% -*- mode: noweb; noweb-default-code-mode: R-mode; -*-
%\VignetteIndexEntry{simpleaffy primer}
%\VignetteKeywords{Analysis, Affymetrix}
%\VignetteDepends{simpleaffy, affy}
%\VignettePackage{simpleaffy}
%documentclass[12pt, a4paper]{article}
\documentclass[12pt]{article}
\usepackage{amsmath}
\usepackage{hyperref}
\usepackage[authoryear,round]{natbib}
\textwidth=6.2in
\textheight=8.5in
%\parskip=.3cm
\oddsidemargin=.1in
\evensidemargin=.1in
\headheight=-.3in
\newcommand{\scscst}{\scriptscriptstyle}
\newcommand{\scst}{\scriptstyle}
\newcommand{\Rfunction}[1]{{\texttt{#1}}}
\newcommand{\Robject}[1]{{\texttt{#1}}}
\newcommand{\Rpackage}[1]{{\textit{#1}}}
\author{Crispin J Miller}
\begin{document}
\title{Description of simpleaffy: easy analysis routines for Affymetrix data}
\maketitle
\tableofcontents
\section{Introduction}
The \Rpackage{simpleaffy} package is part of the
Bioconductor\footnote{\url{http://www.bioconductor.org/}} project. It
it was written to provide a starting point for exploring
Affymetrix data, and to provide functions for some of the most common
tasks we found ourselves doing over and over again. It is based on the
\Rpackage{affy} package, which does does most of the hard work.
\Rpackage{affy} provides a variety of functions for processing
Affymetrix data, with many more in \Rpackage{affycomp}. Even so, some
tasks (such as computing t-tests and fold changes between replicate
groups, plotting scatter-plots and generating tables of annotated
'hits') require a bit of coding, and some of the most commonly used
functions can be a bit slower than we would like. This package aims to
provide high-level methods to perform these routine analysis tasks,
and many of them have been re-implemented in C for speed.
Since \Rpackage{simpleaffy} is written over the top of the
\Rpackage{affy} package, a basic understanding of the library and its
vignette is a good idea. We also assume that the reader knows how the
Affymetrix system works. If not, a brief introduction can be found at
{\url http://bioinf.picr.man.ac.uk/}; a more detailed description is
in the Affymetrix MAS manual at {\url http://www.affymetrix.com}.
\section{Reading in data and generating expression calls }
\label{sec:get.started} The first thing you need to do is to get R to use
the \Rpackage{simpleaffy} package by telling it to load the library:
<<>>=
library(simpleaffy) ##the affy package also gets loaded
library(affy)
@
(NB: The examples in the vignette are hypothetical - we are putting
together a data package containing a complete experimental dataset,
when that's done, we will use these as the basis for the example code
in this document).
Affymetrix data is generated by processing an image of the microarray
(stored in a .DAT file) to produce a .CEL file, which contains, for
each probe on the array, a single number defining its intensity. These
are combined for each {\it probeset} using an algorithm such as RMA or
MAS 5.0, to generate an expression level call for each transcript
represented on the chip. Both the \Rpackage{affy} and
\Rpackage{simpleaffy} packages work on the data in .CEL files, so we
need to load them into R. In order to compute means, fold changes and
t-tests, \Rpackage{simpleaffy} needs to know about the replicates in
your experiment, so we must also load some descriptive data that says
which arrays were replicates and also something about the different
experimental conditions you were testing. This means that
\Rpackage{simpleaffy} needs {\it two things}: \begin{enumerate} \item
your .CEL files, and \item a white-space delimited file describing the
samples that went on them. \end{enumerate}
By default, this file is called {\it covdesc}. The first column should
have no header, and contains the names of the .CEL files you want to
process. Each remaining column is used to describe something in the
experiment you want to study. For example you might have a set of
chips produced by treating a cell line with two drugs. Your {\it
covdesc} file might look like something like this:
\begin{tabular}{rr}
& treatment \\
ctrl1.cel& n \\
ctrl2.cel& n \\
ctrl3.cel& n \\
a1.cel& a \\
a2.cel& a \\
a3.cel& a \\
b1.cel& b \\
b2.cel& b \\
b3.cel& b \\
ab1.cel& a.b \\
ab2.cel& a.b
\end{tabular}
Sooo, the easiest way to get going is it:
\begin{enumerate}
\item Create a directory, move all the relevant
{\it CEL} files to that directory
\item Create a {\it covdesc} file and put it in the same directory
\item If using linux/unix, start R in that directory.
\item If using the Rgui for
Microsoft Windows make sure your working directory contains the {\it
Cel} files (use ``File -> Change Dir'' menu item).
\item Load the library.
<<>>=
library(simpleaffy) ##load the simpleaffy package
##(which loads the affy package too)
@
\item Read in the data and generate expression calls, (using RMA), for example.
\begin{Sinput}
R> library(simpleaffy)
R> raw.data <- read.affy() ##read data in working directory
R> x.rma <- call.exprs(raw.data,"rma")
R> # alternatively, use MAS 5.0: x.mas <- call.exprs(raw.data,"mas5")
\end{Sinput}
\end{enumerate}
Take a look at the help files for a more detailed description of these functions (i.e. ?read.affy or ?call.exprs).
The function \Rfunction{justMAS} provides a faster implementation of
the MAS 5.0 expression summary algorithm written in C. (described in:
Hubbell et al. (2002) Robust Estimators for expression analysis. Bioinformatics 18(12) 1585-1592), and in Affymetrix's 'Statistical Algorithms Description
Document' that can be found on their website at
\url{http://www.affymetrix.com}). As with any implementation of an
algorithm, variations can occur. The simpleaffy website,
\url{http://bioinf.picr.man.ac.uk/simpleaffy}, describes what testing
was done: you should be aware of these differences, and if in any
doubt, use MAS5.0 or GCOS to generate your data.
By default,
\Rfunction{justMAS} is used by the \Rfunction{call.exprs} to generate the MAS5 calls, to use the
\Rfunction{expresso} version specify 'mas5-R' instead of 'mas5' when
you invoke \Rfunction{call.exprs}.
\Rfunction{justMAS} has not been tested on every chip type available -
the majority of development has been on HGU95A arrays and newer. For
more details of what chips it has been tested on, how the testing was
done, and the results of the comparisons, see our website:
\url{http://bioinformatics.picr.man.ac.uk/simpleaffy}.
\section{Quality Control}
One of the nice things about the Affymetrix platform is the collection
of QC metrics (and accompanying guidelines) that are available. These
may help flag up arrays that may have problems, and are described in
detail in the Affymetrix {\it Data Analysis Fundamentals} manual,
which can be found on their website at {\url http://www.affymetrix.com}.
Also look at the document 'QC and simpleaffy', which accompanies this
one. It discusses QC metrics in significantly more detail than
this vignette. A brief introduction can be found below...
The function \Rfunction{qc} generates the most commonly used metrics:
\begin{enumerate}
\item Average background
\item Scale factor
\item Number of genes called present (see the note on detection p-values) described below.
\item 3' to 5' ratios
\end{enumerate}
All of these stats are parameters computed for/from the MAS 5.0
algorithm. Affy's QC requires that the Scale Factor for all chips are
within 3-fold of one another, and that the average background and
percent present calls for each chip are 'similar'. Affy chips also
use probes at the 3' and 5' ends of the (generally) GAPDH and
beta-actin genes to measure RNA quality, and additional probes spiked
in during the latter stages of the sample preparation process are used
to verify hybridisation efficiency. These probes (BioB,BioC,BioD,and
CreX) should be present in increasing intensity.
The function \Rfunction{qc} produces an object of class 'QCStats'
containing QC metrics for each array in a project. It takes both
processed and raw data, since it makes use of numbers generated during
the mas 5.0 expression calling algorithm. Alternatively, it can be
called with just the raw data, in which case it calls
\Rfunction{call.exprs(x,"mas5")} internally. For example,
\begin{Sinput}
R> x <- read.affy("covdesc");
R> x.mas5 <- call.exprs(x,"mas5");
R> qcs <- qc(x,x.mas5);
\end{Sinput}
Note that we have used \Rfunction{call.exprs(x,"mas5")} to normalise the
data using the mas 5.0 algorithm. This is important because
\Rfunction{call.exprs} stores some additional data in the AffyBatch object's
\Rfunction{description@preprocessing} slot:
\begin{Sinput}
R> x.mas5@description@preprocessing
\end{Sinput}
\Rfunction{qc} returns an object containing scale-factors, \% present,
average, mininimum, maximum and mean background intensities,and bioB,
bioC, bioD and creX present calls (1=present;0=not present). It also
stores 3', 5' and M values for the QC probes. \Rfunction{ratios(qc)}
generates a table of qc ratios for these probes. See ?qc for more details.
For some arrays, there are more than one probeset that target the
gapdh and beta-actin genes. In this situation, we've attempted to make
a sensible choice as to which probeset to use. To find out which
probesets are used for your arrays use the methods \Rfunction{qc.get.ratios},
\Rfunction{qc.get.probes} and \Rfunction{qc.get.spikes}.
The data file \Rfunction{qcs} contains an example QCStats object for a
25 array project comparing different protocols for processing MCF7 and
MCF10A data. This dataset is interesting because some of the arrays
have poor qc.
<<>>=
data(qcs)
@
The QCStats object (returned by qc), or what we just loaded with
\Rfunction{data}, has a number of accessor functions to get at its values:
<<results=hide>>=
ratios(qcs)
avbg(qcs)
maxbg(qcs)
minbg(qcs)
spikeInProbes(qcs)
qcProbes(qcs)
percent.present(qcs)
plot(qcs)
sfs(qcs)
target(qcs)
ratios(qcs)
@
Many of these also have 'set' methods - see section \ref{QCconfig} for more details.
The qc functions and detection p value code have not been tested on every
chip type available - the majority of development has been on HGU95A
arrays and newer. Again, for more details of what chips it has been tested
on, how the testing was done, and the results of the comparisons, see
the simpleaffy website.
A plot of qc data can also be obtained by \Rfunction{plot(qc)}.
<<fig=TRUE>>=
plot(qcs)
@
Each array is represented by a seperate line in the figure. The
central vertical line corresponds to 0 fold change, the dotted lines
on either side correspond to up and down regulation. The blue
bar represents the region in which all arrays have scale factors
within, by default, three-fold of each other. Its position is found by
calculating the mean scale factor for all chips and placing the center
of the region such that the borders are -1.5 fold up or down from the
mean value.
Each array is plotted as a line from the 0-fold line to the point
that corresponds to its scale factor. If the ends of all of the lines
are in the blue region, their scale-factors are compatible. The lines
are coloured blue if OK, red if not.
The figure also shows (for most arrays) GAPDH and beta-actin 3\'/5\'
ratios. These are represented as a pair of points for each chip. Affy
state that beta actin should be within 3, gapdh around 1. Any that
fall outside these thresholds (1.25 for gapdh) are coloured red; the
rest are blue.
Written along the left hand side of the figure are the number of genes called
present on each array and the average background. These will vary
according to the samples being processed, and Affy's QC suggests
simply that they should be similar. If any chips have significantly
different values this is flagged in red, otherwise the numbers are
displayed in blue. By default, 'significant' means that \%-present are
within 10\% of each other; background intensity, 20 units. These last
numbers are somewhat arbitrary and may need some tweaking to find
values that suit the samples you're dealing with, and the overall
nature of your setup.
Finally, if BioB is not present on a chip, this will be flagged by
printing 'BioB' in red.
In short, everything in the figure should be blue - red highlights a problem!
\section{Filtering by expression measures}
When R loaded the .CEL files, it also used the data in the covdesc
file to define which experimental groups the chips belonged to. The
\Rfunction {get.array.subset} function makes it easy to select a
subest of arrays from the experiment. For example,
\begin{Sinput}
R> get.array.subset(x.rma,"treatment",c("a","n"))
\end{Sinput}
will return an \Robject{ExpressionSet} containing just the chips
corresponding to treatment with drug 'a' or or with no drug at all.
The function \Rfunction{pairwise.comparison} allows you to take a
subset of chips and perform the following analyses on it:
\begin{enumerate}
\item find means of the data,
\item compute log2 fold changes between the means,
\item compute a t-test between the groups,
\item possibly compute MAS5.0 style detection {\it p} values and Present/Marginal/Absent calls.
\end{enumerate}
it returns a 'PairComp' object containing the results of the analysis:
\begin{Sinput}
R> results <- pairwise.comparison(
x.rma, ## processed data
"treatment", ## the factor in covdesc
## to use to group arrays
c("n","a"), ## groups to compare
raw.data ## for PMA calls
)
\end{Sinput}
This function is implemented in C for speed - and does everything in
one go. There are no individual functions for fold-change and p-value,
because it works out quicker simply to get everything and to discard
the stuff you don't need.
Note that detection p values are computed using the function
\Rfunction{detection.p.val}. This makes use of two parameters (alpha1
and alpha2) that are, like the control probes described above,
dependent on the array type you're analysing. Use
\Rfunction{getAlpha1} and \Rfunction{getAlpha2} to find out what values are being used for your array.
If you only have two groups of replicates - a simple control
v. treatment experiment, for example, things are even easier, you do
not have to specify the members to compare since the function can work it out
from the group:
\begin{Sinput}
R> results <- pairwise.comparison(
x.rma, ## processed data
"treatment", ## the factor in covdesc
spots=raw.data ## for PMA calls
)
R> ## Find the 100 most changing genes
R> sort(abs(fc(results)),decreasing=TRUE)[1:100]
\end{Sinput}
Since \Rfunction{call.exprs} {\it always} returns logged data, \Rfunction{pairwise.comparison}
expects logged data by default -- this can be changed with the \Rfunction{logged} parameter.
Averages can be calculated 3 ways:
\begin{enumerate}
\item From the unlogged values -- i.e. log2(mean(replicates)).
\item From the logged values -- i.e. mean(log2(replicates)).
\item From the median -- i.e log2(median(replicates))
\end{enumerate}
..and then the log2(fold change) values are simply worked out from these.
By default, unlogged values are used, but the parameter, \Rfunction{method},
allows you to change this by specifying method='unlogged','logged' or 'median'.
{\it Note} that if you have no replicates in one or both of your
experimental groups - so that you have only one control or treatment
chip, for example, the function returns a p-score of 0.0 for
each t-test comparison {\it rather than generating an error} complaining that
there are not enough observations.
The function \Rfunction{pairwise.filter} takes the output of
\Rfunction{pairwise.comparison} and filters it for significantly
changing genes:
\begin{Sinput}
R> # find genes expressed with an intensity
R> # greater than 10 on at least 6 chips,
R> # that show a fold change greater than 1.5
R> # and are significantly different
R> # between groups with a t-test p.value
R> # of 0.001 or better
R> significant <- pairwise.filter(
results,
min.exp=log2(10),
min.exp.no=6, fc=log2(1.5),
tt= 0.001)
\end{Sinput}
If the pairwise comparison object was created from MAS 5.0 data, it
can be filtered by Present Marginal Absent calls. Each gene is tested
to see how many arrays it is called present on and this value used to
decide whether the transcript passes the filter. Two ways of doing
this are possible: the first simply sees if the transcript is Present
on more than \Rfunction{min.present.no} arrays in the experiment. To
do this, the parameter \Rfunction{present.by.group} must be set to
FALSE.
The second makes use of the replicate groups used to calculate the
\Rfunction{pairwise.comparison} object. In this case, the transcript
must be present by more than \Rfunction{min.present.no} arrays in at
least one of the two experimental groups. For this
\Rfunction{present.by.group} must be TRUE. This is useful because it
provides a way of trying to identify transcripts that have gone from
'off' to 'on', i.e. from being called Absent in one set to Present in
the other, as well as those that are Present on all arrays.
For both methods of filtering, if \Rfunction{min.present.no="all"} is
specified instead of a number , the transcript must be present in
"all" arrays of a replicate group (or the whole experiment, depending
on \Rfunction{present.by.group}).
For example:
\begin{Sinput}
R> # find genes present on all chips,
R> # that show a fold change greater than 1.5
R> # and are significantly different
R> # between groups with a t-test p.value
R> # of 0.001 or better
R> significant <- pairwise.filter(
results,
min.present.no="all",
present.by.group=F, fc=log2(1.5),
tt= 0.001)
\end{Sinput}
\begin{Sinput}
R> # find genes present on all chips in at least one replicate group,
R> # that show a fold change greater than 1.5
R> # and are significantly different
R> # between groups with a t-test p.value
R> # of 0.001 or better
R> significant <- pairwise.filter(
results,
min.present.no="all",
present.by.group=T, fc=log2(1.5),
tt= 0.001)
\end{Sinput}
\subsection{Paired replicates}
In the above example, we simply lumped all the replicates together
when we calculated the t-tests. Ideally, we would like to design
experiments so that each treatment is matched by a control sample that
mirrors the protocols and processes it was subjected to as closely as
possible.
This is particularly important if we are going to be processing
replicates on separate days, or perhaps with different operators. A
protocol that requires incubation on the bench for a couple of hours,
say, might behave differently on a hot day and we would
like the control samples to pick this sort of thing up.
The \Rfunction{pairwise.comparison} function lets us do a t-test with
paired replicates, by specifying the ordering of replicates within
groups.
\begin{Sinput}
R> results <- pairwise.comparison(
x.rma, ## processed data
"treatment", c("n","a"),## groups to compare
raw.data,
a.order=c(1,3,2), ## a.1 matches b.1
b.order=c(1,2,3)) ## a.3 matches b.2 etc...
R> ## Find the 100 most changing genes
R> sort(abs(fc(results)),decreasing=TRUE)[1:100]
\end{Sinput}
Here, we have specified the order the replicate samples should be
compared to each other - so that replicate 3 in group a is compared to
replicate 2 in group b, and so on.
Again, the help pages for these functions explain more about them and
the values they return.
\section{Viewing results}
The function \Rfunction{trad.scatter.plot} does a scatter plot between a pair of vectors:
\begin{Sinput}
R> trad.scatter.plot(exprs(x.rma)[,1],exprs(x.rma)[,3],
fc.line.col="lightblue",col="blue");
R> trad.scatter.plot(exprs(x.rma)[,2],exprs(x.rma)[,4],
add=T,col="red");
R> legend(2,12,c("Control v. treatment rep 1","Control v. treatment rep 2"),
col=c("blue","red"),pch=20)
\end{Sinput}
Generic plot functions also exist for \Rfunction{pairwise.comparison} objects.
\Rfunction{plot(pairwise.comparison)} will do a straight scatter plot
of the means of the two replicate groups. In addition the parameter \Rfunction{type} can be used:
\begin{Sinput}
R> plot(results,type="scatter") #scatter plot
R> plot(results,type="ma") #M v A plot
R> plot(results,type="volcano") #volcano plot
\end{Sinput}
If PMA calls are available, the function will colour the points as follows:
\begin{enumerate}
\item Red -- all present
\item Orange -- all present in one group or the other
\item Yellow -- all that remain
\end{enumerate}
If a second \Rfunction{pairwise.comparison} object is supplied, then these points are drawn (by default) as blue circles. This allows the results of a pairwise.filter to be identified on the graph - eg.:
\begin{Sinput}
R> # find genes present on all chips in at least one replicate group,
R> # that show a fold change greater than 1.5
R> # and are significantly different
R> # between groups with a t-test p.value
R> # of 0.001 or better
R> results <- pairwise.comparison(
x.rma, ## processed data
"treatment", c("n","a"),## groups to compare
raw.data,
a.order=c(1,3,2), ## a.1 matches b.1
b.order=c(1,2,3)) ## a.3 matches b.2 etc...
R> significant <- pairwise.filter(
results,
min.present.no="all",
present.by.group=T, fc=log2(1.5),
tt= 0.001)
R> plot(results,significant)
\end{Sinput}
\subsection{Heatmaps}
Simpleaffy also has a pair of functions for plotting simple heatmaps,
\Rfunction{hmap.eset} and \Rfunction{hmap.pc} for AffyBatch and
PairComp objects, respectively. The simplest use is as follows:
\begin{Sinput}
R> #generate a heatmap of the first 100 genes in an expression set.
R> eset <- read.affy()
R> eset.rma <- call.exprs(eset)
R> hmap.eset(eset.rma,1:100)
\end{Sinput}
You can specify colours using the 'col' parameter either by giving the function a vector of colors (e.g. rainbow(21)) or by the following strings:
\begin{enumerate}
\item{``bwr'' -- from blue to red via white }
\item{``rbg'' -- from red to green via black }
\item{``ryw'' -- from red to white via yellow }
\end{enumerate}
By default, colouring is such that the colours are scaled so that all
the data in the heatmap fits in the colour range
supplied. Alternatively, minimum and maximum values for the colour
range can be specified. Clustering functions or trees generated by
hclust or dendrogram can also be provided -- by default 1-Pearson correlation
is used. See ?hmap.eset for more details.
If you have the results of a pairwise.filter, you can use this to
select the samples and probesets to plot:
\begin{Sinput}
R> #generate a heatmap of the first 100 probesets in an expression set.
R> eset <- read.affy()
R> eset.rma <- call.exprs(eset)
R> pc <- pairwise.comparison(eset.rma,''group'',c(``a'',''b''))
R> pc.f <- pairwise.filter(pc,tt=0.001,fc=1)
R> hmap.pc(pc.f,eset.rma)
\end{Sinput}
By default, this scales the colouring for each probeset in terms of its standard deviation. This is done as follows:
\begin{enumerate}
\item{The replicate groups used for the pairwise comparison are found. The s.d. for each probeset is calculated for each group, and the results averaged to give a mean standard deviation for each probeset.}
\item{Data are clustered as before}
\item{Each probeset is scaled by it standard deviation, and coloured according to how many s.d.s it is from its mean.}
\item{The samples used for the pariwise comparison are plotted (by default)}
\end{enumerate}
The parameter 'scale' can be used to turn this scaling off, and spread
defines how many standard deviations above and below the mean should
be shown. see ?hmap.pc for more details.
\subsection{Printing}
Another thing that can sometimes be tricky is producing figures for
papers and presentations. Two utility functions,
\Rfunction{journalpng} and \Rfunction{screenpng} make it easy to
generate .png files at 300dpi (huge) and 72dpi. Most journals accept
.pngs, and they can be converted into other formats using a
decent graphics package.
R uses the concept of a {\it device} to deal with graphics. When you
start to plot a graph, it looks for a graphics device to print it
on. If it can't find one that's already there, it opens a new one,
which by default corresponds to a window on the screen. In order to
generate a {\it file} (rather than a window) containing our figure, we
can use \Rfunction{journalpng} and \Rfunction{screenpng} to
create a new graphics device that R can use instead. This device
results in our graph being plotted to a file on disk - when you type
something like:
\begin{Sinput}
R> journalpng(file="results/figure1.png");
R> trad.scatter.plot(exprs(x.rma)[,1],exprs(x.rma)[,3],fc.line.col="lightblue");
R> dev.off();
\end{Sinput}
a new png file is created, the scatterplot is printed into that file
and then, when dev.off() is called, the data is saved. Note that
because R is plotting to our \Rfunction{journalpng} device rather than
the screen, we won't see any pictures (or anything happen at all).
\subsection{Generating a table of results}
It would also be nice to know what our changing genes actually do. The
function \Rfunction{get.annotation} takes a list of probe set names
along with a string specifying the type of array we are looking at,
and uses these to look up annotation for our data. The resulting
dataframe, when saved as a tab delimited .XLS file, loads into excel,
with hyperlinks to the NCBI's Unigene and LocusLink databases.
Two functions \Rfunction{results.summary} and
\Rfunction{write.annotation} are also useful - the former generates a
summary table with expression data and associated annotation, the
latter spits it out in tab delimited format (see
\Rfunction{?get.annotation} for more details):
\begin{Sinput}
R> x <- read.affy()
R> x.rma <- call.exprs(x,"rma")
R> pw <- pairwise.comparison(x.rma,"drug",c("formulaX","nothing"))
R> pw.filtered <- pairwise.filter(pw)
R> summary <- results.summary(pw.filtered,"hgu133a")
R> write.annotation(file="spreadsheet.xls",summary)
\end{Sinput}
\subsection{ Specifying alternate QC configurations } \label{QCconfig}
If when running \Rfunction{qc}, one of the other qc functions, or
\Rfunction{detection.p.val}, you get an error because simpleaffy
doesn't recgnize the type of array in your project, or because, for example, you are using custom arrays, you need to
specify your own QC parameters. These include alpha1 and alpha2 values
for the array, the name of the spike control probesets and the pairs
of probesets used to calculate the 5'/3' ratios. These can be then specified in a config file. This, for example, is the configuration for the hgu133 plus2 array:
\begin{Sinput}
array hgu133plus2cdf
alpha1 0.05
alpha2 0.065
spk bioB AFFX-r2-Ec-bioB-3_at
spk bioC AFFX-r2-Ec-bioC-3_at
spk bioD AFFX-r2-Ec-bioD-3_at
spk creX AFFX-r2-P1-cre-3_at
ratio actin3/actin5 AFFX-HSAC07/X00351_3_at AFFX-HSAC07/X00351_5_at
ratio actin3/actinM AFFX-HSAC07/X00351_3_at AFFX-HSAC07/X00351_M_at
ratio gapdh3/gapdh5 AFFX-HUMGAPDH/M33197_3_at AFFX-HUMGAPDH/M33197_5_at
ratio gapdh3/gapdhM AFFX-HUMGAPDH/M33197_3_at AFFX-HUMGAPDH/M33197_M_at
\end{Sinput}
The file must be called <cdfname>.qcdef, where cdfname is the 'clean' cdfname
produced by calling \Rfunction{cleancdfname} on the \Rfunction{cdfName} of the appropriate AffyBatch
object. For example:
\begin{Sinput}
R> n <- cdfName(eset)
R> cleancdfname(n)
\end{Sinput}
<<echo=FALSE>>=
cleancdfname("HGU_133_plus_2")
@
The first token on each line specfies what the rest of the line
describes. Allowable tokens are: 'array' (array name), 'alpha1',
'alpha2' (alpha values), 'spk' (the name of a spike probeset -- must be
followed by one of 'bioB','bioC', 'bioD' or 'creX'), or 'ratio' (to
specify pairs to comput 3'/5' or 3'/M metrics). 'ratio' must be
followed by the name of the ratio, which must be of the form
'name1/name2', where name1 and name2 are textual identifiers suitable
for displaying in the qc plot, for example. This must then be followed by two probeset
ids.
Once the file is created, the QC environment can be set up with a call
to \Rfunction{setQCEnvironment(array,path)} specifying the appropriate
cdfname and the path to the directory
containing the file.
MORE on changes to follow...
\bibliography{simpleaffy}
\end{document}