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The hierarchical classification of nature initiated by Carl Linnaeus today consists of eight major “ranks”, namely species, genus, family, order, class, phylum, kingdom and domain:

01_14ClassifyingLife_L

In the microbial world it makes sense to refine the standard taxonomy by subdividing species into strains. An important reason to do so is that bacterial taxonomy must reflect not only phylogeny but also pathogenicity, and small differences in genomes can translate to large pathogenic differences. This has implications for metagenomic analyses of microbial communities: for many biomedical applications it is desirable to characterize individuals strains.

Metagenomics has its roots in culture-independent retrieval and sequencing of 16S rRNA genes, and while variations in 16S can sometimes distinguish between strains, a single gene is not always sufficient. This limitation of 16S can be overcome with whole genome shotgun sequencing of microbial communities, an approach to metagenomics that became popular in the early 2000s and  that opened the door to higher resolution characterization of communities. In 2005 Kevin Chen and I wrote a review on the bioinformatics challenges that would have to be overcome to walk through the door. One of the things we did was to emphasize “problems and their connections to other areas of bioinformatics, such as… gene expression analysis”, and throughout the past decade I’ve always hoped for deeper connections to be established between metagenomics and gene expression bioinformatics. I’ve noticed interesting connections pop up from time to time (e.g. Paulson et al. 2013)  and have occasionally entertained the thought with my students and collaborators, especially as work in my group became more focused on RNA-Seq since the development of Cufflinks in 2008.

However connection modern transcriptome analysis methodology, specifically bioinformatics of RNA-Seq to metagenomics has been difficult to do until recently. One major reason is that until just a few years ago, there was no reference genome database for metagenomics analogous to the reference annotation databases available for use in transcriptomics. Another way to put this is that metagenomics has, until recently, been “de novo” bioinformatics. By this I mean that the analysis of communities from whole genome shotgun data had to largely proceed via de novo analyses of the data (e.g. de novo assembly of genomes), “binning” of reads according to sequence characteristics or hits to gene databases was required because it was impossible to compare sequences to references genomes. While de novo methods have also been developed for RNA-Seq, the scale of transcriptome analysis is much smaller than that of most metagenomic analyses, and as has been well documented, de novo transcriptomics is already very difficult (e.g. Amin et al. 2014).

The de novo state of metagenomics has changed in recent years, as (relatively) low-cost sequencing has been a boon for microbial genomics. The graph below, extracted from NCBI and published in a recent review, shows that in just the past few years thousands of bacterial genomes have been sequences, enabling, for the first time, reference based metagenomics:

Number_genomes

This observation is reflected in the recent development of many methods for a variety of metagenomic applications that take advantage of reference genome databases.  Specifically, the problem of read assignment, which is fundamental for abundance estimation, has benefited from the possibility of metagenomic read alignment to reference databases.

The figure below, reproduced from the preprint “An evaluation of the accuracy and speed of metagenome analysis tools” by Stinus Lindgreen, Karen L. Adair and Paul Gardner, bioRxiv May 15, 2015 shows a benchmark of the accuracy and runtime of 14 programs developed for metagenomic read assignment for whole genome shotgun data:

Performance_Lindgreen

The problem these methods are solving is really similar to the problem of read assignment in RNA-Seq. In RNA-Seq, instead of originating from strains, reads originate from transcripts. Just as strains are present in different abundances in a community, so are RNA transcripts in a cell (or in bulk). The analogy of taxonomy in metagenomics, i.e. the grouping of strains into species, genus etc. is also present in RNA-Seq, where transcripts are grouped into genes. The fragment (or read) assignment problem in RNA-Seq is closely related to the quantification problem in RNA-Seq and is a problem that has been thoroughly researched and for which many algorithms have been developed. I discussed the importance of the fragment assignment problem for RNA-Seq in my 2013 Genome Informatics Keynote.

In response to the development of reference-based bioinformatics possibilities for metagenomics, about three years ago my student Lorian Schaeffer started looking at the suitability of RNA-Seq tools for metagenomic read assignment. Although the metagenomic and RNA-Seq assignment problems are conceptually similar and methodologically related, there are various technical issues involved in applying RNA-Seq tools in the metagenomic setting (e.g. the need to carefully account for taxonomy in the metagenomics setting). After developing the computational infrastructure to benchmark RNA-Seq programs in the metagenomic setting, she proceeded to evaluate the accuracy of eXpress, a streaming algorithm for RNA-Seq quantification. Although the quantification of eXpress was specifically designed to be suitable for large numbers of reads, the program requires read alignments to a reference transcriptome (or in Lorian’s experiments a genome) database. In the metagenomic setting realistic databases are huge, and she found that it took days just to map the reads. Nevertheless, her initial benchmarks revealed that eXpress was significantly more accurate than the available metagenomic read assignment tools of the time.

When Kraken (Wood and Salzberg 2014), and later CLARK (Ounit et al. 2015) were published in 2014 and 2015 respectively, we took note because by circumventing the alignment step they dramatically altered the tractability of metagenomic read assignment. In parallel, in my group, Nicolas Bray and later Páll Melsted and Harold Pimentel were developing what is now kallisto (Bray et al. 2015). Like Kraken, kallisto avoided the need for aligning reads, but with the introduction of the concept of pseudoalignment, allowed for accurate read assignments based on joint analysis of exact k-mer matches. What we showed earlier this year is that unlike naïve k-mer based approaches to quantification, kallisto is as accurate as eXpress and other read alignment based quantification tools, and this observation led Lorian to immediately proceed to benchmark it on metagenomic data. The result of her work was just posted as a preprint:

Lorian Schaeffer, Harold Pimentel, Nicolas Bray, Páll Melsted and Lior Pachter, Pseudoalignment for metagenomic read assignment, arXiv 1510.07371, 2015.

With this paper we demonstrate a “technology transfer” from RNA-Seq bioinformatics to metagenomics, one that achieves dramatic improvements in read assignment accuracy in the metagenomics setting. The main result of her work is Table 1 in our preprint:

Table1_kallisto_metagenomics

Using a published simulated Illumina dataset from Mende et al. 2012 (based on 100 genomes and containing 53.33 million reads), and augmenting it with another 2,308 genomes for the purpose of testing, she shows that kallisto significantly outperforms the best quantification methods (as benchmarked by Lindgreen et al., see figure above). “Significant” here refers to what I think is fair to characterize as an extraordinary improvement: at the genus level, a level that programs such as CLARK have been optimized for, kallisto’s RRMSE (relative root mean squared error)  is 0.13 compared to 17.05 for Kraken and 18.58 for CLARK. The improvement is based on two ideas: first, the results show that the model based approach for read assignment, the concept that underlies GASiC and eXpress, outperforms direct taxonomic read assignment as implemented by MEGAN and Kraken and CLARK (in the latter approach reads are aligned to the lowest rank to which they align unambigously). Second, pseudoalignment is not just faster than traditional alignment but also accurate.

The upshot: the accuracy and efficiency of kallisto make strain level analysis of metagenomes possible. In fact kallisto is more accurate at the strain level than other programs are at the genus level. Just as we have been advocating for transcript level analysis from RNA-Seq data, we believe that strain level analysis should become commonplace in metagenomics.

In digging deeply into the bioinformatics of metagenomics bioinformatics we noticed a few other areas that could benefit from RNA-Seq technology transfer. For example, the standard of RNA-Seq methods benchmarking appears to be higher than in metagenomics. Both the Kraken and CLARK papers benchmarked their programs on simulations with 10 genomes (the number ten is not a typo). CLARK did test on one dataset with 20 genomes, although using only 10,000 reads. To be fair to the authors of those papers, their standards were much higher than others in the field. The paper

Yu-Wei Wu and Yuzhen Ye, A novel abundance-based algorithm for binning metagenomic sequences using l-tuples, Journal of Computational Biology 2011.

benchmarked their method on simulations of reads from 2 (two!!) organisms. Biologists frequently complain that simulations of bioinformaticians are completely non-informative and unfortunately these cases provide fodder for such prejudice. Having said that, the RNA-Seq community also has much to learn from the metagenomics community. The previously mentioned paper by Paulson et al. 2013 addresses missing data in a way that should translate directly to missing data in single-cell RNA-Seq (the paper also makes performance comparisons with their comparative metagenomics approach to the RNA-Seq programs DESeq and edgeR) . One paper (McDavid et al. 2012) does take a look at modeling single-cell data with zero inflated distributions but I think this is a good example where metagenomics is ahead of RNA-Seq. Our immediate plans are to develop the kallisto application to metagenomics to include the ability to perform metagenome comparisons using sleuth. Conversely, inspired by the taxonomy hierarchy fundamental to metagenomics we’re going to explore RNA-Seq quantification with groups of transcripts that go beyond just genes.

Horizontal transfer is good.

[Update July 15, 2016: A preprint describing sleuth is available on BioRxiv]

Today my student Harold Pimentel released the beta version of his new RNA-Seq analysis method and software program called sleuth. A sleuth for RNA-Seq begins with the quantification of samples with kallisto, and together a sleuth of kallistos can be used to analyze RNA-Seq data rigorously and rapidly.

sleuth

Why do we need another RNA-Seq program?

A major challenge in transcriptome analysis is to determine the transcripts that have changed in their abundance across conditions.  This challenge is not entirely trivial because the stochasticity in transcription both within and between cells (biological variation), and the randomness in the experiment (RNA-Seq) that is used to determine transcript abundances (technical variation), make it difficult to determine what constitutes “significant” change. 

Technical variation can be assessed by performing technical replicates of experiments. In the case of RNA-Seq, this can be done by repeatedly sequencing from one cDNA library. Such replicates are fundamentally distinct from biological replicates designed to assess biological variation. Biological replicates are performed by sequencing different cDNA libraries that have been constructed from repeated biological experiments performed under the same (or in practice near-same) conditions. Because biological replicates require sequencing different cDNA libraries, a key point is that biological replicates include technical variation.

In the early days of RNA-Seq a few papers (e.g. Marioni et al. 2008, Bullard et al. 2010) described analyses of technical replicates and concluded that they were not really needed in practice, because technical variation could be predicted statistically from the properties of the Poisson distribution. The point is that in an idealized RNA-Seq experiment counts of reads are multinomial (according to abundances of the transcripts they originate from), and therefore approximately Poisson distributed. Their variance is therefore approximately equal to the mean, so that it is possible to predict the variance in counts across technical replicates based on the abundance of the transcripts they originate from. There is, however, one important subtlety here: “counts of reads” as used above refers to the number of reads originating from a transcript, but in many cases, especially in higher eukaryotes, reads are frequently ambiguous as to their transcript of origin because of the presence of multi-isoform genes and genes families. In other words, transcript counts cannot be precisely measured. However, the statement about the Poisson distribution of counts in technical replicates remain true when considering counts of reads by genomic features because then reads are no longer ambiguous. 

This is why, in so-called “count-based methods” for RNA-Seq analysis, there is an analysis only at the gene level. Programs such as DESeq/DESeq2, edgeR and a literally dozens of other count-based methods first require counting reads across genome features using tools such as HTSeq or featureCounts. By utilizing read counts to genomic features, technical replicates are unnecessary in lieu of the statistical assumption that they would reveal Poisson distributed data, and instead the methods focus on modeling biological variation. The issue of how to model biological variation is non-trivial because typically very few biological replicates are performed in experiments. Thus, there is a need for pooling information across genes to obtain reliable variance estimates via a statistical process called shrinkage. How and what to shrink is a matter of extensive debate among statisticians engaged in the development of count-based RNA-Seq methods, but one theme that has emerged is that shrinkage approaches can be compatible with general and generalized linear models, thus allowing for the analysis of complex experimental designs.

Despite these accomplishments,  count-based methods for RNA-Seq have two major (related) drawbacks: first, the use of counts to gene features prevents inference about the transcription of isoforms, and therefore with most count-based methods it is impossible to identify splicing switches and other isoform changes between conditions. Some methods have tried to address this issue by restricting genomic features to specific exons or splice junctions (e.g. DEXSeq) but this requires throwing out a lot of data, thereby reducing power for identifying statistically significant differences between conditions. Second, because of the fact that in general \frac{a}{b} + \frac{c}{d} \neq \frac{a+b}{c+d} it is mathematically incorrect to estimate gene abundances by adding up counts to their genomic region. One consequence of this, is that it is not possible to accurately measure fold change between conditions by using counts to gene features. In other words, count-based methods are problematic even at the gene-level and it is necessary to estimate transcript-level counts.

While reads might be ambiguous as to exactly which transcripts they originated from, it is possible to statistically infer an estimate of the number of reads from each transcript in an experiment. This kind of quantification has its origin in papers of Jiang and Wong, 2009 and Trapnell et al. 2010. However the process of estimating transcript-level counts introduces technical variation. That is to say, if multiple technical replicates were performed on a cDNA library and then transcript-level counts were to be inferred, those inferred counts would no longer be Poisson distributed. Thus, there appears to be a need for performing technical replicates after all. Furthermore, it becomes unclear how to work within the shrinkage frameworks of count-based methods. 

There have been a handful of attempts to develop methods that combine the uncertainty of count estimates at the transcript level with biological variation in the assessment of statistically significant changes in transcript abundances between conditions. For example, the Cuffdiff2 method generalizes DESeq while the bitSeq method relies on a Bayesian framework to simultaneously quantify abundances at the transcript level while modeling biological variability. Despite showing improved performance over count-based methods, they also have significant shortcomings. For example the methods are not as flexible as those of general(ized) linear models, and bitSeq is slow partly because it requires MCMC sampling.

Thus, despite intensive research on both statistical and computational methods for RNA-Seq over the past years, there has been no solution for analysis of experiments that allows biologists to take full advantage of the power and resolution of RNA-Seq.

The sleuth model

The main contribution of sleuth is an intuitive yet powerful model for RNA-Seq that bridges the gap between count-based methods and quantification algorithms in a way that fully exploits the advantages of both.

To understand sleuth, it is helpful to start with the general linear model:

Y_t = X_t\beta_t + \epsilon_t.

Here the subscript t refers to a specific transcript, Y_t is a vector describing transcript abundances (of length equal to the number of samples), X_t is a design matrix (of size number of samples x number of confounders), \beta_t is a parameter vector (of size the number of confounders) and \epsilon_t is a noise vector (of size the number of samples). In this model the abundances Y_t are normally distributed. For the purposes of RNA-Seq data, the Y_t may be assumed to be the logarithm of the counts (or normalized counts per million) from a transcript, and indeed this is the approach taken in a number of approaches to RNA-Seq modeling, e.g. in limma-voom. A common alternative to the general linear model is the generalized linear model, which postulates that some function of Y_t has a distribution with mean equal to g^{-1}(X_t \beta_t) where g is a link function, such as log, thereby allowing for distributions other than the normal to be used for the observed data. In the RNA-Seq context, where the negative binomial distribution may make sense because it is frequently a good distribution for modeling count data, the generalized model is sometimes preferred to the standard general model (e.g. by DESeq2). There is much debate about which approach is “better”.

In the sleuth model the Y_t in the general linear model are modeled as unobserved. They can be thought of us the unobserved logarithms of true counts for each transcript across samples and are assumed to be normally distributed. The observed data D_t is the logarithm of estimated counts for each transcript across samples, and is modeled as

D_t = Y_t + \zeta_t

where the \zeta_t vector parameterizes a perturbation to the unobserved Y_t. This can be understood as the technical noise due to the random sequencing of fragments from a cDNA library and the uncertainty introduced in estimating transcript counts.

The sleuth model incorporates the assumptions that the response error is additive, i.e. if  the variance of transcript in sample is V(D_{t,i}) then V(D_{t,i}) = \sigma^2_t + \tau^2_t where the variance V(\epsilon_{t,i}|y_{t,i}) = \sigma^2_t and the variance V(\zeta_{t,i}|y_{t,i}) = \tau^2_t. Intuitively, sleuth teases apart the two sources of variance by examining both technical and biological replicates, and in doing so directly estimates “true” biological variance, i.e. the variance in biological replicates that is not technical.  In lieu of actual technical replicates, sleuth takes advantage of the bootstraps of kallisto which serve as accurate proxies.

In a test of sleuth on data simulated according to the DESeq2 model we found that sleuth significantly outperforms other methods:

3_3_1_1_1

In this simulation transcript counts were simulated according to a negative binomial distribution, following closely the protocol of the DESeq2 paper simulations. Reference parameters for the simulation were first estimated by running DESeq2 on a the female Finnish population from the GEUVADIS dataset (59 individuals). In the simulation above size factors were set to be equal in accordance with typical experiments being performed, but we also tested sleuth with size factors drawn at random with geometric mean of 1 in accordance with the DESeq2 protocol (yielding factors of 1, 0.33, 3, 3, 0.33 and 1) and sleuth still outperformed other methods.

There are many details in the implementation of sleuth that are crucial to its performance, e.g. the approach to shrinkage to estimate the biological variance \sigma^2_t. A forthcoming preprint, together with Nicolas Bray and Páll Melsted that also contributed to the project along with myself, will provide the details.

Exploratory data analysis with sleuth

One of the design goals of sleuth was to create a simple and efficient workflow in line with the principles of kallisto. Working with the Shiny web application framework we have designed an html interface that allows users to interact with sleuth plots allowing for real time exploratory data analysis.

The sleuth Shiny interface is much more than just a GUI for making plots of kallisto processed data. First, it allows for the exploration of the sleuth fitted models; users can explore the technical variation of each transcript, see where statistically significant differential transcripts appear in relationship to others in terms of abundance and variance and much more. Particularly useful are interactive features in the plots. For example, when examining an MA plot, users can highlight a region of points (dynamically created box in upper panel) and see their variance breakdown of the transcripts the points correspond to, and the list of the transcripts in a table below:

 

MA_plot

The web interface contains diagnostics, summaries of the data, “maps” showing low-dimensional representations of the data and tools for analysis of differential transcripts. The interactivity via Shiny can be especially useful for diagnostics; for example, in the diagnostics users can examine scatterplots of any two samples, and then select outliers to examine their variance, including the breakdown of technical variance. This allows for a determination of whether outliers represent high variance transcripts, or specific samples gone awry. Users can of course make figures showing transcript abundances in all samples, including boxplots displaying the extent of technical variation. Interested in the differential transcribed isoform ENST00000349155 of the TBX3 gene shown in Figure 5d of the Cuffdiff2 paper? It’s trivial to examine using the transcript viewer:

TBX3

One can immediately see visually that differences between conditions completely dominate both the technical and biological variation within conditions. The sleuth q-value for this transcript is 3*10^(-68).

Among the maps, users can examine PCA projections onto any pair of components, allowing for rapid exploration of the structure of the data. Thus, with kallisto and sleuth raw RNA-Seq reads can be converted into a complete analysis in a matter of minutes. Experts will be able to generate plots and analyses in R using the sleuth library as they would with any R package. We plan numerous improvements and developments to the sleuth interface in the near future that will further facilitate data exploration; in the meantime we welcome feedback from users.

How to try out sleuth

Since sleuth requires the bootstraps and quantifications output by kallisto we recommend starting by running kallisto on your samples. The kallisto program is very fast, processing 30 million reads on a laptop in a matter of minutes. You will have to run kallisto with bootstraps- we have been using 100 bootstraps per sample but it should be possible to work with many fewer. We have yet to fully investigate the minimum number of bootstraps required for sleuth to be accurate.

To learn how to use kallisto start here. If you have already run kallisto you can proceed to the tutorial for sleuth. If you’re really eager to see sleuth without first learning kallisto, you can skip ahead and try it out using pre-computed kallisto runs of the Cuffdiff2 data- the tutorial explains where to obtain the data for trying out sleuth.

For questions, suggestions or help see the program websites and also the kallisto-sleuth user group. We hope you enjoy the tools!

The Genotype-Tissue Expression (GTEx) project is an NIH initiative to catalog human tissue-specific expression patterns in order to better understand gene regulation (see initial press release). The project is an RNA-Seq tour-de-force: RNA extracted from multiple tissues from more than 900 individuals is been quantified with more than 1,800 RNA-Seq experiments. An initial paper describing the experiments was published in Nature Genetics earlier this year and the full dataset is currently being analyzed by a large consortium of scientists.

I have been thinking recently about how to analyze genotype-tissue expression data, and have been looking forward to testing some ideas. But I have not yet become involved directly with the data, and in fact have not even submitted a request to analyze it. Given the number of samples, I’d been hoping that some basic mapping/quantification had already been done so that I could build on the work of the consortium. But, alas, this past week I got some bad news.

In a recent twitter conversation, I discovered that the program that is being used by several key GTEx consortium members to quantify the data is Flux Capacitor developed by Michael Sammeth while he was in Roderic Guigós group at the CRG in Barcelona.

What is Flux Capacitor?

Strangely, the method has never been published, despite the fact that it has been used in ten publications over the course of four years, including high profile papers from consortia such as ENCODE, GENCODE, GEUVADIS and GTEx. There is no manuscript on the author’s website or in a preprint archive. There is a website for the program but it is incomplete and unfinished, and contains no coherent explanation of what the program does. Papers using the method point to the article S. B. Montgomery, … , E. T. DermitzakisTranscriptome genetics using second generation sequencing in a Caucasian population, Nature 464 (2010) and/or the website http://sammeth.net/confluence/display/FLUX/Home for a description of the method. Here is what these citations amount to:

The Montgomery et al. paper contains one figure providing the “FluxCapacitor outline”. It is completely useless in actually providing insight into what Flux Capacitor does:

Splicing_graph

Modification of the top half of Supplementary Figure 23 from Montgomery et al (2010) titled “Flux Capacitor Outline” (although it actually shows a splice graph if one corrects the errors as I have done in red).

The methods description in the Online Methods of Montgomery et al. can only be (politely) described as word salad. Consider for example the sentence:

In our approach we estimate the biases characteristic of each experiment by collecting read distribution profiles in non-overlapping transcripts, binned by several transcript lengths and expression levels. From these profiles, we estimate for each edge and transcript a flux correction factor b^j_i that following the language of hydro-dynamic flow networks, we denote as the capacity of the edge, as the area under the transcript profile between the edge boundaries (Supplementary Fig. 23).

The indices and j for b^j_i are never defined, but more importantly its completely unclear what the the correction factor actually is, how it is estimated, and how it is used (this should be compared to the current sophistication of other methods). On the program website there is no coherent information either. Here is an example:

The resulting graph with edges labelled by the number of reads can be interpreted as a flow network where each transcript representing a transportation path from its start to its end and consequently each edge a possibly shared segment of transportation along which a certain number of reads per nucleotide — i.e., a flux — is observed.

I downloaded the code and it is undocumented- even to the extent that it is not clear what the input needs to be or what the output means. There is no example provided with the software to test the program.

I therefore became curious why GTEx chose Flux Capacitor instead of many other freely available tools for RNA-Seq (e.g. ALEXA-SeqCLIIQCufflinks, eXpress, iReckon IsoEM, IsoformExMISO, NEUMARSEM, rSEQrQuantSLIDE, TIGAR, …). Although many of these programs are not suitable for production-scale analysis, Cufflinks and RSEM certainly are, and eXpress was specifically designed for efficient quantification (linear in the number of mapped reads and constant memory). I looked around and no benchmark of Flux Capacitor has ever been performed–there is literally not even a mention of it in any paper other than in manuscripts by Sammeth, Guigó or Dermitzakis. So I thought that after four years of repeated use of the program in high profile projects, I would take a look for myself:

After fumbling about with the barely usable Flux Capacitor software, I finally managed to run it on simulated data generated for my paper: Adam Roberts and Lior Pachter, Streaming fragment assignment for real time analysis of sequencing experiments, Nature Methods 10 (2013), 71–73. One example of the state of the software is the example page (the required sorted file is posted there but its download requires the realization that is is linked to from the non-obviously placed paperclip). Fortunately, I was using my own reads and the UCSC annotation. The Roberts-Pachter simulation is explained in the Online Methods of our paper (section “Simulation RNA-Seq study”). It consists of 75bp paired-end reads simulated according to parameters mimicking real data from an ENCODE embryonic stem cell line. I tested Flux Capacitor with both 10 million and 100 million simulated reads; the results are shown in the figure below:

fc_plots

Flux Capacitor accuracy on simulations with 10 million and 100 million reads. The top panels show scatterplots of estimated transcript abundance vs. true transcript abundance. The lower panels show the same data with both axes logged.

For comparison, the next figure shows the results of RSEM, Cufflinks and eXpress on a range of simulations (up to a billion reads) from the Roberts-Pachter paper (Figure 2a):

Roberts-Pachter_Fig2a

Modification of Figure 2a from A. Roberts and L. Pachter, Nature Methods (2013) showing the performance of Flux Capacitor in context.

Flux Capacitor has very poor performance. With 100 million reads, its performance is equivalent to other software programs at 10 million reads, and similarly, with 10 million reads, it has the performance of other programs at 1 million reads. I think its fair to say that

Using Flux Capacitor is equivalent to throwing out 90% of the data!

The simulation is a best case scenario. It adheres to the standard model for RNA-Seq in which fragments are generated uniformly at random with lengths chosen from a distribution, and with errors. As explained above, all these parameters were set according to an actual ENCODE dataset, so that the difficulty of the problem corresponds to realistic RNA-Seq data. I can’t explain the poor performance of Flux Capacitor because I don’t understand the method. However my best guess is that it is somehow solving min-flow using linear programming along the lines of the properly fomulated ideas in E. Bernard, L. Jacob, J. Mairal and J.-P. VertEfficient RNA isoform identification and quantification from RNA-seq data with network flows, Technical Report HAL-00803134, March 2013. If this is the case, the poor performance might be a result of some difficulties resulting from the minimization of isoforms and reflected in the (incorrectly estimated) stripes on the left and bottom of the log-log plots. That is not to say the conclusions of the papers where Flux Capacitor is used are wrong. As can be seen from our benchmark, although performance is degraded with Flux Capacitor, the quantifications are not all wrong. For example, abundant transcripts are less likely to be affected by Flux Capacitor’s obviously poor quantification. Still, the use of Flux Capacitor greatly reduces resolution of low-expressed genes and, as mentioned previously, is effectively equivalent to throwing out 90% of the data.

As far as GTEx is concerned, I’ve been told that a significant amount of the analysis is based on raw counts obtained from reads uniquely mapping to the genome (this approach appears to have also been used in many of the other papers where Flux Capacitor was used). Adam Roberts and I examined the performance of raw counts in the eXpress paper (Figure S8, reproduced below):

Raw_reads_comparison

Figure S8 from A. Roberts and L. Pachter, Nature Methods (2013) showing the limits of quantification when ignoring ambiguous reads. NEUMA (Normalization by Expected Uniquely Mappable Areas) calculates an effective length for each transcript in order to normalize counts based on uniquely mappable areas of transcripts. We modified NEUMA to allow for errors, thereby increasing the accuracy of the method considerably, but its accuracy remains inferior to eXpress, which does consider ambiguous reads. Furthermore, NEUMA is unable to produce abundance estimates for targets without sufficient amounts of unique sequence. The EM algorithm is superior because it can take advantage of different combinations of shared sequence among multiple targets to produce estimates. The accuracy was calculated using only the subset of transcripts (77% of total) that NEUMA quantifies.

Quantification with raw counts is even worse than Flux Capacitor. It is not even possible to quantify 23% of transcripts  (due to insufficient uniquely mapping reads). This is why in the figure above the eXpress results are better than on the entire transcriptome (third figure of this post). The solid line shows that on the (raw count) quantifiable part of the transcriptome, quantification by raw counting is again equivalent to throwing out about 90% of the data. The dashed line is our own improvement of NEUMA (which required modifying the source code) to allow for errors in the reads. This leads to an improvement in performance, but results still don’t match eXpress (and RSEM and Cufflinks), and are worse than even Flux Capacitor if the unquantifiable transcripts are taken into account. In the recent Cufflinks 2 paper, we show that raw counts also cannot be used for differential analysis (as “wrong does not cancel out wrong”–  see my previous post on this).

One criticism of my simulation study could be that I am not impartial. After all, Cufflinks and eXpress were developed in my group, and the primary developer of RSEM, Bo Li, is now my postdoc. I agree with this criticism! This study should have been undertaken a long time ago and subjected to peer review by the author(s?) of Flux Capacitor and not by me. The fact that I have had to do it is a failure on their part, not mine. Moreover, it is outrageous that multiple journals and consortia have published work based on a method that is essentially a black box. This degrades the quality of the science and undermines scientists who do work hard to diligently validate, benchmark and publish their methods. Open source (the Flux Capacitor source code is, in fact, available for download) is not open science. Methods matter.

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