---
title: "An end to end workflow for differential gene expression using Affymetrix microarrays"
author: 
  - name: Bernd Klaus
    affiliation: EMBL Heidelberg, Meyerhofstrasse 1, 69117 Heidelberg, Germany, bernd.klaus@embl.de
  - name: Stefanie Reisenauer
    affiliation: EMBL Heidelberg, Meyerhofstrasse 1, 69117 Heidelberg, Germany, steffi.reisenauer@tum.de
header-includes:
  - \usepackage{float}
abstract: In this article, we walk through an end-to-end Affymetrix microarray differential expression workflow using Bioconductor packages. This workflow is directly applicable to current "Gene" type arrays, e.g. the HuGene or MoGene arrays, but can easily be adapted to similar platforms. The data analyzed here is a typical clinical microarray data set that compares inflamed and non-inflamed colon tissue in two disease subtypes. For each disease, the differential gene expression between inflamed- and non-inflamed colon tissue was analyzed. We will start from the raw data CEL files, show how to import them into a Bioconductor ExpressionSet, perform quality control and normalization and finally differential gene expression (DE) analysis, followed by some enrichment analysis.
keywords: Microarray, Gene Expression
bibliography: MAEndToEnd.bib
output:
  BiocStyle::html_document:
    self_contained: yes
    toc: true
    toc_float: true
    toc_depth: 2
    code_folding: show
csl: f1000-research.csl
date: 14 September 2018
vignette: >
  %\VignetteIndexEntry{An end to end workflow for differential gene expression using Affymetrix microarrays}
  %\VignetteEngine{knitr::rmarkdown}
  %\VignetteEncoding{UTF-8} 
---
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```{r biocSetup, include = FALSE, results="hide", warning=FALSE, echo=FALSE}
## Changing the YAML to the following changes the output to 'latex'
## output:
## BiocWorkflowTools::f1000_article
## More at:
## https://stackoverflow.com/questions/35144130/in-knitr-how-can-i-test-for-if-the-output-will-be-pdf-or-word

## add figures to current directory if Rmd is converted to LaTeX
## This simplifies processing on overleaf

if (!is.null(knitr::opts_knit$get("rmarkdown.pandoc.to"))) {
    
    to_html <- knitr::opts_knit$get("rmarkdown.pandoc.to") != 'latex'
        if (!to_html) {
          knitr::opts_chunk$set(fig.path = "", out.width = '.8\\linewidth')
        } 

}
## here, pdf specific adapations can be made
## however, the option "rmarkdown.pandoc.to" is only set by rmarkdown::render,
## so it returns NULL if run interactively!

```

```{r testAlternativeToChunkAbove, eval=FALSE, include=FALSE, echo=FALSE}
# this does not work as it will only test the output formats
# specified in the YAML header
any(grepl("f1000", rmarkdown::all_output_formats(knitr::current_input()))) 
```


```{r knitrOptions, include=FALSE}
library(BiocStyle)
library(knitr)
options(digits = 4, width = 80)
opts_chunk$set(echo = TRUE, 
               tidy = FALSE, 
               include = TRUE,
               dev = c('png'),
               fig.width = 6, fig.height = 3.5,
               comment = '  ', 
               dpi = 300, cache = TRUE,
               fig.pos = "H")

```


<p>
**R version**: `r R.version.string`
<br />
**Bioconductor version**: `r BiocManager::version()`
<br />
**Package version**: `r packageVersion("maEndToEnd")`
</p>


# Introduction

In this article we introduce a complete workflow for a typical (Affymetrix)
microarray analysis. Data import, preprocessing, differential expression and
enrichment analysis are discussed.

The data set used [@Palmieri_2015] is from a paper studying the differences in 
gene expression in inflamed and non-inflamed tissue. 
14 patients suffering from Ulcerative colitis (UC) and 15 patients with Crohn's 
disease (CD) were tested, and from each patient inflamed and non-inflamed 
colonic mucosa tissue was obtained via a biopsy. 
This is a typical clinical data set consisting of 58 arrays in total.
Our aim is to analyze differential expression (DE) between the tissues. 
Our results show a substantial overlap with the results of the original paper.

# Workflow package installation

The workflow is wrapped in a package called `maEndToEnd`. 

The `maEndToEnd` package can currently be obtained from GitHub and is 
available via the current development version of Bioconductor (3.8) (see here: <http://bioconductor.org/packages/devel/workflows/html/maEndToEnd.html>).

## Workflow package installation from Bioconductor

You can install the package via the `r CRANpkg("BiocManager")`.

```{r installBioc, eval=FALSE}
if (!require("BiocManager"))
    install.packages("BiocManager")
BiocManager::install("maEndToEnd", version = "devel")
```

Currently, the workflow is available in the development version of Bioconductor
(3.8), which will become the release version in October 2018.

For details on how to use this version of Bioconductor see:

<http://bioconductor.org/developers/how-to/useDevel/>

## Workflow package installation from Github

```{r, echo=TRUE, results="hide", warning=FALSE, eval=FALSE}

#In order to download the package from GitHub, we need the "install_github" 
#function from the "remotes" package. We download the latest developer 
#version of "remotes" from GitHub with the devtool::install_github
#function; note that this is necessary as the current "remotes" version on
#CRAN doesn't allow us to correctly download the "maEndToEnd" package: 

install.packages("devtools")
library(devtools)

devtools::install_github("r-lib/remotes")
library(remotes)
packageVersion("remotes") # has to be 1.1.1.9000 or later

remotes::install_github("b-klaus/maEndToEnd", ref="master")

```

## Workflow package import

Once the workflow package has been successfully installed, we can use 
a call to `library()` in order to load it. This will also load
all the other packages neccessary to run the workflow.

```{r maEndToEndImport}
suppressPackageStartupMessages({library("maEndToEnd")})
```

## List of packages required for the workflow

Below, you find a list of packages that are required by the workflow.
Some Helper/Styling packages have been commented here, as they are
not strictly neccesary to execute the workflow.

```{r pkgList, results="hide"}
#General Bioconductor packages
    library(Biobase)
    library(oligoClasses)
     
#Annotation and data import packages
    library(ArrayExpress)
    library(pd.hugene.1.0.st.v1)
    library(hugene10sttranscriptcluster.db)
     
#Quality control and pre-processing packages
    library(oligo)
    library(arrayQualityMetrics)
     
#Analysis and statistics packages
    library(limma)
    library(topGO)
    library(ReactomePA)
    library(clusterProfiler)
     
#Plotting and color options packages
    library(gplots)
    library(ggplot2)
    library(geneplotter)
    library(RColorBrewer)
    library(pheatmap)
     
#Formatting/documentation packages
   #library(rmarkdown)
   #library(BiocStyle)
    library(dplyr)
    library(tidyr)

#Helpers:
    library(stringr)
    library(matrixStats)
    library(genefilter)
    library(openxlsx)
   #library(devtools)

```


# Downloading the raw data from ArrayExpress

The first step of the analysis is to download the raw data CEL files. These 
files are produced by the array scanner software and contain the measured probe intensities. The data we use have been deposited at [ArrayExpress](https://www.ebi.ac.uk/arrayexpress/)
and have the accession code **E-MTAB-2967**.

We will store these files in the directory **raw\_data\_dir** which defaults to 
a temporary directory.

```{r generateFolderForRawData}
raw_data_dir <- tempdir()

if (!dir.exists(raw_data_dir)) {
    dir.create(raw_data_dir)
}

```


Each ArrayExpress data set has a landing page summarizing the data set, 
and we use the `getAE`function from the `r Biocpkg("ArrayExpress") ` Bioconductor package to obtain 
the ftp links to the raw data files ([Data from Palmieri et. al. on ArrayEpress](https://www.ebi.ac.uk/arrayexpress/experiments/E-MTAB-2967/)).


With the code below, we download the raw data (also including annotation data) from
[ArrayExpress](https://www.ebi.ac.uk/arrayexpress/) [@Kolesnikov_2014] 
by using the `getAE`-function. The data are saved in the `raw_data_dir`
created above. The names of the downloaded files are returned as a list.


```{r getDataEBI, eval=TRUE, results='hide', message=FALSE}
anno_AE <- getAE("E-MTAB-2967", path = raw_data_dir, type = "raw")
```

We will now have a closer look at the data we downloaded from ArrayExpress

# Background information on the data

## Information stored in ArrayExpress

Each dataset at ArrayExpress is stored according to the MAGE-TAB 
(MicroArray Gene Expression Tabular) specifications as a collection of
tables bundled with the raw data. The MAGE-TAB format specifies up to five 
different types of files:


* **Investigation Description Format (IDF)**
* **Array Design Format (ADF)**
* **Sample and Data Relationship Format (SDRF)**
* **raw data files** 
* **processed data files**

Other than the raw data files, the IDF and the SDRF file are important for us. 
The IDF file contains top level information
about the experiment including title, description, submitter contact details 
and protocols. The SDRF file contains essential information on the experimental 
samples, e.g. the experimental group(s) they belong to. 

Before we move on to the actual raw data import, we will briefly introduce the
`ExpressionSet` class contained in the `r Biocpkg("Biobase")` package.
It is commonly used to store microarray data in Bioconductor.

## Bioconductor ExpressionSets

Genomic data can be very complex,
usually consisting of a number of different components, e.g. information
on the experimental samples, annotation of genomic features measured as well
as the experimental data itself.
In Bioconductor, the approach is taken that these components should be 
stored in a single structure to easily manage the data.

The package `r Biocpkg("Biobase")` contains standardized data structures
to represent genomic data. The `ExpressionSet` class is designed
to combine several different sources of information (i.e. as contained in the 
various MAGE-TAB files) into a single convenient
structure. An ExpressionSet can be manipulated (e.g., subsetted, copied),
and is the input to or output of many Bioconductor functions.

The data in an ExpressionSet consist of:

* **assayData**: Expression data from microarray experiments
with microarray probes in rows and sample identifiers in columns
* **metaData**
    + **phenoData**: A description of the samples in the experiment with sample identifiers in rows and description elements in columns; holds the content of the SDRF file
    + **featureData**: metadata about the features on the chip   
    or technology used for the experiment with same rows as assayData by default and freely assignable columns
    + further annotations for the features,
    for example gene annotations from biomedical databases (annotation).

* **experimentData**: A flexible structure to describe the experiment.

The ExpressionSet class coordinates all of these data, so that one does not 
have to worry about the details. However, one should keep in mind that the rownames of the `phenoData` have to match the column names of the assay data, while the row names of the assay data have to match the 
row names of the `featureData`. 
This is  illustrated in Figure \@ref(fig:sumexp).


```{r sumexp, fig.cap = "Structure of Bioconductor’s ExpressionSet class.", echo=FALSE, fig.show="asis", fig.width = 7, fig.height = 4.5}
par(mar=c(0,0,0,0))
plot(1,1,xlim=c(0,100),ylim=c(0, 150),bty="n",
     type="n",xlab="",ylab="",xaxt="n",yaxt="n")

polygon(c(50,85,85,50),c(70,70,130,130),col="lightcoral",border=NA)
polygon(c(55, 56, 56, 55),c(70,70,130,130),col=rgb(1,0,0,.5),border=NA)
polygon(c(50,85,85,50),c(118,118,120,120),col=rgb(1,0,0,.5),border=NA)
text(67.5,100,"assay(s)", cex = 1)
text(67.5,90,"e.g. 'exprs'", cex = 1)

polygon(c(45,49,49,45),c(70,70,130,130),col="honeydew2",border=NA)
text(47,100, "microarray probes", srt=90, cex=1)

polygon(c(50,85,85,50),c(131,131,140,140),col="darkseagreen3",border=NA)
text(67.5,135.5,"sample IDs", cex = 1)


polygon(c(20,40,40,20),c(70,70,130,130),col=rgb(0,0,1,.5),border=NA)
polygon(c(20,40,40,20),c(118,118,120,120),col=rgb(0,0,1,.5),border=NA)
text(30,100,"featureData", cex = 1)

polygon(c(15,19,19,15),c(70,70,130,130),col="honeydew2",border=NA)
text(17,100, "microarray probes", srt=90, cex=1)

polygon(c(20,40,40,20),c(131,131,140,140),col="bisque3",border=NA)
text(30,135.5,"features", cex = 1)

polygon(c(50,65, 65, 50),c(55, 55, 0, 0),col=rgb(.5,0,.5,.5),border=NA)
polygon(c(55, 56, 56, 55),c(55, 55, 0, 0),col=rgb(.5,0,.5,.5),border=NA)
text(57.5, 30,"phenoData", cex = 1, srt=270)

polygon(c(50,65, 65, 50),c(56,56,65,65),col="darkseagreen3",border=NA)
text(57.5,60.5,"sample IDs", cex = 1)
```

You can use the functions ` pData ` and ` fData ` to extract
the sample and feature annotation, respectively, from an ` ExpressionSet `.
The function ` exprs ` will return the expression data itself as a matrix.

# Import of annotation data and microarray expression data as "ExpressionSet"

We import the SDRF file with the `read.delim` function from the raw data 
folder in order to obtain the sample annotation.

The sample names are given in the column Array.Data.File of the SDRF data table and will be used as rownames for the SDRF file.

We turn the SDRF table into an `AnnotatedDataFrame` 
from the `r Biocpkg("Biobase")` package that we
will need later to create an `ExpressionSet` for our data [@Bioc].

```{r getSDRF}
sdrf_location <- file.path(raw_data_dir, "E-MTAB-2967.sdrf.txt")
SDRF <- read.delim(sdrf_location)

rownames(SDRF) <- SDRF$Array.Data.File
SDRF <- AnnotatedDataFrame(SDRF)
```


We now create the Expression Set object 
`raw_data`, which contains array data, pheno data
(from the SDRF file) as well as the information 
of the chip annotation package used.


The analysis of Affymetrix arrays starts with CEL files. These are the result
of the processing of the raw image files using the Affymetrix software 
and contain estimated probe intensity values. Each CEL file additionally 
contains some metadata, such as a chip identifier. 

We use the function `read.celfiles` from the `r Biocpkg("oligo") ` 
package [@oligo] to import the files:

```{r importCelfiles, results="hide", eval=TRUE, dependson="getSDRF", warning = FALSE }
raw_data <- oligo::read.celfiles(filenames = file.path(raw_data_dir, 
                                                SDRF$Array.Data.File),
                                    verbose = FALSE, phenoData = SDRF)
stopifnot(validObject(raw_data))
```

This automatically creates an ExpressionSet,
fills the sections "array data" with the data from the CEL files 
and uses the correct chip annotation package, in this case
`r Biocannopkg("pd.hugene.1.0.st.v1") ` (the 
chip-type is also stored in the .CEL files). 

Furthermore, we specified our `AnnotatedDataFrame` "`SDRF`" 
created earlier from the SDRF file as `phenoData`. Thus, 
we had to make sure to import the CEL files 
in the order that corresponds to the SDRF table --- to enforce this, 
we used the column `Array.Data.File` of the `SDRF` table as the `filenames` 
argument.

Finally, we checked whether the object created is valid (e.g. sample names 
match between the different tables).

We now have a first look on the raw data.

The `pData` function of the Biobase package directly accesses the phenoData in the ExpressionSet `raw_data`. With the `head()` function, we can view the 
first six lines of the table. We have a look at the columns included 
and retain only those columns that are related to the experimental 
factors of interest.

```{r inspectPhenoData, eval=TRUE }
head(Biobase::pData(raw_data))
```


The columns of interest for us are the following:

  * identifiers of the individuals, i.e. columns "Source.Name", "Characteristics.individual."
  * disease of the individual, i.e. "Factor.Value.disease."
  * mucosa type, i.e. "Factor.Value.phenotype."

We now subselect the corresponding columns: 

```{r reassignpData, eval=TRUE}
Biobase::pData(raw_data) <- Biobase::pData(raw_data)[, c("Source.Name",
                                     "Characteristics.individual.",
                                     "Factor.Value.disease.",
                                     "Factor.Value.phenotype.")]
```

# Quality control of the raw data

The first step after the initial data import is the quality control of the data.
Here we check for outliers and try to see whether the data clusters as expected, 
e.g. by the experimental conditions. The expression intensity values are in the 
assayData sub-object "exprs" and can be accessed by the `exprs(raw_data)` 
function. The rows represent the microarray probes, i.e. 
the single DNA locations on the chip, while the columns 
represent one microarray, i.e. a sample of inflamed 
and non-inflamed tissue of every patient, respectively. 

```{r inspectAssayData, eval=TRUE}
Biobase::exprs(raw_data)[1:5, 1:5]
```


For quality control, we take the log2 of `Biobase::exprs(raw_data)`, 
as expression data is commonly analyzed on a logarithmic scale.

We then perform a principal component analysis (PCA) and plot it (Figure 
\@ref(fig:qualityControlRawDataPCA)). Every point in the plot 
represents one sample, with the colour indicating the mucosa type 
(inflamed vs non-inflamed) and the shape  indicating the disease (UC or CD).



```{r qualityControlRawDataPCA, fig.cap="PCA plot of the log–transformed raw expression data."}
exp_raw <- log2(Biobase::exprs(raw_data))
PCA_raw <- prcomp(t(exp_raw), scale. = FALSE)

percentVar <- round(100*PCA_raw$sdev^2/sum(PCA_raw$sdev^2),1)
sd_ratio <- sqrt(percentVar[2] / percentVar[1])

dataGG <- data.frame(PC1 = PCA_raw$x[,1], PC2 = PCA_raw$x[,2],
                    Disease = pData(raw_data)$Factor.Value.disease.,
                    Phenotype = pData(raw_data)$Factor.Value.phenotype.,
                    Individual = pData(raw_data)$Characteristics.individual.)

ggplot(dataGG, aes(PC1, PC2)) +
      geom_point(aes(shape = Disease, colour = Phenotype)) +
  ggtitle("PCA plot of the log-transformed raw expression data") +
  xlab(paste0("PC1, VarExp: ", percentVar[1], "%")) +
  ylab(paste0("PC2, VarExp: ", percentVar[2], "%")) +
  theme(plot.title = element_text(hjust = 0.5))+
  coord_fixed(ratio = sd_ratio) +
  scale_shape_manual(values = c(4,15)) + 
  scale_color_manual(values = c("darkorange2", "dodgerblue4"))
```

The PCA plot (Figure \@ref(fig:qualityControlRawDataPCA), performed on 
the log-intensity scale) of the raw data shows that the 
first principal component differentiates between the diseases. This means that 
the disease type is a major driver of gene expression differences.
This might hinder our analysis, as we want to analyze the differential expression between inflamed and non-inflamed 
tissues, independently of the disease a person suffers from. 

We also represent the probe intensities via a boxplot graph with one box per
individual microarray. (Figure \@ref(fig:qualityControlRawDataBox)). Note that the 
`oligo::boxplot` function, i.e. the boxplot function of the oligo package, can take 
expression sets as argument. It accesses the expression data and performs a 
log2-transformation by default. We therefore can use `raw_data` as argument here.

```{r qualityControlRawDataBox, fig.cap="Intensity boxplots of the log2–transformed raw data."}
oligo::boxplot(raw_data, target = "core", 
               main = "Boxplot of log2-intensitites for the raw data")
```

When looking at the boxplot (Figure 
\@ref(fig:qualityControlRawDataBox)), we see that the intensity distributions of the 
individual arrays are quite different, indicating the need for an appropriate 
normalization, which we will discuss next.

Until now, we have only performed a very basic quality control; 
more elaborate quality control plots are available in the package
`r Biocpkg("arrayQualityMetrics")` [@AQM]. The package produces an html report, 
containing the quality control plots together with a description of their
aims and an identification of possible outliers. We do not discuss this tool in 
detail here, but simply provide the code below, 
which creates a report for our raw data. 

```{r arrayQualityMetricsRaw, eval = TRUE, warning=FALSE, message=FALSE}
arrayQualityMetrics(expressionset = raw_data,
    outdir = tempdir(),
    force = TRUE, do.logtransform = TRUE,
    intgroup = c("Factor.Value.disease.", "Factor.Value.phenotype."))
```

# Background adjustment, calibration, summarization and annotation

## Background adjustment

After the initial import and quality assessment, the next step in processing of
microarray data is background adjustment. This is essential because 
a proportion of the measured probe intensities are due to non-specific 
hybridization and the noise in the optical detection system. 
Therefore, observed intensities need to be adjusted to 
give accurate measurements of specific hybridization.

## Across-array normalization (calibration)

Normalization across arrays is needed in order to be able
to compare measurements from different array hybridizations 
due to many obscuring sources of variation.
These include different efficiencies of
reverse transcription, labeling or hybridization
reactions, physical problems with the arrays, reagent batch effects, 
and laboratory conditions. 

## Summarization

After normalization, summarization is necessary to be done because on 
the Affymetrix platform, transcripts are represented
by multiple probes, that is multiple locations on the array. 
For each gene, the background-adjusted and normalized intensities of all probes
need to be summarized into one quantity that estimates an amount proportional to
the amount of RNA transcript.

After the summarization step, the summarized data can be annotated with various
information, e.g. gene symbols and ENSEMBL gene identifiers. There is an
annotation database available from Bioconductor 
for our platform, namely the package
`r Biocannopkg("hugene10sttranscriptcluster.db") `.

You can view its content like this:

```{r annotationDataBaseContent, eval = TRUE}
head(ls("package:hugene10sttranscriptcluster.db"))
```

Additional information is available from the reference manual of the package.
Essentially, the package provides a mapping from the transcript cluster 
identifiers to the various annotation data.

## Old and new "probesets" of Affymetrix microarrays

Traditionally, Affymetrix arrays (the so-called 3' IVT arrays)
were probeset based: a certain fixed group of probes were part of a probeset
which represented a certain gene or transcript (note however, that a
gene can be represented by multiple probesets).

The more recent "Gene" and "Exon" Affymetrix arrays are exon based and hence 
there are two levels of summarization to get to the gene level. The "probeset" summarization leads to the exon level. The gene / transcript level is given by "transcript clusters". Hence, the appropriate annotation package for 
our chip type is called
`r Biocpkg("hugene10sttranscriptcluster.db") `.

"Gene" arrays were created as affordable versions of the "Exon" arrays, by only 
taking the "good" probes from the Exon array. Initially on the Exon array, 
at least four probes were part of one "Exon". 
With the thinned out "Gene" array, many probesets were made up of three or fewer 
probes. This is visualized in Figure 
\@ref(fig:DifferenceBetweenExonAndGeneTypeArrays): Single probesets 
are indicated by single colours; probesets representing one gene 
are indicated by a colour shade: e.g., all yellow probes belong to one Exon, 
and all yellow, orange and red probesets belong to one gene: 

```{r DifferenceBetweenExonAndGeneTypeArrays, fig.width=7, fig.height=6, echo=FALSE, fig.cap="Visualization of the difference between \"Exon\" type array (left) and \"Gene\" type array (right)."}
library(grid)

par(mar=c(0,0,0,0))
plot(1,1,xlim=c(0,100),ylim=c(0, 20),bty="n",
     type="n",xlab="",ylab="",xaxt="n",yaxt="n")
dat <- data.frame(x = rep(seq(0, 2.9, 0.1), 30), 
                  y = rep(seq(0, 2.9, 0.1), each = 30))


col <- c("blue", "darkslategray1", "goldenrod1", "yellow", "royalblue1", "coral")
dat$colours<-sample(rep(col, 150))
dat$colours2 <- sample(c(rep("yellow", 2), rep("goldenrod1", 5), rep("coral",7), 
                  rep("blue", 5), rep ("darkslategray1",3), rep("royalblue1", 8), rep(gray(0:9/10), 87)))

# opening the graphic device and
# setting up a viewport with borders:
vp1 <- viewport(x = 0.1, y = 0.1, w = 0.12, h = 0.12, 
                just = c("left", "bottom"), name = "vp1")


# plotting rectangles using x/y positions
grid.rect(x=dat$x,y=dat$y, height=0.1, width=0.1, hjust=0,vjust=0,vp=vp1,
          gp=gpar(col=1, fill=as.character(dat$colours)))

##########################

vp2 <- viewport(x = 0.5, y = 0.1, w = 0.12, h = 0.12,
                just = c("left", "bottom"), name = "vp1")


# plotting rectangles using x/y positions
grid.rect(x=dat$x,y=dat$y, height=0.1, width=0.1, hjust=0,vjust=0,vp=vp2,
          gp=gpar(col=1, fill=as.character(dat$colours2)))
```


On the left side, we see plenty of probes for each Exon / probeset 
(i.e. each colour): therefore, a summarization on the probeset / exon level 
makes sense. In the gene type array, however, only a small proportion of the 
original probes per probeset is included. Thus, a summarization on the 
probeset / exon level  is not recommended for "Gene" arrays 
but nonetheless possible by using the `r Biocannopkg("hugene10stprobeset.db") ` 
annotation package.

Note that furthermore, there are also no longer designated match/mismatch probes 
present on "Gene" and "Exon" type chips. The mismatch probe was initially 
intended as base-level for background correction, but hasn't prevailed 
due to more elaborate background correction techniques that 
do not require a mismatch probe. 

## One-step preprocessing in oligo

The package `r Biocpkg("oligo") ` allows us to perform background correction, 
normalization and summarization in one single step using a deconvolution 
method for background correction, quantile normalization and
the RMA (robust multichip average) algorithm for summarization.

This series of steps as a whole is commonly referred to as RMA algorithm,
although strictly speaking RMA is merely a summarization method
[@Irizarry_2003; @Bolstad_2003; @Irizarry_2003a].

# Relative Log Expression data quality analysis

Before calibrating and evaluating the data, we want to perform another quality 
control procedure, namely Relative Log Expression (RLE), as described in 
the article by Gandolfo et al [@Gandolfo_2018]. To this end, we first 
perform an RMA without prior normalization: 

```{r RMAcalibrationForRLE, eval=TRUE}
palmieri_eset <- oligo::rma(raw_data, target = "core", normalize = FALSE)
```

Further details on the RMA algorithm will be provided after RLE analysis, when 
the "full" RMA is carried out, including normalization. 

## Computing the RLE

The RLE is performed by calculating the median log2 intensity of every 
transcript across all arrays. 

We do this by calculating the row medians of `exprs(palmieri_eset)`, 
as the transcripts are represented by the rows and the single microarrays 
by the columns. 

Note that we do not have to apply the log2 manually, 
as the output data of the RMA function is in log2 scale by default. 

We then substract this transcript median intensity from every 
transcript intensity via the `sweep` function. 

## Plotting the RLE

We finally reshape the data into a format in which we can use to create a boxplot 
for each array, as before:


```{r boxplotDataForRLETidy, fig.cap="Boxplot for the RLE values", warning=FALSE}
row_medians_assayData <- 
  Biobase::rowMedians(as.matrix(Biobase::exprs(palmieri_eset)))

RLE_data <- sweep(Biobase::exprs(palmieri_eset), 1, row_medians_assayData)

RLE_data <- as.data.frame(RLE_data)
RLE_data_gathered <- 
  tidyr::gather(RLE_data, patient_array, log2_expression_deviation)

ggplot2::ggplot(RLE_data_gathered, aes(patient_array,
                                       log2_expression_deviation)) + 
  geom_boxplot(outlier.shape = NA) + 
  ylim(c(-2, 2)) + 
  theme(axis.text.x = element_text(colour = "aquamarine4", 
                                  angle = 60, size = 6.5, hjust = 1 ,
                                  face = "bold"))
```

Note that the y-axis now displays for each microarray the deviation 
of expression intensity from the median expression of the respective single 
transcripts across arrays.

Boxes with a larger extension therefore indicate an unusually high deviation 
from the median in a lot of transcripts, suggesting that these arrays
are different from most of the others in some way.

Boxes that are shifted in y-direction indicate a systematically higher or lower 
expression of the majority of transcripts in comparison to most of the other 
arrays. This could be caused by quality issues or batch effects. 

Therefore, if shape and median of a given box varies too much from the 
bulk, they should be inspected and potentially removed.

By inspecting the boxplot in Figure \@ref(fig:boxplotDataForRLETidy), 
five arrays could be considered as outliers:
2826_I, 2826_II, 3262_II, 3302_II and 3332_II are negatively y-shifted. 

We will keep these samples in mind for heatmap cluster analysis later on in 
the workflow. Arrays that are confirmed to be outliers by heatmap 
analysis could be removed for subsequent analysis.


# RMA calibration of the data

Now, we can apply the full RMA algorithm to our data in order to 
background-correct, normalize and summarize:

```{r RMAcalibrationWITHnormalization, eval=TRUE}
palmieri_eset_norm <- oligo::rma(raw_data, target = "core")
```

The parameter `target` defines the degree of summarization, the
default option of which is "core", using transcript clusters containing
"safely" annotated genes. For summarization on the exon level (not recommended for Gene
arrays), one can use "probeset" as the target option.
Although other methods for background correction and normalization exist,
RMA is usually a good default choice.
RMA shares information across arrays and
uses the versatile quantile normalization method that
will make the array intensity distributions match. However, it is preferable 
to apply it only after outliers have been removed. 
The quantile normalization algorithm used by RMA
works by replacing values by the average of identically
ranked (within a single chip) values across arrays. A more detailed 
description can be found on the [Wikipedia page](https://en.wikipedia.org/wiki/Quantile_normalization).

An alternative to quantile normalization would be the `r Biocpkg("vsn") ` 
algorithm,that performs background correction and normalization by robustly
shifting and scaling intensity values within arrays before log-transforming
them. This is less "severe" than quantile normalization [@vsn]. 

## Some mathematical background on normalization (calibration) and background correction

A generic model for the value of the intensity \(Y\) of a single probe on
a microarray is given by

\[
    Y = B + \alpha \cdot S
\]
    
where B is a random quantity due to background noise, usually composed of
optical effects and non-specific binding, \(\alpha\) is a gain factor, 
and \(S\) is the amount of measured specific binding. The signal \(S\) 
is considered a random variable as well and accounts for measurement 
error and probe effects.
The measurement error is typically assumed to be multiplicative so we can write:

\[
    \log(S) = \theta + \varphi + \varepsilon
\]
    
Here \(\theta\) represents the logarithm of the true abundance,
\(\varphi\) is a probe-specific effect, and $\varepsilon$ accounts for the 
nonspecific error.
This is the additive-multiplicative-error model for microarray data used 
by RMA and also the `r Biocpkg("vsn") ` algorithm [@vsn]. The algorithms differ 
in the way that \(B\) is removed and an estimate of \(\theta\) is obtained.


## Quality assessment of the calibrated data

We now produce a clustering heatmap and another PCA plot using the 
calibrated data. 

**PCA analysis**

First, we perform a PCA analysis of the calibrated data analogously 
to the one with the raw data:

```{r PCAMetricsCalibrated, fig.cap = "PCA plot of the calibrated, summarized data.", eval = TRUE }
exp_palmieri <- Biobase::exprs(palmieri_eset_norm)
PCA <- prcomp(t(exp_palmieri), scale = FALSE)

percentVar <- round(100*PCA$sdev^2/sum(PCA$sdev^2),1)
sd_ratio <- sqrt(percentVar[2] / percentVar[1])

dataGG <- data.frame(PC1 = PCA$x[,1], PC2 = PCA$x[,2],
                    Disease = 
                     Biobase::pData(palmieri_eset_norm)$Factor.Value.disease.,
                    Phenotype = 
                     Biobase::pData(palmieri_eset_norm)$Factor.Value.phenotype.)


ggplot(dataGG, aes(PC1, PC2)) +
      geom_point(aes(shape = Disease, colour = Phenotype)) +
  ggtitle("PCA plot of the calibrated, summarized data") +
  xlab(paste0("PC1, VarExp: ", percentVar[1], "%")) +
  ylab(paste0("PC2, VarExp: ", percentVar[2], "%")) +
  theme(plot.title = element_text(hjust = 0.5)) +
  coord_fixed(ratio = sd_ratio) +
  scale_shape_manual(values = c(4,15)) + 
  scale_color_manual(values = c("darkorange2", "dodgerblue4"))
```

In comparison to the first PCA analysis before RMA (Figure \@ref(fig:qualityControlRawDataPCA)), we see that now the first principal 
component separates between the tissues types (Figure \@ref(fig:PCAMetricsCalibrated)).
This indicates that now differential expression between the tissue types is the
dominant source of variation. Note that the second principal component
separates the diseases.

**Heatmap clustering analysis**

We want to plot a heatmap with the sample-to-sample distances with the 
sample names as row-names. Also, we want to see how well the samples 
cluster for phenotype (inflamed and non-inflamed tissue) and disease 
(UC and CD), respectively. We use row annotation for that: it means that these features get a colour code and will be displayed to the left of each row. 

```{r rownamesForHeatmap, fig.height = 8.5, fig.width = 7, eval = TRUE, echo=TRUE}
phenotype_names <- ifelse(str_detect(pData
                                    (palmieri_eset_norm)$Factor.Value.phenotype.,
                             "non"), "non_infl.", "infl.")

disease_names <- ifelse(str_detect(pData
                                    (palmieri_eset_norm)$Factor.Value.disease.,
                             "Crohn"), "CD", "UC")

annotation_for_heatmap <- 
  data.frame(Phenotype = phenotype_names,  Disease = disease_names)

row.names(annotation_for_heatmap) <- row.names(pData(palmieri_eset_norm))

```

In order to map the sample-to-sample distances, we first compute the distances 
using the `dist` function. We need to transpose the expression values since
the function computes the distances between the rows (i.e. genes in our case) by
default. The default distance is the Euclidean one. However this can be
changed and we choose the Manhattan distance here (it uses absolute 
distances along rectangular paths instead of squared distances 
of the direct path), as it is more robust. 
We set the diagonal of the distance matrix to ` NA ` in order 
to increase the contrast of the color coding. Those diagonal entries 
do not contain information since the distance of a sample 
to itself is always equal to zero.


```{r HeatmapWithAnnotation, fig.height = 8.5, fig.width = 7, fig.cap="Heatmap of the sample-to-sample distances"}
dists <- as.matrix(dist(t(exp_palmieri), method = "manhattan"))

rownames(dists) <- row.names(pData(palmieri_eset_norm))
hmcol <- rev(colorRampPalette(RColorBrewer::brewer.pal(9, "YlOrRd"))(255))
colnames(dists) <- NULL
diag(dists) <- NA

ann_colors <- list(
  Phenotype = c(non_infl. = "chartreuse4", infl. = "burlywood3"),
  Disease = c(CD = "blue4", UC = "cadetblue2")
                   )
pheatmap(dists, col = (hmcol), 
         annotation_row = annotation_for_heatmap,
         annotation_colors = ann_colors,
         legend = TRUE, 
         treeheight_row = 0,
         legend_breaks = c(min(dists, na.rm = TRUE), 
                         max(dists, na.rm = TRUE)), 
         legend_labels = (c("small distance", "large distance")),
         main = "Clustering heatmap for the calibrated samples")
```
On the heatmap plot (Figure \@ref(fig:HeatmapWithAnnotation)) we also see that 
the samples do not cluster strongly by tissue, confirming the impression 
from the PCA plot (Figure \@ref(fig:PCAMetricsCalibrated)) that the separation 
between the tissues is not perfect. The yellow stripes in the heatmap might correspond
to outliers that could potentially be removed: the ones that could be flagged 
here are 2826_II, 3262_II, 3271_I, 2978_II and 3332_II. 
2826_II, 3262_II and 3332_II were found to be outliers in both RLE and heatmap 
analysis and could therefore potentially be removed; however, in order 
to stay as close as possible to the
original paper, we continue with the complete set of samples.
Note again that more elaborate metrics to identify and remove outliers 
are provided by the `r Biocpkg("arrayQualityMetrics")` package.


# Filtering based on intensity 

We now filter out lowly expressed genes. Microarray data commonly show
a large numberof probes in the background intensity range. These probes 
also do not  change much across arrays. Hence they combine a low variance 
with a low  intensity. Thus, they could end up being detected as differentially 
expressed although they are barely above the "detection" limit and are not very 
informative in general. 

We will perform a "soft" intensity based filtering here, since this 
is recommended by the `r Biocpkg("limma")` [@limma; @Smyth_2004] user guide 
(a package we will use below for the differential 
expression analysis). 

However, note that a variance based filter might exclude a similar set of 
probes in practice. 
For intensity-based filtering, we calculate the row-wise medians from the 
expression data, as they represent the transcript medians, and assign them to 
`palmieri_medians`. From this we create a histogram: 

```{r IntensityBasedManualFiltering, fig.cap="Histogram of the median intensities per gene"}
palmieri_medians <- rowMedians(Biobase::exprs(palmieri_eset_norm))

hist_res <- hist(palmieri_medians, 100, col = "cornsilk1", freq = FALSE, 
            main = "Histogram of the median intensities", 
            border = "antiquewhite4",
            xlab = "Median intensities")
```

In the histogram of the gene-wise medians (Figure 
\@ref(fig:IntensityBasedManualFiltering)), we 
can clearly see an enrichment of low medians on the left hand side. 
These represent the genes we want to filter. In order to infer a cutoff from the 
data, we inspect the histogram: We visually set a cutoff line `man_threshold` 
to the left of the histogram peak in order not to exclude too many genes. In our 
example, we choose a threshold of 4. We plot the same histogram as before 
and add the threshold line with the `abline()` function:

```{r setManualThreshold, fig.cap="Histogram of the median intensities per gene with manual intensity filtering threshold (red line)."}
man_threshold <- 4

hist_res <- hist(palmieri_medians, 100, col = "cornsilk", freq = FALSE, 
            main = "Histogram of the median intensities",
            border = "antiquewhite4",
            xlab = "Median intensities")

abline(v = man_threshold, col = "coral4", lwd = 2)
```

Transcripts that do not have intensities larger than the threshold in 
at least as many arrays as the smallest experimental group are excluded. 

In order to do so, we first have to get a list with the number 
of samples (=arrays) (`no_of_samples`) in the experimental groups: 

```{r expGroups, dependson="PCAMetricsCalibrated"}
no_of_samples <- 
  table(paste0(pData(palmieri_eset_norm)$Factor.Value.disease., "_", 
                  pData(palmieri_eset_norm)$Factor.Value.phenotype.))
no_of_samples 
```


We now filter out all transcripts 
that do not have intensities greater than the threshold in at least 
as many arrays as the smallest experimental group (`r min(no_of_samples)`) 
which we  define as `samples_cutoff`. 

A function `idx_man_threshold` is then applied to each row, i.e. 
to each transcript  across all arrays.
It evaluates whether the number of arrays where the median intensity passes the
threshold (`sum(x > man_threshold)`) is greater than the `samples_cutoff` and 
returns TRUE or FALSE for each row, i.e. each transcript.

We then create a table of `idx_man_threshold` to summarize the results and
get an overview over how many genes are filtered out. 
In the last step, we subset our expression set to `palmieri_manfiltered` 
and keep the TRUE elements of `idx_man_threshold`.

```{r filteringOfLowIntensity_transcripts}
samples_cutoff <- min(no_of_samples)

idx_man_threshold <- apply(Biobase::exprs(palmieri_eset_norm), 1,
                           function(x){
                          sum(x > man_threshold) >= samples_cutoff})
                          table(idx_man_threshold)

palmieri_manfiltered <- subset(palmieri_eset_norm, idx_man_threshold)
```


# Annotation of the transcript clusters

Before we continue with the linear models for microarrays and differential
expression, we first add "feature data", i.e. annotation
information to the transcript cluster identifiers stored in the featureData of
our ExpressionSet: 

```{r annotateData, eval=TRUE, dependson="intensityBasedFiltering", message = FALSE}
anno_palmieri <- AnnotationDbi::select(hugene10sttranscriptcluster.db,
                                  keys = (featureNames(palmieri_manfiltered)),
                                  columns = c("SYMBOL", "GENENAME"),
                                  keytype = "PROBEID")

anno_palmieri <- subset(anno_palmieri, !is.na(SYMBOL))
```

We used the function `select` from `r Biocpkg("AnnotationDbi") ` to query 
the gene symbols and associated short descriptions for the transcript clusters. 
For each cluster, we added the gene symbol (`SYMBOL`) and a short description
of the gene the cluster represents (`GENENAME`). 

In a second step, we filtered out the probes that do not map to a gene,
i.e. that do not have a gene symbol assigned. 

## Removing multiple mappings 

Many transcript-cluster identifiers will map to multiple gene symbols, i.e. 
they can't be unambigously assigned.

We compute a summary table in the code below to see how many there are:


```{r multipleMappings, dependson="annotateData"}
anno_grouped <- group_by(anno_palmieri, PROBEID)
anno_summarized <- 
  dplyr::summarize(anno_grouped, no_of_matches = n_distinct(SYMBOL))

head(anno_summarized)

anno_filtered <- filter(anno_summarized, no_of_matches > 1)

head(anno_filtered)

probe_stats <- anno_filtered 

nrow(probe_stats)
```

First, we grouped `anno_palmieri` by their PROBEID; that way, 
the subsequent operations are not carried through for each single row, 
but for each  group, i.e. each PROBEID.
We then summarized the groups and indicate the number of different genes 
assigned to a transcript cluster in the column `no_of_matches`. 
Finally, we filtered for PROBEIDs with multiple matches, 
i.e. `no_of_matches > 1`.

With `dim(probe_stats)`, we could see how many 
probes have been mapped to multiple genes.

We have close to 2000 transcript clusters that map to multiple gene symbols.
It is difficult to decide which mapping is "correct". Therefore,
we exclude these transcript clusters. 

We want to remove those probe IDs that match the ones in `probe_stats`, 
as those are the probes with multiple mappings. We assign these IDs 
to the variable `ids_to_exclude`. Then, we generate `palmieri_final`, 
an expression set without the `ids_to_exclude`. 

```{r excludeMultipleMappingsFromAssayData, dependson="multipleMappings"}
ids_to_exlude <- (featureNames(palmieri_manfiltered) %in% probe_stats$PROBEID)

table(ids_to_exlude)

palmieri_final <- subset(palmieri_manfiltered, !ids_to_exlude)

validObject(palmieri_final)
```

As we have just excluded probe IDs from the assay data, we now have 
to also exclude them from the feature data `anno_palmieri`:

```{r recallAnnoPalmieri}
head(anno_palmieri)
```

Recall that `fData` enables us to access the feature data of an expression set.
Until now, no feature data whatsoever is stored in the `fData(palmieri_final)`.
Only the row names are the row names of the assay data by default, which are the 
PROBEIDs of the transcripts.

Therefore, we generate a column PROBEID in `fData(palmieri_final)` and 
assign the row names of `fData(palmieri_final)` to it:

```{r excludeMultipleMappingsFromFeatureData}
fData(palmieri_final)$PROBEID <- rownames(fData(palmieri_final))
```

Then, we left-join `fData(palmieri_final)`with `anno_palmieri`, which already contains 
the columns "SYMBOL" and "GENENAME".
A left-join keeps the rows and columns of the first argument 
and adds the corresponding column entries of the second argument:

```{r excludeMultipleMappingsFromFeatureData2}
fData(palmieri_final) <- left_join(fData(palmieri_final), anno_palmieri)


# restore rownames after left_join
rownames(fData(palmieri_final)) <- fData(palmieri_final)$PROBEID 
    
validObject(palmieri_final)
```

By left-joining with `anno_palmieri`, we thus add the "SYMBOL" and "GENENAME" 
columns from `anno_palmieri` for only the PROBEIDs that are in 
`fData(palmieri_final)` and thus get the feature Data for the filtered probes.


## Building custom annotations

Alternatively, one can re-map the probes of the array
to a current annotation. A workflow to do this for Illumina arrays is given in
@Arloth_2015.
Essentially, the individual probe sequences are re-aligned to an in-silico
"exome" that consists of all annotated transcript exons.

In any case, the package `r Biocpkg("pdInfoBuilder") ` can be used to build 
customannotation packages for use with `r Biocpkg("oligo") `. 
In order to do this, PGF / CLF files (called "Library files" on the Affymetrix 
website) as well as the probeset annotations are required. 
The probesets typically represent small stretches 
of the genome (such as a single exon) and multiple probesets
are then used to form a transcript cluster.

The CLF file contains information about the location of
individual probes on the array. The PGF file then contains the individual probe
sequences and shows the probeset they belong to. Finally, the probeset 
annotation .csv then contains information about which probesets are used
in which transcript cluster. Commonly, multiple probesets are used in one
transcript cluster and some probesets are contained in multiple transcript
clusters.


# Linear models

In order to analyse which genes are differentially expressed between 
inflamed and non-inflamed tissue, we have to fit a linear model to our 
expression  data. Linear models are the "workhorse" for the analysis of 
experimental data. They can be used to analyse almost arbitrarily 
complex designs, however they also take a while to learn and understand
and a thorough description is beyond the scope of this workflow.

Mike Love's and Rafael Irizzary's ["Biomedical Data Science"](http://genomicsclass.github.io/book/) [@Irizarry_2015] is a very good resource, especially the section on [interactions and contrasts](http://genomicsclass.github.io/book/pages/interactions_and_contrasts.html).
It might also be helpful to learn some linear algebra to better understand
the concepts here. The Khan Academy offers helpful (and free) [online courses](https://www.khanacademy.org/math/linear-algebra).

## Linear models for microarrays

We now apply linear models to microarrays. Specifically, we discuss
how to use the `r Biocpkg("limma") ` package for differential expression 
analysis. The package is designed to analyze complex experiments involving 
comparisons between many experimental groups simultaneously while remaining 
reasonably easy to use for simple experiments. The main idea is to fit 
a linear model to the expression data for each gene. Empirical Bayes and other 
methods are used to borrow information across genes for the residual variance 
estimation leading to "moderated" \(t\)-statistics, and stabilizing 
the analysis for experiments with just a small number of arrays [@Smyth_2004]. 
Conceptually, the final per gene variance is a mix of a prior variance and 
the per gene variance.

Typical experimental designs are disussed in chapter 9 of `r Biocpkg("limma") `
"User Guide", which can be found on the Bioconductor landing page 
of `r Biocpkg("limma") `. 

In the following, we use appropriate design and contrast matrices for
our linear models and fit a linear model to each gene separately.

## A linear model for the data

For the subsequent linear modelling of the data, we introduce the abbreviations 
"UC" and "CD" for the disease types, and "non_infl." and "infl."
for the phenotypes, respectively: 

```{r introductionOfAbbreviations}
individual <- 
  as.character(Biobase::pData(palmieri_final)$Characteristics.individual.)

tissue <- str_replace_all(Biobase::pData(palmieri_final)$Factor.Value.phenotype.,
                  " ", "_")

tissue <- ifelse(tissue == "non-inflamed_colonic_mucosa",
                 "nI", "I")

disease <- 
  str_replace_all(Biobase::pData(palmieri_final)$Factor.Value.disease.,
                  " ", "_")
disease <- 
  ifelse(str_detect(Biobase::pData(palmieri_final)$Factor.Value.disease., 
                    "Crohn"), "CD", "UC")
```

The original paper is interested in changes in transcription that occur
between inflamed and adjacent non-inflamed mucosal areas of the colon.
This is studied in both inflammatory bowel disease types.

For building our linear model, we have to think about 
which experimental variables we want to consider.
As we want to find differential expression 
between non-inflamed and inflamed tissue, in principle, 
those are the only two variables 
we would have to consider.

However, since we have two arrays per individual patient, we have a
"Paired Samples" design (see section 9.4 of the `r Biocpkg("limma")` 
user guide). This means that the samples might be biased by the person they come 
from. Whenever a feature in an experimental setup 
is expected to have a systematic influence on the result, blocking factors 
on these features should be introduced.

Thus, the first factor we need is a blocking factor for the individuals 
that will absorb differences in expression between
them. Therefore, we block on patients, which means that the 
patient IDs become variables of the linear model.

Then we create factors that give us the grouping for
the tissue types (non-inflamed and inflamed).

Finally, we create two design matrices, 
one for each of the two diseases as we will analyze them separately 
in order to follow the analysis strategy of the original paper 
closely (one could also fit a joint model to 
the complete data set; however, the two diseases might show very different
characteristics so that a joint fit might not be appropriate).

```{r createDesign, eval=TRUE, dependson="excludeMultipleMappings" }
i_CD <- individual[disease == "CD"]
design_palmieri_CD <- model.matrix(~ 0 + tissue[disease == "CD"] + i_CD)
colnames(design_palmieri_CD)[1:2] <- c("I", "nI")
rownames(design_palmieri_CD) <- i_CD 

i_UC <- individual[disease == "UC"]
design_palmieri_UC <- model.matrix(~ 0 + tissue[disease == "UC"] + i_UC )
colnames(design_palmieri_UC)[1:2] <- c("I", "nI")
rownames(design_palmieri_UC) <- i_UC 
```

We can inspect the design matrices:

```{r inspectDesignMatrix, eval = TRUE, dependson="createDesign"}
head(design_palmieri_CD[, 1:6])

head(design_palmieri_UC[, 1:6])
```

In the design matrix, the rows represent the patient array, 
and the columns are the variables we include in our linear model. The variables 
correspond to our blocking factors: there are two for non-inflamed and inflamed 
tissue, respectively, and one for each patient. "`i_UC2424`" for example is the 
blocking variable of patient 2424; UC stands for the disease the patient is 
suffering from. 
For example, the first two rows of the design matrix `design_palmieri_CD` 
correspond to the two arrays for individual "164". 

The design matrix entries are 0 or 1 and thus tell us 
which variables are "active" for which sample: 

It can be seen as a switch that turns the single variables "on" (with a 1 at the 
corresponding position) or "off" (with a 0 at the corresponding position) for 
each row, i. e. each patient array. If we take a closer look at the single rows,
we see that for each sample, there are two "ones" assigned: one to one of the 
variables "nI" or "I" corresponding to the tissue the array came from, 
and one to the corresponding patient-specific blocking variable.

Note that in the linear model, individual 164 serves as the baseline for all 
other individuals and thus isn't included in the sample variables.

Note that instead of blocking on individuals, it would also be possible to use a 
"mixed model" approach with the `duplicateCorrelation()`function from the 
`r Biocpkg("limma")` package. 
It has advantages over the "fixed patient effect model" presented here 
in terms of applicability to more complicated experimental designs, 
where we want to perform comparisons both within and between the patients (e.g. 
comparing between the two diseases; "split-plot-designs"). 

More information on it can be found in the [limma User's Guide](https://www.bioconductor.org/packages/release/bioc/vignettes/limma/inst/doc/usersguide.pdf) (section 17.3.6). 
However, the above explained is more intuitive and is therefore used here.

Before heading on to find all differentially expressed genes for both diseases, 
we will first have a look at how this works in principle for one gene.
We will fit the linear model for one gene and run a t-test in order 
to see whether the gene is differentially expressed or not.

## Analysis of differential expression based on a single gene

For linear model fitting and subsequent testing for differential expression 
by t-test, we will pick the gene with the PROBEID 8164535. It has the gene 
symbol  *CRAT* and will be named as such in the following code. 

**Illustration of the fitted linear model on the *CRAT* gene**

Before fitting the linear model, we have a look at the expression intensities
of this gene for each patient in non-inflamed and inflamed tissue, 
respectively: 


```{r visualizeExpressionChanges, fig.cap="Visualization of expression changes"}
tissue_CD <- tissue[disease == "CD"]
crat_expr <- Biobase::exprs(palmieri_final)["8164535", disease == "CD"]
crat_data <- as.data.frame(crat_expr)
colnames(crat_data)[1] <- "org_value"
crat_data <- mutate(crat_data, individual = i_CD, tissue_CD)

crat_data$tissue_CD <- factor(crat_data$tissue_CD, levels = c("nI", "I"))

ggplot(data = crat_data, aes(x = tissue_CD, y = org_value, 
                             group = individual, color = individual)) +
      geom_line() +
      ggtitle("Expression changes for the CRAT gene")
```

We see that overall, this gene is expressed less in inflamed tissue 
(Figure \@ref(fig:visualizeExpressionChanges)). We also see that the absolute 
expression intensities vary greatly between patients. 
However, we have already taken care of this 
problem by introducing blocking factors based on the individuals, which allows 
us to compare the tissues within each individual as represented by
the single lines.

If we had not blocked for individuals, the linear model would 
treat them interchangably and a graphical depiction would only
include a single line. Since the individuals haver very different
baseline expression levels, this would lead to a very high variance of
the estimated fold changes.

We now compute the variable coefficients by fitting a linear model. 
We get a vector `crat_coef` with one entry for each variable.

```{r obtainFitCRAT}
crat_coef <- lmFit(palmieri_final[,disease == "CD"],
                design = design_palmieri_CD)$coefficients["8164535",]

crat_coef
```


In order to now obtain the expression values fitted by the model, 
we multiply the design matrix with this vector of coefficients `crat_coef`:

```{r 2obtainFitCRAT}
crat_fitted <- design_palmieri_CD %*% crat_coef
rownames(crat_fitted) <- names(crat_expr)
colnames(crat_fitted) <- "fitted_value"

crat_fitted
```

Recall that for every row in the design matrix (i.e. every patient sample) 
only the variables with a 1 in the design matrix are taken into account for 
calculating the fitted expression value. 

This means that as output of the multiplication, we get 
a vector `crat_fitted` whose entries are the sum of relevant 
variable coefficients for each sample, respectively.

For example, the fitted value for patient sample `2114_I.CEL` 
is `r crat_coef["nI"] + crat_coef["i_CD2114"]`: it is the sum of the respective activated variable coefficients "nI" (`r crat_coef["nI"]`) 
and "i_CD2114" (`r crat_coef["i_CD2114"]`).


Let's visualize the difference between non-inflamed and inflamed tissue 
again after fitting: 

```{r 3obtainFitCRAT, fig.cap="Expression changes for the CRAT gene"}
crat_data$fitted_value <- crat_fitted

ggplot(data = crat_data, aes(x = tissue_CD, y = fitted_value, 
                             group = individual, color = individual)) +
      geom_line() +
      ggtitle("Fitted expression changes for the CRAT gene")
```


Note that the difference of the fitted expression values between inflamed and 
non-inflamed samples of one patient is the same for all patients and 
is determined by the difference between the variable coefficients of 
`I` (`r crat_coef["I"]`) and `nI` (`r crat_coef["nI"]`), 
which is  `r crat_coef["I"] - crat_coef["nI"]` 
(Figure \@ref(fig:3obtainFitCRAT)). 

This is the case because the same blocking variable is activated by the design 
matrix for both samples from a single patient, leading to a comparison within
patients only. These blocking variables correct the fitted tissue specific 
expression values towards the expression levels of the individual patients.
Therefore the final estimate is like an average of all the within-individual
differences.

The "difference" between non-inflamed and inflamed tissue of 
`r crat_coef["I"] -  crat_coef["nI"]` is actually a log2 fold change, as our 
expression data is on the log2 scale. `r crat_coef["I"] - crat_coef["nI"]`
therefore is our log2 fold change for the CRAT gene.

**Differential expression analysis of the CRAT gene**

In order to test whether the gene is differentially expressed or not, 
a \(t\)-test with the null hypothesis that there is no difference 
in the expression between non-inflamed and inflamed tissue is carried out. 
Our blocking design is conceptually similar to a paired t-test for 
which the statistic is given by:


\[
 t = \frac{\bar{d} }{s/\sqrt{n}}
\]

Where, \(\bar{d}\) is the mean difference in expression values
between the individuals. 
The paired t-test computes the variance \(s^2\) from the paired differences.
This is lower than the variance of a standard t-test and thus the paired 
t-test has higher power as long as the expression values 
for the same individual are correlated ([see e.g. the article on Wikipedia](https://en.wikipedia.org/wiki/Paired_difference_test)).

We thus have improved the power of the ordinary \(t\)-test 
by reducing the variance via blocking on individuals.

We now conduct the \(t\)-test on the linear model in order to find 
out whether the difference between non-inflamed and inflamed 
tissue differs significantly from 0: 

```{r tTest}
crat_noninflamed <- na.exclude(crat_data$org_value[tissue == "nI"])
crat_inflamed <- na.exclude(crat_data$org_value[tissue == "I"])
res_t <- t.test(crat_noninflamed ,crat_inflamed , paired = TRUE)
res_t
```

We get a low p-value close to `r round(res_t[["p.value"]], 4) `
and thus can conclude that the *CRAT* gene is differentially 
expressed between non-inflamed and inflamed tissue.

Note that the p-value isn't exactly the same one as below when analyzing the 
differential expression of all genes. This is due to the variance moderation 
performed by `r Biocpkg("limma")` .

## Contrasts and hypotheses tests

We now fit the linear model for all genes and
define appropriate contrasts to test hypotheses of interest.

We want to compare the inflamed to the non-inflamed tissue.
Thus, we create a contrast matrix consisting of only one contrast "I-nI": 
`r Biocpkg("limma")`'s function
`makeContrasts` creates this matrix from 
a symbolic description of the contrast of
interest.

We now fit a linear model to our data and apply the `contrasts.fit()` function 
to it in order to find genes with significant differential expression between non-inflamed and inflamed tissue:



```{r createContrastMatrixAndFitModel, eval=TRUE, dependson="createDesign" }
contrast_matrix_CD <- makeContrasts(I-nI, levels = design_palmieri_CD)

palmieri_fit_CD <- eBayes(contrasts.fit(lmFit(palmieri_final[,disease == "CD"],
                                design = design_palmieri_CD),
                                contrast_matrix_CD))

contrast_matrix_UC <- makeContrasts(I-nI, levels = design_palmieri_UC)

palmieri_fit_UC <- eBayes(contrasts.fit(lmFit(palmieri_final[,disease == "UC"],
                                design = design_palmieri_UC),
                                contrast_matrix_UC))
```

We applied the empirical Bayes variance moderation method to the model 
via the `eBayes()` function, which
computes moderated \(t\)-statistics. In microarray analysis, 
the number of arrays often is quite small, 
and thus variance estimation is difficult.
Using a combination of the per-gene-variance 
and a prior variance we can improve the 
variance estimate, hence the term "moderation". "Empirical Bayes" 
means that the prior is estimated from the data.

The result of the `eBayes()` step is that the individual variances are
shrunken towards the prior value. 

## Extracting results

Finally, we extract the number of differentially expressed genes. 
Results can be extracted by use of the `topTable` function. 
We extract the results for both Crohn's disease and ulcerative colitis, and the 
results are sorted by their absolute \(t\)-statistics.
As a diagnostic check, we also plot the p-value histogram: We expect a uniform 
distribution for the p-values that correspond to true null hypotheses, while 
a peak near zero shows an enrichment for low p-values corresponding to differentially 
expressed (DE) genes.
 
Note that if the p-value distribution for a dataset is very different 
from the ones in the histograms below, this might lead to quality loss in the 
subsequent analysis. Reasons for a divergent p-value-distribution might be
batch effects or a lack of consideration of other blocking factors in the design 
matrix. Thus, if the p-value is not as expected, 
try to include possible blocking factors and batches and rerun the analysis. 
If this does not help, empirical Bayes / null estimation methods 
for multiple testing are useful. 

A good starting point to learn about these methods is the article on 
false discovery rate estimation by Korbininan Strimmer [@Strimmer_2008] 
and chapter 1--6 of Efron's book on Large-Scale Inference [@Efron_2010], 
as well as the blog-post on "How to interpret a p-value histogram" by David 
Robinson [@Robinson_2014].

```{r extractResultsCD, eval = TRUE, dependson="createContrastMatrixAndFitModel", message=FALSE, fig.cap="Histogram of the p–values for Crohn’s disease."}
table_CD <- topTable(palmieri_fit_CD, number = Inf)
head(table_CD)

hist(table_CD$P.Value, col = brewer.pal(3, name = "Set2")[1],
     main = "inflamed vs non-inflamed - Crohn's disease", xlab = "p-values")

```

```{r extractResultsUC, eval = TRUE, dependson="createContrastMatrixAndFitModel", message=FALSE, fig.cap="Histogram of the p–values for ulcerative colitis."}

table_UC <- topTable(palmieri_fit_UC, number = Inf)
head(table_UC)

hist(table_UC$P.Value, col = brewer.pal(3, name = "Set2")[2],
     main = "inflamed vs non-inflamed - Ulcerative colitis", xlab = "p-values")

```

## Multiple testing FDR, and comparison with results from the original paper

In the original paper, a p-value of 0.001 was used as a significance
cutoff. Using this we get `r nrow(subset(table_UC, P.Value < 0.001))`
genes identified as differentially expressed for UC:


```{r pCutForCD}
nrow(subset(table_UC, P.Value < 0.001))
```


However, it is impossible
to determine a precise bound on the number of false positive genes
in this list. All that we can say using p-values is that we have 
at most `r nrow(table_UC)` (total number of tests) * 0.001
= `r nrow(table_UC)* 0.001` false positive genes in our list. Therefore, 
by choosing a p-value cutoff of 0.001, as much as
`r round(nrow(table_UC)* 0.001 / nrow(subset(table_UC, P.Value < 0.001))*100, 2)`% 
of our genes
identified as differentially expressed might be false positives. 

Thus, we can see that the "raw" p-values are very "liberal" when
looking at many tests simultaneously. We therefore need error
rates adapted to the multiple testing situation. By far the 
most popular one in molecular biology is the **false discovery
rate** or FDR for short. It is the percentage of false positives 
among all positives. As we have seen, the FDR of our genes list
using a simple p-value cutoff might be quite high.

On the other hand, we can see a clear peak in the p-value histogram 
(Figure \@ref(fig:extractResultsUC)), caused by the differentially
expressed genes. There we expect the actual FDR of our list 
to be lower.

The FDR at a given cutoff is given by the "adjusted" p-value in
the results table.

```{r FDRforUC}
tail(subset(table_UC, P.Value < 0.001))
```

The adjusted p-value for a raw p-value of 0.001 in 
the table is `r max(subset(table_UC, P.Value < 0.001)$adj.P.Val)`,
which is an order of magnitude lower than the FDR we can infer 
from p-values alone.

So although this is not recommended in general,
we also use a p-value cutoff at 0.001 in the following
in order to be able to compare our workflow results to the paper results. 

The paper results can be downloaded as excel files from
[http://links.lww.com/IBD/A795](http://links.lww.com/IBD/A795) and 
should be saved as an .xlsx file named `palmieri_DE_res.xlsx` in 
your working directory. 

Note that while the paper uses p-value cutoffs, it also reports the
corresponding FDRs (just as we did for the UC data here).

For a p-value cutoff of 0.001, the corresponding FDRs are 0.05 in 
Crohn's disease  and 0.02 in ulcerative colitis. There are four tables in total, 
giving the list of up and downregulated genes in CD and UC, respectively.
We calculate the overlap between our results and the ones from the 
paper as the ratio of the genes that were found in both analyses 
and the genes that were only found in the paper.

We also calculate the total number of diffentially expressed
genes that we find in our workflow analysis.
 
```{r compareDEgenes, dependson=c("extractResultsUC", "extractResultsCD")}
fpath <- system.file("extdata", "palmieri_DE_res.xlsx", package = "maEndToEnd")
palmieri_DE_res <- sapply(1:4, function(i) read.xlsx(cols = 1, fpath, 
                                                     sheet = i, startRow = 4))

names(palmieri_DE_res) <- c("CD_UP", "CD_DOWN", "UC_UP", "UC_DOWN")
palmieri_DE_res <- lapply(palmieri_DE_res, as.character)
paper_DE_genes_CD <- Reduce("c", palmieri_DE_res[1:2])
paper_DE_genes_UC <- Reduce("c", palmieri_DE_res[3:4])

overlap_CD <- length(intersect(subset(table_CD, P.Value < 0.001)$SYMBOL,  
                               paper_DE_genes_CD)) / length(paper_DE_genes_CD)


overlap_UC <- length(intersect(subset(table_UC, P.Value < 0.001)$SYMBOL,
                               paper_DE_genes_UC)) / length(paper_DE_genes_UC)
overlap_CD
overlap_UC 

total_genenumber_CD <- length(subset(table_CD, P.Value < 0.001)$SYMBOL)
total_genenumber_UC <- length(subset(table_UC, P.Value < 0.001)$SYMBOL)

total_genenumber_CD
total_genenumber_UC
```

We find `r total_genenumber_CD` (CD) and `r total_genenumber_UC` (UC) 
differentially expressed genes ("DE-genes").

In the paper, 298 (CD) and 520 (UC) DE-genes were found for the two diseases 
at the same cutoff. This higher number of DE-genes identified is probably
due to the increased power of the blocking according to the individuals
and the moderated variance estimation that `r Biocpkg("limma") ` performs.

We see that we get a moderate overlap of `r overlap_CD` for CD and
`r overlap_UC` for UC, showing that both analyses lead to
somewhat comparable results. 

## Visualization of DE analysis results - volcano plot

For a visualization of the differentially expressed genes, we create a volcano 
plot, which is commonly used to summarize the results of a differential expression analysis in a single figure.

For a better overview, we only show gene symbols of genes with 
a fold change greater than 1, which we define in the `volcano_names` object. The 
`highlight` option in the `volcanoplot` function is set to 100 and thus only 
labels the 100 genes with the lowest p-values. 

```{r VolcanoPlot, fig.cap="Volcano plot of the DE-genes", fig.height=8, fig.width=7}

volcano_names <- ifelse(abs(palmieri_fit_CD$coefficients)>=1, 
                        palmieri_fit_CD$genes$SYMBOL, NA)
             
             
volcanoplot(palmieri_fit_CD, coef = 1L, style = "p-value", highlight = 100, 
            names = volcano_names,
            xlab = "Log2 Fold Change", ylab = NULL, pch=16, cex=0.35)
```

We can now do a little research on the biological function of genes that show a
high foldchange, for example the gene with the symbol S100A8 on the right side
of the plot (Figure \@ref(fig:VolcanoPlot)). If we search for this gene symbol on
[genecards.org](https://www.genecards.org), we find that it encodes for a
protein that builds a pro-inflammatory complex in association with another
protein.

# Gene ontology (GO) based enrichment analysis

As discussed above, it is recommended to use an FDR cutoff in differential 
expression analysis rather than a p-value cutoff, 
since this way you control an explicitly defined error rate 
and the results are easier to interpret and
to compare. For the following enrichment analysis, we create tables with 
differentially expressed genes for CD and UC, respectively, 
and choose an FDR cutoff of 10\%. Here, we focus on the CD subset of the data.


```{r FDRcontrolledDEgenes, dependson=c("extractResultsUC", "extractResultsCD"), eval=TRUE}
DE_genes_CD <- subset(table_CD, adj.P.Val < 0.1)$PROBEID
```

We can now try to characterize the identified differentially expressed genes
more in detail by performing a GO enrichment analysis. Essentially the
gene ontology ([http://www.geneontology.org/](http://www.geneontology.org/)) is 
a hierarchically organized collection of functional gene sets
[@Ashburner_2000; @GO_2015; @du_Plessis_2011]. 

## Matching the background set of genes

The function ` genefinder ` from the `r Biocpkg("genefilter") ` 
package [@Bourgon_2010] 
will be used to find a background set of genes that are similar in expression 
to the differentially expressed genes. We then check whether 
the background has roughly the same distribution
of average expression strength as the foreground.

We do this in order not to select a biased background since the gene set testing
is performed by a simple Fisher test on a 2x2 table. Note that this approach
is very similar to commonly used web tools like GOrilla [@Eden_2009]. 

For every differentially expressed gene, we try to find genes with similar
expression with `genefinder`. The `genefinder` function returns a list with two 
elements for each gene: one with the indices of the 
background genes found and one with the distances to the DE-genes:

```{r GOAnalysisCreateBackgrounds, eval=TRUE, warning=FALSE, message=FALSE}
back_genes_idx <- genefilter::genefinder(palmieri_final, 
                                        as.character(DE_genes_CD), 
                                        method = "manhattan", scale = "none")
```

We have to extract the PROBEIDs, which correspond to the indices. 
We do that by using the `sapply` 
function, which gives us a single matrix with the DE-genes as column names 
and the  PROBEIDs of the corresponding background genes in the cells below:

```{r GOAnalysisCreateBackgrounds2, eval=TRUE, warning=FALSE, message=FALSE}
back_genes_idx <- sapply(back_genes_idx, function(x)x$indices)
```

We then create a vector `back_genes` containing all background gene PROBEIDs:



In order to eliminate foreground genes, i.e. DE-genes, from the `back_genes` 
set, we use the `setdiff` function. It returns all elements from the first 
argument (`back_genes`) that are not part of the second argument 
(`DE_genes_CD`). 
With the `intersect` function, we verify that we were successful: 
it should return 0, as there shouldn't be any intersect anymore between
`back_genes`and `DE_genes_CD`:

```{r GOAnalysisCreateBackgrounds3, eval=TRUE, warning=FALSE, message=FALSE}
back_genes <- featureNames(palmieri_final)[back_genes_idx]
back_genes <- setdiff(back_genes, DE_genes_CD)

    
intersect(back_genes, DE_genes_CD)
length(back_genes)
```

We create a multidensity plot with mean expression on the x-axis 
and curves for all genes, foreground genes and background genes, respectively 
(Figure \@ref(fig:multidensityPlot)). We want to see whether the background 
genes show a plot similar to the foreground genes so that the background 
is not biased for the gene enrichment analysis:

```{r multidensityPlot, fig.cap="Selecting a background set of genes for the gene ontology analysis."}
multidensity(list(
        all = table_CD[,"AveExpr"] ,
        fore = table_CD[DE_genes_CD , "AveExpr"],
        back = table_CD[rownames(table_CD) %in% back_genes, "AveExpr"]),
        col = c("#e46981", "#ae7ee2", "#a7ad4a"),
     xlab = "mean expression",
   main = "DE genes for CD-background-matching")
```

When comparing the "background gene" curve to the "foreground gene" curve, 
we see a  similar curve shape, indicating a sensible background matching 
(Figure \@ref(fig:multidensityPlot)). Note that the right-shift of the 
"foreground-gene" curve in comparison to the "background-gene" curve 
indicates that DE-genes are generally very
highly expressed, so that it wasn't possible to find background-genes 
with exactly equal overall expression distribution.

The "all gene" curve has the leftmost curve maximum; 
this can be explained by a high number of lowly expressed genes 
in all samples and shows that a background matching is sensible 
in order to avoid biases.

For the actual testing of which GO gene sets are enriched in inflamed tissue, 
we use the `r Biocpkg("topGO")` package which implements a nice interface 
to Fisher testing and also has additional algorithms
taking the GO structure into account, by e.g. only reporting the most specific
gene set in the hierarchy [@Alexa_2006].

The GO has three top ontologies: Cellular component (CC), biological processes
(BP), and molecular function (MF). For illustrative purposes we limit ourselves 
to the BP category here. 

## Running topGO

topGO requires a topGOdata object containing the necessary information for the 
analysis. We follow the steps described in the topGO vignettes: First, 
we will create a named vector `all_genes` with all genes to be analyzed, i.e. 
DE-genes and background genes: 

```{r createFactorOfInterestingGenes, dependson="GOAnalysisCreateBackgrounds", eval=TRUE}
gene_IDs <- rownames(table_CD)
in_universe <- gene_IDs %in% c(DE_genes_CD, back_genes)
in_selection <- gene_IDs %in% DE_genes_CD 

all_genes <- in_selection[in_universe]
all_genes <- factor(as.integer(in_selection[in_universe]))
names(all_genes) <- gene_IDs[in_universe] 
```

The following steps were carried through: 

  1. we created an `in_universe` vector by using the `%in%` matching function. 
  We want to know which elements from `gene_IDs` are also contained in 
  `DE_genes_CD` and `back_genes`, as the latter two are our gene universe 
  we use for enrichment analysis.
  We got a vector `in_universe` with the length of `gene_IDs` that has the 
  entry `TRUE` when the corresponding gene in `gene_IDs` could be also found in 
  `DE_genes_CD` or `back_genes`, and `FALSE` otherwise.
  
  2. We did the same for our DE-genes and call this vector `in_selection`.
  
  3. We created the `all_genes` vector:
    a) First, we selected all the elements from `in_selection` 
    that are `TRUE` in `in_universe` by applying
    `all_genes <- in_selection[in_universe]`.
    b) Then, we converted the elements in `all_genes` from `TRUE` and `FALSE` 
    to 0 and 1 by converting the vector to an integer vector. 
    This way, each element in the vector is a 0 if the corresponding gene is a
    background gene and a 1 if the corresponding gene is a DE-gene. Also, we 
    converted the vector to a factor.
    c) We named the vector elements with the corresponding `gene_IDs`.


We now initialize the `r Biocpkg("topGO") ` data set, using the GO annotations 
contained in the annotation data base for the chip we are using. The `nodeSize`
parameter specifies a minimum size of a GO category we want to use: i.e. here,
categories with less than 10 genes are not included in the testing.

```{r createTopGODataSet, dependson="createFactorOfInterestingGenes", eval=TRUE, message = FALSE, warning=FALSE}
top_GO_data <- new("topGOdata", ontology = "BP", allGenes = all_genes,
 nodeSize = 10, annot = annFUN.db, affyLib = "hugene10sttranscriptcluster.db")
```

Now the tests can be run. `r Biocpkg("topGO") ` offers a wide range of options,
for details see the paper [@Alexa_2006] or the package vignette.

We run two common tests: an ordinary Fisher test for every GO category, and the
"elim" algorithm, which tries to incorporate the hierarchical structure of the
GO and tries to "decorrelate" it in order to report the most specific 
significant term in the hierarchy.

The algorithm starts processing the nodes / GO categories
from the highest (bottommost) level and then iteratively
moves to nodes from a lower level. If a node is scored as significant,
all of its genes are marked as removed in all ancestor nodes.
This way, the "elim" algorithm aims at finding the most specific node
for every gene.

The test uses a 0.01 p-value cutoff by default.

```{r runtopGOTests, results='hide', eval=TRUE, dependson = "createTopGODataSet", message = FALSE}
result_top_GO_elim <- 
  runTest(top_GO_data, algorithm = "elim", statistic = "Fisher")
result_top_GO_classic <- 
  runTest(top_GO_data, algorithm = "classic", statistic = "Fisher")
```

We can now inspect the results. We look at the top 100 GO categories according
to the "Fisher elim" algorithm. The function `GenTable` produces
a table of significant GO categories, the function `printGenes`
gives genes annotated to them; the significant ones are denoted with a "2"
in the "raw  p-value" column, the non-significant ones with a "1". 
We therefore select `raw p-value == 2`. 

Note that we do not get the actual p-values here because our `all_genes` vector 
doesn't contain this information; it only tells us
whether a gene is differentially  expressed or not.

```{r processtopGOResults, eval=TRUE, dependson="runtopGOTests"}
res_top_GO <- GenTable(top_GO_data, Fisher.elim = result_top_GO_elim,
        Fisher.classic = result_top_GO_classic,
        orderBy = "Fisher.elim" , topNodes = 100)

genes_top_GO <- printGenes(top_GO_data, whichTerms = res_top_GO$GO.ID,
    chip = "hugene10sttranscriptcluster.db", geneCutOff = 1000)

res_top_GO$sig_genes <- sapply(genes_top_GO, function(x){
                str_c(paste0(x[x$'raw p-value' == 2, "Symbol.id"],";"), 
                      collapse = "")
    })

head(res_top_GO[,1:8], 20)
```

## Visualization of the GO-analysis results

A graph of the results can also be produced. Here we visualize the three most
significant nodes according to the Fisher elim algorithm in the context of
the GO hierarchy.


```{r graphOfResults, fig.height = 6, eval=TRUE, results='hide', dpi=600, fig.cap="Significantly enriched GO nodes in the GO hierarchy"}
showSigOfNodes(top_GO_data, score(result_top_GO_elim), firstSigNodes = 3,
               useInfo = 'def')
```

We can see that indeed GO categories related to inflammation, signalling and 
immune response come up as significant (Figure \@ref(fig:graphOfResults))
Gene set enrichment analysis has been a field of very extensive
research in bioinformatics. For additional approaches
see the `r Biocpkg("topGO") ` vignette and the references therein 
and also in the [GeneSetEnrichment view](http://bioconductor.org/packages/release/BiocViews.html#___GeneSetEnrichment). 


# A pathway enrichment analysis using reactome

The package `r Biocpkg("ReactomePA") ` offers the possibility to test enrichment
of specific pathways using the free, open-source, curated and peer reviewed 
[Reactome](http://www.reactome.org/) pathway database [@Croft_2013; @Fabregat_2017]. 
The package requires entrez identifiers, so we convert our PROBEIDs 
(transcript cluster identifiers) to entrez identifiers 
using the function `mapIDs` from the package `r Biocpkg("AnnotationDbi")`. 
This will create a named vector that maps the PROBEIDs to the entrez ones, 
with the PROBEIDs as names and the entrez ids as vector elements.


```{r mapIDsToEntrez, dependson="createFactorOfInterestingGenes", message = FALSE}
entrez_ids <- mapIds(hugene10sttranscriptcluster.db, 
      keys = rownames(table_CD), 
      keytype = "PROBEID",
      column = "ENTREZID")
```

We can now run the enrichment analysis that performs a statistical test
based on the hypergeoemtric distribution that is the same as a one sided 
Fisher-test, which `r Biocpkg("topGO")` calls "Fisher-classic".
Details can be found in the vignette of the `r Biocpkg("DOSE")` 
package [@Yu_2014].

```{r runReactomeEnrichment, fig.cap="Enriched Reactome pathways and their p–values as a bar chart.", eval = TRUE, warning=FALSE}
reactome_enrich <- enrichPathway(gene = entrez_ids[DE_genes_CD], 
                                universe = entrez_ids[c(DE_genes_CD, 
                                                        back_genes)],
                                organism = "human",
                                pvalueCutoff = 0.05,
                                qvalueCutoff = 0.9, 
                                readable = TRUE)

reactome_enrich@result$Description <- paste0(str_sub(
                                    reactome_enrich@result$Description, 1, 20),
                                    "...")

head(as.data.frame(reactome_enrich))[1:6]
```

Note that we trimmed pathway names to 20 characters.

## Visualizing the reactome based analysis results 

The top pathways can be displayed as a bar chart that displays all categories
with a p-value below the specified cutoff (Figure \@ref(fig:reactomeBar)).

```{r reactomeBar, dependson="runReactomeEnrichment", fig.cap="Enriched Reactome pathways and their p–values as a bar chart."}
barplot(reactome_enrich)
```

The "enrichment map" from the package `r Biocpkg("enrichplot")` 
displays the results of the enrichment analysis as 
a graph, where the color represents the p-value of the pathway and the
edge-thickness (that is the line connecting two pathways) is proportional 
to the number of overlapping genes between two pathways.

```{r emapplot, dependson="runReactomeEnrichment", fig.width=6, fig.height = 7, fig.cap="Enriched Reactome pathways enrichment results as a graph."}
emapplot(reactome_enrich, showCategory = 10)
```

Again, the graph in Figure \@ref(fig:emapplot) shows pathways related 
to signalling and immune response. 

The package `r Biocpkg("clusterProfiler") ` [@Yu_2012] can also perform these 
analyses using downloaded KEGG data. Furthermore, 
the package `r Biocpkg("EnrichmentBrowser")`
[@Geistlinger_2016] additionally offers network-based enrichment analysis of individual 
pathways. This allows the mapping of the expression data at hand to known
regulatory interactions.


# Session information

As the last part of this document, we call the function *sessionInfo*,
which reports the version numbers of R and all the packages used in
this session. It is good practice to always keep such a record of this
as it will help to track down what has happened in case an R script
ceases to work or gives different results because the functions have
been changed in a newer version of one of your packages. By including
it at the bottom of a script, your reports will become more reproducible.

The session information should also *always*
be included in any emails to the
[Bioconductor support site](https://support.bioconductor.org) along
with all code used in the analysis.

```{r}
gc()

length(getLoadedDLLs())

sessionInfo()
```


# Acknowledgements

The authors would like to thank Vladislava Milchevskaya, 
Julian Gehring and Mike Smith for helpful comments on and small contributions 
to the workflow. They also would like to thank Frederik Ziebell, Holly Giles and 
Jennifer Huellein for proof-reading.

This workflow draws a lot of inspiration from the Bioconductor 
books [@Bioc2005; @useRbook2008] as well as Love et. al.'s workflow 
for gene level analysis of RNA-Seq data [@Love_2016]. James W. MacDonald 
provided valuable information on the evolution of Affymetrix arrays in some of 
his posts of on the Biocondctor support site. 
BK would also like to thank him for some friendly personal 
correspondence about the annotation resources available
for microarrays in Bioconductor.



# Author Contributions

BK produced the first version of this article. 
BK implemented the main scaffold of the workflow and designed 
the workflow steps. SR implemented additional workflow steps and 
improved existing ones from version 1. BK and SR wrote the article.


# References