% NOTE -- ONLY EDIT THE .Rnw FILE!!! The .tex file is % likely to be overwritten. % %\VignetteEngine{knitr::knitr} %\VignetteIndexEntry{Basic Functions for Flow Cytometry Data} %\VignetteDepends{flowCore, flowViz} %\VignetteKeywords{} %\VignettePackage{flowCore} \documentclass[11pt]{article} \usepackage{times} \usepackage{hyperref} \usepackage[authoryear,round]{natbib} \usepackage{times} \usepackage{comment} \usepackage{graphicx} \usepackage{subfigure} \textwidth=6.2in \textheight=8.5in \oddsidemargin=.1in \evensidemargin=.1in \headheight=-.3in \newcommand{\scscst}{\scriptscriptstyle} \newcommand{\scst}{\scriptstyle} \newcommand{\Rfunction}[1]{{\texttt{#1}}} \newcommand{\Rcode}[1]{{\texttt{#1}}} \newcommand{\Robject}[1]{{\texttt{#1}}} \newcommand{\Rpackage}[1]{{\textsf{#1}}} \newcommand{\Rclass}[1]{{\textit{#1}}} \setkeys{Gin}{width=0.5\textwidth} \setkeys{Gin}{height=0.5\textwidth} \title{flowCore: data structures package for flow cytometry data} \author{N. Le Meur F. Hahne B. Ellis P. Haaland} \begin{document} %\SweaveOpts{concordance=TRUE} <>= library(knitr) opts_chunk$set( concordance=TRUE, warning = FALSE ) @ \maketitle \begin{abstract} \noindent \textbf{Background} The recent application of modern automation technologies to staining and collecting flow cytometry (FCM) samples has led to many new challenges in data management and analysis. We limit our attention here to the associated problems in the analysis of the massive amounts of FCM data now being collected. From our viewpoint, see two related but substantially different problems arising. On the one hand, there is the problem of adapting existing software to apply standard methods to the increased volume of data. The second problem, which we intend to address here, is the absence of any research platform which bioinformaticians, computer scientists, and statisticians can use to develop novel methods that address both the volume and multidimensionality of the mounting tide of data. In our opinion, such a platform should be Open Source, be focused on visualization, support rapid prototyping, have a large existing base of users, and have demonstrated suitability for development of new methods. We believe that the Open Source statistical software R in conjunction with the Bioconductor Project fills all of these requirements. Consequently we have developed a Bioconductor package that we call \Rpackage{flowCore}. The \Rpackage{flowCore} package is not intended to be a complete analysis package for FCM data. Rather, we see it as providing a clear object model and a collection of standard tools that enable R as an informatics research platform for flow cytometry. One of the important issues that we have addressed in the \Rpackage{flowCore} package is that of using a standardized representation that will insure compatibility with existing technologies for data analysis and will support collaboration and interoperability of new methods as they are developed. In order to do this, we have followed the current standardized descriptions of FCM data analysis as being developed under NIH Grant xxxx [n]. We believe that researchers will find flowCore to be a solid foundation for future development of new methods to attack the many interesting open research questions in FCM data analysis.\\ \noindent \textbf{Methods} We propose a variety of data structures. We have implemented the classes and methods in the Bioconductor package flowCore. We illustrate their use with X case studies.\\ \noindent \textbf{Results} We hope that those proposed data structures will be the base for the development of many tools for the analysis of high throughput flow cytometry.\\ \noindent \textbf{keywords} Flow cytometry, high throughput, software, standard \end{abstract} <>= library(flowCore) @ \section{Introduction} Traditionally, flow cytometry has been a tube-based technique limited to small-scale laboratory and clinical studies. High throughput methods for flow cytometry have recently been developed for drug discovery and advanced research methods \citep{Gasparetto2004}. As an example, the flow cytometry high content screening (FC-HCS) can process up to a thousand samples daily at a single workstation, and the results have been equivalent or superior to traditional manual multi-parameter staining and analysis techniques. The amount of information generated by high throughput technologies such as FC-HCS need to be transformed into executive summaries (which are brief enough) for creative studies by a human researcher \citep{Brazma2001}. Standardization is critical when developing new high throughput technologies and their associated information services \citep{Brazma2001, Chicurel2002, Boguski2003}. Standardization efforts have been made in clinical cell analysis by flow cytometry \citep{Keeney2004}, however data interpretation has not been standardized for even low throughput FCM. It is one of the most difficult and time consuming aspects of the entire analytical process as well as a primary source of variation in clinical tests, and investigators have traditionally relied on intuition rather than standardized statistical inference \citep{Bagwell2004,Braylan2004,Parks1997,Suni2003}. In the development of standards in high throughput FCM, little progress has been made in terms of Open Source software. In this article we propose R data structures to handle flow cytometry data through the main steps of preprocessing: compensation, transformation, filtering. The aim is to merge both \Rpackage{prada} and \Rpackage{rflowcyt} \citep{LeMeur2006} into one core package which is compliant with the data exchange standards that are currently developed in the community \citep{Spidlen2006}. Visualization as well as quality control will then be part of the utility packages that depend on the data structures defined in the flowCore package. \section{Representing Flow Cytometry Data} \Rpackage{flowCore}'s primary task is the representation and basic manipulation of flow cytometry (or similar) data. This is accomplished through a data model very similar to that adopted by other Bioconductor packages using the \Robject{expressionSet} and \Robject{AnnotatedDataFrame} structures familiar to most Bioconductor users. \subsection{The \Rclass{flowFrame} Class} The basic unit of manipulation in \Rpackage{flowCore} is the \Rclass{flowFrame}, which corresponds roughly to a single ``FCS'' file exported from the flow cytometer's acquisition software. At the moment we support FCS file versions 2.0 through 3.1, and we expect to support FCS4/ACS1 as soon as the specification has been ratified. \subsubsection{Data elements} The primary elements of the \Rclass{flowFrame} are the \Robject{exprs} and \Robject{parameters} slots, which contain the event-level information and column metadata respectively. The event information, stored as a single matrix, is accessed and manipulated via the \Rfunction{exprs()} and \Rfunction{exprs<-} methods, allowing \Rclass{flowFrame}s to be stitched together if necessary (for example, if the same tube has been collected in two acquisition files for memory reasons). The \Robject{parameters} slot is an \Rclass{AnnotatedDataFrame} that contains information derived from an FCS file's ``\$Pn'' keywords, which describe the detector and stain information. The entire list is available via the \Rfunction{parameter()} method, but more commonly this information is accessed through the \Rfunction{names}, \Rfunction{featureNames} and \Rfunction{colnames} methods. The \Rfunction{names} function returns a concatenated version of \Rfunction{colnames} and \Rfunction{featureNames} using a format similar to the one employed by most flow cytometry analysis software. The \Rfunction{colnames} method returns the detector names, often named for the fluorochrome detected, while the \Rfunction{featureNames} method returns the description field of the parameters, which will typically be an identifier for the antibody. The \Rfunction{keyword} method allows access to the raw FCS keywords, which are a mix of standard entries such as ``SAMPLE ID,'' vendor specific keywords and user-defined keywords that add more information about an experiment. In the case of plate-based experiments, there are also one or more keywords that identify the specific well on the plate. Most vendor software also includes some sort of unique identifier for the file itself. The specialized method \Rfunction{identifier} attempts to locate an appropriate globally unique identifier that can be used to uniquely identify a frame. Failing that, this method will return the original file name offering some assurance that this frame is at least unique to a particular session. \subsection{The \Rclass{flowSet} Class} Most experiments consist of several \Rclass{flowFrame} objects, which are organized using a \Rclass{flowSet} object. This class provides a mechanism for efficiently hosting the \Rclass{flowFrame}s with minimal copying. This reduces memory requirements and ensures that experimental metadata stay properly attached to their corresponding \Rclass{flowFrame}s. \section{Reading and Manipulating \Rpackage{flowCore} Data Classes} \subsection{Reading an FCS file into a \Rclass{flowFrame}} FCS files are read into the R environment via the \Rfunction{read.FCS} function using the standard connection interface---allowing for the possibility of accessing FCS files hosted on a remote resource as well as those that have been compressed or even retrieved as a blob from a database interface. FCS files (version 2.0, 3.0, and 3.1) and LMD (List Mode Data) extensions are currently supported. There are also several immediate processing options available in this function, the most important of which is the \Rfunction{transformation} parameter, which can ``linearize'' (the default), ``linearize-with-PnG-scaling'', or ``scale'' our data. To see how this works, first we will examine an FCS file without any transformation at all: <>= file.name <- system.file("extdata","0877408774.B08", package="flowCore") x <- read.FCS(file.name, transformation=FALSE) summary(x) @ As we can see, in this case the values from each parameter seem to run from $0$ to $1023$ ($2^{10}-1$). However, inspection of the documentation of the ``exponentiation'' keyword (\$PnE) reveals that some of the parameters (3 and 4) have been stored in a format specifying that the channel values should be scaled as $y=f_2\times 10^{f_1\cdot x/R}$ where $x$ is the original channel value, $y$ is the scaled value, $f_1$ and $f_2$ are given respectively by the first and second element of the value of the \$PnE key and $R$ is the channel range given by the value of the \$PnR key. The special \$PnE value of ``$0,0$'' corresponds to a scale that is already linear. <>= keyword(x,c("$P1E", "$P2E", "$P3E", "$P4E")) @ The default ``linearize'' transformation option will convert these to, effectively, have a \$PnE value of ``$0,0$'': <>= summary(read.FCS(file.name)) @ The ``linearize-with-PnG-scaling'' option will perform the previous transformation and it will also apply a ``division by gain'' to parameters stored on linear scale with specified gain. The gain is specified in the \$PnG keyword. This option has been introduced as part of Gating-ML 2.0 compliance. Finally, the ``scale'' option will both linearize values as well as ensure that output values are contained in $[0,1]$, which is the proposed method of data storage for the ACS1.0/FCS4.0 specification: <>= summary(read.FCS(file.name,transformation="scale")) @ Another parameter of interest is the \Rfunction{alter.names} parameter, which will convert the parameter names into more ``R friendly'' equivalents, usually by replacing ``-'' with ``.'': <>= read.FCS(file.name,alter.names=TRUE) @ When only a particular subset of parameters is desired, the \Rfunction{column.pattern} parameter allows for the specification of a regular expression and only parameters that match the regular expression will be included in the frame. For example, to include on the Height parameters: <>= x <- read.FCS(file.name, column.pattern="-H") x @ \textit{Note that \Rfunction{column.pattern} is applied after \Rfunction{alter.names} if it is used.} Finally, only a sample of lines can be read in case you need a quick overview of a large series of files. <>= lines <- sample(100:500, 50) y <- read.FCS(file.name, which.lines = lines) y @ \subsection{Creating a \Rclass{flowSet}} To facilitate the creation of \Rclass{flowSet} objects from a variety of sources, we provide a means to coerce \Rclass{list} and \Rclass{environment} objects to a \Rclass{flowSet} object using the usual coercion mechanisms. For example, if we have a directory containing FCS files, we can read in a list of those files and create a \Rclass{flowSet} out of them: <>= fcs.dir <- system.file("extdata", "compdata", "data", package="flowCore") frames <- lapply(dir(fcs.dir, full.names=TRUE), read.FCS) fs <- as(frames, "flowSet") fs sampleNames(fs) @ Note that the original list is unnamed and that the resulting sample names are not particularly meaningful. If the list is named, the list constructed is much more meaningful. One such approach is to employ the \Rfunction{keyword} method for \Rclass{flowFrame} objects to extract the ``SAMPLE ID'' keyword from each frame: <>= names(frames) <- sapply(frames, keyword, "SAMPLE ID") fs <- as(frames, "flowSet") fs sampleNames(fs) @ \subsubsection{Working with experimental metadata} Like most Bioconductor organizational classes, the \Rclass{flowSet} has an associated \Rclass{AnnotatedDataFrame} that provides metadata not contained within the \Rclass{flowFrame} objects themselves. This data frame is accessed and modified via the usual \Rfunction{phenoData} and \Rfunction{phenoData<-} methods. You can also generally treat the phenotypic data as a normal data frame to add new descriptive columns. For example, we might want to track the original filename of the frames from above in the phenotypic data for easier access: <>= phenoData(fs)$Filename <- fsApply(fs,keyword, "$FIL") pData(phenoData(fs)) @ Note that we have used the \Rclass{flowSet}-specific iterator, \Rfunction{fsApply}, which acts much like \Rfunction{sapply} or \Rfunction{lapply} and will be discussed shortly. Additionally, we should also note that the \Rfunction{phenoData} data frame \textbf{must} have row names that correspond to the original names used to create the \Rclass{flowSet}. \subsubsection{Reading multiple FCS files with \Rfunction{read.FlowSet}} The process of reading multiple FCS files in to a \Rclass{flowSet} is simplified by using the \Rfunction{read.flowSet} function. In its simplest incarnation, this function takes a \Rfunction{path}, which defaults to the current working directory, and an optional \Rfunction{pattern} argument that allows only a subset of files contained within the working directory to be selected. For example, to read a \Rclass{flowSet} of the files read in as \Robject{frames} above: <>= fs <- read.flowSet(path = fcs.dir) @ \Rfunction{read.flowSet} will pass on additional arguments meant for the underlying \Rfunction{read.FCS} function, such as \Rfunction{alter.names} and \Rfunction{column.pattern}, but also supports several other interesting arguments for conducting initial processing: \begin{description} \item[files]{ An alternative to the \Rfunction{pattern} argument, you may also supply a vector of filenames to read.} \item[name.keyword]{ Like the example in the previous section, you may specify a particular keyword to use in place of the filename when creating the \Rclass{flowSet}.} \item[phenoData]{ If this is an \Rclass{AnnotatedDataFrame}, then this will be used in place of the data frame that is ordinarily created. Additionally, the row names of this object will be taken to be the filenames of the FCS files in the directory specified by \Robject{path}. This argument may also be a named list made up of a combination of \Robject{character} and \Robject{function} objects that specify a keyword to extract from the FCS file or a function to apply to each frame that will return a result. } \end{description} To recreate the \Rclass{flowSet} that we created by hand from the last section we can use \Rfunction{read.flowSet}'s advanced functionality: <>= fs <- read.flowSet(path=system.file("extdata", "compdata", "data", package="flowCore"), name.keyword="SAMPLE ID", phenoData=list(name="SAMPLE ID", Filename="$FIL")) fs pData(phenoData(fs)) @ \subsubsection{Manipulating a \Rclass{flowSet}} You can extract a \Rclass{flowFrame} from a \Rclass{flowSet} object in the usual way using the \Rfunction{[[} or \Rfunction{\$} extraction operators. On the other hand using the \Rfunction{[} extraction operator returns a new \Rclass{flowSet} by \textbf{copying} the environment. However, simply assigning the \Rclass{flowFrame} to a new variable will \textbf{not} copy the contained frames. The primary iterator method for a \Rclass{flowSet} is the \Rfunction{fsApply} method, which works more-or-less like \Rfunction{sapply} or \Rfunction{lapply} with two extra options. The first argument, \Robject{simplify}, which defaults to \Robject{TRUE}, instructs \Rfunction{fsApply} to attempt to simplify its results much in the same way as \Rfunction{sapply}. The primary difference is that if all of the return values of the iterator are \Rclass{flowFrame} objects, \Rfunction{fsApply} will create a new \Rclass{flowSet} object to hold them. The second argument, \Robject{use.exprs}, which defaults to \Robject{FALSE} instructs \Rfunction{fsApply} to pass the expression matrix of each frame rather than the \Rclass{flowFrame} object itself. This allows functions to operate directly on the intensity information without first having to extract it. As an aid to this sort of operation we also introduce the \Rfunction{each\_row} and \Rfunction{each\_col} convenience functions that take the place of \Rfunction{apply} in the \Rfunction{fsApply} call. For example, if we wanted the median value of each parameter of each \Rclass{flowFrame} we might write: <>= fsApply(fs, each_col, median) @ % which is equivalent to the less readable <>= fsApply(fs,function(x) apply(x, 2, median), use.exprs=TRUE) @ % In this case, the \Robject{use.exprs} argument is not required in the first case because \Rfunction{each\_col} and \Rfunction{each\_row} are methods and have been defined to work on \Rclass{flowFrame} objects by first extracting the intensity data. \section{Visualizing Flow Cytometry Data} Much of the more sophisticated visualization of \Rclass{flowFrame} and \Rclass{flowSet} objects, including an interface to the \Rpackage{ggplot2} graphics system is implemented by the \Rpackage{ggcyto} package, also included as part of Bioconductor. Here, we will only introduce the \Rfunction{autoplot} function for the purpose of demonstrating \Rpackage{flowCore} operations. See the vignettes of \Rpackage{ggcyto} for more examples of how to visualize flow data. \subsection{Visualizing a single \Rclass{flowFrame}} It is frequently helpful to view the density of events in a \Rclass{flowFrame} across one channel or two. To create a bivariate density plot, simply provide the appropriate channels: <>= library(ggcyto) autoplot(x, "FL1-H", "FL2-H") @ Similarly, to get a univariate densityplot: <>= autoplot(x, "FL1-H") @ \subsection{Visualizing a \Rclass{flowSet}} The syntax is basically the same for \Rclass{flowSet} objects, with the output now being a grid of plots corresponding to each \Rclass{flowFrame} (which may or may not be useful depending on context). The grid layout can be adjusted with additional arguments to \Rfunction{autoplot} <>= fs <- read.flowSet(path = system.file("extdata", package = "flowCore"), pattern = "\\.") autoplot(fs, "FL1-H", "FL2-H") @ \section{Compensation} Before proceeding with further analysis of the data in a \Rclass{flowFrame} or \Rclass{flowSet}, it is important to properly compensate the data for spectral overlap between fluorescence channels. \subsection{Extracting and applying a pre-calculated spillover matrix} If the original FCS file contains a pre-calculated spillover matrix as the value of the \$SPILLOVER or \$SPILL keywords, this can be accessed from the \Rclass{flowFrame}'s \Robject{description} slot using the \Rfunction{spillover} method: <>= fcsfiles <- list.files(pattern = "CytoTrol", system.file("extdata", package = "flowWorkspaceData"), full = TRUE) fs <- read.flowSet(fcsfiles) x <- fs[[1]] comp_list <- spillover(x) comp_list comp <- comp_list[[1]] @ Given a \Rclass{flowFrame}, \Rfunction{spillover} will check a few valid spillover matrix keywords. This will result in a list, usually only one of which will be valid (in this case, the first one, corresponding to the \$SPILL keyword). This spillover matrix can then be applied to the channels of the \Rclass{flowFrame} using the \Rfunction{compensate} function. <>= x_comp <- compensate(x, comp) @ The \Rfunction{compensate} function can also take a \Rclass{flowSet} as its first argument and a list of named spillover matrices to compensate the corresponding \Rclass{flowFrame}s. Here the \Rcode{simplify = FALSE} option is necessary so that \Rfunction{fsApply} returns a list of named matrices. <>= comp <- fsApply(fs, function(x) spillover(x)[[1]], simplify=FALSE) fs_comp <- compensate(fs, comp) @ While transformation will be discussed in the next section, it is helpful to visualize the effect of compensation on transformed parameters. Quick inspection of the compensation matrix above reveals large overlap between the 450nm and 545nm channels, so we would expect to see a significant difference in each of those channels after compensation. This is in fact the case: <>= library(gridExtra) transList <- estimateLogicle(x, c("V450-A","V545-A")) p1 <- autoplot(transform(x, transList), "V450-A", "V545-A") + ggtitle("Before") p2 <- autoplot(transform(x_comp, transList), "V450-A", "V545-A") + ggtitle("Before") grid.arrange(as.ggplot(p1), as.ggplot(p2), ncol = 2) @ Often the spillover matrix is not provided in the FCS file for each analyzed sample and instead it is necessary to work with a set of FCS files representing the controls from which the spillover matrix is to be calculated, as follows. \subsection{Computing a spillover matrix from a set of compensation controls} The spillover matrix can also be calculated using a set of FCS files that contain data for an unstained sample and singly-stained samples for each of the measurement channels. If these files are not already incorporated as \Rclass{flowFrame}s in a \Rclass{flowSet} object, this must be done first. Here we also are assigning the appropriate channel name for each control file before coercion of the frames to a \Rclass{flowSet}. This spillover calculation method is now part of the \Rpackage{flowStats} package. <>= library(flowStats) fcs.dir <- system.file("extdata", "compdata", "data", package="flowCore") frames <- lapply(dir(fcs.dir, full.names=TRUE), read.FCS) names(frames) <- c("UNSTAINED", "FL1-H", "FL2-H", "FL4-H", "FL3-H") frames <- as(frames, "flowSet") comp <- spillover(frames, unstained="UNSTAINED", patt = "-H", fsc = "FSC-H", ssc = "SSC-H", stain_match = "ordered") comp @ Note that when using the \Rcode{stain\_match = "ordered"} option of \Rfunction{spillover}, we are specifying that the method should assume the singly-stained \Rclass{flowFrame}s in the \Rclass{flowSet} are in the order of the channel columns. In this case this behavior is incorrect as can be seen by quick inspection of the resulting matrix. To override this behavior and use the names we have provided, use the \Rcode{stain\_match = "regexpr"} option instead. This will use regular expression matching of the channel names to the \Rclass{flowFrame} names to achieve the proper ordering. <>= sampleNames(frames) comp <- spillover(frames, unstained="UNSTAINED", patt = "-H", fsc = "FSC-H", ssc = "SSC-H", stain_match = "regexpr") comp @ As an alternative way to match channels to their corresponding compensation controls, we provide the \Rfunction{spillover\_match} method. This allows for the matching of control files to specific channels via a simple csv file, resulting in a \Rclass{flowSet} ready for \Rfunction{spillover}. Lastly, it is acceptable to omit the \Rclass{flowSet} argument \Rcode{x} to both \Rcode{spillover} and \Rcode{spillover\_match} if you provide the \Rcode{path} argument to point to a directory containing the files to be used in the compensation \Rclass{flowSet}: <>= comp_match <- system.file("extdata", "compdata", "comp_match", package="flowCore") # The matchfile has a simple format writeLines(readLines(comp_match)) control_path <- system.file("extdata", "compdata", "data", package="flowCore") # Using path rather than pre-constructed flowSet matched_fs <- spillover_match(path=control_path, fsc = "FSC-H", ssc = "SSC-H", matchfile = comp_match) comp <- spillover(matched_fs, fsc = "FSC-H", ssc = "SSC-H", prematched = TRUE) @ % Note for the \Rfunction{spillover\_match} function % to work correctly, the sample names of each of the \Rclass{flowFrame}s % should be their corresponding file names. Recall that this is the default % behavior of \Rfunction{read.flowSet}. This spillover matrix can then be used to compensate a sample \Rclass{flowFrame} or an entire \Rclass{flowSet}. % PICKUP FOLLOWING LEAD OF http://127.0.0.1:23643/library/flowWorkspace/doc/flowWorkspace-Introduction.html \section{Transformation} \Rpackage{flowCore} features two methods of transforming parameters within a \Rclass{flowFrame} or \Rclass{flowSet}: inline and out-of-line. The inline method, discussed in the next section, has been developed primarily to support filtering features and is strictly more limited than the out-of-line transformation method, which uses R's \Rfunction{transform} function to accomplish the filtering. Like the normal \Rfunction{transform} function, the \Rclass{flowFrame} is considered to be a data frame with columns named for parameters of the FCS file. For example, if we wished to plot our first \Rclass{flowFrame}'s first two fluorescence parameters on the log scale we might write: <>= fs <- read.flowSet(path=system.file("extdata", "compdata", "data", package="flowCore"), name.keyword="SAMPLE ID", phenoData=list(name="SAMPLE ID", Filename="$FIL")) autoplot(transform(fs[[1]], `FL1-H`=log(`FL1-H`), `FL2-H`=log(`FL2-H`) ), "FL1-H","FL2-H") @ Like the usual \Rfunction{transform} function, we can also create new parameters based on the old parameters, without destroying the old <>= autoplot(transform(fs[[1]], log.FL1.H=log(`FL1-H`), log.FL2.H=log(`FL2-H`) ), "log.FL1.H", "log.FL2.H") @ \subsection{Standard Transforms} Though any function can be used as a transform in both the out-of-line and inline transformation techniques, \Rpackage{flowCore} provides a number of parameterized transform generators that correspond to the transforms commonly found in flow cytometry and defined in the Transformation Markup Language (Transformation-ML, see \url{http://www.ficcs.org/} and \cite{Spidlen2006} for more details). Briefly, the predefined transforms are: \begin{description} \item[truncateTransform]{ $y = \left\{\begin{array}{ll} a & x < a \\ x & x \geq a \\ \end{array}\right.$ } \item[scaleTransform]{$ f(x) = \frac{x-a}{b-a}$} \item[linearTransform]{ $ f(x) = a+bx$ } \item[quadraticTransform]{ $ f(x) = ax^2 + bx + c$ } \item[lnTransform]{$ f(x) = \log\left(x\right)\frac{r}{d} $} \item[logTransform]{$ f(x) = \log_b\left(x\right)\frac{r}{d} $} \item[biexponentialTransform]{ $ f^{-1}(x) = ae^{bx}-ce^{dx}+f$} \item[logicleTransform]{ A special form of the biexponential transform with parameters selected by the data.} \item[arcsinhTransform]{ $ f(x) = asinh\left(a+bx\right)+c$} \end{description} To use a standard transform, first we create a transform function via the constructors supplied by \Rpackage{flowCore}: <>= aTrans <- truncateTransform("truncate at 1", a=1) aTrans @ which we can then use on the parameter of interest in the usual way, in this case for the full \Rclass{flowSet} using the same syntax <>= transform(fs,`FL1-H`=aTrans(`FL1-H`)) @ However this form of transform call is not intended to be used in the programmatic context because a locally defined transform function (e.g. 'aTrans') may not always be visible to the non-standard evaluation environment. .e.g <>= f1 <- function(fs,...){ transform(fs, ...)[,'FL1-H'] } f2 <- function(fs){ aTrans <- truncateTransform("truncate at 1", a=1) f1(fs, `FL1-H` = aTrans(`FL1-H`)) } res <- try(f2(fs), silent = TRUE) res @ So this form of usage of 'transform' method is only useful for the interactive exploratory. we highly recommend the usage of \Robject{transformList} instead for more robust and reproducible code. <>= myTrans <- transformList('FL1-H', aTrans) transform(fs, myTrans) @ \section{Gating} The most common task in the analysis of flow cytometry data is some form of filtering operation, also known as gating, either to obtain summary statistics about the number of events that meet certain criteria or to perform further analysis on a subset of the data. Most filtering operations are a composition of one or more common filtering operations. The definition of gates in \Rpackage{flowCore} follows the Gating Markup Language Candidate Recommendation \cite{Spidlen2008}, thus any \Rpackage{flowCore} gating strategy can be reproduced by any other software that also adheres to the standard and \textit{vice versa}. \subsection{Standard gates and filters} Like transformations, \Rpackage{flowCore} includes a number of built-in common flow cytometry gates. The simplest of these gates are the geometric gates, which correspond to those typically found in interactive flow cytometry software: \begin{description} \item[rectangleGate]{Describes a cubic shape in one or more dimensions--a rectangle in one dimension is simply an interval gate.} \item[polygonGate]{Describes an arbitrary two dimensional polygonal gate.} \item[polytopeGate]{Describes a region that is the convex hull of the given points. This gate can exist in dimensions higher than $2$, unlike the \Robject{polygonGate}.} \item[ellipsoidGate]{Describes an ellipsoidal region in two or more dimensions} \end{description} These gates are all described in more or less the same manner (see man pages for more details): <>= rectGate <- rectangleGate(filterId="Fluorescence Region", "FL1-H"=c(0, 12), "FL2-H"=c(0, 12)) @ In addition, we introduce the notion of data-driven gates, or filters, not usually found in flow cytometry software. In these approaches, the necessary parameters are computed based on the properties of the underlying data, for instance by modeling data distribution or by density estimation: \begin{description} \item[norm2Filter]{ A robust method for finding a region that most resembles a bivariate Normal distribution. Note that this method now resides in the \Rpackage{flowStats} package} \item[kmeansFilter]{ Identifies populations based on a one dimensional k-means clustering operation. Allows the specification of \textbf{multiple} populations. } \end{description} \subsection{Count Statistics} When we have constructed a filter, we can apply it in two basic ways. The first is to collect simple summary statistics on the number and proportion of events considered to be contained within the gate or filter. This is done using the \Rfunction{filter} method. The first step is to apply our filter to some data <>= result = filter(fs[[1]],rectGate) result @ As we can see, we have returned a \Rclass{filterResult} object, which is in turn a filter allowing for reuse in, for example, subsetting operations. To obtain count and proportion statistics, we take the summary of this \Rclass{filterResult}, which returns a list of summary values: <>= summary(result) summary(result)$n summary(result)$true summary(result)$p @ A filter which contains multiple populations, such as the \Rclass{kmeansFilter}, can return a list of summary lists: <>= summary(filter(fs[[1]], kmeansFilter("FSC-H"=c("Low", "Medium", "High"), filterId="myKMeans"))) @ A filter may also be applied to an entire \Rclass{flowSet}, in which case it returns a list of \Rclass{filterResult} objects: <>= filter(fs,rectGate) @ \subsection{Subsetting} To subset or split a \Rclass{flowFrame} or \Rclass{flowSet}, we use the \Rfunction{Subset} and \Rfunction{split} methods respectively. The first, \Rfunction{Subset}, behaves similarly to the standard R \Rfunction{subset} function, which unfortunately could not be used. For example, the morphology parameters, Forward Scatter and Side Scatter, contain a large more-or-less ellipse shaped population: <>= autoplot(fs[[1]], "FSC-H", "SSC-H") @ If we wished to deal only with that population, we might use \Rfunction{Subset} along with a \Rclass{norm2Filter} object as follows: <>= library(flowStats) morphGate <- norm2Filter("FSC-H", "SSC-H", filterId="MorphologyGate", scale=2) smaller <- Subset(fs, morphGate) fs[[1]] smaller[[1]] @ Notice how the \Robject{smaller} \Rclass{flowFrame} objects contain fewer events. Now imagine we wanted to use a \Rclass{kmeansFilter} as before to split our first fluorescence parameter into three populations. To do this we employ the \Rfunction{split} function: <>= split(smaller[[1]], kmeansFilter("FSC-H"=c("Low","Medium","High"), filterId="myKMeans")) @ or for an entire \Rclass{flowSet} <>= split(smaller, kmeansFilter("FSC-H"=c("Low", "Medium", "High"), filterId="myKMeans")) @ \subsection{Combining Filters} Of course, most filtering operations consist of more than one gate. To combine gates and filters we use the standard R Boolean operators: \&, | and ! to construct an intersection, union and complement respectively: <>= rectGate & morphGate rectGate | morphGate !morphGate @ We also introduce the notion of the subset operation, denoted by either \%subset\% or \%\&\%. This combination of two gates first performs a subsetting operation on the input \Rclass{flowFrame} using the right-hand filter and then applies the left-hand filter. For example, <>= summary(filter(smaller[[1]],rectGate %&% morphGate)) @ first calculates a subset based on the \Robject{morphGate} filter and then applies the \Robject{rectGate}. \subsection{Transformation Filters} Finally, it is sometimes desirable to construct a filter with respect to transformed parameters. To allow for this in our filtering constructs we introduce a special form of the \Rfunction{transform} method along with another filter combination operator \%on\%, which can be applied to both filters and \Rclass{flowFrame} or \Rclass{flowSet} objects. To specify our transform filter we must first construct a transform list using a simplified version of the \Rfunction{transform} function: <>= tFilter <- transform("FL1-H"=log,"FL2-H"=log) tFilter @ Note that this version of the transform filter does not take parameters on the right-hand side--the functions can only take a single vector that is specified by the parameter on the left-hand side. In this case those parameters are ``FL1-H'' and ``FL2-H.'' The function also does not take a specific \Rclass{flowFrame} or \Rclass{flowSet} allowing us to use this with any appropriate data. We can then construct a filter with respect to the transform as follows: <>= rect2 <- rectangleGate(filterId="Another Rect", "FL1-H"=c(1,2), "FL2-H"=c(2,3)) %on% tFilter rect2 @ Additionally, we can use this construct directly on a \Rclass{flowFrame} or \Rclass{flowSet} by moving the transform to the left-hand side and placing the data on the right-hand side: <>= autoplot(tFilter %on% smaller[[1]], "FL1-H","FL2-H") @ which has the same effect as the $\log$ transform used earlier. \section{Extensions from \Rpackage{flowWorkspace}: \Rclass{GatingHierarchy} and \Rclass{GatingSet}} The now-defunct \Rclass{filterSet} class was very limited in its use for complex analysis work flows. It was result-centric and it was hard to access intermediate results. \Rpackage{flowWorkspace} and the \Rpackage{openCyto} framework (http://opencyto.org) offer much more versatile tools for such tasks through the \Rclass{GatingSet} class. The general idea is to let the software handle the organization of intermediate results and operations and to provide a unified API to access and summarize these operations. \subsection{Abstraction of GatingSet} There are two classes in \Rpackage{flowWorkspace} that are used to abstract work flows: \Rclass{GatingSet} objects are the basic container holding all the necessary bits and pieces and they are the main structure for user interaction. It is the container storing multiple \Rclass{GatingHierarchy} objects which are associated with individual samples. One can think of \Rclass{GatingSet} as corresponding to \Rclass(flowSet) and \Rclass{GatingHierarchy} as corresponding to \Rclass(flowFrame). It is important to know that \Rclass{GatingSet} objects use an external pointer to store the gating tree and thus most of its accessors have a reference semantic instead of the pass-by-value semantic that is usually found in the R language. The main consequence on the user-level is the fact that direct assignments to a \Rclass{GatingSet} object are usually not necessary; i.e., functions that operate on the \Rclass{GatingSet} have the potential side-effect of modifying the object. \subsection{Creating \Rclass{GatingSet} objects} Before creating a \Rclass{GatingSet}, we need to have flow data loaded into R as a \Rclass{flowSet} or \Rclass{ncdfFlowSet} (the disk-based version of \Rclass{flowSet}, to handle large data sets that are too big for memory; see \Rpackage{ncdfFlow} package). <>= library(flowWorkspace) fcsfiles <- list.files(pattern = "CytoTrol", system.file("extdata", package = "flowWorkspaceData"), full = TRUE) fs <- read.flowSet(fcsfiles) @ Then a \Rclass{GatingSet} can be created using the constructor \Rfunction{GatingSet}. <>= gs <- GatingSet(fs) gs @ Normally, we want to compensate the data first by using a user-supplied compensation matrix: <>= gs_manual <- load_gs(list.files(pattern = "gs_manual", system.file("extdata", package = "flowWorkspaceData"), full = TRUE)) comp <- gh_get_compensations(gs_manual[[1]]) @ <>= comp gs <- compensate(gs, comp) fs_comp <- getData(gs) @ We can query the available nodes in the \Rclass{GatingSet} using the \Rfunction{gs\_get\_pop\_paths} method: <>= gs_get_pop_paths(gs) @ It shows the only node ``root'' which corresponds to the raw flow data just added. \subsection{transform the data} Transformation can be either done on a \Rclass{flowSet} before using it to construct a \Rclass{GatingSet} or a \Rclass{transformerList} can be directly added to a \Rclass{GatingSet}: <>= biexpTrans <- flowJo_biexp_trans(channelRange=4096, maxValue=262144 , pos=4.5,neg=0, widthBasis=-10) chnls <- parameters(comp) tf <- transformerList(chnls, biexpTrans) # or use estimateLogicle directly on GatingHierarchy # object to generate transformerList automatically: # tf <- estimateLogicle(gs[[1]], chnls) gs <- transform(gs, tf) @ <>= p1 <- autoplot(fs_comp[[1]], "B710-A") + ggtitle("raw") p2 <- autoplot(gs_cyto_data(gs)[[1]], "B710-A") + ggtitle("trans") + ggcyto_par_set(limits = "instrument") grid.arrange(as.ggplot(p1), as.ggplot(p2), ncol = 2) @ Note that we did assign the return value of \Rfunction{transform} back to \Robject{gs}. This is because the flow data is stored as an R object and thus transforming the data still follows the pass-by-value semantics. \subsection{Add the gates} Some basic flowCore \Rclass{filter}s can be added to a \Rclass{GatingSet}: <>= rg1 <- rectangleGate("FSC-A"=c(50000, Inf), filterId="NonDebris") gs_pop_add(gs, rg1, parent = "root") gs_get_pop_paths(gs) # gate the data recompute(gs) @ As we see, here we don't need to assign \Rclass{GatingSet} back because all the modifications are made in place to the \Rclass{external pointer} rather than the R object itself. And now there is one new population node under the ``root'' node called ``NonDebris''. The node is named after the \Robject{filterId} of the gate if not explictly supplied. After the gates are added, the actual gating process is done by explictly calling the \Rfunction{recompute} method. Note that the numeric value it returns is the internal ID for the new population just added, which can normally be ignored since the gating path is the recommended way to refer to population nodes later. To view the gate we just added, <>= autoplot(gs, "NonDebris") @ Since They are 1-D gates, we can also display them in density plots <>= ggcyto(gs, aes(x = `FSC-A`)) + geom_density() + geom_gate("NonDebris") @ To get population statistics for the given populatuion <>= gh_pop_get_stats(gs[[1]], "NonDebris")#counts gh_pop_get_stats(gs[[1]], "NonDebris", type = "percent")#proportion @ Now we add two more gates: <>= # add the second gate mat <- matrix(c(54272,59392,259071.99382782 ,255999.994277954,62464,43008,70656 ,234495.997428894,169983.997344971,34816) , nrow = 5) colnames(mat) <-c("FSC-A", "FSC-H") mat pg <- polygonGate(mat) gs_pop_add(gs, pg, parent = "NonDebris", name = "singlets") # add the third gate rg2 <- rectangleGate("V450-A"=c(2000, Inf)) gs_pop_add(gs, rg2, parent = "singlets", name = "CD3") gs_get_pop_paths(gs) @ We see two more nodes are added to 'GatingSet' and the population names are explicitly specified during the addition this time. A \Rclass{quadrantGate}, which results in four sub-populations, is also supported. <>= qg <- quadGate("B710-A" = 2000, "R780-A" = 3000) gs_pop_add(gs, qg, parent="CD3", names = c("CD8", "DPT", "CD4", "DNT")) gs_pop_get_children(gs[[1]], "CD3") # gate the data from "singlets" recompute(gs, "singlets") @ Here we see four child nodes are added to the ``CD3'' parent node. Four quadrants are named explicitly through the \Robject{names} argument by clock-wise order (start from top-left quadrant). Recomputing only needs to be done once from the first ungated node, which will automatically compute all of its descendants. To plot the underlying tree: <>= plot(gs) @ <>= if(suppressWarnings(require(Rgraphviz))){ plot(gs) }else{ plot(1,1, type="n", axes=FALSE, ann=FALSE) text(1,1,"Need to install Rgraphviz") } @ To plot all gates for one sample: <>= autoplot(gs[[1]]) @ To retreive the underlying flow data for a gated \Rclass{population}: <>= fs_nonDebris <- getData(gs, "NonDebris") fs_nonDebris nrow(fs_nonDebris[[1]]) nrow(fs[[1]]) @ To get all the population statistics: <>= gs_pop_get_count_fast(gs) @ \subsection{Removing nodes from a \Rclass{GatingSet} object} There are dependencies between \Rclass{nodes} in the hierarchical structure of a \Rclass{GatingSet} object. Thus, removing a particular node means also removing all of its associated child \Rclass{nodes}. <>= Rm('CD3', gs) gs_get_pop_paths(gs) Rm('NonDebris', gs) gs_get_pop_paths(gs) @ Now for the larger data set, it would be either inaccurate to apply the same hard-coded gate to all samples or impractical to manually set the gate coordinates for each individual sample. \Rpackage{openCyto}(\citep{Finak2014}) provides some data-driven gating functions to automatically generate these gates. For example, \Rfunction{mindensity} can be used for estimating a ``nonDebris'' gate for each sample. <>= if(require(openCyto)){ thisData <- getData(gs) nonDebris_gate <- fsApply(thisData, function(fr) openCyto:::.mindensity(fr, channels = "FSC-A")) gs_pop_add(gs, nonDebris_gate, parent = "root", name = "nonDebris") recompute(gs) } @ \Rfunction{singletGate} can be used for estimating ``singlets" <>= if(require(openCyto)){ thisData <- getData(gs, "nonDebris") #get parent data singlet_gate <- fsApply(thisData, function(fr) openCyto:::.singletGate(fr, channels =c("FSC-A", "FSC-H"))) gs_pop_add(gs, singlet_gate, parent = "nonDebris", name = "singlets") recompute(gs) } @ and then \Rfunction{mindensity} can be used again for the "CD3" gate <>= if(require(openCyto)){ thisData <- getData(gs, "singlets") #get parent data CD3_gate <- fsApply(thisData, function(fr) openCyto:::.mindensity(fr, channels ="V450-A")) gs_pop_add(gs, CD3_gate, parent = "singlets", name = "CD3") recompute(gs) } @ and then we can use the more adanced version of \Rfunction{quadrantGate}: \Rfunction{quadGate.seq} for gating "CD4" and "CD8" sequentially: <>= if(require(openCyto)){ thisData <- getData(gs, "CD3") #get parent data Tsub_gate <- fsApply(thisData, function(fr) openCyto::quadGate.seq(fr, channels = c("B710-A", "R780-A"), gFunc = 'mindensity') ) gs_pop_add(gs, Tsub_gate, parent = "CD3", names = c("CD8", "DPT", "CD4", "DNT")) recompute(gs) } @ and then plot the gates <>= autoplot(gs[[1]]) @ Note that in order to get parent gated data by \Rfunction{getData}, we have to call \Rfunction{recompute} after adding each gate. The result is very similar to the manual gates but the gating process is more data-driven and more consistent across samples. To further automate the process, a gating pipeline can be established through \Rpackage{OpenCyto}(\citep{Finak2014}) that defines the hierarchical gating template in a text-based csv file. (See more details from http://openCyto.org) \clearpage \bibliographystyle{plainnat} \bibliography{../inst/doc/cytoref} \end{document}