \Biocpkg{mzR}'s \Rfunction{openMSfile} (\texttt{mzML},
\texttt{mzXML}, ...). Below we first download a raw data file from the
PRIDE repository and create\footnote{This code chunk is not evaluated
  to avoid repeated downloaded of the raw data file. The \Robject{M}
  map is provided with the package and loaded to evaluate subsequent
  code chunks.} an \Rclass{MSmap} containing all the MS$^1$ spectra
between acquired between 30 and 35 minutes and peaks between 521 and
523 $m/z$. See \Rfunction{?MSmap} for details.

<<msmap, eval=FALSE>>=
## downloads the data
library("rpx")
px1 <- PXDataset("PXD000001")
mzf <- pxget(px1, 6)
     
## reads the data 
ms <- openMSfile(mzf)
hd <- header(ms)
     
## a set of spectra of interest: MS1 spectra eluted
## between 30 and 35 minutes retention time
ms1 <- which(hd$msLevel == 1)
rtsel <- hd$retentionTime[ms1] / 60 > 30 &
    hd$retentionTime[ms1] / 60 < 35
     
## the map
M <- MSmap(ms, ms1[rtsel], 521, 523, .005, hd, zeroIsNA = TRUE)
@

<<msmaplaod, echo=FALSE>>=
mrda <- dir(system.file(package = "MSnbase", dir = "extdata"),
            full.names = TRUE, pattern = "M.rda$")
mrda2 <- dir(system.file(package = "MSnbase", dir = "extdata"),
            full.names = TRUE, pattern = "M2.rda$")
load(mrda)
load(mrda2)
@

<<msmaphow>>=
M
@

The \Robject{M} map object can be rendered as a heatmap with
\Rfunction{plot}, as shown on figure \ref{fig:mapheat}.

\begin{figure}[!ht]
\centering
<<mapheat, dev='pdf', fig.width=5, fig.height=5>>=
plot(M, aspect = 1, allTicks = FALSE)
@
\caption{Heat map of a chunk of the MS data.}
\label{fig:mapheat}
\end{figure}

One can also render the data in 3 dimension with the
\Rfunction{plot3D} function, as show on figure \ref{fig:map3d}.

\begin{figure}[!ht]
\centering
<<map3d1, dev='pdf', fig.width=4, fig.height=4>>=
plot3D(M)
@
\caption{3 dimensional represention of MS map data.}
\label{fig:map3d}
\end{figure}

To produce figure \ref{fig:map3d2}, we create a second \Rclass{MSmap}
object containing the first two MS$^1$ spectra of the first map
(object \Robject{M} above) and all intermediate MS$^2$ spectra and
display $m/z$ values between 100 and 1000.

<<msmap2, eval=FALSE>>=
i <- ms1[which(rtsel)][1]
j <- ms1[which(rtsel)][2]
M2 <- MSmap(ms, i:j, 100, 1000, 1, hd)
@

<<m2>>=
M2
@

\begin{figure}[!ht]
\centering
<<map3d2, dev='pdf', fig.width=4, fig.height=4>>=
plot3D(M2)
@
\caption{3 dimensional represention of MS map data. MS$^1$ and MS$^2$
  spectra are coloured in blue and magenta respectively.}
\label{fig:map3d2}
\end{figure}


\subsection{MS Spectra}\label{sec:specplots}

Spectra can be plotted individually or as part of (subset) experiments
with the \Rfunction{plot} method. Full spectra can be plotted (using
\texttt{full=TRUE}), specific reporter ions of interest (by specifying
with reporters with \texttt{reporters=iTRAQ4} for instance) or both
(see figure \ref{fig:spectrum-plot}).

\begin{figure}[!ht]
\centering
<<spectrumPlot, dev='pdf', fig.width=6, fig.height=4, fig.keep='high'>>=
plot(sp, reporters = iTRAQ4, full = TRUE)
@
\caption{Raw MS2 spectrum with details about reporter ions.}
\label{fig:spectrum-plot}
\end{figure}


<<bsaSelect, eval=TRUE, echo=FALSE>>=
## bsasel <- fData(itraqdata)$ProteinAccession == "BSA"
bsasel <- 1:3
@
%% $

It is also possible to plot all spectra of an experiment (figure
\ref{fig:msnexp-plot}).  Lets start by subsetting the
\Robject{itraqdata} experiment using the protein accession numbers
included in the feature metadata, and keep the \Sexpr{sum(bsasel)}
from the \textit{BSA} protein.

<<subset, echo=TRUE>>=
sel <- fData(itraqdata)$ProteinAccession == "BSA"
bsa <- itraqdata[sel]
bsa
as.character(fData(bsa)$ProteinAccession)
@

%% Lets start by extracting all spectra that have the same precursor MZ value than \Robject{sp}
%% into a separate experiment, using the \Rfunction{extractPrecSpectra} method.
%% This method takes an \Robject{MSnExp} experiment and a precursor MZ value as parameters.
%% <<extractPrecSpec,echo=T,cache=F>>=
%% exp2 <- extractPrecSpectra(raw.experiment,precursorMz(sp))
%% exp2
%% @

These can then be visualised together by plotting the
\Rclass{MSnExp} object, as illustrated on figure \ref{fig:msnexp-plot}.

\begin{figure}[p]
\centering
<<msnexpPlot, dev='pdf', fig.width=3.7, fig.height=4.5, fig.keep='last'>>=
plot(bsa, reporters = iTRAQ4, full = FALSE) + theme_gray(8)
@
\caption{Experiment-wide raw MS2 spectra. The y-axes of the individual
  spectra are automatically rescaled to the same range.
  See section \ref{sec:norm} to rescale peaks identically. }
\label{fig:msnexp-plot}
\end{figure}

\paragraph{Customising your plots}\label{sec:customplots}

The \Biocpkg{MSnbase} \Rfunction{plot} methods have a logical
\Rfunction{plot} parameter (default is \Robject{TRUE}), that specifies
if the plot should be printed to the current device. A plot object is
also (invisibly) returned, so that it can be saved as a variable for
later use or for customisation.

\Biocpkg{MSnbase} uses the \CRANpkg{ggplot2} package to generate
plots, which can subsequently easily be customised.  More details
about \CRANpkg{ggplot2} can be found in \cite{ggplot2} (especially
chapter 8) and on
\href{http://had.co.nz/ggplot2/}{http://had.co.nz/ggplot2/}.  Finally,
if a plot object has been saved in a variable \Robject{p}, it is
possible to obtain a summary of the object with
\Rfunction{summary(p)}.  To view the data frame used to generate the
plot, use \Rfunction{p@data}.

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Tandem MS identification data}\label{sec:id}

\subsection{Adding identification data}

\Biocpkg{MSnbase} is able to integrate identification data from
\texttt{mzIdentML} \cite{Jones2012} files.

We first load two example files shipped with the \Biocpkg{MSnbase}
containing raw data (as above) and the corresponding identification
results respectively. The raw data is read with the
\Rfunction{readMSData}, as demonstrated above. As can be seen, the
default feature data only contain spectra numbers\footnote{More data
  about the spectra is of course available in an \Rclass{MSnExp}
  object, as illustrated in the previous sections. See also
  \Rfunction{?pSet} and \Rfunction{?MSnExp} for more details.}.

<<msnexpIdentification, echo=TRUE, cache=FALSE, tidy=FALSE>>=
## find path to a mzXML file
quantFile <- dir(system.file(package = "MSnbase", dir = "extdata"),
                 full.name = TRUE, pattern = "mzXML$")
## find path to a mzIdentML file
identFile <- dir(system.file(package = "MSnbase", dir = "extdata"),
                 full.name = TRUE, pattern = "dummyiTRAQ.mzid")
## create basic MSnExp
msexp <- readMSData(quantFile, verbose = FALSE)
head(fData(msexp), n = 2)
@


The \Rfunction{addIdentificationData} method takes an \Rclass{MSnExp}
instance (or an \Rclass{MSnSet} instance storing quantitation data,
see section \ref{sec:quant}) as first argument and one or multiple
\texttt{mzIdentML} file names (as a character vector) as second one and
updates the \Rclass{MSnExp} feature data using the identification
data read from the \texttt{mzIdentML} file(s).

<<msnexpIdentification2, echo=TRUE, cache=FALSE, tidy=FALSE>>=
## add identification information
msexp <- addIdentificationData(msexp, id = identFile,
                               verbose = FALSE)
head(fData(msexp), n = 2)
@

Finally we can use \Rfunction{idSummary} to summarise the percentage
of identified features per quantitation/identification pairs.

<<msnexpIdentification3, echo=TRUE, cache=FALSE, tidy=FALSE>>=
idSummary(msexp)
@

When identification data is present, and hence peptide sequences, one
can annotation fragment peaks on the MS2 figure by passing the peptide
sequence to the \Rfunction{plot} method.

<<fragplot>>=
itraqdata2 <- pickPeaks(itraqdata, verbose=FALSE)
i <- 14
s <- as.character(fData(itraqdata2)[i, "PeptideSequence"])
@

\begin{figure}[!ht]
\centering
<<fragplotfig, dev='pdf', fig.width=5, fig.height=5>>=
plot(itraqdata2[[i]], s, main = s)
@
\caption{Annotated MS2 spectrum.}
\label{fig:fragplot}
\end{figure}


The fragment ions are calculated with the
\Rfunction{calculateFragments}, described in section
\ref{sec:calcfrag} on page \pageref{calcfrag}.

\subsection{Filtering identification data}

One can remove the features that have not been identified using
\Rfunction{removeNoId}. This function uses by default the
\Robject{pepseq} feature variable to search the presence of missing data
(\Robject{NA} values) and then filter these non-identified spectra.

<<msnexpIdentification4, echo=TRUE, cache=FALSE, tidy=FALSE>>=
fData(msexp)$pepseq
msexp <- removeNoId(msexp)
fData(msexp)$pepseq
idSummary(msexp)
@

Similarly, the \Rfunction{removeMultipleAssignment} method can be used
to filter out non-unique features, i.e. that have been assigned to
protein groups with more than one member. This function uses by
default the \Robject{nprot} feature variable.

\bigskip

Note that \Rfunction{removeNoId} and
\Rfunction{removeMultipleAssignment} methods can also be called on
\Rclass{MSnExp} instances.


\subsection{Calculate Fragments}\label{sec:calcfrag}

\Biocpkg{MSnbase} is able to calculate theoretical peptide fragments via
\Rfunction{calculateFragments}.

<<calculateFragments, echo=TRUE, cache=FALSE, tidy=FALSE>>=
calculateFragments("ACEK",
                   type = c("a", "b", "c", "x", "y", "z"))
@

It is also possible to match these fragments against an \Rclass{Spectrum2}
object.
<<msnexpcalculateFragments, echo=TRUE, cache=FALSE, tidy=FALSE>>=
pepseq <- fData(msexp)$pepseq[1]
calculateFragments(pepseq, msexp[[1]], type=c("b", "y"))
@

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Quality control}\label{sec:qc}

The current section is not executed dynamically for package size and
processing time constrains.  The figures and tables have been
generated with the respective methods and included statically in the
vignette for illustration purposes.

\bigskip

\Biocpkg{MSnbase} allows easy and flexible access to the data, which
allows to visualise data features to assess it's quality. Some methods
are readily available, although many QC approaches will be experiment
specific and users are encourage to explore their data.


The \Rfunction{plot2d} method takes one \Rclass{MSnExp} instance as
first argument to produce retention time \textit{vs.} precursor MZ
scatter plots. Points represent individual MS2 spectra and can be
coloured based on precursor charge (with second argument
\Robject{z="charge"}), total ion count (\Robject{z="ionCount"}),
number of peaks in the MS2 spectra \Robject{z="peaks.count"}) or, when
multiple data files were loaded, file \Robject{z="file"}), as
illustrated on figure \ref{fig:plot2d}.  The lower right panel is
produced for only a subset of proteins.  See the method documentation
for more details.

\begin{figure}[htb]
    \includegraphics[width=\linewidth]{./plot2d-figure.png}
\caption{Illustration of the \Rfunction{plot2d} output. }
\label{fig:plot2d}
\end{figure}

The \Rfunction{plotDensity} method illustrates the distribution of
several parameters of interest (see figure \ref{fig:plotDensity}).
Similarly to \Rfunction{plot2d}, the first argument is an
\Rclass{MSnExp} instance.  The second is one of
\Robject{precursor.mz}, \Robject{peaks.count} or \Robject{ionCount},
whose density will be plotted.  An optional third argument specifies
whether the x axes should be logged.

\begin{figure}[htb]
    \includegraphics[width=\linewidth]{./plotDensity-figure.png}
\caption{ Illustration of the \Rfunction{plotDensity} output. }
\label{fig:plotDensity}
\end{figure}

%% xtable(preprocSelectionTable(velos))
%% % latex table generated in R 2.14.0 by xtable 1.5-6 package
%% % Thu May 19 10:22:29 2011
%% \begin{table}[ht]
%% \begin{center}
%% \begin{tabular}{rr}
%%   \hline
%%  & x \\
%%   \hline
%% 1 & 6836 \\
%%   2 &  59 \\
%%   3 &   5 \\
%%    \hline
%% \end{tabular}
%% \end{center}
%% \end{table}

%% matplot on iTRAQ5 quantitation (see allquant code chunk)

\bigskip

The \Rfunction{plotMzDelta} method\footnote{The code to generate the
  histograms has been contributed by Guangchuang Yu from Jinan
  University, China.} implements the $m/z$ delta plot from
\cite{Foster11} The $m/z$ delta plot illustrates the suitability of MS2
spectra for identification by plotting the $m/z$ differences of the most
intense peaks. The resulting histogram should optimally shown
outstanding bars at amino acid residu masses. More details and
parameters are described in the method documentation
(\Rfunction{?plotMzDelta}).  Figure \ref{fig:plotMzDelta-pride12011}
has been generated using the PRIDE experiment 12011, as in
\cite{Foster11}.

\begin{figure}[htb]
  \begin{center}
    \includegraphics[width=.7\linewidth]{./plotMzDelta-pride12011.pdf}
    \caption{Illustration of the \Rfunction{plotMzDelta} output for
      the PRIDE experiment 12011, as in figure 4A from
      \cite{Foster11}. }
\label{fig:plotMzDelta-pride12011}
  \end{center}
\end{figure}


\bigskip

In section \ref{sec:incompdissoc} on page \pageref{sec:incompdissoc},
we illustrate how to assess incomplete reporter ion dissociation.


\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Raw data processing}\label{sec:rawprocessing}

\subsection{Cleaning spectra}\label{sec:clean}

There are several methods implemented to perform basic raw data
processing and manipulation. Low intensity peaks can be set to 0 with
the \Rfunction{removePeaks} method from spectra or whole
experiments. The intensity threshold below which peaks are removed is
defined by the \Robject{t} parameter. \Robject{t} can be specified
directly as a numeric. The default value is the character
\texttt{"min"}, that will remove all peaks equal to the lowest non
null intensity in any spectrum.  We observe the effect of the
\Rfunction{removePeaks} method by comparing total ion count (i.e. the
total intensity in a spectrum) with the \Rfunction{ionCount} method
before (object \Robject{itraqdata}) and after (object
\Robject{experiment}) for spectrum \texttt{X55}. The respective
spectra are shown on figure \ref{fig:spectrum-clean-plot} (page
\pageref{fig:spectrum-clean-plot}).


<<removePeaks, echo=TRUE, cache=FALSE>>=
experiment <- removePeaks(itraqdata, t = 400, verbose = FALSE)
## total ion current
ionCount(itraqdata[["X55"]])
ionCount(experiment[["X55"]])
@

\begin{figure}[!ht]
<<spectrum-clean-plot, dev='pdf', fig.width=6, fig.height=3, echo=FALSE, fig.keep='high'>>=
p1 <- plot(itraqdata[["X55"]], full = TRUE, plot = FALSE) + theme_gray(5)
p2 <- plot(experiment[["X55"]], full = TRUE, plot = FALSE) + theme_gray(5)
grid.newpage()
pushViewport(viewport(layout = grid.layout(1, 2)))
vplayout <- function(x, y)
  viewport(layout.pos.row = x, layout.pos.col = y)
print(p1,vp=vplayout(1,1))
print(p2,vp=vplayout(1,2))
@
\caption{Same spectrum before (left) and after setting peaks <= 400 to 0.}
\label{fig:spectrum-clean-plot}
\end{figure}

Unlike the name might suggest, the \Rfunction{removePeaks} method does
not actually remove peaks from the spectrum; they are set to 0. This
can be checked using the \Rfunction{peaksCount} method, that returns
the number of peaks (including 0 intensity peaks) in a spectrum.  To
effectively remove 0 intensity peaks from spectra, and reduce the size
of the data set, one can use the \Rfunction{clean} method.  The effect
of the \Rfunction{removePeaks} and \Rfunction{clean} methods are
illustrated on figure \ref{fig:preproc} on page \pageref{fig:preproc}.

<<clean, echo=TRUE, cache=FALSE>>=
## number of peaks
peaksCount(itraqdata[["X55"]])
peaksCount(experiment[["X55"]])
experiment <- clean(experiment, verbose = FALSE)
peaksCount(experiment[["X55"]])
@


<<preprosp, cache=FALSE, echo=FALSE>>=
int <- c(0,1,1,3,1,1,0,0,0,1,3,7,3,1,0)
mz <- c(113.9,114.0,114.05,114.1,114.15,114.2,114.25,
        114.3,114.35,114.4,114.42,114.48,114.5,114.55,114.6)
ppsp <- new("Spectrum2",intensity=int,mz=mz,centroided=FALSE)
p1 <- plot(ppsp, full = TRUE, plot = FALSE) + theme_gray(5) +
  geom_point(size=3,alpha=I(1/3)) +
  geom_hline(yintercept=3,linetype=2) +
  ggtitle("Original spectrum")
p2 <- plot(removePeaks(ppsp,t=3), full=TRUE, plot = FALSE) +
  theme_gray(5) +
  geom_point(size=3,alpha=I(1/3)) +
  geom_hline(yintercept=3,linetype=2) +
  ggtitle("Peaks < 3 removed")
p3 <- plot(clean(removePeaks(ppsp,t=3)), full = TRUE, plot = FALSE) +
  theme_gray(5) +
  geom_point(size=3,alpha=I(1/3)) +
  geom_hline(yintercept=3,linetype=2) +
  ggtitle("Peaks < 3 removed and cleaned")
@

\begin{figure}[p]
\centering
<<preprocPlot, dev='pdf', fig.width=3, fig.height=6, echo=FALSE>>=
grid.newpage()
pushViewport(viewport(layout = grid.layout(3, 1)))
print(p1, vp=vplayout(1, 1))
print(p2, vp=vplayout(2, 1))
print(p3, vp=vplayout(3, 1))
@
\caption{This figure illustrated the effect or the \Rfunction{removePeaks}
  and \Rfunction{clean} methods. The left-most spectrum displays two peaks,
  of max height 3 and 7 respectively. The middle spectrum shows the result
  of calling \Rfunction{removePeaks} with argument \Robject{t=3}, which sets
  all data points of the first peak, whose maximum height is smaller or equal
  to \Robject{t} to 0. The second peak is unaffected. Calling \Rfunction{clean}
  after \Rfunction{removePeaks} effectively deletes successive 0 intensities from
  the spectrum, as shown on the right plot. }
\label{fig:preproc}
\end{figure}

\subsection{Focusing on specific MZ values}\label{sec:trim}

Another useful manipulation method is \Rfunction{trimMz}, that takes
as parameters and \Rclass{MSnExp} (or a \Rclass{Spectrum}) and a
numeric \Robject{mzlim}. MZ values smaller then \texttt{min(mzlim)} or
greater then \texttt{max(mzmax)} are discarded. This method is
particularly useful when one wants to concentrate on a specific MZ
range, as for reporter ions quantification, and generally results in
substantial reduction of data size. Compare the size of the full
trimmed experiment to the original \Sexpr{sz} Mb.
\label{trimMz-example}

<<trimMz, echo=TRUE, cache=FALSE>>=
range(mz(itraqdata[["X55"]]))
experiment <- filterMz(experiment, mzlim = c(112,120))
range(mz(experiment[["X55"]]))
experiment
@

As can be seen above, all processing performed on the experiment is
recorded and displayed as integral part of the experiment object.

\subsection{Spectrum processing}\label{sec:specproc}

\Rclass{MSnExp} and \Rclass{Spectrum2} instances also support standard
MS data processing such as smoothing and peak picking, as described in
the \Rfunction{smooth} and \Rfunction{pickPeak} manual pages. The
methods that either single spectra of experiments, process the
spectrum/spectra, and return a updated, processed, object. The
implementations originate from the \CRANpkg{MALDIquant} package
\cite{Gibb2012}.

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{MS$^2$ isobaric tagging quantitation}\label{sec:isoquant}

\subsection{Reporter ions quantitation}\label{sec:quant}

Quantitation is performed on fixed peaks in the spectra, that are
specified with an \Rclass{ReporterIons} object. A specific peak is
defined by it's expected \Robject{mz} value and is searched for within
\Robject{mz} $\pm$ \Robject{width}.  If no data is found, \Robject{NA}
is returned.

<<reporters, echo=TRUE, cache=FALSE>>=
mz(iTRAQ4)
width(iTRAQ4)
@

The \Rfunction{quantify} method takes the following parameters: an
\Rclass{MSnExp} experiment, a character describing the quantification
\Robject{method}, the \Robject{reporters} to be quantified and a
\Robject{strict} logical defining whether data points ranging outside
of \Robject{mz} $\pm$ \Robject{width} should be considered for
quantitation.  Additionally, a progress bar can be displaying when
setting the \Robject{verbose} parameter to \Robject{TRUE}.  Three
quantification methods are implemented, as illustrated on figure
\ref{fig:quant-methods}: \Robject{trapezoidation} returns the area
under the peak of interest, \Robject{max} returns the apex of the peak
and \Robject{sum} returns the sum of all intensities of the peak.  See
\Rfunction{?quantify} for more details.

<<simplesp, cache=FALSE, echo=FALSE, fig.keep='none'>>=
int <- c(0, 1, 1, 3, 1, 1, 0)
mz <- c(113.9, 114.0, 114.05, 114.1, 114.15, 114.2, 114.25)
ssp <- new("Spectrum2", intensity = int, mz = mz, centroided = FALSE)
p <- plot(ssp, full = TRUE, plot = FALSE)
p <- p + theme_gray(5)
@

\begin{figure}[p]
\centering
<<quantitationPlot, dev='pdf',fig.width=5, fig.height=5, echo=FALSE>>=
grid.newpage()
pushViewport(viewport(layout = grid.layout(2, 2)))
vplayout <- function(x, y)
  viewport(layout.pos.row = x, layout.pos.col = y)
print(p + ggtitle("Quantitation using 'sum'") +
      geom_point(size = 3, alpha = I(1/3), colour = "red"),
      vp = vplayout(1, 1))
print(p + ggtitle("Quantitation using 'max'") +
      geom_point(aes(x = 114.1, y = 3),
                 alpha = I(1/18),
                 colour = "red", size = 3),
      vp = vplayout(1, 2))
print(p +
      ggtitle("Trapezoidation and strict=FALSE") +
      geom_polygon(alpha = I(1/5), fill = "red"),
      vp = vplayout(2, 1))
print(p +
      ggtitle("Trapezoidation and strict=TRUE") +
      geom_polygon(aes(x = c(NA, 114.05, 114.05, 114.1, 114.15, 114.15, NA),
                       y=c(NA, 0, 1, 3, 1, 0, NA)), fill = "red", alpha = I(1/5)),
      vp = vplayout(2,2))
@
\caption{The different quantitation methods are illustrated above.
  Quantitation using \Robject{sum} sums all the data points in the
  peaks to produce, for this example,
  \Sexpr{quantify(ssp,iTRAQ4[1],method="sum")[[1]]}, whereas method
  \Robject{max} only uses the peak's maximum intensity,
  \Sexpr{quantify(ssp,iTRAQ4[1],method="max")[[1]]}.
  \Robject{Trapezoidation} calculates the area under the peak taking
  the full with into account (using \Robject{strict=FALSE} gives
  \Sexpr{round(quantify(ssp,iTRAQ4[1],method="trap",strict=FALSE)[[1]],3)})
  or only the width as defined by the reporter (using
  \Robject{strict=TRUE} gives
  \Sexpr{round(quantify(ssp,iTRAQ4[1],method="trap",strict=TRUE)[[1]],3)}). }
\label{fig:quant-methods}
\end{figure}

\bigskip

The \Rfunction{quantify} method returns \Rclass{MSnSet} objects, that
extend the well-known \Rclass{eSet} class defined in the
\Biocpkg{Biobase} package. \Rclass{MSnSet} instances are very
similar to \Rclass{ExpressionSet} objects, except for the experiment
meta-data that captures MIAPE specific information.  The assay data is
a matrix of dimensions $n \times m$, where $m$ is the number of
features/spectra originally in the \Rclass{MSnExp} used as parameter
in \Rfunction{quantify} and $m$ is the number of reporter ions, that can
be accessed with the \Rfunction{exprs} method.  The meta data is
directly inherited from the \Rclass{MSnExp} instance.

<<quantify, echo=TRUE, cache=FALSE, tidy=FALSE>>=
qnt <- quantify(experiment,
                method = "trap",
                reporters = iTRAQ4,
                strict = FALSE,
                verbose = FALSE)
qnt
head(exprs(qnt))
@

Figure \ref{fig:tmt10} illustrated the quantitation of the TMT 10-plex
isobaric tags using the \Rfunction{quantify} method and the
\texttt{TMT10} reporter instance. The data on the $x$ axis has been
quantified using \texttt{method = "max"} and centroided data (as
generated using ProteoWizard's \texttt{msconvert} with vendor
libraries' peak picking); on the $y$ axis, the quantitation method was
\texttt{trapezoidation} and \texttt{strict = TRUE} (that's important
for TMT 10-plex) and the profile data. We observe a very good
correlation.

\begin{figure}[htb]
  \centering
  \includegraphics[width=.6\linewidth]{./tmt10comp.pdf}
\caption{TMT 10-plex quantitation. }
\label{fig:tmt10}
\end{figure}


If no peak is detected for a reporter ion peak, the respective
quantitation value is set to \texttt{NA}. In our case, there
is \Sexpr{sum(is.na(exprs(qnt)))} such case in row
\Sexpr{which(is.na(exprs(qnt))) %% nrow(qnt)}.
We will remove the offending line using the \Rfunction{filterNA} method.
The \Robject{pNA} argument defines the percentage of accepted missing
values per feature. As we do not expect any missing peaks, we set it
to be 0 (which is also the detault value).

<<filterNA, echo=TRUE>>=
table(is.na(qnt))
qnt <- filterNA(qnt, pNA = 0)
sum(is.na(qnt))
@

The filtering criteria for \Rfunction{filterNA} can also be defined as
a pattern of columns that can have missing values and columns that
must not exhibit any.  See \Rfunction{?filterNA} for details and
examples.

The infrastructure around the \Rclass{MSnSet} class allows flexible
filtering using the \Rfunction{[} sub-setting operator. Below, we
mimic the behaviour of \Rfunction{filterNA(, pNA = 0)} by calculating
the row indices that should be removed, i.e. those that have at least
one \Robject{NA} value and explicitly remove these rows.  This method
allows one to devise and easily apply any filtering strategy.

<<removeNa, echo=TRUE, eval=FALSE>>=
whichRow <- which(is.na((qnt))) %% nrow(qnt)
qnt <- qnt[-whichRow, ]
@

See also the \Rfunction{plotNA} method to obtain a graphical overview of
the completeness of a data set.

\subsection{Importing quantitation data}\label{sec:io2}

If quantitation data is already available as a spreadsheet, it can be
imported, along with additional optional feature and sample (pheno) meta data,
with the \Rfunction{readMSnSet} function. This function takes the
respective text-based spreadsheet (comma- or tab-separated) file names
as argument to create a valid \Rclass{MSnSet} instance.

Note that the quantitation data of \Rclass{MSnSet} objects can also
be exported to a text-based spreadsheet file using the
\Rfunction{write.exps} method.

\Biocpkg{MSnbase} also supports the \texttt{mzTab}
format\footnote{\url{https://github.com/HUPO-PSI/mzTab}}, a
light-weight, tab-delimited file format for proteomics
data. \texttt{mzTab} files can be read into \R with
\Rfunction{readMzTabData} to create and \Rclass{MSnSet} instance.

\bigskip

See the \texttt{MSnbase-io} vignette for a general overview of
\Biocpkg{MSnbase}'s input/ouput capabilites.

\subsection{Peak adjustments}\label{sec:purcor}

\paragraph{Single peak adjustment} In certain cases, peak intensities
need to be adjusted as a result of peak interferance. For example, the
$+1$ peak of the phenylalanine (F, Phe) immonium ion (with $m/z$ 120.03)
inteferes with the 121.1 TMT reporter ion.  Below, we calculate the
relative intensity of the +1 peaks compared to the main peak using the
\Biocpkg{Rdisop} package.

<<pheplus1, echo=TRUE, cache=FALSE>>=
library(Rdisop)
## Phenylalanine immonium ion
Fim <- getMolecule("C8H10N")
getMass(Fim)
isotopes <- getIsotope(Fim)
F1 <- isotopes[2, 2]
F1
@

If desired, one can thus specifically quantify the F immonium ion in
the MS2 spectrum, estimate the intensity of the +1 ion
(\Sexpr{round(F1,4)}\% of the F peak) and substract this calculated
value from the 121.1 TMT reporter intensity.

The above principle can also be generalised for a set of overlapping
peaks, as described below.

\paragraph{Reporter ions purity correction} Impurities in the reporter
reagents can also bias the results and can be corrected when
manufacturers provide correction coefficients.  These generally come
as percentages of each reporter ion that have masses differing by -2,
-1, +1 and +2 Da from the nominal reporter ion mass due to isotopic
variants.  The \Rfunction{purityCorrect} method applies such
correction to \Rclass{MSnSet} instances. It also requires a square
matrix as second argument, \Robject{impurities}, that defines the
relative percentage of reporter in the quantified each peak.  See
\Rfunction{?purityCorrect} for more details.

<<purityCorrect, echo=TRUE, cache=FALSE, tidy = FALSE>>=
impurities <- matrix(c(0.929, 0.059, 0.002, 0.000,
                       0.020, 0.923, 0.056, 0.001,
                       0.000, 0.030, 0.924, 0.045,
                       0.000, 0.001, 0.040, 0.923),
                     nrow = 4)
qnt.crct <- purityCorrect(qnt, impurities)
head(exprs(qnt))
head(exprs(qnt.crct))
@

The \Rfunction{makeImpuritiesMatrix} can be used to create impurity
matrices.  It opens a rudimentary spreadsheet that can be directly
edited.

\clearpage

\section{Processing quantitative data}\label{sec:qproc}

\subsection{Data imputation}\label{sec:imp}

A set of imputation methods are available in the \Rfunction{impute}
method: it takes an \Rclass{MSnSet} instance as input, the name of the
imputation method to be applied (one of \Sexpr{paste(imputeMethods(),
  collapse=", ")}), possible additional parameters and returns an
updated for \Rclass{MSnSet} without any missing values. Below, we
apply a deterministic minimum value imputation on the \Robject{naset}
example data:

<<impute>>=
## an example MSnSet containing missing values
data(naset)
table(is.na(naset))
## number of NAs per protein
table(fData(naset)$nNA) 
x <- impute(naset, "min")
processingData(x)
table(is.na(x))
@


There are two types of mechanisms resulting in missing values in
LC/MSMS experiments.

\begin{itemize}
\item Missing values resulting from absence of detection of a feature,
  despite ions being present at detectable concentrations.  For
  example in the case of ion suppression or as a result from the
  stochastic, data-dependent nature of the MS acquisition
  method. These missing value are expected to be randomly distributed
  in the data and are defined as \emph{missing at random} (MAR) or
  \emph{missing completely at random} (MCAR).
\item Biologically relevant missing values, resulting from the
  \emph{absence} of the low abundance of ions (below the limit of
  detection of the instrument). These missing values are not expected
  to be randomly distributed in the data and are defined as
  \emph{missing not at random} (MNAR).
\end{itemize}

MAR and MCAR values can be reasonably well tackled by many imputation
methods. MNAR data, however, requires some knowledge about the
underlying mechanism that generates the missing data, to be able to
attempt data imputation. MNAR features should ideally be imputed with
a \emph{left-censor} (for example using a deterministic or
probabilistic minimum value) method. Conversely, it is recommended to
use \emph{hot deck} methods (for example nearest neighbour, maximum
likelihood, etc) when data are missing at random. 

\begin{figure}[h]
  \centering
  <<miximpfig, echo=FALSE, dev='pdf', fig.width=8, fig.width=8>>=
  x <- impute(naset, "zero")
  exprs(x)[exprs(x) != 0] <- 1

  suppressPackageStartupMessages(library("gplots"))

  heatmap.2(exprs(x), col = c("lightgray", "black"),
              scale = "none", dendrogram = "none",
              trace = "none", keysize = 0.5, key = FALSE,
              RowSideColors = ifelse(fData(x)$randna, "orange", "brown"),
              ColSideColors = rep(c("steelblue", "darkolivegreen"), each = 8))
  @
\caption{Mixed imputation method. Black cells represent presence of
  quantitation values and light grey corresponds to missing data. The
  two groups of interest are depicted in green and blue along the
  heatmap columns. Two classes of proteins are annotated on the left:
  yellow are proteins with randomly occurring missing values (if any)
  while proteins in brown are candidates for non-random missing value
  imputation.}
\label{fig:miximp}
\end{figure}


It is anticipated that the identification of both classes of missing
values will depend on various factors, such as feature intensities and
experimental design. Below, we use perform mixed imputation, applying
nearest neighbour imputation on the \Sexpr{sum(fData(naset)$randna)}
features that are assumed to contain randomly distributed missing
values (if any) (yellow on figure \ref{fig:miximp}) and a
deterministic minimum value imputation on the
\Sexpr{sum(!fData(naset)$randna)} proteins that display a non-random
pattern of missing values (brown on figure \ref{fig:miximp}).

<<miximp>>=
x <- impute(naset, method = "mixed",
            randna = fData(naset)$randna,
            mar = "knn", mnar = "min")
x
@


Please read \Rfunction{?impute} for a description of the different
methods.

\clearpage

\subsection{Normalisation}\label{sec:norm}

A \Rclass{MSnSet} object is meant to be compatible with further
downstream packages for data normalisation and statistical
analysis. There is also a \Rfunction{normalise} (also available as
\Rfunction{normalize}) method for expression sets. The method takes
and instance of class \Rclass{MSnSet} as first argument, and a
character to describe the \Robject{method} to be used:

\begin{description}
\item{\Robject{quantiles}} Applies quantile normalisation
  \cite{Bolstad03} as implemented in the
  \Rfunction{normalize.quantiles} function of the
  \Biocpkg{preprocessCore} package.
\item{\Robject{quantiles.robust}} Applies robust quantile
  normalisation \cite{Bolstad03} as implemented in the
  \Rfunction{normalize.quantiles.robust} function of the
  \Biocpkg{preprocessCore} package.
\item{\Robject{vsn}} Applies variance stabilisation normalization
  \cite{Huber2002} as implemented in the \Rfunction{vsn2} function of
  the \Biocpkg{vsn} package.
\item{\Robject{max}} Each feature's reporter intensity is divided by
  the maximum of the reporter ions intensities.
\item{\Robject{sum}} Each feature's reporter intensity is divided by
  the sum of the reporter ions intensities.
\end{description}

See \Rfunction{?normalise} for more methods. A \Rfunction{scale}
method for \Rclass{MSnSet} instances, that relies on the
\Rfunction{base::scale} function.

<<normalise, echo=TRUE, cache=FALSE>>=
qnt.max <- normalise(qnt, "max")
qnt.sum <- normalise(qnt, "sum")
qnt.quant <- normalise(qnt, "quantiles")
qnt.qrob <- normalise(qnt, "quantiles.robust")
qnt.vsn <- normalise(qnt, "vsn")
@

The effect of these are illustrated on figure \ref{fig:norm-plot} and
figure \ref{fig:cv-plot} reproduces figure 3 of \cite{Karp2010} that
described the application of vsn on iTRAQ reporter data.

\begin{figure}[p]
\centering
<<normPlot, dev='pdf', fig.width=5, fig.height=7, echo=FALSE>>=
.plot <- function(x,ttl=NULL) {
  boxplot(exprs(x),
          main=ifelse(is.null(ttl),processingData(x)@processing[2],ttl),
          cex.main=.8,
          cex.lab=.5,
          cex.axis=.5,
          cex=.8)
  grid()
}
oldmar <- par()$mar
par(mfrow=c(3,2),mar=c(2.9,2.9,2.9,1))
.plot(qnt, ttl = "Non-normalised data")
.plot(qnt.max, ttl = "Maximum")
.plot(qnt.sum, ttl = "Sum")
.plot(qnt.quant, ttl = "Quantile")
.plot(qnt.qrob, ttl = "Robust quantile")
.plot(qnt.vsn, ttl = "vsn")
@
\caption{ Comparison of the normalisation \Rclass{MSnSet} methods.
  Note that vsn also glog-transforms the intensities. }
\label{fig:norm-plot}
\end{figure}

<<cvdata, echo=FALSE, cache=FALSE>>=
sd1 <- apply(log2(exprs(qnt))+10,1,sd)
mn1 <- apply(log2(exprs(qnt))+10,1,mean)
cv1 <- sd1/mn1
sd2 <- apply(exprs(qnt.vsn)+10,1,sd)
mn2 <- apply(exprs(qnt.vsn)+10,1,mean)
cv2 <- sd2/mn2
dfr <- rbind(data.frame(rank=order(mn1),cv=cv1,norm="raw"),
             data.frame(rank=order(mn2),cv=cv2,norm="vsn"))
library("zoo")
## rmed1 <- rollapply(cv1,7,function(x) median(x,na.rm=TRUE))
## rmed2 <- rollapply(cv2,7,function(x) median(x,na.rm=TRUE))
##
## Calling directly rollapply.zoo to make it zoo_1.6-4 compatible.
## The above requires zoo >= 1.7-0, which is as of 15 March 2011
## not yet available on CRAN (only on r-forge).
rmed1 <- zoo:::rollapply.zoo(cv1,7,function(x) median(x,na.rm=TRUE))
rmed2 <- zoo:::rollapply.zoo(cv2,7,function(x) median(x,na.rm=TRUE))
dfr2 <-
  rbind(data.frame(x=seq(1,70,by=(70/length(rmed1))),
                   y=rmed1,norm="raw"),
        data.frame(x=seq(1,70,by=(70/length(rmed1))),
                   y=rmed2,norm="vsn"))

p <- qplot(rank,cv,data=dfr,col=norm) +
  geom_line(data=dfr2,aes(x=x,y=y,colour=norm)) +
  theme_gray(7)
@

\begin{figure}[!ht]
\centering
<<cvPlot, dev='pdf', fig.width=5, fig.height=4, echo=FALSE>>=
print(p)
@
\caption{CV versus signal intensity comparison for log2 and vsn
  transformed data. Lines indicate running CV medians.}
\label{fig:cv-plot}
\end{figure}

Note that it is also possible to normalise individual spectra or whole
\Rclass{MSnExp} experiments with the \Rfunction{normalise} method
using the \Robject{max} method.  This will rescale all peaks between 0
and 1.  To visualise the relative reporter peaks, one should this
first trim the spectra using method \Rfunction{trimMz} as illustrated
in section \ref{sec:rawprocessing}, then normalise the \Rclass{MSnExp}
with \Rfunction{normalise} using \Robject{method="max"} as illustrated
above and plot the data using \Rfunction{plot} (figure
\ref{fig:msnexp-norm-plot}).

<<prepareMsnsetNormPlot, cache=FALSE, echo=FALSE, keep.fig='none'>>=
p <- plot(normalise(experiment[bsasel], "max"),
          reporters = iTRAQ4, full = FALSE, plot = FALSE)
p <- p + theme_gray(7)
@

\begin{figure}[p]
\centering
<<msnexpNormPlot, dev='pdf', fig.width=5, fig.height=6, echo=FALSE>>=
print(p)
@
\caption{Experiment-wide normalised MS2 spectra. The y-axes of the individual
  spectra is now rescaled between 0 and 1 (highest peak), as opposed to
  figure \ref{fig:msnexp-plot}. }
\label{fig:msnexp-norm-plot}
\end{figure}

\bigskip

Additional dedicated normalisation method are available for MS$^2$
label-free quantitation, as described in section \ref{sec:lf} and in
the \Rfunction{quantify} documentation.

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Feature aggregation}\label{sec:feataggregation}

The above quantitation and normalisation has been performed on
quantitative data obtained from individual spectra. However, the
biological unit of interest is not the spectrum but the peptide or the
protein. As such, it is important to be able to summarise features
that belong to a same group, i.e. spectra from one peptide, peptides
that originate from one protein, or directly combine all spectra that
have been uniquely associated to one protein.

\Biocpkg{MSnbase} provides one function, \Rfunction{combineFeatures},
that allows to aggregate features stored in an \Rclass{MSnSet} using
build-in or user defined summary function and return a new
\Rclass{MSnSet} instance.  The three main arguments are described
below.  Additional details can be found in the method documentation.

<<makeGroups1,echo=FALSE,cache=FALSE>>=
gb <- fData(qnt)$ProteinAccession
@

\Rfunction{combineFeatures}'s first argument, \Robject{object}, is an
instance of class \Rclass{MSnSet}, as has been created in the section
\ref{sec:quant} for instance.  The second argument, \Robject{groupBy},
is a \Robject{factor} than has as many elements as there are features
in the \Rclass{MSnSet} \Robject{object} argument. The features
corresponding to the \Robject{groupBy} levels will be aggregated so
that the resulting \Rclass{MSnSet} output will have
\Rfunction{length(levels(groupBy))} features. Here, we will combine
individual MS2 spectra based on the protein they originate from.  As
shown below, this will result in \Sexpr{length(table(gb))} new
aggregated features.

<<makeGroups2,echo=TRUE,cache=FALSE>>=
gb <- fData(qnt)$ProteinAccession
table(gb)
length(unique(gb))
@

The third argument, \Robject{fun}, defined how to combine the
features. Predefined functions are readily available and can be
specified as strings (\Robject{fun="mean"}, \Robject{fun="median"},
\Robject{fun="sum"}, \Robject{fun="weighted.mean"} or
\Robject{fun="medianpolish"} to compute respectively the mean, media,
sum, weighted mean or median polish of the features to be aggregated).
Alternatively, is is possible to supply user defined functions with
\Robject{fun=function(x) \{ ... \}}. We will use the \Robject{median}
here.

<<combineFeatures, echo=TRUE, cache=FALSE>>=
qnt2 <- combineFeatures(qnt, groupBy = gb, fun = "median")
qnt2
@

%% Other intended ways to define aggregation groups is to import
%% identification data into R and collate it to the \Robject{MSnSet}
%% feature data, and subsequently use peptide sequences or protein assignments
%% to combine features.

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Label-free MS$^2$ quantitation}\label{sec:lf}

\subsection{Peptide counting}

Note that if samples are not multiplexed, label-free MS$^2$
quantitation by spectral counting is possible using
\Biocpkg{MSnbase}. Once individual spectra have been assigned to
peptides and proteins (see section \ref{sec:id}), it becomes
straightforward to estimate protein quantities using the simple
peptide counting method, as illustrated in section
\ref{sec:feataggregation}.

<<count>>=
sc <- quantify(msexp, method = "count")
## lets modify out data for demonstration purposes
fData(sc)$accession[1] <- fData(sc)$accession[2]
fData(sc)$accession
sc <- combineFeatures(sc, groupBy = fData(sc)$accession,
                      fun = "sum")
exprs(sc)
@

Such count data could then be further analyses using dedicated count
methods (originally developed for high-throughput sequencing) and
directly available for \Rclass{MSnSet} instances in the
\Biocpkg{msmsTests} Bioconductor package.

\subsection{Spectral counting and intensity methods}

The spectral abundance factor (SAF) and the normalised form (NSAF)
\cite{Paoletti2006} as well as the spectral index (SI) and other
normalised variations (SI$_{GI}$ and SI$_N$) \cite{Griffin2010} are
also available. Below, we illustrate how to apply the normalised
SI$_N$ to the experiment containing identification data produced in
section \ref{sec:id}.

The spectra that did not match any peptide have already been remove
with the \Rfunction{removeNoId} method. As can be seen in the
following code chunk, the first spectrum could not be matched to any
single protein. Non-identified spectra and those matching multiple
proteins are removed automatically prior to any label-free
quantitation. Once can also remove peptide that do not match uniquely
to proteins (as defined by the \Robject{nprot} feature variable column)
with the \Rfunction{removeMultipleAssignment} method.

<<labelfree>>=
fData(msexp)[, c("accession", "nprot")]
@

Note that the label-free methods implicitely apply feature aggregation
(section \ref{sec:feataggregation}) and normalise (section
\ref{sec:norm}) the quantitation values based on the total sample
intensity and or the protein lengths (see \cite{Paoletti2006} and
\cite{Griffin2010} for details).

Let's now proceed with the quantitation using the
\Rfunction{quantify}, as in section \ref{sec:quant}, this time however
specifying the method of interest, \Robject{SIn} (the \Robject{reporters}
argument can of course be ignored here). The required peptide-protein
mapping and protein lengths are extracted automatically from the
feature meta-data using the default \Robject{accession} and \Robject{length}
feature variables.

<<SIn>>=
siquant <- quantify(msexp, method = "SIn")
processingData(siquant)
exprs(siquant)
@

Other label-free methods can be applied by specifiying the appropriate
\Rfunction{method} argument. See \Rfunction{?quantify} for more details.

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Spectra comparison}

\subsection{Plotting two spectra}

\Biocpkg{MSnbase} provides functionality to compare spectra against
each other. The first notable function is \Rfunction{plot}. If two
\Rclass{Spectrum2} objects are provided \Rfunction{plot} will draw
two plots: the upper and lower panel contain respectively the first
and second spectrum. Common peaks are drawn in a slightly darker
colour.

\begin{figure}[!ht]
\centering
<<plotSpectrumSpectrum, dev='pdf', fig.width=6.5, fig.height=6.5>>=
centroided <- pickPeaks(itraqdata, verbose = FALSE)
(k <- which(fData(centroided)[, "PeptideSequence"] == "TAGIQIVADDLTVTNPK"))
mzk <- precursorMz(centroided)[k]
zk <- precursorCharge(centroided)[k]
mzk * zk
plot(centroided[[k[1]]], centroided[[k[2]]])
@
\caption{Comparing two MS$^2$ spectra.}
\label{fig:compms2plot}
\end{figure}

\subsection{Comparison metrics}

Currently \Biocpkg{MSnbase} supports three different metrics to
compare spectra against each other: \Robject{common} to calculate the
number of common peaks, \Robject{cor} to calculate the Pearson
correlation and \Robject{dotproduct} to calculate the dot product. See
\Rfunction{?compareSpectra} to apply other arbitrary metrics.

<<compareSpectra, tidy=FALSE>>=
compareSpectra(centroided[[2]], centroided[[3]],
               fun = "common")
compareSpectra(centroided[[2]], centroided[[3]],
               fun = "cor")
compareSpectra(centroided[[2]], centroided[[3]],
               fun = "dotproduct")
@

\Rfunction{compareSpectra} supports \Rclass{MSnExp} objects as well.

<<msnexpcompareSpectra>>=
compmat <- compareSpectra(centroided, fun="cor")
compmat[1:10, 1:5]
@

Below, we illustrate how to compare a set of spectra using a
hierarchical clustering.

\begin{figure}[!ht]
\centering
<<hclustcompareSpectra, dev='pdf', fig.width=5, fig.height=6>>=
plot(hclust(as.dist(compmat)))
@
\label{fig:dendo}
\end{figure}

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Quantitative assessment of incomplete dissociation}
\label{sec:incompdissoc}

Quantitation using isobaric reporter tags assumes complete
dissociation between the reporter group (red on figure
\ref{fig:itraqchem}), balance group (blue) and peptide (the peptide
reactive group is drawn in green).  However, incomplete dissociation
does occur and results in an isobaric tag (i.e reporter and balance
groups) specific peaks.

\begin{figure}[h]
  \begin{center}
    \includegraphics[width=4cm]{./itraqchem.pdf}
    \caption{ iTRAQ 4-plex isobaric tags reagent consist of three
      parts: (1) a charged reporter group (MZ of 114, 115, 116 and
      117) that is unique to each of the four reagents (red), (2) an
      uncharged mass balance group (28-31 Da) (blue)and (3) a peptide
      reactive group (NHS ester) that binds to the peptide.  In case
      of incomplete dissociation, the reporter and balance groups
      produce a specific peaks at MZ 145.  }
\label{fig:itraqchem}
  \end{center}
\end{figure}

\Biocpkg{MSnbase} provides, among others, a \Rclass{ReporterIons}
object for iTRAQ 4-plex that includes the 145 peaks, called
\Rclass{iTRAQ5}. This can then be used to quantify the experiment as
show in section \ref{sec:quant} to estimate incomplete dissociation
for each spectrum.

<<incompdiss, echo=TRUE, cache=FALSE, tidy=FALSE>>=
iTRAQ5
incompdiss <- quantify(itraqdata,
                       method = "trap",
                       reporters = iTRAQ5,
                       strict = FALSE,
                       verbose = FALSE)
head(exprs(incompdiss))
@

Figure \ref{fig:incompdiss} compares these intensities for the whole experiment.

\begin{figure}[!hbt]
\centering
<<incompdissPlot, dev='pdf', fig.width=5, fig.height=2.5, echo=FALSE, warning=FALSE>>=
dfr <- melt(exprs(incompdiss))
colnames(dfr) <- c("feature", "reporters", "intensity")
dfr$reporters <- sub("iTRAQ5.", "", dfr$reporters)
repsum <- apply(exprs(incompdiss), 1, function(x) sum(x[1:4]))
dfr2 <- data.frame(iTRAQ1to4 = repsum, iTRAQ5 = exprs(incompdiss)[,5])
p1 <- ggplot(data = dfr, aes(x = reporters,y = log10(intensity))) +
  geom_boxplot() +
  theme_gray(6)
p2 <- ggplot(data = dfr2, aes(x = log10(iTRAQ1to4), y = log10(iTRAQ5))) +
  geom_point(alpha = I(1/2)) +
  geom_abline(intercept = 0, slope = 1, linetype = "dotted") +
  stat_smooth(method = lm, se = FALSE) +
  xlab(label = expression(log[10]~sum~114~to~117)) +
  ylab(label = expression(log[10]~145)) +
  theme_gray(6)
grid.newpage()
pushViewport(viewport(layout = grid.layout(1, 2)))
print(p1, vp = vplayout(1, 1))
print(p2, vp = vplayout(1, 2))
@
\caption{Boxplot and scatterplot comparing intensities of the 4
  reporter ions (or their sum, on the right) and the incomplete
  dissociation specific peak. }
\label{fig:incompdiss}
\end{figure}

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{Combining \Rclass{MSnSet} instances}\label{sec:combine}


Combining mass spectrometry runs can be done in two different ways
depending on the nature of these runs. If the runs represent repeated
measures of identical samples, for instance multiple fractions, the
data has to be combine along the row of the quantitation matrix: all
the features (along the rows) represent measurements of the same set
of samples (along the columns). In this situation, described in
section \ref{sec:comb1}, two experiments of dimensions $n_1$ (rows) by
$m$ (columns and $n_2$ by $m$ will produce a new experiment of
dimensions $n_1 + n_2$ by $m$.

When however, different sets of samples have been analysed in
different mass spectrometry runs, the data has to be combined along
the columns of the quantitation matrix: some features will be shared
across experiments and should thus be aligned on a same row in the new
data set, whereas unique features to one experiment should be set as
missing in the other one.  In this situation, described in section
\ref{sec:comb2}, two experiments of dimensions $n_1$ by $m_1$ and
$n_2$ by $m_2$ will produce a new experiment of dimensions
$unique_{n_1} + unique_{n_2} + shared_{n_1, n_2}$ by $m_1 + m_2$.  The
two first terms of the first dimension will be complemented by
\Robject{NA} values.

Default \Rclass{MSnSet} feature names (\Robject{X1}, \Robject{X2},
\ldots) and sample names (\Robject{iTRAQ4.114}, \Robject{iTRAQ4.115},
\Robject{iTRAQ4.116}, \ldots) are not informative.  The features and
samples of these anonymous quantitative data-sets should be updated
before being combined, to guide how to meaningfully merge them.


\subsection{Combining identical samples}\label{sec:comb1}

To simulate this situation, let us use quantiation data from the
\Robject{itraqdata} object that is provided with the package as
experiment 1 and the data from the \Robject{rawdata} \Rclass{MSnExp}
instance created at the very beginning of this document. Both
experiments share the \emph{same} default iTRAQ 4-plex reporter names
as default sample names, and will thus automatically be combined along
rows.

<<makeexp12, echo=TRUE, cache=FALSE, tidy = FALSE>>=
exp1 <- quantify(itraqdata, reporters = iTRAQ4,
                 verbose = FALSE)
sampleNames(exp1)
centroided(rawdata) <- FALSE
exp2 <- quantify(rawdata, reporters = iTRAQ4,
                 verbose = FALSE)
sampleNames(exp2)
@

It important to note that the features of these independent
experiments share the same default anonymous names: X1, X2, X3,
\ldots, that however represent quantitation of distinct physical
analytes.  If the experiments were to be combined as is, it would
result in an error because data points for the same \emph{feature}
name (say \Robject{X1}) and the same \emph{sample name} (say
\Robject{iTRAQ4.114}) have different values.  We thus first update the
feature names to explicitate that they originate from different
experiment and represent quantitation from different spectra using the
convenience function \Rfunction{updateFeatureNames}.  Note that
updating the names of one experiment would suffice here.

<<updateFnames, echo=TRUE, cache=FALSE>>=
head(featureNames(exp1))
exp1 <- updateFeatureNames(exp1)
head(featureNames(exp1))
head(featureNames(exp2))
exp2 <- updateFeatureNames(exp2)
head(featureNames(exp2))
@

The two experiments now share the same sample names and have different
feature names and will be combined along the row. Note that all
meta-data is correctly combined along the quantitation values.

<<comb1, echo=TRUE, cache=FALSE>>=
exp12 <- combine(exp1, exp2)
dim(exp1)
dim(exp2)
dim(exp12)
@

\subsection{Combine different samples}\label{sec:comb2}

Lets now create two \Rclass{MSnSet}s from the same raw data to
simulate two different independent experiments that share some
features.  As done previously (see section \ref{sec:feataggregation}),
we combine the spectra based on the proteins they have been identified
to belong to.  Features can thus naturally be named using protein
accession numbers.  Alternatively, if peptide sequences would have
been used as grouping factor in \Rfunction{combineFeatures}, then
these would be good feature name candidates.

<<make2exps2, echo=TRUE, cache=FALSE, tidy=FALSE>>=
set.seed(1)
i <- sample(length(itraqdata), 35)
j <- sample(length(itraqdata), 35)
exp1 <- quantify(itraqdata[i], reporters = iTRAQ4,
                 verbose = FALSE)
exp2 <- quantify(itraqdata[j], reporters = iTRAQ4,
                 verbose = FALSE)
exp1 <- droplevels(exp1)
exp2 <- droplevels(exp2)
table(featureNames(exp1) %in% featureNames(exp2))
exp1 <- combineFeatures(exp1,
                        groupBy = fData(exp1)$ProteinAccession)
exp2 <- combineFeatures(exp2,
                        groupBy = fData(exp2)$ProteinAccession)
head(featureNames(exp1))
head(featureNames(exp2))
@

The \Rfunction{droplevels} drops the unused \Robject{featureData}
levels.  This is required to avoid passing absent levels as
\Robject{groupBy} in \Rfunction{combineFeatures}.  Alternatively, one
could also use \Robject{factor(fData(exp1)\$ProteinAccession)} as
\Robject{groupBy} argument.

The feature names are updated automatically by
\Rfunction{combineFeatures}, using the \Robject{groupBy} argument.
Proper feature names, reflecting the nature of the features (spectra,
peptides or proteins) is critical when multiple experiments are to be
combined, as this is done using common features as defined by their
names (see below).

\bigskip

Sample names should also be updated to replace anonymous reporter
names with relevant identifiers; the individual reporter data is
stored in the \Robject{phenoData} and is not lost.  A convenience
function \Rfunction{updateSampleNames} is provided to append the
\Rclass{MSnSet}'s variable name to the already defined names,
although in general, biologically relevant identifiers are preferred.

<<renameSamples, echo=TRUE, cache=FALSE>>=
sampleNames(exp1)
exp1 <- updateSampleNames(exp1)
sampleNames(exp1)
sampleNames(exp1) <- c("Ctrl1", "Cond1", "Ctrl2", "Cond2")
sampleNames(exp2) <- c("Ctrl3", "Cond3", "Ctrl4", "Cond4")
@

At this stage, it is not yet possible to combine the two experiments,
because their feature data is not compatible yet; they share the same
feature variable labels, i.e. the feature data column names
(\Sexpr{paste(head(fvarLabels(exp1), n = 3), collapse=", ")}, \ldots),
but the part of the content is different because the original data was
(in particular all the spectrum centric data: identical peptides in
different runs will have different retention times, precursor
intensities, \ldots).  Feature data with identical labels (columns in
the data frame) and names (row in the data frame) are expected to have
the same data and produce an error if not conform.

<<fdatanames, echo=TRUE, cache=FALSE>>=
stopifnot(all(fvarLabels(exp1) == fvarLabels(exp2)))
fData(exp1)["BSA", 1:4]
fData(exp2)["BSA", 1:4]
@

Instead of removing these identical feature data columns, one can use
a second convenience function, \Rfunction{updateFvarLabels}, to update
feature labels based on the experiements variable name and maintain
all the metadata.

<<renameFvars, echo=TRUE, cache=FALSE>>=
exp1 <- updateFvarLabels(exp1)
exp2 <- updateFvarLabels(exp2)
head(fvarLabels(exp1))
head(fvarLabels(exp2))
@

It is now possible to combine \Robject{exp1} and \Robject{exp2},
including all the meta-data, with the \Rfunction{combine} method.  The
new experiment will contain the union of the feature names of the
individual experiments with missing values inserted appropriately.

<<combine, echo=TRUE, cache=FALSE>>=
exp12 <- combine(exp1, exp2)
dim(exp12)
pData(exp12)
exprs(exp12)[25:28, ]
exp12
@

\bigskip

In summary, when experiments with different samples need to be
combined (along the columns), one needs to (1) clarify the sample
names using \Rfunction{updateSampleNames} or better manually, for
biological relevance and (2) update the feature data variable labels
with \Rfunction{updateFvarLabels}.  The individual experiments (there
can be more than 2) can then easily be combined with the
\Rfunction{combine} method while retaining the meta-data.

If runs for the same sample (different fractions for example) need to
be combines, one needs to (1) differentiate the feature provenance
with \Rfunction{updateFeatureNames} prior to use \Rfunction{combine}.

\subsection{Splitting and unsplitting \Rclass{MSnSet} instances}\label{sec:split}

A single \Rclass{MSnSet} can also be split along the features/rows or
samples/columns using the \Rfunction{split} method and a factor
defining the splitting groups, resulting in an instance of class
\Rclass{MSnSetList}:

<<split>>=
data(dunkley2006)
head(pData(dunkley2006))
split(dunkley2006, dunkley2006$replicate)
## or, defining the appropriate annotation variable name
dun <- split(dunkley2006, "replicate")
@

Above, we split along the columns/samples, but the function would
equally work with a factor of length equal to the number of rows of
the \Rclass{MSnSet} (or a feature variable name) to split along the
rows/features.

Finally, the effect of \Rfunction{split} can be reverted by
\Rfunction{unsplit}.

<<unsplit>>=
dun2 <- unsplit(dun, pData(dunkley2006)$replicate)
compareMSnSets(dunkley2006, dun2)
@

See \Rfunction{?MSnSetList} for more details about the class,
\Rfunction{split} and \Rfunction{unsplit} and comments about storing
multiple assays pertaining the same experiment.

\subsection{Averaging \Rclass{MSnSet} instances}\label{sec:avg}

It is sometimes useful to average a set of replicated experiments to
facilitate their visualisation. This can be easily achieved with the
\Rfunction{averageMSnSet} function, which takes a list of valid
\Rclass{MSnSet} instances as input and creates a new object whose
expression values are an average of the original values. A value of
dispersion (\Robject{disp}) and a count of missing values (\Robject{nNA}) is
recorded in the feature metadata slot. The average and dispersion are
computed by default as the median and (non-parametric) coefficient of
variation (see \Rfunction{?npcv} for details), although this can easily be
parametrised, as described in \Rfunction{?averageMSnSet}.

The next code chunk illustrates the averaging function using three
replicated experiments from \cite{Tan2009} available in the
\Biocpkg{pRolocdata} package.

<<avg>>=
library("pRolocdata")
data(tan2009r1)
data(tan2009r2)
data(tan2009r3)
msnl <- MSnSetList(list(tan2009r1, tan2009r2, tan2009r3))
avgtan <- averageMSnSet(msnl)
head(exprs(avgtan))
head(fData(avgtan)$disp)
head(fData(avgtan)$nNA)
@

We are going to visualise the average data on a principle component
(PCA) plot using the \Rfunction{plot2D} function from the
\Biocpkg{pRoloc} package \cite{Gatto2014}. In addition, we are going
to use the measure of dispersion to highlight averages with high
variability by taking, for each protein, the maximum observed
dispersion in the 4 samples. Note that in the default implementation,
dispersions estimated from a single measurement (i.e. that had 2
missing values in our example) are set to 0; we will set these to the
overal maximum observed dispersion.

\begin{figure}[!ht]
\centering
<<avg2, dev='pdf', fig.height=5, fig.width=5>>=
disp <- rowMax(fData(avgtan)$disp)
disp[disp == 0] <- max(disp)
range(disp)
library("pRoloc")
plot2D(avgtan, cex = 3 * disp)
@
\caption{PCA plot of the averaged \Rclass{MSnSet}. The point sizes
  are proportional to the dispersion of the protein quantitation
  across the averaged data.}
\label{fig:plot2Davg}
\end{figure}

\clearpage
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
\section{MS$^E$ data processing}\label{sec:mse}

\Biocpkg{MSnbase} can also be used for MS$^E$ data independent
acquisition from Waters instrument.  The MS$^E$ pipeline depends on
the Bioconductor \Biocpkg{synapter} package \cite{Bond2013} that
produces \Rclass{MSnSet} instances for indvidual acquisitions. The
\Biocpkg{MSnbase} infrastructure can subsequently be used to further
combine experiments, as shown in section \ref{sec:comb2} and apply
\textit{top3} quantitation using the \Rfunction{topN} method.

%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% Section
%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%%
%% \section{Conclusion}\ref{sec:ccl}

\section{Session information}\label{sec:sessionInfo}
<<sessioninfo, results='asis', echo=FALSE, cache=FALSE>>=
toLatex(sessionInfo())
@

\bibliography{MSnbase}

\end{document}

