Bi/BE/CS183 is a computational biology class at Caltech with a mix of undergraduate and graduate students. Matt Thomson and I are co-teaching the class this quarter with help from teaching assistants Eduardo Beltrame, Dongyi (Lambda) Lu and Jialong Jiang. The class has a focus on the computational biology of single-cell RNA-seq analysis, and as such we recently taught an introduction to single-cell RNA-seq technologies. We thought the slides used would be useful to others so we have published them on figshare:

Eduardo Beltrame, Jase Gehring, Valentine Svensson, Dongyi Lu, Jialong Jiang, Matt Thomson and Lior Pachter, Introduction to single-cell RNA-seq technologies, 2019. doi.org/10.6084/m9.figshare.7704659.v1

Thanks to Eduardo Beltrame, Jase Gehring and Valentine Svensson for many extensive and helpful discussions that clarified many of the key concepts. Eduardo Beltrame and Valentine Svensson performed new analysis (slide 28) and Jase Gehring resolved the tangle of “doublet” literature (slides 17–25). The 31 slides were presented in a 1.5 hour lecture. Some accompanying notes that might be helpful to anyone interested in using them are included for reference (numbered according to slide) below:

1. The first (title) slide makes the point that single-cell RNA-seq is sufficiently complicated that a deep understanding of the details of the technologies, and methods for analysis, is required. The #methodsmatter.
2. The second slide presents an overview of attributes associated with what one might imagine would constitute an ideal single-cell RNA-seq experiment. We tried to be careful about terminology and jargon, and therefore widely used terms are italicized and boldfaced.
3. This slide presents Figure 1 from Svensson et al. 2018. This is an excellent perspective that highlights key technological developments that have spurred the growth in popularity of single-cell RNA-seq. At this time (February 2019) the largest single-cell RNA-seq dataset that has been published consists of 690,000 Drop-seq adult mouse brain cells (Saunders, Macosko et al. 2018). Notably, the size and complexity of this dataset rivals that of a large-scale genome project that until recently would be undertaken by hundreds of researchers. The rapid adoption of single-cell RNA-seq is evident in the growth of records in public sequence databases.
4. The Chen et al. 2018 review on single-cell RNA-seq is an exceptionally useful and thorough review that is essential reading in the field. The slide shows Figure 2 which is rich in information and summarizes some of the technical aspects of single-cell RNA-seq technologies. Understanding of the details of individual protocols is essential to evaluating and assessing the strengths and weaknesses of different technologies for specific applications.
5. Current single-cell RNA-seq technologies can be broadly classified into two groups: well-based and droplet-based technologies. The Papalexi and Satija 2017 review provides a useful high-level overview and this slide shows a part of Figure 1 from the review.
6. The details of the SMART-Seq2 protocol are crucial for understanding the technology. SMART is a clever acronym for Switching Mechanism At the 5′ end of the RNA Transcript. It allows the addition of an arbitrary primer sequence at the 5′ end of a cDNA strand, and thus makes full length cDNA PCR possible. It relies on the peculiar properties of the reverse transcriptase from the Moloney murine leukemia virus (MMLV), which, upon reaching the 5’ end of the template, will add a few extra nucleotides (usually Cytosines). The resultant overhang is a binding site for the “template switch oligo”, which contains three riboguanines (rGrGrG). Upon annealing, the reverse transcriptase “switches” templates, and continues transcribing the DNA oligo, thus adding a constant sequence to the 5’ end of the cDNA. After a few cycles of PCR, the full length cDNA generated is too long for Illumina sequencing (where a maximum length of 800bp is desirable). To chop it up into smaller fragments of appropriate size while simultaneously adding the necessary Illumina adapter sequences, one can can use the Illumina tagmentation Nextera™ kits based on Tn5 tagmentation. The SMART template switching idea is also used in the Drop-seq and 10x genomics technologies.
7. While it is difficult to rate technologies exactly on specific metrics, it is possible to identify strengths and weaknesses of distinct approaches. The SMART-Seq2 technology has a major advantage in that it produces reads from across transcripts, thereby providing “full-length” information that can be used to quantify individual isoforms of genes. However this superior isoform resolution requires more sequencing, and as a result makes the method less cost effective. Well-based methods, in general, are not as scalable as droplet methods in terms of numbers of cells assayed. Nevertheless, the tradeoffs are complex. For example robotics technologies can be used to parallelize well-based technologies, thereby increasing throughput.
8. The cost of a single-cell technology is difficult to quantify. Costs depend on number of cells assayed as well as number of reads sequenced, and different technologies have differing needs in terms of reagents and library preparation costs. Ziegenhain et al. 2017 provide an in-depth discussion of how to assess cost in terms of accuracy and power, and the table shown in the slide is reproduced from part of Table 1 in the paper.
9. A major determinant of single-cell RNA-seq cost is sequencing cost. This slide shows sequencing costs at the UC Davis Genome Center and its purpose is to illustrate numerous tradeoffs, relating to throughput, cost per base, and cost per fragment that must be considered when selecting which sequencing machine to use. In addition, sequencing time frequently depends on core facilities or 3rd party providers multiplexing samples on machines, and some sequencing choices are likely to face more delay than others.
10. Turning to droplet technologies based on microfluidics, two key papers are the Drop-seq and inDrops papers which were published in the same issue of a single journal in 2015. The papers went to great lengths to document the respective technologies, and to provide numerous text and video tutorials to facilitate adoption by biologists. Arguably, this emphasis on usability (and not just reproducibility) played a major role in the rapid adoption of single-cell RNA-seq by many labs over the past three years. Two other references on the slide point to pioneering microfluidics work by Rustem Ismagilov, David Weitz and their collaborators that made possible the numerous microfluidic single-cell applications that have since been developed.
11.  This slide displays a figure showing a monodispersed emulsion from the recently posted preprint “Design principles for open source bioinstrumentation: the poseidon syringe pump system as an example” by Booeshaghi et al., 2019. The generation of such emulsions is a prerequisite for successful droplet-based single-cell RNA-seq. In droplet based single-cell RNA-seq, emulsions act as “parallelizing agents”, essentially making use of droplets to parallelize the biochemical reactions needed to capture transcriptomic (or other) information from single-cells.
12. The three objects that are central to droplet based single-cell RNA-seq are beads, cells and droplets. The relevance of emulsions in connection to these objects is that the basis of droplet methods for single-cell RNA-seq is the encapsulation of single cells together with single beads in the emulsion droplest. The beads are “barcode” delivery vectors. Barcodes are DNA sequences that are associated to transcripts, which are accessible after cell lysis in droplets. Therefore, beads must be manufactured in a way that ensures that each bead is coated with the same barcodes, but that the barcodes associated with two distinct beads are different from each other.
13. The inDrops approach to single-cell RNA-seq serves as a useful model for droplet based single-cell RNA-seq methods. The figure in the slide is from a protocol paper by Zilionis et al. 2017 and provides a useful overview of inDrops. In panel (a) one sees a zoom-in of droplets being generated in a microfluidic device, with channels delivering cells and beads highlighted. Panel (b) illustrates how inDrops hydrogel beads are used once inside droplets: barcodes (DNA oligos together with appropriate priming sequences) are released from the hydrogel beads and allow for cell barcoded cDNA synthesis. Finally, panel (c) shows the sequence construct of oligos on the beads.
14. This slide is analogous to slide 6, and shows an overview of the protocols that need to be followed both to make the hydrogel beads used for inDrops, and the inDrops protocol itself.  In a clever dual use of microfluidics, inDrops makes the hydrogel beads in an emulsion. Of note in the inDrops protocol itself is the fact that it is what is termed a “3′ protocol”. This means that the library, in addition to containing barcode and other auxiliary sequence, contains sequence only from 3′ ends of transcripts (seen in grey in the figure). This is the case also with other droplet based single-cell RNA-seq technologies such as Drop-seq or 10X Genomics technology.
15. The significance of 3′ protocols it is difficult to quantify individual isoforms of genes from the data they produce. This is because many transcripts, while differing in internal exon structure, will share a 3′ UTR. Nevertheless, in exploratory work aimed at investigating the information content delivered by 3′ protocols, Ntranos et al. 2019 show that there is a much greater diversity of 3′ UTRs in the genome than is currently annotated, and this can be taken advantage of to (sometimes) measure isoform dynamics with 3′ protocols.
16. To analyze the various performance metrics of a technology such as inDrops it is necessary to understand some of the underlying statistics and combinatorics of beads, cells and drops. Two simple modeling assumptions that can be made is that the number of cells and beads in droplets are each Poisson distributed (albeit with different rate parameters). Specifically, we assume that
$\mathbb{P}(\mbox{droplet has } k \mbox{ cells}) = \frac{e^{-\lambda}\lambda^k}{k!}$ and $\mathbb{P}(\mbox{droplet has } k \mbox{ beads}) = \frac{e^{-\mu}\mu^j}{j!}$. These assumptions are reasonable for the Drop-seq technology. Adjustment of concentrations and flow rates of beads and cells and oil allows for controlling the rate parameters of these distributions and as a result allow for controlling numerous tradeoffs which are discussed next.
17. The cell capture rate of a technology is the fraction of input cells that are assayed in an experiment. Droplets that contain one or more cells but no beads will result in a lost cells whose transcriptome is not measured. The probability that a droplet has no beads is $e^{-\mu}$ and therefore the probability that a droplet has at least one bead is $1-e^{-\mu}$. To raise the capture rate it is therefore desirable to increase the Poisson rate $\mu$ which is equal to the average number of beads in a droplet. However increasing $\mu$ leads to duplication, i.e. cases where a single droplet has more than one bead, thus leading .a single cell transcriptome to appear as two or more cells. The duplication rate is the fraction of assayed cells which were captured with more than one bead. The duplication rate can be calculated as $\frac{\mathbb{P}(\mbox{droplet has 2 or more beads})}{\mathbb{P}(\mbox{droplet has 1 or more beads})}$ (which happens to be equivalent to a calculation of the probability that we are alone in the universe). The tradeoff, shown quantitatively as capture rate vs. duplication rate, is shown in a figure I made for the slide.
19. This slide shows a comparison of bead technologies. In addition to inDrops, 10x Genomics single-cell RNA-seq technology also uses hydrogel beads. There are numerous technical details associated with beads including whether or not barcodes are released (and how).
20. Barcode collisions arise when two cells are separately encapsulated with beads that contain identical barcodes. The slide shows the barcode collision rate formula, which is $1-\left( 1-\frac{1}{M} \right)^{N-1}$. This formula is derived as follows: Let $p=\frac{1}{M}$. The probability that a barcodes is associated with k cells is given by the binomial formula ${N \choose k}p^k(1-p)^{N-k}$. Thus, the probability that a barcode is associated to exactly one cell is $Np(1-p)^{N-1} = \frac{N}{M}\left(1-\frac{1}{M}\right)^{N-1}$. Therefore the expected number of cells with a unique barcode is $N\left(1-\frac{1}{M}\right)^{N-1}$ and the barcode collision rate is $\left(1-\frac{1}{M}\right)^{N-1}$. This is approximately $1-\left( \frac{1}{e} \right)^{\frac{N}{M}}$. The term synthetic doublet is used to refer to the situation when two or more different cells appear to be a single cell due to barcode collision.
21. In the notation of the previous slide, the barcode diversity is $\frac{N}{M}$, which is an important number in that it determines the barcode collision rate. Since barcodes are encoded in sequence, a natural question is what sequence length is needed to ensure a desired level of barcode diversity. This slide provides a lower bound on the sequence length.
22. Technical doublets are multiple cells that are captured in a single droplet, barcoded with the same sequence, and thus the transcripts that are recorded from them appear to originate from a single-cell.  The technical doublet rate can be estimated using a calculation analogous to the one used for the cell duplication rate (slide 17), except that it is a function of the Poisson rate $\lambda$ and not $\mu$. In the single-cell RNA-seq literature the term “doublet” generally refers to technical doublets, although it is useful to distinguish such doublets from synthetic doublets and biological doublets (slide 25).
23. One way to estimate the technical doublet rate is via pooled experiments of cells from different species. The resulting data can be viewed in what has become known as a “Barnyard” plot, a term originating from the Macosko et al. 2015 Drop-seq paper. Despite the authors’ original intention to pool mouse, chicken, cow and pig cells, typical validation of single-cell technology now proceeds with a mixture of human and mouse cells. Doublets are readily identifiable in barnyard plots as points (corresponding to droplets) that display transcript sequence from two different species. The only way for this to happen is via the capture of a doublet of cells (one from each species) in a single droplet. Thus, validation of single-cell RNA-seq rigs via a pooled experiment, and assessment of the resultant barnyard plot, has become standard for ensuring that the data is indeed single cell.
24.  It is important to note that while points on the diagonal of barnyard plots do correspond to technical doublets, pooled experiments, say of human and mouse cells, will also result in human-human and mouse-mouse doublets that are not evident in barnyard plots. To estimate such within species technical doublets from a “barnyard analysis”, Bloom 2018 developed a simple procedure that is described in this slide. The “Bloom correction” is important to perform in order to estimate doublet rate from mixed species experiments.
25. Biological doublets arise when two cells form a discrete unit that does not break apart during disruption to form a suspension. Such doublets are by definition from the same species, and therefore will not be detected in barnyard plots. One way to avoid biological doublets is via single nucleus RNA-seq (see, Habib et al. 2017). Single nuclear RNA-seq has proved to be important for experiments involving cells that are difficult to dissociate, e.g. human brain cells. In fact, the formation of suspensions is a major bottleneck in the adoption of single-cell RNA-seq technology, as techniques vary by tissue and organism, and there is no general strategy. On the other hand, in a recent interesting paper, Halpern et al. 2018 show that biological doublets can sometimes be considered a feature rather than a bug. In any case, doublets are more complicated than initially appears, and we have by now seen that there are three types of doublets: synthetic doublets, technical doublets, and biological doublets .
26. Unique molecular identifiers (UMIs) are part of the oligo constructs for beads in droplet single-cell RNA-seq technologies. They are used to identify reads that originated from the same molecule, so that double counting of such molecules can be avoided. UMIs are generally much shorter than cell barcodes, and estimates of required numbers (and corresponding sequence lengths) proceed analogously to the calculations for cell barcodes.
27. The sensitivity of a single-cell RNA-seq technology (also called transcript capture rate) is the fraction of transcripts captured per cell. Crucially, the sensitivity of a technology is dependent on the amount sequenced, and the plot in this slide (made by Eduardo Beltrame and Valentine Svensson by analysis of data from Svensson et al. 2017) shows that dependency.  The dots in the figure are cells from different datasets that included spike-ins, whose capture is being measured (y-axis). Every technology displays an approximately linear relationship between number of reads sequenced and transcripts captured, however each line is describe by two parameters: a slope and an intercept. In other words, specification of sensitivity for a technology requires reporting two numbers, not one.
28. Having surveyed the different attributes of droplet technologies, this slide summarizes some of the pros and cons of inDrops similarly to slide 7 for SMART-Seq2.
29. While individual aspects of single-cell RNA-seq technologies can be readily understood via careful modeling coupled to straightforward quality control experiments, a comprehensive assessment of whether a technology is “good” or “bad” is meaningless. The figure in this slide, from Zhang et al. 2019, provides a useful overview of some of the important technical attributes.
30. In terms of the practical question “which technology should I choose for my experiment?”, the best that can be done is to offer very general decision workflows. The flowchart shown on this slide, also from Zhang et al. 2019, is not very specific but does provide some simple to understand checkpoints to think about. A major challenge in comparing and contrasting technologies for single-cell RNA-seq is that they are all changing very rapidly as numerous ideas and improvements are now published weekly.
31. The technologies reviewed in the slides are strictly transcriptomics methods (single-cell RNA-seq). During the past few years there has been a proliferation of novel multi-modal technologies that simultaneously measures the transcriptome of single cells along with other modalities. Such technologies are reviewed in Packer and Trapnell 2018 and the slide mentions a few of them.

Five years ago on this day, Nicolas Bray and I wrote a blog post on The network nonsense of Manolis Kellis in which we described the paper Feizi et al. 2013 from the Kellis lab as dishonest and fraudulent. Specifically, we explained that:

“Feizi et al. have written a paper that appears to be about inference of edges in networks based on a theoretically justifiable model but

1. the method used to obtain the results in the paper is completely different than the idealized version sold in the main text of the paper and
2. the method actually used has parameters that need to be set, yet no approach to setting them is provided. Even worse,
3. the authors appear to have deliberately tried to hide the existence of the parameters. It looks like
4. the reason for covering up the existence of parameters is that the parameters were tuned to obtain the results. Moreover,
5. the results are not reproducible. The provided data and software is not enough to replicate even a single figure in the paper. This is disturbing because
6. the performance of the method on the simplest of all examples, a correlation matrix arising from a Gaussian graphical model, is poor.”

A second point we made is that the justification for the method, which the authors called “network deconvolution” was nonsense. For example, the authors wrote that “The model assumes that networks are “linear time-invariant flow-preserving operators.” Perhaps I take things too literally when I read papers but I have to admit that five years later I still don’t understand the sentence. However just because a method is ad-hoc, heuristic, or perhaps poorly explained, doesn’t mean it won’t work well in practice. In the blog post we compared network deconvolution to regularized partial correlation on simulated data, and found network deconvolution performed poorly. But in a responding comment, Kellis noted that “in our experience, partial correlation performed very poorly in practice.” He added that “We have heard very positive feedback from many other scientists using our software successfully in diverse applications.”

Fortunately we can now evaluate Kellis’ claims in light of an independent analysis in Wang, Pourshafeie, Zitnik et al. 2018, a paper from the groups of Serafim Batzoglou and Jure Leskovec (in collaboration with Carlos Bustamante) at Stanford University. There are three main results presented in Wang, Pourshafeie and Zitnik et al. 2018 that summarize the benchmarking of network deconvolution and other methods, and I reproduce figures showing the results below. The first shows the performance of network deconvolution and some other network denoising methods on a problem of butterfly species identification (network deconvolution is abbreviated ND and is shown in green). RAW (in blue) is the original unprocessed network. Network deconvolution is much worse than RAW:

The second illustrates the performance of network denoising methods on Hi-C data. The performance metric in this case is normalized mutual information (NMI) which Wang, Pourshafeie, Zitnik et al. described as “a fair representation of overall performance”. Network deconvolution (ND, dark green) is again worse than RAW (dark blue):

Finally, in an analysis of gene function from tissue-specific gene interaction networks, ND (blue) does perform better than RAW (pink) although barely. In four cases out of eight shown it is the worst of the four methods benchmarked:

Network deconvolution was claimed to be applicable to any network when it was published. At the time, Feizi stated that “We applied it to gene networks, protein folding, and co-authorship social networks, but our method is general and applicable to many other network science problems.” A promising claim, but in reality it is difficult to beat the nonsense law: Nonsense methods tend to produce nonsense results.

The Feizi et al. 2013 paper now has 178 citations, most of them drive by citations. Interestingly this number, 178 is exactly the number of citations of the Barzel et al. 2013 network nonsense paper, which was published in the same issue of Nature Biotechnology. Presumably this reflects the fact that authors citing one paper feel obliged to cite the other. These pair of papers were thus an impact factor win for the journal. For the first authors on the papers, the network deconvolution/silencing work is their most highly cited first author papers respectively. Barzel is an assistant professor at Bar-Ilan University where he links to an article about his network nonsense on his “media page”. Feizi is an assistant professor at the University of Maryland where he lists Feizi et al. 2013 among his “selected publications“. Kellis teaches the “network deconvolution” and its associated nonsense in his computational biology course at MIT. And why not? These days truth seems to matter less and less in every domain. A statement doesn’t have to be true, it just has to work well on YouTube, Twitter, Facebook, or some webpage, and as long as some people believe it long enough, say until the next grant cycle, promotion evaluation, or election, then what harm is done? A win-win for everyone. Except science.

The encapsulation of beads together with cells in droplets is the basis of microfluidic based single-cell RNA-seq technologies. Ideally droplets contain exactly one bead and one cell, however in practice the number of beads and cells in droplets is stochastic and encapsulation of cells in droplets produces an approximately Poisson distribution of number of cells per droplet:

Specifically, the probability of observing k cells in a droplet is approximated by

$\mathbb{P}(\mbox{k cells in a droplet}) = \frac{e^{-\lambda}\lambda^k}{k!}$.

The rate parameter $\lambda$ can be controlled and the average number of cells per droplet is equal to it. Therefore, setting $\lambda$ to be much less than 1 ensures that two or more cells are rarely encapsulated in a single droplet. A consequence of this is that the number of empty droplets, given by $e^{-\lambda}$, is large. Importantly, one of the properties of the Poisson distribution is that variance is equal to the mean so the number of cells per droplet is also equal to $\lambda$.

Along with cells, beads must also be captured in droplets, and when plastic beads are used the occupancy statistics follow a Poisson distribution as well. This means that with technologies such as Drop-seq (Macosko et al. 2015), which uses polystyrene beads, many droplets are either empty, contain a bead and no cell, or a cell and no bead. The latter situation (cell and no bead) leads to a low “capture rate”, i.e. not many of the cells are assayed in an experiment.

One of the advantages of the inDrops method (Klein et al. 2015) over other single-cell RNA-seq methods is that it uses hydrogel beads which allow for a reduction in the variance of the number of beads per cell. In an important paper Abate et al. 2009 showed that close packing of hydrogel beads allows for an almost degenerate distribution where the number of beads per droplet is exactly one 98% of the time. The video below shows how close to degeneracy the distribution can be squeezed (in the example two beads are being encapsulated per droplet):

A discrete distribution defined over the non-negative integers with variance less than the mean is called sub-Poisson. Similarly, a discrete distribution defined over the non-negative integers with variance greater than the mean is called super-Poisson. This terminology dates back to at least the 1940s (e.g., Berkson et al. 1942) and is standard in many fields from physics (e.g. Rodionov and Cherkin 2004) to biology (e.g. Pitchiaya et al. 2014 ).

Figure 5.26 from Adrian Jeantet, Cavity quantum electrodynamics with carbon nanotubes, 2017.

Using this terminology, the close packing of hydrogel beads can be said to enable sub-Poisson loading of beads into droplets because the variance of beads per droplet is reduced in comparison to the Poisson statistics of plastic beads.

Unfortunately, in a 2015 paper, Bose et al. used the term “super-Poisson” instead of “sub-Poisson” in discussing an approach to reducing bead occupancy variance in the single-cell RNA-seq context. This sign error in terminology has subsequently been propagated and recently appeared in a single- cell RNA-seq review (Zhang et al. 2018) and in 10x Genomics advertising materials.

When it comes to single-cell RNA-seq we already have people referring to the number of reads sequenced as “the library size” and calling trees “one-dimensional manifolds“. Now sub-Poisson is mistaken for super-Poisson. Before you know it we’ll have professors teaching students that cell clusters obtained by k-means clustering are “cell types“…

Supper poisson (not to be confused with super-Poisson (not to be confused with sub-Poisson)).

The title of this blog post is a phrase coined by Paul Wouters and Rodrigo Costas in their 2012 publication Users, Narcissism and Control—Tracking the Impact of Scholarly Publications in the 21st Century. By “technologies of narcissism”, Wouters and Costas mean tools that allow individuals to rapidly assess the impact, usage and influence of their publications without much effort.  One of the main points that Wouters and Costas try to convey is that individuals using technologies of narcissism must exercise “great care and caution” due to the individual level focus of altmetrics.

I first recall noticing altmetrics associated to one of my papers after publishing the paper “Bioinformatics for Whole-Genome Shotgun Sequencing of Microbial Communities” in PLoS Computational Biology in 2005. Public Library of Science (PLoS) was one of the first publishers to collect and display views and downloads of papers, and I remember being intrigued the first time I noticed my paper statistics. Eventually I developed the habit of revisiting the paper website frequently to check “how we were doing”. I’m probably responsible for at least a few dozen of the downloads that led to the paper making the “top ten” list the following year. PLoS Computational Biology even published a paper where they displayed paper rankings (by downloads). Looking back, while PLoS was a pioneer in developing technologies of narcissism, it was the appetite for them from individuals such as myself that drove a proliferation of new metrics and companies devoted to disseminating them. For example, a few years later in 2012, right when Wouters and Costas were writing about technologies of narcissism, Altmetric.com was founded and today is a business with millions of dollars in revenue, dozens of employees, and a name that is synonymous with the metrics they measure.

Today Altmetric.com’s “Attention Score”  is prominently displayed alongside articles in many journals (for example all of the publications of the Microbial Society) and even bioRxiv displays them. In fact, the importance of altmetrics to bioRxiv is evident as they are mentioned in the very first entry of the bioRxiv FAQ, which states that “bioRxiv provides usage metrics for article views and PDF downloads, as well as altmetrics relating to social media coverage. These metrics will be inaccurate and underestimate actual usage in article-to-article comparisons if an article is also posted elsewhere.” Altmetric.com has worked hard to make it easy for anyone to embed the “Altmetric Attention Score” on their website, and in fact some professors now do so:

What does Altmetric.com measure? The details may surprise you. For example, Altmetric tracks article mentions in Wikipedia, but only in the English, Finnish and Swedish Wikipedias. Tweets, retweets and quoted tweets of articles are tracked, but not all of them. This is because Altmetric.com was cognizant of the possibility of “gaming the system” from the outset and it therefore looks for “evidence of gaming” to try to defend against manipulation of its scores. The “Gaming altmetrics” blogpost by founder Euan Adie in 2013 is an interesting read. Clearly there have been numerous attempts to manipulate the metrics they measure. He writes “We flag up papers this way and then rely on manual curation (nothing beats eyeballing the data) to work out exactly what, if anything, is going on.”

Is anything going on? It’s hard to say. But here are some recent comments on social media:

Perhaps this exchange is tongue-in-cheek but notice that by linking to a bioRxiv preprint the second tweet in the series above actually affected the Altmetric Attention Score of that preprint:

Here is another exchange:

Apparently they are not the first to have this idea:

The last tweet is by a co-founder of bioRxiv. These recent “jokes” (I suppose?) about altmetrics are in response to a recent preprint by Abdill and Blekhman (note that I’ve just upped the Altmetric Attention Score of the preprint by linking to it from this blog; Altmetric.com tracks links from manually curated lists of blogs). The Abdill-Blekhman preprint included an analysis showing a strong correlation between paper downloads and the impact factor of journals where they are published:

The analogous plot showing the correlation between tweets and citations per preprint (averaged by journal where they ended up being published) was made by Sina Booeshaghi and Lynn Yi last year (GitHub repository here):

There are some caveats to the Booeshaghi-Yi analysis (the number of tweets per preprint is capped at 100) but it shows a similar trend to the Abdill-Blekhman analysis. One question which these data raise (and the convenient API by Abdill and Blekhman makes possible to study) is what is the nature of this correlation? Are truly impactful results in preprints being recognized as such (independently) by both journal reviewers/editors and scientists via twitter, or are the altmetrics of preprints directly affecting where they end up being published? The latter possibility is disturbing if true. Twitter activity is highly biased and associated with many factors that have nothing to do with scientific interest or relevance. For example, women are less influential on twitter than men (men are twice as likely to be retweeted as women). Thus, the question of causation in this case is not just of academic interest, it is also of career importance for individuals, and important for science as a whole. The data of Abdill and Blekhman will be useful in studying the question, and are an important starting point to assimilate and build on. I am apparently not the only person who thinks so; Abdill and Blekhman’s preprint is already “highly downloaded”, as can be seen on a companion website to the preprint called Rxivist.

The Rxivist website is useful for browsing the bioRxiv, but it does something else as well. For the first time, it makes accessible two altmetric statistics (paper downloads and tweets) via “author leaderboards”. Unlike Altmetric.com, which appears to carefully (and sometimes) manually curates the statistics going into its final score, and which acts against manipulation of tweets by filtering for bots, the Rxivist leaderboards are based on raw data. This is what a leaderboard of “papers downloaded” looks like:

The fact is that Stephen Floor is right; it is already accepted that the number of times a preprint has been downloaded is relevant and important:

But this raises a question, are concerns about gaming the system overblown? A real problem? How hard is it, really, to write a bot to boost one’s download statistics? Has someone done it already?

Here is a partial answer to the questions above in the form of a short script that downloads any preprint (also available on the blog GitHub repository where the required companion chromedriver binary is also available):

The scientific community would be remiss to ignore the proliferation of technologies of narcissism. These technologies can have real benefit, primarily by providing novel ways to help researchers identify interesting and important work that is relevant to the questions they are pursuing. Furthermore, one of the main advantages of the open licensing of resources such as bioRxiv or PLoS is that they permit natural language processing to facilitate automatic prioritization of articles, search for relevant literature, and mining for specific scientific terms (see e.g., Huang et al. 2016). But I am loathe to accept a scientific enterprise that rewards winners of superficial, easily gamed, popularity contests.

The long-standing practice of data sharing in genomics can be traced to the Bermuda principles, which were formulated during the human genome project (Contreras, 2010). While the Bermuda principles focused on open sharing of DNA sequence data, they heralded the adoption of other open source standards in the genomics community. For example, unlike many other scientific disciplines, most genomics software is open source and this has been the case for a long time (Stajich and Lapp, 2006). The open principles of genomics have arguably greatly accelerated progress and facilitated discovery.

While open sourcing has become de rigueur in genomics dry labs, wet labs remain beholden to commercial instrument providers that rarely open source hardware or software, and impose draconian restrictions on instrument use and modification. With a view towards joining others who are working to change this state of affairs, we’ve posted a new preprint in which we describe an open source syringe pump and microscope system called poseidon:

A. Sina Booeshaghi, Eduardo da Veiga Beltrame, Dylan Bannon, Jase Gehring and Lior Pachter,

The poseidon system consists of

• A syringe pump that can operate at a wide range of flow rates. The bulk cost per pump is $37.91. A system of three pumps that can be used for droplet based single-cell RNA-seq experiments can be assembled for$174.87
• A microscope system that can be used to evaluate the quality of emulsions produced using the syringe pumps. The cost is $211.69. • Open source software that can be used to operate four pumps simultaneously, either via a Raspberry Pi (that is part of the microscope system) or directly via a laptop/desktop. Together, these components can be used to build a Drop-seq rig for under$400, or they can be used piecemeal for a wide variety of tasks. Along with describing benchmarks of poseidon, the preprint presents design guidelines that we hope can accelerate both development and adoption of open source bioinstruments. These were developed while working on the project; some were borrowed from our experience with bioinformatics software, while others emerged as we worked out kinks during development. As is the case with software, open source is not,  in and of itself, enough to make an application usable.  We had to optimize many steps during the development of poseidon, and in the preprint we illustrate the design principles we converged on with specific examples from poseidon.

The complete hardware/software package consists of the following components:

We benchmarked the system thoroughly and it has similar performance to a commercial Harvard Apparatus syringe pump; see panel (a) below. The software driving the pumps can be used for infusion or withdrawl, and is easily customizable. In fact, the ability to easily program arbitrary schedules and flow rates without depending on vendors who frequently charge money and require firmware upgrades for basic tasks, was a major motivation for undertaking the project. The microscope is basic but usable for setting up emulsions. Shown in panel (b) below is a microfluidic droplet generation chip imaged with the microscope. Panel (c) shows that we have no trouble generating uniform emulsions with it.

Together, the system constitutes a Drop-seq rig (3 pumps + microscope) that can be built for under 400: We did not start the poseidon project from scratch. First of all, we were fortunate to have some experience with 3D printing. Shortly after I started setting up a wet lab, Shannon Hateley, a former student in the lab, encouraged me to buy a 3D printer to reduce costs for basic lab supplies. The original MakerGear M2 we purchased has served us well saving us enormous amounts of money just as Shannon predicted, and in fact we now have added a Prusa printer: The printer Shannon introduced to the lab came in handy when, some time later, after starting to perform Drop-seq in the lab, Jase Gehring became frustrated with the rigidity commercial syringe pumps he was using. With a 3D printer available in-house, he was able to print and assemble a published open source syringe pump design. What started as a hobby project became more serious when two students joined the lab: Sina Booeshaghi, a mechanical engineer, and Eduardo Beltrame, an expert in 3D printing. We were also encouraged by the publication of a complete Drop-seq do-it-yourself design from the Satija lab. Starting with the microscope device from the Stephenson et al. paper, and the syringe pump from the Wijnen et al. paper, we worked our way through numerous hardware design optimizations and software prototypes. The photo below shows the published work we started with at the bottom, the final designs we ended up with at the top, and intermediate versions as we converged on design choices: In the course of design we realized that despite a lot of experience developing open source software in the lab, there were new lessons we were learning regarding open-source hardware development, and hardware-software integration. We ended up formulating six design principles that we explain in detail in the preprint via example of how they pertained to the poseidon design: We are hopeful that these principles we adhered to will serve as useful guidelines for others interested in undertaking open source bioinstrumentation projects. This post is a review of a recent preprint on an approach to testing for RNA-seq gene differential expression directly from transcript compatibility counts: Marek Cmero, Nadia M Davidson and Alicia Oshlack, Fast and accurate differential transcript usage by testing equivalence class counts, bioRxiv 2018. To understand the preprint two definitions are important. The first is of gene differential expression, which I wrote about in a previous blog post and is best understood, I think, with the following figure (reproduced from Supplementary Figure 1 of Ntranos, Yi, et al., 2018): In this figure, two isoforms of a hypothetical gene are called primary and secondary, and two conditions in a hypothetical experiment are called “A” and “B”. The black dots labeled conditions A and B have x-coordinates $x_A$ and $x_B$ corresponding to the abundances of the primary isoform in the respective conditions, and y-coordinates $y_A$ and $y_B$ corresponding to the abundance of the secondary isoforms. In data from the hypothetical experiment, the black dots represent the mean level of expression of the constituent isoforms as derived from replicates. Differential transcript expression (DTE) refers to change in one of the isoforms. Differential gene expression (DGE) refers to change in overall gene expression (i.e. expression as the sum of the expression of the two isoforms). Differential transcript usage (DTU) refers to change in relative expression between the two isoform and gene differential expression (GDE) refers to change in expression along the red line. Note that DGE, DTU and DGE are special cases of GDE. The Cmero et al. preprint describes a method for testing for GDE, and the method is based on comparison of equivalence classes of reads between conditions. There is a natural equivalence relation $\sim$ on the set of reads in an RNA-seq experiment, where two reads $r_1$ and $r_2$ are related by $\sim$ when $r_1$ and $r_2$ align (ambiguously) to exactly the same set of transcripts (see, e.g. Nicolae et al. 2011). The equivalence relation $\sim$ partitions the reads into equivalence classes, and, in a slight abuse of notation, the term “equivalence class” in RNA-seq is used to denote the set of transcripts corresponding to an equivalence class of reads. Starting with the pseudoalignment program kallisto published in Bray et al. 2016, it became possible to rapidly obtain the (transcript) equivalence classes for reads from an RNA-seq experiment. In previous work (Ntranos et al. 2016) we introduced the term transcript compatibility counts to denote the cardinality of the (read) equivalence classes. We thought about this name carefully; due to the abuse of notation inherent in the term “equivalence class” in RNA-seq, we felt that using “equivalence class counts” would be confusing as it would be unclear whether it refers to the cardinalities of the (read) equivalence classes or the (transcript) “equivalence classes”. With these definitions at hand, the Cmero et al.’s preprint can be understood to describe a method for identifying GDE between conditions by directly comparing transcript compatibility counts. The Cmero et al. method is to perform Šidák aggregation of p-values for equivalence classes, where the p-values are computed by comparing transcript compatibility counts for each equivalence class with the program DEXSeq (Anders et al. 2012). A different method that also identifies GDE by directly comparing transcript compatibility counts was previously published by my student Lynn Yi in Yi et al. 2018. I was curious to see how the Yi et al. method, which is based on Lancaster aggregation of p-values computed from transcript compatibility counts compares to the Cmero et al. method. Fortunately it was really easy to find out because Cmero et al. released code with their paper that can be used to make all of their figures. I would like to note how much fun it is to reproduce someone else’s work. It is extremely empowering to know that all the methods of a paper are pliable at the press of a button. Below is the first results figure, Figure 2, from Cmero et al.’s paper: Below is the same figure reproduced independently using their code (and downloading the relevant data): It’s beautiful to see not only apples-to-apples, but the exact same apple! Reproducibility is obviously important to facilitate transparency in papers and to ensure correctness, but its real value lies in the fact that it allows for modifying and experimenting with methods in a paper. Below is the second results figure, Figure 3, from Cmero et al.’s paper: The figure below is the reproduction, but with an added analysis in Figure 3a, namely the method of Yi et al. 2018 included (shown in orange as “Lancaster_equivalence_class”). The additional code required for the extra analysis is just a few lines and can be downloaded from the Bits of DNA Github repository: library(aggregation) library(dplyr) dm_dexseq_results <- as.data.frame(DEXSeqResults(dm_ec_resultsdexseq_object))
dm_lancaster_results <- dm_dexseq_results %>% group_by(groupID) %>% summarize(pval = lancaster(pvalue, log(exonBaseMean)))
dm_lancaster_results$gene_FDR <- p.adjust(dm_lancaster_results$pval, ‘BH’)
dm_lancaster_results <- data.frame(gene = dm_lancaster_results$groupID, FDR = dm_lancaster_results$gene_FDR)

hs_dexseq_results <- as.data.frame(DEXSeqResults(hs_ec_results$dexseq_object)) hs_lancaster_results <- hs_dexseq_results %>% group_by(groupID) %>% summarize(pval = lancaster(pvalue, log(exonBaseMean))) hs_lancaster_results$gene_FDR <- p.adjust(hs_lancaster_results$pval, ‘BH’) hs_lancaster_results <- data.frame(gene = hs_lancaster_results$groupID,
FDR = hs_lancaster_results$gene_FDR) A zoom-in of Figure 3a below shows that the improvement of Yi et al.’s method in the hsapiens dataset over the method of Cmero et al. is as large as the improvement of aggregation (of any sort) over GDE based on transcript quantifications. Importantly, this is a true apples-to-apples comparison because Yi et al.’s method is being tested on exactly the data and with exactly the metrics that Cmero et al. chose: The improvement is not surprising; an extensive comparison of Lancaster aggregation with Šidák aggregation is detailed in Yi et al. and there we noted that while Šidák aggregation performs well when transcripts are perturbed independently, it performs very poorly in the more common case of correlated effect. Furthermore, we also examined in detail DEXSeq’s aggregation (perGeneQvalue) which appears to be an attempt to perform Šidák aggregation but is not quite right, in a sense we explain in detail in Section 2 of the Yi et al. supplement. While DEXSeq’s implementation of Šidák aggregation does control the FDR, it will tend to report genes with many isoforms and consumes the “FDR budget” faster than Šidák aggregation. This is one reason why, for the purpose of comparing Lancaster and Šidák aggregation in Yi et al. 2018, we did not rely on DEXSeq’s implementation of Šidák aggregation. Needless to say, separately from this issue, as mentioned above we found that Lancaster aggregation substantially outperforms Šidák aggregation. The figures below complete the reproduction of the results of Cmero et al. The reproduced figures are are very similar to Cmero et al.’s figures but not identical. The difference is likely due to the fact that the Cmero paper states that a full comparison of the “Bottomly data” (on which these results are based) is a comparison of 10 vs. 10 samples. The reproduced results are based on downloading the data which consists of 10 vs. 11 samples for a total of 21 samples (this is confirmed in the Bottomly et al. paper which states that they “generated single end RNA-Seq reads from 10 B6 and 11 D2 mice.”) I also noticed one other small difference in the Drosophila analysis shown in Figure 3a where one of the methods is different for reasons I don’t understand. As for the supplement, the Cmero et al. figures are shown on the left hand side below, and to their right are the reproduced figures: The final supplementary figure is a comparison of kallisto to Salmon: the Cmero et al. paper shows that Salmon results are consistent with kallisto results shown in Figure 3a, and reproduces the claim I made in a previous blog post, namely that Salmon results are near identical to kallisto: The final paragraph in the discussion of Cmero et al. states that “[transcript compatibility counts] have the potential to be useful in a range of other expression analysis. In particular [transcript compatibility counts] could be used as the initial unit of measurement for many other types of analysis such as dimension reduction visualizations, clustering and differential expression.” In fact, transcript compatibility counts have already been used for all these applications and have been shown to have numerous advantages. See the papers: Many of these papers were summarized in a talk I gave at Cold Spring Harbor in 2017 on “Post-Procrustean Bioinformatics”, where I emphasized that instead of fitting methods to the predominant data types (in the case of RNA-seq, gene counts), one should work with data types that can support powerful analysis methods (in the case of RNA-seq, transcript compatibility counts). Three years ago, when my coauthors (Páll Melsted, Nicolas Bray, Harold Pimentel) and I published the “kallisto paper” on the arXiv (later Bray et al. “Near-optimal probabilistic RNA-seq quantification“, 2016), we claimed that kallisto removed a major computational bottleneck from RNA-seq analysis by virtue of being two orders of magnitude faster than other state-of-the-art quantification methods of the time, without compromising accuracy. With kallisto, computations that previously took days, could be performed as accurately in minutes. Even though the speedup was significant, its relevance was immediately questioned. Critics noted that experiments, library preparations and sequencing take at least months, if not years, and questioned the relevance of a speedup that would save only days. One rebuttal we made to this legitimate point was that kallisto would be useful not only for rapid analysis of individual datasets, but that it would enable analyses at previously unimaginable scales. To make our point concrete, in a follow-up paper (Pimentel et al., “The Lair: a resource for exploratory analysis of published RNA-seq data”, 2016) we described a semi-automated framework for analysis of archived RNA-seq data that was possible thanks to the speed and accuracy of kallisto, and we articulated a vision for “holistic analysis of [short read archive] SRA data” that would facilitate “comparison of results across studies [by] use of the same tools to process diverse datasets.” A major challenge in realizing this vision was that although kallisto was fast enough to allow for low cost processing of all the RNA-seq in the short read archive (e.g. shortly after we published kallisto, Vivian et al., 2017 showed that kallisto reduced the cost of processing per sample from$1.30 to $0.19, and Tatlow and Piccolo, 2016 achieved$0.09 per sample with it), an analysis of experiments consists of much more than just quantification. In Pimentel et al. 2016 we struggled with how to wrangle metadata of experiments (subsequently an entire paper was written by Bernstein et al. 2017 just on this problem), how to enable users to dynamically test distinct hypotheses for studies, and how to link results to existing databases and resources. As a result, Pimentel et al. 2016 was more of a proof-of-principle than a complete resource; ultimately we were able to set up analysis of only a few dozen datasets.

Now, the group of Avi Ma’ayan at the Icahn School of Medicine at Mount Sinai has surmounted the many challenges that must be overcome to enable automated analysis of RNA-seq projects on the short read archive, and has published a tool called BioJupies (Torre et al. 2018). To assess BioJupies I began by conducting a positive control in the form of analysis of data from the “Cuffdiff2” paper, Trapnell et al. 2013. The data is archived as GSE37704. This is the dataset I used to initially test the methods of Pimentel et al. 2016 and is also the dataset underlying the Getting Started Walkthrough for sleuth. I thought, given my familiarity with it, that it would be a good test case for BioJupies.

Briefly, in Trapnell et al. 2013, Trapnell and Hendrickson performed a differential analysis of lung fibroblasts in response to an siRNA knockdown of HOXA1 which is a developmental transcription factor. Analyzing the dataset with BioJupies is as simple as typing the Gene Expression Omnibus (GEO) accession on the BioJupies searchbox. I clicked “analyze”, clicked on “+” a few times to add all the possible plots that can be generated, and this opened a window asking for a description of the samples:

I selected “Perturbation” for the HOXA1 knockdown samples and “Control” for the samples that were treated with scrambled siRNA that did not target a specific gene. Finally, I  clicked on “generate notebook”…

and

BioJupies displayed a notebook (Trapnell et al. 2013 | BioJupies) with a complete analysis of the data. Much of the Trapnell et al. 2013 analysis was immediately evident in the notebook. For example, the following figure is Figure 5a in Trapnell et al. 2013. It is a gene set enrichment analysis (GSEA) of the knockdown:

BioJupies presents the same analysis:

It’s easy to match them up. Of course BioJupies presents a lot of other information and analysis, ranging from a useful PCA plot to an interesting L1000 connectivity map analysis (expression signatures from a large database of over 20,000 perturbations applied to various cell lines that match the signatures in the dataset).

One of the powerful applications of BioJupies is the presentation of ARCHS⁴ co-expression data. ARCHS⁴ is the kallisto computed database of expression for the whole and is the primary database that enables BioJupies. One of its features is a list of co-expressed genes (as ascertained via correlation across the whole short read archive). These are displayed in BioJupies making it possible to place the results of an experiment in the context of “global” transcriptome associations.

While the Trapnell et al. 2013 reanalysis was fun, the real power of BioJupies is clear when analyzing a dataset that has not yet been published. I examined the GEO database and found a series GSE60538 that appears to be a partial dataset from what looks like a paper in the works. The data is from an experiment designed to investigate the role of Sox5 and Sox6 in the mouse heart via two single knockout experiments, and a double knockout. The entry originates in 2014 (consistent with the single-end 50bp reads it contains), but was updated recently. There are a total of 8 samples: 4 controls and 4 from the double knockout (the single knockouts are not available yet). I could not find an associated paper, nor was one linked to on GEO, but the abstract of the paper has already been uploaded to the site. Just as I did with the Trapnell et al. 2013 dataset, I entered the accession in the BioJupies website and… four minutes later:

The abstract of the GSE60538 entry states that “We performed RNA deep sequencing in ventricles from DKO and control mice to identify potential Sox5/6 target genes and found altered expression of genes encoding regulators of calcium handling and cation transporters” and indeed, BioJupies verifies this result (see Beetz et al. GSE60538| BioJupies):

Of course, there is a lot more analysis than just this. The BioJupies page includes, in addition to basic QC and datasets statistics, the PCA analysis, a “clustergrammer” showing which genes drive similarity between samples, differentially expressed genes (with associated MA and volcano plots), gene ontology enrichment analysis, pathway enrichment analysis, transcription factor enrichment analysis, kinase enrichment analysis, microRNA enrichment analysis, and L1000 analysis. In a sense, one could say that with BioJupies, users can literally produce the analysis for a paper in four minutes via a website.

The Ma’ayan lab has been working towards BioJupies for some time. The service is essentially a combination of a number of tools, workflows and resources published previously by the lab, including:

With BioJupies, these tools become more than the sum of their parts. Yet while BioJupies is impressive, it is not complete. There is no isoform analysis, which is unfortunate; for example one of the key points of Trapnell et al. 2013 was how informative transcript-level analysis of RNA-seq data can be. However I see no reason why a future release of BioJupies can’t include detailed isoform analyses. Isoform quantifications are provided by kallisto and are already downloadable via ARCHS⁴. It would also be great if BioJupies were extended to organisms other than human and mouse, although some of the databases that are currently relied on are less complete in other model organisms. Still, it should even be possible to create a BioJupies for non-models. I expect the authors have thought of all of these ideas. I do have some other issues with BioJupies: e.g. the notebook should cite all the underlying programs and databases used to generate the results, and while it’s neat that there is an automatically generated methods section, it is far from complete and should include the actual calls made to the programs used so as to facilitate complete reproducibility. Then, there is my pet peeve: “library size” is not the number of reads in a sample. The number of reads sequenced is “sequencing depth”.  All of these issues can be easily fixed.

In summary, BioJupies represents an impressive breakthrough in RNA-seq analysis. It leverages a comprehensive analysis of all (human and mouse) publicly available RNA-seq data to enable rapid and detailed analyses that transcend what has been previously possible. Discoveries await.

The post concerns Yuval Peres, a principal researcher in the Microsoft Theory Group [update Dec. 26, 2018: YP is no longer employed at Microsoft] and a former colleague of mine at UC Berkeley. Below is a copy of an email sent yesterday to numerous theory of computer science professors worldwide, and published on the Stanford Theory Seminar List. It corroborates information I heard about Yuval Peres a number of years ago when I was a mathematics professor at UC Berkeley. At the time I was asked to keep the information I heard confidential, and I did so because the person who discussed it with me was, understandably, afraid of retaliation. Now I wonder to what extent my silence allowed his harassment of women to continue unabated. I also wonder when the leaders of the statistics department at UC Berkeley, where Peres used to work, and where Terry Speed was a professor emeritus before I reported him, will end their culture of silence.

Hello all,

This is an email composed by Irit Dinur, Oded Goldreich and me. The purpose of this email is to share with you concerns that we had regarding the unethical behavior of Yuval Peres. The behavior we are referring to includes several recent incidents from the past few years, on top of the two “big” cases of sexual harassment that led to severe sanctions against him by his employer, Microsoft, and to the termination of his connections with the University of Washington. Together with two colleagues who are highly regarded and trusted by us, we have first and second-hand testimonies (by people we trust without a shed of doubt) of at least five additional cases of him approaching junior female scientists, some of them students, with offers of intimate nature, behavior that has caused its victims quite a bit of distress since these offers were “insistent”. While the examples that we encountered from the last few years do not fall under the category of sexual harassment from a legal point of view, they certainly caused great discomfort to the victims, who were young female scientists, putting them in a highly awkward situation, and creating an atmosphere that they’d rather avoid (i.e., they would rather miss a conference or a lecture than risk being subjected to repeated intimate offers by him). We wish to stress that his aggressive advances toward young women, usually with no previous friendly connections with him, puts them in a vulnerable position of fearing to cross a senior scientist who might have an impact on their career, which is at a fragile stage. We believe that the questions of whether or not Yuval Peres intended to make them uncomfortable, and whether or not he would or could actually harm their scientific status are irrelevant; the fact is that the victims felt very stressed to a point that they’d rather miss professional events than risk encountering the same situation again. Needless to say, it is the responsibility of senior members of our community to avoid putting less senior members in such a position.

Our current involvement with this issue was triggered by an invitation Yuval Peres received to give a plenary talk at an international conference next year. We felt that this invitation sends a highly undesirable message to our community in general, and to the women he harassed in particular, as if his transgressions are considered unimportant.

We sent an email conveying our concern to the organizers of the conference, suggesting that they disinvite him. With our permission, they forwarded a version of our letter (in which we made changes in order to protect the identity of the women involved) to Yuval Peres. They did not reveal our identity, rather they told him that this is a letter from “senior members of the community”. In our letter we included a paragraph describing a general principle that should be followed. The principle is:

A senior researcher should not approach a junior researcher with an invitation that may be viewed as intimate or personal unless such an invitation was issued in the past by this specific junior to that specific senior. The point being that even if the senior researcher has no intimate/personal intentions, such intentions may be read by the junior researcher, placing the junior in an awkward situation and possibly causing them great distress. Examples for such an invitation include any invitation to a personal event in which only the senior and the junior will be present (e.g., a two-person dinner, a meeting in a private home, etc).

Yuval’s reply was rather laconic, in particular, he did not address the issue of his behavior in the past couple of years. However, he did write:

“I certainly embrace the principle described in boldface in the letter. This seems to be the right approach for any senior scientist these days.”

The reason we are copying this to all of you (as opposed, for example, to using bcc) is related to the islanders’ paradox: we believe that the fact that everyone knows that everyone knows is a significant boost to holding Yuval Peres accountable for his future actions. We’re also bcc’ing several young women who already aware of Yuval Peres’s actions, in order to keep them in the know too.

We understand that sending this out to a large number of people without offering Yuval Peres the chance to respond may be considered unfair. However, after weighing the pros and cons carefully we believe this is a good course of action. First of all, because it is clear that the victims did not invent his offers and their ensuing feelings of anxiety and stress. Secondly, we know that Yuval Peres has been confronted in a face to face conversation by a senior colleague, and it did not end his behavior, so we think it’s important to stay vigilant in protecting the younger members of our community. Thirdly, the information in this letter will reach (or has already reached) almost all of you in any case, so the main effect of the letter is making what everyone knows into public knowledge. Finally, although his response to the organizers did include the minimum of declaring he accepts the guiding principle that we stated, it did not include any reference to the ongoing behavior we described- neither regret nor concern nor denial. So it’s not easy to assume that he truly intends to mend his ways.

We hope that our actions will contribute to the future of our community as an environment that offers all a pleasant and non-threatening atmosphere.

Sincerely,
Irit Dinur, Ehud Friedgut, Oded Goldreich

Last year I wrote a blog post on being wrong. I also wrote a blog post about being wrong three years ago. It’s not fun to admit being wrong, but sometimes it’s necessary. I have to admit to being wrong again.

To place this admission in context I need to start with Mordell’s finite basis theorem, which has been on my mind this past week. The theorem, proved in 1922, states the rational points on an elliptic curve defined over the rational numbers form a finitely generated abelian group. There is quite a bit of math jargon in this statement that makes it seem somewhat esoteric, but it’s actually a beautiful, fundamental, and accessible result at the crossroads of number theory and algebraic geometry.

First, the phrase elliptic curve is just a fancy name for a polynomial equation of the form y² = x³ + ax + b (subject to some technical conditions). “Defined over the rationals” just means that and b are rational numbers. For example a=-36, b=0 or a=0, b=-26 would each produce an elliptic curve. A “rational point on the curve” refers to a solution to the equation whose coordinates are rational numbers. For example, if we’re looking at the case where a=0 and b=-26 then the elliptic curve is y² = x³ – 26 and one rational solution would be the point (35,-207). This solution also happens to be an integer solution; try to find some others! Elliptic curves are pretty and one can easily explore them in WolframAlpha. For example, the curve y² = x³ – 36x looks like this:

WolframAlpha does more than just provide a picture. It finds integer solutions to the equation. In this case just typing the equation for the elliptic curve into the WolframAlpha box produces:

One of the cool things about elliptic curves is that the points on them form the structure of an abelian group. That is to say, there is a way to “add” points on the curves. I’m not going to go through how this works here but there is a very good introduction to this connection between elliptic curves and groups in an exposition by Tanuj Nayak, an undergrad at Carnegie Mellon University.

Interestingly, even just the rational points on an elliptic curve form a group, and Mordell’s theorem says that for an elliptic curve defined over the rational numbers this group is finitely generated. That means that for such an elliptic curve one can describe all rational points on the curve as finite combinations of some finite set of points. In other words, we (humankind) has been interested in studying Diophantine equations since the time of Diophantus (3rd century). Trying to solve arbitrary polynomial equations is very difficult, so we restrict our attention to easier problems (elliptic curves). Working with integers is difficult, so we relax that requirement a bit and work with rational numbers. And here is a theorem that gives us hope, namely the hope that we can find all solutions to such problems because at least the description of the solutions can be finite.

The idea of looking for all solutions to a problem, and not just one solution, is fundamental to mathematics. I recently had the pleasure of attending a lesson for 1st and 2nd graders by Oleg Gleizer, an exceptional mathematician who takes time not only to teach children mathematics, but to develop mathematics (not arithmetic!) curriculum that is accessible to them. The first thing Oleg asks young children is what they see when looking at this picture:

Children are quick to find the answer and reply either “rabbit” or “duck”. But the lesson they learn is that the answer to his question is that there is no single answer! Saying “rabbit” or “duck” is not a complete answer. In mathematics we seek all solutions to a problem. From this point of view, WolframAlpha’s “integer solutions” section is not satisfactory (it omits x=6, y=0), but while in principle one might worry that one would have to search forever, Mordell’s finite basis theorem provides some peace of mind for an important class of questions in number theory. It also guides mathematicians: if interested in a specific elliptic curve, think about how to find the (finite) generators for the associated group. Now the proof of Mordell’s theorem, or its natural generalization, the Mordell-Weil theorem, is not simple and requires some knowledge of algebraic geometry, but the statement of Mordell’s theorem and its meaning can be explained to kids via simple examples.

I don’t recall exactly when I learned Mordell’s theorem but I think it was while preparing for my qualifying exam in graduate school, when I studied Silverman’s book on elliptic curves for the cryptography section on my qualifying exam- yes, this math is even related to some very powerful schemes for cryptography! But I do remember when a few years later a (mathematician) friend mentioned to me “the coolest paper ever”, a paper related to generalizations of Mordell’s theorem, the very theorem that I had studied for my exam. The paper was by two mathematicians, Steven Zucker and David Cox, and it was titled Intersection Number of Sections of Elliptic Surfaces. The paper described an algorithm for determining whether some sections form a basis for the Mordell-Weil group for certain elliptic surfaces. The content was not why my friend thought this paper was cool, and in fact I don’t think he ever read it. The excitement was because of the juxtaposition of author names. Apparently David Cox had realized that if he could coauthor a paper with his colleague Steven Zucker, they could publish a theorem, which when named after the authors, would produce a misogynistic and homophobic slur. Cox sought out Zucker for this purpose, and their mission was a “success”. Another mathematician, Charles Schwartz, wrote a paper in which he built on this “joke”. From his paper:

So now, in the mathematics literature, in an interesting part of number theory, you have the Cox-Zucker machine. Many mathematicians think this is hilarious. I thought this was hilarious. In fact, when I was younger I frequently boasted about this “joke”, and how cool mathematicians are for coming up with clever stuff like this.

I was wrong.

I first started to wonder about the Zucker and Cox stunt when a friend pointed out to me, after I had used the term C-S to demean someone, that I had just spouted a misogynistic and homophobic slur. I started to notice the use of the C-S phrase all around me and it made me increasingly uncomfortable. I stopped using it. I stopped thinking that the Zucker-Cox stunt was funny (while noticing the irony that the sexual innuendo they constructed was much more cited than their math), and I started to think about the implications of this sort of thing for my profession. How would one explain the Zucker-Cox result to kids? How would undergraduates write a term paper about it without sexual innuendo distracting from the math? How would one discuss the result, the actual math, with colleagues? What kind of environment emerges when misogynistic and homophobic language is not only tolerated in a field, but is a source of pride by the men who dominate it?

These questions have been on my mind this past week as I’ve considered the result of the NIPS conference naming deliberation. This conference was named in 1987 by founders who, as far as I understand, did not consider the sexual connotations (they dismissed the fact that the abbreviation is a racial slur since they considered it all but extinct). Regardless of original intentions I write this post to lend my voice to those who are insisting that the conference change its name. I do so for many reasons. I hear from many of my colleagues that they are deeply offended by the name. That is already reason enough. I do so because the phrase NIPS has been weaponized and is being used to demean and degrade women at one of the main annual machine learning conferences. I don’t make this claim lightly. Consider, for example, TITS 2017 (the (un)official sister event to NIPS). I’ve thought about this specific aggression a lot because in mathematics there is a mathematician by the name of Tits who has many important objects named after him (e.g. Tits buildings). So I have worked through the thought experiment of trying to understand why I think it’s wrong to name a conference NIPS but I’m fine talking about the mathematician Tits. I remember when I first learned of Tits buildings I was taken aback for a moment. But I learned to understand the name Tits as French and I pronounce it as such in my mind and with my voice when I use it. There is no problem there, nor is there a problem with many names that clash across cultures and languages. TITS 2017 is something completely different. It is a deliberate use of NIPS and TITS in a way that can and will make many women uncomfortable. As for NIPS itself perhaps there is a “solution” to interpreting the name that doesn’t involve a racial slur or sexual innuendo (Neural Information Processing Systems). Maybe some people see a rabbit. But others see a duck. All the “solutions” matter. The fact is many women are uncomfortable because instead of being respected as scientists, their bodies and looks have become a subtext for the science that is being discussed. This is a longstanding problem at NIPS (see e.g., Lenna). Furthermore, it’s not only women who are uncomfortable. I am uncomfortable with the NIPS name for the reasons I gave above, and I know many other men are as well. I’m not at ease at conferences where racial slurs and sexual innuendo are featured prominently, and if there are men who are (cf. NIPS poll data) then they should be ignored.

I think this is an extremely important issue not only for computer science, but for all of science. It’s about much more than a name of some conference. This is about recognizing centuries of discriminatory and exclusionary practices against women and minorities, and about eliminating such practices when they occur now rather than encouraging them. The NIPS conference must change their name. #protestNIPS

A few years ago I wrote a post arguing that it is time to end ordered authorship. However that time has not yet arrived, and it appears that it is unlikely to arrive anytime soon. In the meantime, if one is writing a paper with 10 authors, a choice for authorship ordering and equal contribution designation must be made from among the almost 2 billion possibilities (1857945600 to be exact). No wonder authorship arguments are commonplace! The purpose of this short post is to explain the number 1857945600.

At first glance the enumeration of authorship orderings seems to be straightforward, namely that in a paper with n authors there are n! ways to order the authors. However this solution fails to account for designation of authors as “equal contributors”. For example, in the four author paper Structural origin of slow diffusion in protein folding, the first two authors contributed equally, and separately from that, so did the last two (as articulated via a designation of “co-corresponding” authorship). Another such example is the paper PRDM/Blimp1 downregulates expression of germinal center genes LMO2 and HGAL. Equal contribution designations can be more complex. In the recent preprint Connect-seq to superimpose molecular on anatomical neural circuit maps the first and second authors contributed equally, as did the third and fourth (though the equal contributions of the first and second authors was distinct from that of the third and fourth). Sometimes there are also more than two authors who contributed equally. In SeqVis: Visualization of compositional heterogeneity in large alignments of nucleotides the first eight authors contributed equally. A study on “equal contribution” designation in biomedical papers found that this type of designation is becoming increasingly common and can be associated with nearly every position in the byline.

To account for “equal contribution” groupings, I make the assumption that a set of authors who contributed equally must be consecutive in the authorship ordering. This assumption is certainly reasonable in the biological sciences given that there are two gradients of “contribution” (one from the front and one from the end of the authorship list), and that contributions for those in the end gradient are fundamentally distinct from those in the front. An authorship designation for a paper with n authors therefore consists of two separate parts: the n! ways to order the authors, and then the $2^{n-1}$ ways of designating groups of equal contribution for consecutive authors. The latter enumeration is simple: designation of equal authorship is in one-to-one correspondence with placement of dividers in the n-1 gaps between the authors in the authorship list. In the extreme case of placement of no dividers the corresponding designation is that all authors contributed equally. Similarly, the placement of dividers between all consecutive pairs of authors corresponds to all contributions being distinct. Thus, the total number of authorship orderings/designations is given by $n! \cdot 2^{n-1}$. These numbers also enumerate the number of ways to lace a shoe. Other examples of objects whose enumeration results in these numbers are given in the Online Encyclopedia of Integer Sequences entry for this sequence (A002866). The first twenty numbers are:

1, 4, 24, 192, 1920, 23040, 322560, 5160960, 92897280, 1857945600, 40874803200, 980995276800, 25505877196800, 714164561510400, 21424936845312000, 685597979049984000, 23310331287699456000, 839171926357180416000, 31888533201572855808000, 1275541328062914232320000.

In the case of a paper with 60 authors, the number of ways to order authors and designate equal contribution is much larger than the number of atoms in the universe. Good luck with your next consortium project!