The Mammal’s very own Hox genes (excite! Woo!)

It’s kind of hard to begin this post. First of all, let’s get the important news out of the way: I’ve just published a paper. In a moment, I’ll get around to discussing it at even more than my usual length, but I feel that I can’t do my excited puppy act without at least trying to capture how bloody much this paper means to me. The following may get a little personal; if you want to jump straight to the Cool Stuff, feel free to scroll a couple of paragraphs down.

<personal bit>

As you may have guessed from the long silence here, it’s not been a good handful of years, Real Life and mental health-wise. After my PhD, the prospect of the research career I’d dreamed of since I first began to grasp the meaning of the word “scientist” no longer seemed so dreamlike. It may surprise you to hear this from someone who finished a PhD with four published papers and spent the years of said PhD blathering regularly on the internet, but I find writing things for other people to read very, very stressful. In the case of a job application or a thesis chapter, that becomes “I’m not eating or sleeping properly” stressful. (Don’t ask me how I survived 20+ years of formal education.)

Long story short, for the last 3 years I’ve been getting by with a minimum wage job for which I’m both vastly overqualified and singularly ill-suited. I started the research project that culminated in the paper you can now read (for free, yay!) in BMC Evolutionary Biology (Szabó and Ferrier, 2018) while unemployed and broke, and I did most of it in my free time around work. This paper is a hard-won victory over myself and my circumstances. It’s a tiny glint of self-worth in the depth of the tunnel. In some ways, it was harder than my thesis: no funding body to satisfy, no lab mates to gripe at, no deadlines to spur me on. The only constant was my ex-supervisor turned co-author, who took my hobby project under his wings for the slim reward of having his name on a paper and nudged me into finishing it with unending patience. Here’s to Dave Ferrier, champion of non-model organisms, homeobox guy extraordinaire and all-round excellent human being. Dave, I hope you know you’re an absolute star.

</personal bit>

With that out of the way, it’s time for the Cool Stuff. There are Hox genes! More Hox genes than anyone ever imagined! (That is kind of the point, in fact!)

Apologies for the word count. I thought it would be a good idea to explain a few things, but also, I think I enjoy waffling about my baby far too much 😊

Hox therapy

The story of my Hox paper begins with an unemployed biologist with an overabundance of free time and a desperate need to do something scientific. Since I have a slightly odd idea of “fun”, back in 2015 I decided to catalogue Hox gene (or rather, protein) diversity in the animal kingdom, with particular focus on obscure and poorly studied groups. (I didn’t get very far, as we’ll see.)

Since it’s hard to discuss the paper without dropping some arcane zoological nomenclature, here’s my trusty old animal phylogeny to (re)acquaint us with the general outlines of the animal kingdom (I might need to update this in light of the Great Ctenophore Controversy some day, but we’re not dealing with anything outside the Bilateria today):

animalPhylogeny

For the purposes of my paper, we’re zooming into the deuterostome branch, which looks something like this on the inside (borrowing my own rather lacklustre last-minute figure from Szabó and Ferrier [2018]):

12862_2018_1307_Fig1_HTML

Everything on this tree apart from chordates (that’s us) belongs to a group called Ambulacraria, which contains two phyla, hemichordates (top two branches) and echinoderms (the next five). Echinoderms are the more familiar of the two – starfish and sea urchins and suchlike – and also the focus of my project. (I could find no Hox gene data from pterobranchs, which puts a slight caveat on everything I say about hemichordates)

Back to Hox genes.

Hox genes were kind of my gateway drug into evolutionary developmental biology. A few decades earlier, they had served the same purpose for developmental biology as a whole, since they were among the first genes to be discovered that (1) directed embryonic development (2) were comparable between very disparate animal groups. The short version, which will suffice for our purposes here, is that Hox genes are important in what we eggheads call anteroposterior patterning, or determining what body parts go where along the head (anterior) to tail (posterior) axis of a (bilaterian) animal.

In (I think, I haven’t counted) the majority of animals that have them, Hox genes are clustered to a greater or lesser extent. Rather than being scattered haphazardly across the genome, they sit close to one another along the same stretch of DNA. (Duboule [2007] is an excellent – albeit now slightly out of date – review of the various known configurations.)

Since my study is about echinoderms, the schematic Hox cluster shown below is the neatest known example from an echinoderm, the crown-of-thorns starfish Acanthaster planci (source: Baughman et al., 2014):

baughman2014_1

In this image, Hox genes are colour-coded according to a commonly used classification scheme. This classification is mostly based on the homeodomain, or the “business” end of the protein that a Hox gene encodes. A homeodomain makes up a relatively small portion (maybe 1/5th on average) of a typical Hox protein, but it’s the part that interacts with the DNA switches through which Hoxes control their target genes, and it’s often the only part that is similar enough to be compared between different Hox types.

The important genes for us today are the “posterior” Hox genes shown in pink and red above, especially the last two. The four posterior Hox genes seen here represent the “standard” set for ambulacrarians, although it’s uncertain whether Hox11/13b-c were already separate genes or just a single precursor gene in the ambulacrarian ancestor.

Eureka… or WTF?

“The most exciting phrase to hear in science, the one that heralds new discoveries, is not Eureka! (I found it!) but rather, ‘hmm… that’s funny…”Almost certainly not Isaac Asimov

In creating my grand catalogue, I’d quickly breezed through vertebrates (which are all essentially the same for my purposes) and other chordates (for which the data I could find were rather limited). I thought echinoderms would be an easy job, too: there were good in-depth studies of a few species, and they hadn’t revealed anything terribly unusual other than a rearrangement of the Hox cluster in sea urchins (Cameron et al., 2006).

In fact, through comparison with their sister group, the hemichordates (Freeman et al., 2012), it seemed likely that the ancestral echinoderm had a nice, ordered Hox cluster with few if any oddities (Baughman et al., 2014). So I clicked my way to the wonderful Echinobase, which has searchable draft genomes from four of the five living classes of echinoderms (crinoids, a.k.a sea lilies and feather stars, are missing, although a genome in a very early, fragmentary stage exists here). I expected to double-check the published data, collect the same genes from the groups for which Hox papers hadn’t been published, and be off to protostomes in a day or two. Two years later, I still haven’t made it to protostomes, but I’ve gone rather deeper than expected in echinoderms…

(Below: my cast. The main characters are Strongylocentrotus purpuratus [photo: Kirt L. Onthank] and Lytechinus variegatus [photo: Hans Hillewaert] representing sea urchins, Patiria miniata [photo: Jerry Kirkhart] and Acanthaster planci [photo: JSLUCAS75] for sea stars, Parastichopus parvimensis [from here] and Apostichopus japonicus [photo: OpenCage] for sea cucumbers, Metacrinus rotundus [photo: OpenCage] and Anneissia japonica [photo: OpenCage] for crinoids, Ophiothrix spiculata [photo: Jerry Kirkhart] for brittle stars, with supporting acts from Peronella japonica [sea urchins, photo: Endo et al., 2018], Ophiopsila aranea [brittle stars, photo: Bernard Picton], Balanoglossus simodensis [photo: Misaki Marine Biological Station, U of Tokyo], Saccoglossus kowalevskii [photo: Lowe lab] and Ptychodera flava [photo: Moorea BioCode via CalPhotos] for hemichordates, and Branchiostoma floridae [photo via JGI genome portal], Latimeria menadoensis [photo: Claudio Martino] and Callorhinchus milii [photo: fir0002/Flagstaffotos] for chordates. I sourced the photos through Wikipedia/Wikimedia Commons where I could; other sources are linked where applicable.)

cast

You see, I didn’t want to stop at just homeodomains. Homeodomains are cool and important and all, but one thing I’d learned from my earlier forays into the world of Hox genes was that valuable information hid in small patches of conserved sequence elsewhere in their proteins. Besides, I am a pathological perfectionist. I felt a terrible need to collect complete Hox sequences wherever possible.

I already mentioned that sequence similarity between Hoxes outside the homeodomain can be weak to non-existent. I ran into this problem with Echinobase’s brittle star, Ophiothrix spiculata. Using the known sea urchin Hoxes to search its genome, I’d found believable matches for many of them, but the 11/13s defeated me. I had two homeodomains that I thought represented 11/13b and c, but I couldn’t for the life of me recover the rest of the proteins.

The problem with genome databases (or their great advantage depending on your perspective) is that they contain all of the DNA that could be sequenced from the owner of the genome. The problem with Hox genes – most of our genes, in fact – is that they aren’t continuous stretches of DNA. Your typical gene exists in multiple segments (exons) separated by a whole lot of DNA that leaves no trace in the protein product of the gene. (Hox genes normally have two or three exons, the first of which is devoid of homeodomain parts.)

When a gene is expressed, the cell first makes an RNA copy of all that, which is edited to throw out the introns and splice the exons together. That intron-less RNA copy is then carried off to be translated into a protein. Transcriptomes are derived from the RNA copies of active genes. Introns lie forgotten on the cutting room floor: in the sequenced transcripts, one exon continues straight into the next. Therefore, if I could find a brittle star transcriptome, and the 11/13b-c homeodomains in it, perhaps there would be enough of the rest in there to reconstruct those elusive first exons.

Luckily, Delroisse et al. (2016) had published exactly what I needed. In one of their transcriptomes, I found a homeodomain that looked like my Ophiothrix Hox11/13c, as part of a near-complete sequence. Excited, I did the reciprocal search against the Ophiothrix genome…

… and hit neither 11/13b nor 11/13c.

So here I am, staring at a beautiful match between this transcript and a part of the Ophiothrix genome that I hadn’t examined before. The match contains sequence from the first exon, which, given my previous experience with these buggers, is a sure sign that they’re the same gene. And it’s neither of the ones I’d expected.

A bit later in a different database, I hit upon an automatically predicted sea urchin protein that definitely isn’t 11/13b or c either. This is the model sea urchin, S. purpuratus, the one I thought we knew inside out when it came to Hoxes. I check the genome on Echinobase, and lo and behold, there’s the third 11/13b-c type gene, and it’s nowhere near the Hox cluster.

If memory serves, it’s roughly at this point that the words, “What. The. Actual. Fuck. Is. Going. On.” occur in my research notes. (Complete with punctuation.)

I checked the other species on Echinobase. Three 11/13b-c genes again, every time. Over on Genbank, I found a complete protein sequence from a sand dollar that Tsuchimoto and Yamaguchi (2014) had previously classified as 11/13c by exclusion. The Japanese duo had a clear b, but this other sequence was behaving oddly in their phylogenetic analyses. Now I had the obvious explanation: it wasn’t 11/13c at all.*

I wrote to Dave and found out that this was also news to him. By all appearances, I had stumbled on something truly new, in a gene family that’s both iconic in our field, and dear to my obsessive little heart.

We decided to try to turn it into a paper.

In search of the alphabet’s end

Once we’d made that decision, and following Dave’s advice, I had a few tasks ahead of me. I had to check how far back in evolution our new gene (which we called Hox11/13d) went. I had to test whether it had truly escaped the Hox cluster in all of our study species. I had to refresh my memory on deuterostome posterior Hox genes in general, both for paper-writing purposes and in case there was a forgotten reference to our “new” gene lurking somewhere in the literature.

There wasn’t, but.

In a figure legend in Thomas-Chollier et al., 2010), there is a brief mention of an unnamed “Hox11/13c-like” sequence in sea urchins. When I saw that, I damn near soiled myself, but the authors couldn’t definitively identify this sequence as a Hox gene, so they left it at that throwaway comment and a few bits of supplementary data. Luckily, they had a gene ID that I could look up on Echinobase.

Gods help me, it turned out to be another new Hox. When the shock of Hox11/13d had barely worn off, I was confronted with a possible Hox11/13e. And this one wasn’t in the Hox cluster either.

Aside from not being part of the Hox cluster, Hox11/13d is a pretty good echinoderm Hox gene. The homeodomain it encodes is reminiscent of Hox11/13b and c, and, although they are hard for automated searches to find, there are similarities outside the homeodomain that place it firmly in the same group as b-c.

Unlike d, Thomas-Chollier’s “11/13c-like” sequence isn’t that 11/13c-like at all, as you might have guessed from the fact that they weren’t even sure it’s a Hox. The region immediately following the homeodomain (sometimes known as the C-peptide) is very similar to the same part of Hox11/13d. These kinds of motifs can sometimes be used to tell different Hox genes apart. Two C-peptides being strongly similar is a clue that we’re dealing with related genes. However, the homeodomain of Hox11/13e, as we indeed dubbed Thomas-Chollier’s sequence, is really, really weird. It isn’t just unlike 11/13c, it’s unlike anything else I’d seen before. It groups with posterior Hoxes when we test it against a variety of homeodomains, but you wouldn’t know that simply from looking at it.

It is, however, an oddball with a history. As strange as that homeodomain is, once I knew what I was looking for, I found examples in all my other echinoderms. This combination of strong conservation of one Hox gene with considerable differences from other Hox genes just screams “study me more!”, especially when you realise that Hox11/13e appears to be limited to echinoderms (unless something like it is hiding in protostomes…). I looked quite carefully in the hemichordates available to me (Simakov et al., 2015), but the only thing I found that wasn’t one of the “canonical” four posteriors is something called “Abdominal B-like”, which is weird in its own way and not obviously connected to either of our two new genes.

Tangled histories and unhelpful clues

I alluded to the question of Hox11/13b-c origins earlier on. Posterior Hox genes in deuterostomes are notoriously difficult to classify (Ferrier et al., 2000; Thomas-Chollier et al., 2010). When you try to use traditional tree-building methods on them, you get a big unresolved mess, as if the twigs on the tree emerged from an impenetrable mist that hides the arrangement of the older branches from view. Ambulacrarians are definitely the better-behaved half of the Deuterostomia in this regard, since we can say with some confidence that Hox9/10, 11/13a and at least a single precursor to 11/13b-c were present in their last common ancestor.

Nonetheless, two new genes, at least one of which is clearly close to 11/13b-c, complicate matters (Abdominal B-like, as they say in scientist-speak, is beyond the scope of this work). Were they lost in hemichordates? Did echinoderms undergo extra gene duplications, and if so, was it from one or two ancestral genes? Where on earth does Hox11/13e fit? I did a lot of exploratory tree-building for this paper, none of which was particularly helpful in answering those questions.

My other hope was to look at the parts of the protein sequence that led me to my new Hoxes in the first place: all the stuff other than the homeodomain. Using a program called MEME, I found a fair few conserved motifs, but they only seemed to add to the confusion. Hox11/13e, for which I only had first exons (and tentative ones at that) from sea urchins and sea stars, yielded nothing of use apart from its striking C-peptide. In the others, the distribution of motifs created a patchwork of similarities that didn’t neatly align with any one possible history. Echinoderm Hox11/13c mostly did its own thing, while b and d each shared a different subset of motifs with one or both of the hemichordate b-c proteins.

I’m almost inclined to think that there was a single, “prototype” Hox11/13b+ sequence in the ambulacrarian ancestor, which contained all of the motifs I found. In that scenario, separate b and c (and d and maybe e) genes would have evolved independently in hemichordates and echinoderms, and each descendant gene would have lost some of the original motifs more or less at random. Duplicated genes can split the functions of their single ancestor between them (Force et al., 1999), so why not motifs? Short sequence motifs like the ones I was looking for can have important functions, after all. It’s a possibility, but we may never know for sure.

Hox genes gone rogue

I mentioned before that Hox11/13d was outside the Hox cluster. Well, so is Hox11/13e. As far as I can tell, Hox 11/13d and e always reside on separate chunks of the genome form any other Hox gene, including each other. They are always accompanied by neighbouring genes that aren’t Hoxes. Although detachment of a posterior gene from an otherwise apparently intact Hox cluster also happened in ragworms (Hui et al., 2012), it’s still a surprise in echinoderms. Since the relationship between the organisation of Hox genes and their regulation in space and time is… kinda complicated, we can’t really tell what, if anything, all this wandering implies without actually looking at some gene expression.

What are they for?

Then there’s the question of what on earth these genes do. Thanks to Tsuchimoto and Yamaguchi (2014), we know that Hox11/13d is active in later embryonic stages of some sea urchins. It even looks like it might be working with Hox11/13b in a Hox-like fashion, the two of them having adjacent expression domains. We have some transcriptomic evidence that this gene is also active in other sea urchins, brittle stars and starfish, but no idea what it’s doing in any of the above.

We know even less about Hox11/13e. The only evidence for expression I’m aware of is from starfish testicles, and testicles will express any old piece of DNA with an “on” switch. If it’s somehow involved in development, it must be either at very low levels that are difficult to capture in a transcriptome, or at developmental stages that weren’t included in the data I encountered.

If it does have a role in adult echinoderm development, that would be crazy exciting, as both adult echinoderm anatomy and Hox11/13e are so weird and unique. Although they develop from bilaterally symmetrical larvae, adult echinoderms have dispensed with the symmetry that gave Bilateria its name. Instead, like a sea anemone (or a regular anemone…), they are radially symmetrical. Hox genes are involved in both larval and adult development in echinoderms, but from what little I’ve been able to glean from the existing literature, it’s different subsets in larvae and adults rather than the entire Hox cluster together. Is Hox11/13e in the “adult” subset, missed until now due to its unusual sequence? I really hope someone with a lab and a ready supply of baby echinoderms investigates in the near future…

A lesson about expectations

I could go on for a lot longer about this project, but it’s probably time to form some sort of conclusion. For me, perhaps the most important take-home message of this adventure is not what I found, but how and where and why I found it.

I didn’t set out to discover anything. All I wanted to do was collect and organise information already out there. (If a genie popped out of my desk lamp, I might just wish for a full-time job where I get to build my Hox directory… given the volume of genome data already out there and coming out every time I look, continuing this as a hobby project in my free time seems hopelessly Sisyphean now.)

The discovery of Hox11/13d and all that followed was an accidental side effect of my penchant for perfectionism. If I’d contented myself with the homeodomains most students of Hox evolution focus on, I would never have seen a Hox that wasn’t in the books, a Hox I hadn’t expected to exist.

Expectations are important. I’d told myself that I wanted to make sure I had everything, but when my searches spat out a hundred different results, I started to slack off soon after I ticked off the Hoxes I knew. I gave the rest of the hit list a half-hearted effort at best. Hox11/13d has a homeodomain that’s split across two exons, and Hox11/13e is weird. In a search that scores both the closeness and the length of a match, that pushes them to the bottom of the results, where a casual observer, or an observer who thinks they know what they’re looking for, will most likely miss them. I thought I knew that sea urchins had a single, intact(ish) Hox cluster with 11 genes. I’d read a pretty good paper on it. Only the paper wasn’t quite right, after all.

To me, this study stands as a reminder to keep looking. In an era when new genomes are popping up left and right and Big Data with automated analyses is the scientific zeitgeist, it’s still worth rolling your sleeves up, picking up the old magnifying glass and taking a closer look – even in organisms you think you know. You might just chance upon some real treasure.

***

Note:

*A “Hox11/13c” behaving oddly should be immediately suspicious based on what I saw in my own trees, where echinoderm Hox11/13c consistently formed a strongly supported group. But that’s hindsight for you…

***

References:

Baughman KW et al. (2014) Genomic organization of Hox and ParaHox clusters in the echinoderm, Acanthaster planci. Genesis 52:952-958

Cameron RA et al. (2006) Unusual gene order and organization of the sea urchin hox cluster. JEZ B 306:45-58

Delroisse J et al. (2016) De novo adult transcriptomes of two European brittle stars: spotlight on opsin-based photoreception. PLoS ONE 11: e0152988

Duboule D (2007) The rise and fall of Hox gene clusters. Development 134:2549-2560

Endo M et al. (2018) Hidden genetic history of the Japanese sand dollar Peronella (Echinoidea: Laganidae) revealed by nuclear intron sequences. Gene 659:37-43

Ferrier DEK et al. (2000) The amphioxus Hox cluster: deuterostome posterior flexibility and Hox14. Evol Dev 2:284-293

Force A et al. (1999) Preservation of duplicate genes by complementary, degenerative mutations. Genetics 151:1531-1545

Freeman R et al. (2012) Identical genomic organization of two hemichordate Hox clusters. Curr Biol 22:2053-2058

Hui JH et al. (2012) Extensive chordate and annelid macrosyntheny reveals ancestral homeobox gene organization. Mol Biol Evol 29:157-165

Simakov O et al. (2015) Hemichordate genomes and deuterostome origins. Nature 527:459-465

Szabó R and Ferrier DEKF (2018) Two more Posterior Hox genes and Hox cluster dispersal in echinoderms. BMC Evol Biol 18:203

Thomas-Chollier M et al. (2010) A non-tree-based comprehensive study of metazoan Hox and ParaHox genes prompts new insights into their origin and evolution. BMC Evol Biol 10:73

Tsuchimoto J and Yamaguchi M (2014) Hox expression in the direct-type developing sea urchin Peronella japonica. Dev Dyn 243:1020-1029

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In which a “living fossil’s” genome delights me

I promised myself I wouldn’t go on for thousands and thousands of words about the Lingula genome paper (I’ve got things to do, and there is a LOT of stuff in there), but I had to indulge myself a little bit. Four or five years ago when I was a final year undergrad trying to figure out things about Hox gene evolution, I would have killed for a complete brachiopod genome. Or even a complete brachiopod Hox cluster. A year or two ago, when I was trying to sweat out something resembling a PhD thesis, I would have killed for some information about the genetics of brachiopod shells that amounted to more than tables of amino acid abundances. Too late for my poor dissertations, but a brachiopod genome is finally sequenced! The paper is right here, completely free (Luo et al., 2015). Yay for labs who can afford open-access publishing!

In case you’re not familiar with Lingula, it’s this guy (image from Wikipedia):

In a classic case of looks being deceiving, it’s not a mollusc, although it does look a bit like one except for the weird white stalk sticking out of the back of its shell. Brachiopods, the phylum to which Lingula belongs, are one of those strange groups no one really knows where to place, although nowadays we are pretty sure they are somewhere in the general vicinity of molluscs, annelid worms and their ilk. Unlike bivalve molluscs, whose shell valves are on the left and right sides of the animal, the shells of brachiopods like Lingula have top and bottom valves. Lingula‘s shell is also made of different materials: while bivalve shells contain calcium carbonate deposited into a mesh of chitin and silk-like proteins,* the subgroup of brachiopods Lingula belongs to uses calcium phosphate, the same mineral that dominates our bones, and a lot of collagen (again like bone). But we’ll come back to that in a moment…

One of the reasons the Lingula genome is particularly interesting is that Lingula is a classic “living fossil”. In the Paleobiology Database, there’s even an entry for a Cambrian fossil classified as Lingula, and there are plenty of entries from the next geological period. If the database is to be believed, the genus Lingula has existed for something like 500 million years, which must be some kind of record for an animal.** Is its genome similarly conservative? Or did the DNA hiding under a deceptively conservative shell design evolve as quickly as anyone’s?

In a heroic feat of self-control, I’m not spending all night poring over the paper, but I did give a couple of interesting sections a look. Naturally, the first thing I dug out was the Hox cluster hiding in the rather large supplement. This was the first clue that Lingula‘s genome is definitely “living” and not at all a fossil in any sense of the word. If it were, we’d expect one neat string of Hox genes, all in the order we’re used to from other animals. Instead, what we find is two missing genes, one plucked from the middle of the cluster and tacked onto its “front” end, and two genes totally detached from the rest. It’s not too bad as Hox cluster disintegration goes – six out of nine genes are still neatly ordered – but it certainly doesn’t look like something left over from the dawn of animals.

The bigger clue that caught my eye, though, was this little family tree in Figure 2:

Luo_etal2015-fig2

The red numbers on each branch indicate the number of gene families that expanded or first appeared in that lineage, and the green numbers are the families shrunk or lost. Note that our “living fossil” takes the lead in both. What I find funny is that it’s miles ahead of not only the animals generally considered “conservative” in terms of genome evolution, like the limpet Lottia and the lancelet Branchiostoma, but also the sea squirt (Ciona). Squirts are notorious for having incredibly fast-evolving genomes; then again, most of that notoriety was based on the crazily divergent sequences and often wildly scrambled order of its genes. A genome can be conservative in some ways and highly innovative in others. In fact, many of the genes involved in basic cellular functions are very slow-evolving in Lingula. (Note also: humans are pretty slow-evolving as far as gene content goes. This is not the first study to find that.)

So, Lingula, living fossil? Not so much.

The last bit I looked at was the section about shell genetics. Although it’s generally foolish to expect the shell-forming gene sets of two animals from different phyla to be similar (see my first footnote), if there are similarities, they could potentially go at least two different ways. First, brachiopods might be quite close to molluscs, which is the hypothesis Luo et al.‘s own treebuilding efforts support. Like molluscs, brachiopods also have a specialised mantle that secretes shell material, though having the same name doesn’t mean the two “mantles” actually share a common origin. So who knows, some molluscan shell proteins, or shell regulatory genes, might show up in Lingula, too.

On the other hand, the composition of Lingula’s shell is more similar to our skeletons’. So, since they have to capture the same mineral, could the brachiopods share some of our skeletal proteins? The answer to both questions seems to be “mostly no”.

Molluscan shell matrix proteins, those that are actually built into the structure of the shell, are quite variable even within Mollusca. It’s probably not surprising, then, that most of the relevant genes that are even present in Lingula are not specific to the mantle, and those that are are the kinds of genes that are generally involved in the handling of calcium or the building of the stuff around cells in all kinds of contexts. Some of the regulatory mechanisms might be shared – Luo et al. report that BMP signalling seems to be going on around the edge of the mantle in baby Lingula, and this cellular signalling system is also involved in molluscan shell formation. Then again, a handful of similar signalling systems “are involved” in bloody everything in animal development, so how much we can deduce from this similarity is anyone’s guess.

As for “bone genes” – the ones that are most characteristically tied to bone are missing (disappointingly or reassuringly, take your pick). The SCPP protein family is so far known only from vertebrates, and its various members are involved in the mineralisation of bones and teeth. SCPPs originate from an ancient protein called SPARC, which seems to be generally present wherever collagen is (IIRC, it’s thought to help collagen fibres arrange themselves correctly). Lingula has a gene for SPARC all right, but nothing remotely resembling an SCPP gene.

I mentioned that the shell of Lingula is built largely on collagen, but it turns out that it isn’t “our” kind of collagen. “Collagen” is just a protein with a particular kind of repetitive sequence. Three amino acids (glycine-proline-something else, in case you’re interested) are repeated ad nauseam in the collagen chain, and these repetitive regions let the protein twist into characteristic rope-like fibres that make collagen such a wonderfully tough basis for connective tissue. Aside from the repeats they all share, collagens are a large and diverse bunch. The ones that form most of the organic matrix in bone contain a non-repetitive and rather easily recognised domain at one end, but when Luo et al. analysed the genome and the proteins extracted from the Lingula shell, they found that none of the shell collagens possessed this domain. Instead, most of them had EGF domains, which are pretty widespread in all kinds of extracellular proteins. Based on the genome sequence, Lingula has a whole little cluster of these collagens-with-EGF-domains that probably originated from brachiopod-specific gene duplications.

So, to recap: Lingula is not as conservative as its looks would suggest (never judge a living fossil by its cover, right?) We also finally have actual sequences for lots of its shell proteins, which reveal that when it comes to building shells, Lingula does its own thing. Not much of a surprise, but still, knowing is a damn sight better than thinkin’ it’s probably so. We are scientists here, or what.

I am Very Pleased with this genome. (I just wish it was published five years ago 😛 )

***

Notes:

*This, interestingly, doesn’t seem to be the general case for all molluscs. Jackson et al. (2010) compared the genes building the pearly layer of snail (abalone, to be precise) and bivalve (pearl oyster) shells, and found that the snail showed no sign of the chitin-making enzymes and silk type proteins that were so abundant in its bivalved cousins. It appears that even within molluscs, different groups have found different ways to make often very similar shell structures. However, all molluscs shells regardless of the underlying genetics are predominantly composed of calcium carbonate.

**You often hear about sharks, or crocodiles, or coelacanths, existing “unchanged” for 100 or 200 or whatever million years, but in reality, 200-million-year-old crocodiles aren’t even classified in the same families, let alone the same genera, as any of the living species. Again, the living coelacanth is distinct enough from its relatives in the Cretaceous, when they were last seen, to warrant its own genus in the eyes of taxonomists. I’ve no time to check up on sharks, but I’m willing to bet the situation is similar. Whether Lingula‘s jaw-dropping 500-million-year tenure on earth is a result of taxonomic lumping or the shells genuinely looking that similar, I don’t know. Anyway, rant over.

***

References:

Jackson DJ et al. (2010) Parallel evolution of nacre building gene sets in molluscs. Molecular Biology and Evolution 27:591-608

Luo Y-J et al. (2015) The Lingula genome provides insights into brachiopod evolution and the origin of phosphate biomineralization. Nature Communications 6:8301

Precambrian muscles??? Oooooh!

Okay, consider this a cautious squee. I wish at least some of those Ediacaran fossils were a little more obvious. I mean, I might love fossils, but I’m trained to squirt nasty chemicals on bits of dead worm and play with protein sequences, not to look at faint impressions in rock and see an animal.

Most putative animals from the Ediacaran period, the “dark age” that preceded the Cambrian explosion, are confusing to the actual experts, not just to a lab/computer biologist with a fondness for long-dead things. The new paper by Liu et al. (2014) this post is about lists a “but see” for pretty much every interpretation they cite. The problem is twofold: one, as far as I can tell, most Ediacaran fossils don’t actually preserve that much interpretable detail. Two, Ediacaran organisms lived at a time when the kinds of animal body plans we’re familiar with today were just taking shape. The Ediacaran is the age of ancestors, and it would be more surprising to find a creature we can easily categorise (e.g. a snail) than a weird beastie that isn’t quite anything we know.

Having said that, Liu et al. think they are able to identify the new fossil they named Haootia quadriformis. Haootia comes from the well-known Fermeuse Formation of New Foundland, and is estimated to be about 560 million years old. The authors say its body plan – insofar as it can be made out on a flat image pressed into the rock – looks quite a lot like living staurozoan jellyfish, with a four-part symmetry and what appear to be branching arms or tentacles coming off the corners of its body. The most obvious difference is that Haootia seems to show the outline of a huge circular holdfast that’s much wider than usual for living staurozoans.

However, the most exciting thing about this fossil is not its shape, but the fact that most of it is made up of fine, highly organised parallelish lines – what the authors interpret as the impressions of muscle fibres. The fibres run in different directions according to their position in the body; for example, they seem to follow the long axes of the arms.

(Below: the type specimen of Haootia with some of the fibres visible, and various interpretive drawings of the same fossil. Liu et al. is a free paper, so anyone can go and look at the other pictures, which include close-ups of the fibres and an artistic reconstruction of the living animal.)

If the lines do indeed come from muscle fibres, then regardless of its precise affinities, Haootia is certainly an animal, and it is probably at least related to the group called eumetazoans, which includes cnidarians like jellyfish and bilaterians like ourselves (and maybe comb jellies, but let’s not open that can of jellies just now). Non-eumetazoans – sponges and Trichoplax – do not have muscles, and unless comb jellies really are what some people think they are, we can be almost certain that the earliest animals didn’t either.

Finding Ediacaran muscles is also interesting because it gives us further evidence that things capable of the kinds of movement attributed to some Ediacaran fossils really existed back then. Of course, it would have been nicer to find evidence of muscle and evidence of movement in the same fossils, but hey, this is the Precambrian. You take what you get.

(P.S.: Alex Liu is cool and I heart him. OK, I saw him give one short talk, interviewing for a job at my department that he didn’t get *sniffles*, so maybe I shouldn’t be pronouncing such fangirlish judgements, but that talk was awesome. As I’ve said before, my fangirlish affections are not very hard to win 🙂 )

***

Reference:

Liu AG et al. (2014) Haootia quadriformis n. gen., n. sp., interpreted as a muscular cnidarian impression from the Late Ediacaran period (approx. 560 Ma). Proceedings of the Royal Society B 281:20141202

Ctenophore nervous systems redux

… and reasons I suddenly find myself liking Joseph Ryan.

Ryan was the first author on the first ctenophore genome paper, published last December, though I’d known his name long before that thanks to his developmental genetic work on jelly creatures of various kinds. As is clear from the genome study, he leans quite strongly towards the controversial idea that ctenophores represent the sister lineage to all other animals.

And here’s reason one that my eyes suddenly have little cartoon hearts pulsing in their irises upon reading his short perspective paper in Zoology (Ryan, 2014). Throughout the paper, not once does he refer to ctenophores as “the” basal animal lineage. Instead, he uses phrases like “most distant relative to all other animals” or “the sister group to the rest of the animals”.

In other words, he’s scrupulously avoiding my giantest pet peeve, and I’m sure he doesn’t do it to please an obscure blogger, but gods, that’s even better. I don’t want to be pleased, I want evolutionary biology to get rid of stupid anthropocentric ladder-thinking nonsense.

Anyway, the little paper isn’t actually about animal phylogeny, it’s about nervous systems.

Both ctenophore genome papers argued that the ancestors of these pretty beasties might have evolved nervous systems independently of ours. The second one seemed positively convinced of this, but, as Ryan’s review points out, there are other possibilities even assuming that the placement of ctenophores outside the rest of the animals is correct.

While it’s possible that nerve cells and nervous systems evolved twice among the animals – it is equally possible that they have been lost twice (i.e. in sponges and blobby little placozoans). Full-fledged nerve cells wouldn’t be the first things that sponges and blobs have lost.

And Ryan basically wrote this short piece just to point that out. The argument that ctenophore nervous systems are their own invention is based on the absence or strange behaviour of many “conserved” nervous system-related genes. Ctenophores appear to completely lack some common neurotransmitters such as dopamine, as well as a lot of genes/proteins that are necessary for nerve synapses to work in us. Other genes that are “neural” in other animals are present but not associated with the nervous system in ctenophores.

BUT, Ryan cautions, there are also commonalities that shouldn’t be dismissed. While ctenophores can’t make dopamine, they do possess several other messenger molecules common in animal nervous systems. Same goes for the proteins involved in making synapses. Likewise, while they completely lack some of the genes responsible for defining various types of nerve cells (see: Hox genes), other genes involved in the same kind of stuff are definitely there.

The key thing, he says, is to take a closer look at more of these genes and find out what they do by manipulating them. Since there are clearly both similarities and differences, we must assess their extent.

And that, my friends, is the question at the heart of every homology argument ever. How similar is similar enough? Greater minds than mine have struggled with the answer, and I imagine they’ll continue to struggle until we invent time machines or find fossils of every single stage in the evolution of everything.

Until then, I’ll leave you with the closing lines of Ryan’s paper. I may not agree that we’ve “revealed” the position of ctenophores, but I’m absolutely on board with the excitement 🙂

One thing is quite clear: something remarkable happened regarding the evolution of the nervous system very early in animal evolution. Either a nervous system existed in the ancestor and was lost in certain lineages, or ctenophores invented their own nervous system independently (Fig. 1). Either possibility is quite extraordinary. The revelation that ctenophores are the sister group to the rest of animals has sparked a truly exciting debate regarding the evolutionary origins of the nervous system, one that will continue as additional genomic and functional data come to the fore.

Reference:

Ryan JF (2014) Did the ctenophore nervous system evolve independently? Zoology in press, available online 11/06/2014, doi: 10.1016/j.zool.2014.06.001

About X-frogs and failing at regeneration

Not the usual mad squee, but here’s a neat little system for studying regeneration that I quite liked today. I normally think about regeneration in terms of amputated limbs, mutilated hearts, decapitated flatworms. But you can induce a kind of “regeneration” in a less drastic and rather more tricksy way, at least in some animals. In newts and salamanders, you can create a small, superficial wound on the side of a limb, then manipulate a nearby nerve into it and add some skin from the other side of the limb.

The poor hurt limb then decides you’ve actually cut something off and tells the wound to grow a new limb. If you don’t add skin, regeneration begins but doesn’t progress very far; if you don’t add a nerve, nothing happens. IIRC you can also make extra heads in some worms in a similar way, but I digress. The figure below from Endo et al. (2004) illustrates just how well the procedure can work. The top row shows stages in the development of the extra limb, while D shows the stained skeletons of the original and new limbs. I’d say that’s a pretty good looking forearm and hand!

Endo_etal2004-ectopicLimb

 

That this trick works is in itself a very interesting insight into the nature of regeneration, as it helps us figure out exactly what it is that triggers various steps of regeneration as opposed to a simple healing process (Endo et al., 2004).

Clawed frogs (Xenopus) have been staples of embryology for a long time, but they are also quite fascinating from a regeneration point of view. One, they can regrow their limbs while they are tadpoles, but mostly lose the ability as they mature. They also have a really weird thing going on with their tadpole tails, which they can regenerate early on, then can’t, then can again (Slack et al., 2004). Huh? O.o

Two, their adult limb regeneration ability is not totally absent: it’s somewhere between salamanders’ (oh, whatever, fine, I can do that!) and ours (uh… as long as I’m a baby and it’s just a fingertip?). In a frog, an amputated arm or leg doesn’t simply heal over, but the… thing that grows out of the stump is just a simple cartilaginous spike with no joints or muscles. It’s as if the system was trying very hard to remember how to form a limb but kind of got distracted.

We are obviously interested in creating superhumans with mad regeneration skillz, which also makes us interested in how and why animals lose this seemingly very useful ability*. (Bely (2010) wrote a lovely piece on this not at all simple question.) So: Xenopus yay!

Now, Mitogawa et al. (2014) have devised a skin wound + nerve deviation system to grow little extra limb buds in adult frogs. As you might expect, it doesn’t work nearly as well as it does in axolotls: you need three nerves rather than one, and it only induces a bud about half the time, but it works well enough for research purposes.

The bud (technically, a blastema when you’re talking about regeneration) looks like a good regeneration blastema: it’s got the seemingly undifferentiated cells inside, it’s got the thickened epidermis at the tip that teams up with the nerves to give developmental instructions to the rest of the thing, and it expresses a whole bunch of genes that are turned on in normal limb blastemas.

(Totally random aside: thanks to Chrome’s spell checker, I have discovered that “blastema” is an anagram for “lambaste”.)

One area where this blastema-by-trickery fails is making cartilage, which is one of the few proper limb things the defective spike regenerates in frogs do contain. There’s no simple way of coaxing a complete spike out of these blastemas. The researchers tried the skin graft thing from axolotls (which can already form cartilage without the skin graft), but they still only got a little blastema with no cartilage. To get a skeleton, however crappy,  you need to cut out muscles and crack the underlying bone, which kind of defeats the purpose of the whole exercise IMO. At that point, you might as well just chop off the arm.

Below: the best a frog can do. Development of blastema-like bumps and “spike limbs” on the upper arm from Mitogawa et al. (2014). Compared to the fully formed accessory limbs of axolotls, the things you can see in B-D here are not terribly impressive, but they may be just the “transitional form” we need!

The failure of skin grafts alone at inducing cartilage, however, does hint at the things that go wrong with regeneration in frogs. Mitogawa et al. speculate that newt and axolotl limbs produce factors that frogs can only get from damaged bone. Broken bones even in us form a cartilaginous callus as they begin to heal, and unlike the cartilage in the extra limbs of axolotls, the cartilage in frog spikes is directly attached to the underlying bone.

They also point out that if you add proteins called BMPs to amputated mouse arms, which are otherwise even shitter at regeneration than frog arms, a surprising amount of bone formation occurs. (“BMP” stands for bone morphogenetic protein, which is a big clue to their function.) So it looks like there may be a kind of regeneration gradient from mammals (need bone damage AND extra BMP), through frogs (need bone damage, take care of BMPs themselves) to salamanders (don’t need either).

I should point out that salamanders and frogs are equally closely related to us, so this isn’t a proper evolutionary gradient, but given all the ways in which we and amphibians are fundamentally similar, our loss of regenerative ability may well have evolved through a similar stage to where frogs are now. Neat!

(I just wish they stopped calling us “higher vertebrates”. That phrase annoys me right up the fucking wall, because, and I may have said this before, EVOLUTION IS NOT A GODDAMNED LADDER. The group they are referring to has a perfectly good name that doesn’t imply ladder thinking. Amniotes, people. Or mammals, if you mean mammals, but I think if they’d meant mammals they would have said mammals. End grump.)

***

*I mean “us” in a very general sense. I think regenerative medicine is the coolest thing in medicine since vaccines and antibiotics, but I personally don’t think that the evolution of regenerative ability needs medical considerations to make it interesting. Whatever. I’m not exactly a practically minded person 😛

***

References:

Bely AE (2010) Evolutionary loss of animal regeneration: pattern and process. Integrative and Comparative Biology 50:515-527

Endo T et al. (2004) A stepwise model system for limb regeneration. Development 270:135-145

Mitogawa K et al. (2014) Ectopic blastema induction by nerve deviation and skin wounding: a new regeneration model in Xenopus laevis. Regeneration 2:11

Slack JMW et al. (2004) Cellular and molecular mechanisms of regeneration in Xenopus. Philosophical Transactions of the Royal Society B 359:745-751

The ctenophore conundrum, by popular demand

So, a new ctenophore genome has just been published in Nature (Moroz et al., 2014), it makes some extraordinary claims, and my resident palaeontologist/web-buddy Dave Bapst wants my opinion 😉

Given that I already planned to have an opinion about the first ctenophore genome back in December (Ryan et al., 2013) and miserably failed to finish the post… the temptation is just too strong. (That thesis chapter draft in the other window of MS Word wasn’t going to be finished today anyway  >_>)

Whatever I might seem from words on the internet, I’m not some kind of expert on phylogenetics, so I’m going to use a crutch. I had this idea back when I first read Ryan et al. (2013), because I remember thinking that it was written almost as if Nosenko et al. (2013) had never happened, and I’d really liked Nosenko et al. (as you can guess from the word count of this post), so I was mildly indignant about that. The Nosenko paper is going to be my crutch. (No offence to Hervé Philippe and friends, but there are only so many papers I’m going to reread for an out of the blue blog post 😉 )

Although I’m obviously not writing a public post specifically for a phylogeny nut, I may get somewhat technical, and I’m definitely going to get verbose.

***

Ctenophores. Comb jellies, sea gooseberries, Venus girdles. They are floaty, ethereal, mesmerizingly beautiful creatures, and I have it on good authority that they are also complete pains in the arse.

Here’s some pretty pictures before it gets too painful 😉 Left: Mnemiopsis leidyi from Ryan et al. (2013); right: Pleurobrachia bachei from Moroz et al. (2014). And a bonus video of a Venus girdle making like an ancient nature spirit. I could watch these beasties all day.

mnemi_pleuro

Venus from Sandrine Ruitton on Vimeo.

The problem(s)

And now, the pain. Let’s pull out my trusty old animal phylogeny, because the question marks are once again highly appropriate. (Also, I’m hell-bent on breaking your bandwidth with PICTURES.)

animalPhylogeny

Ryan et al. (2013) helpfully have a figure distilling the ideas people have had about those question marks so far:

ryan_etal2013-ctenophoreHypotheses

Bi = bilaterians, Cn = cnidarians, Ct = ctenophores, Tr = Trichoplax, and Po = sponges (Porifera).

I say “helpfully,” but it’s not all that helpful after all, since pretty much every possible configuration has been proposed. Why is this such a difficult question? Here’s a quick rundown of the problems Nosenko et al.’s study found to affect the question marks:

  1. Fast-evolving protein sequences – these can cause artefacts because too much change overwrites informative changes and creates chance similarities. Excluding faster-evolving sequences from the analysis changes the tree.
  2. Sequence data that don’t conform to the simplifying assumptions of popular evolutionary models – again, this can result in chance similarities and artefacts, and using a poorer model replicates the effects of using less ideal sequences.
  3. Long-branched outgroups – these are the non-animal groups used to place the root of animals. The more distant from animals and less well-sampled the outgroup, the longer the branches it forms, which can attract fast-evolving animal lineages towards the root. In Nosenko et al.’s analyses, even the closest outgroup seemed to cause problems, and removing the outgroup altogether made the conflicts between different models and datasets disappear completely – but this isn’t exactly helpful when you’re looking for the root of the animal tree!

The problem with ctenophores in particular is illustrated by this one of Nosenko et al.’s trees, made from one of their less error-prone datasets:

Nosenko_etal2013-ribosomalCATtree

The ctenophore branch is not only longer overall than pretty much any other in the tree; its length is also very unevenly distributed between the loooong history common to all species and the short unique lineage of each individual species. That is bad news. And it may stay that way forever, because the last common ancestor of living ctenophores may genuinely be very recent, so there’s no way to divide up that long-ass internal branch without a time machine.

Round 1: Nosenko vs. Ryan

In fairness, the Mnemiopsis genome team probably didn’t have a whole lot of time to specifically deal with Nosenko et al.’s points (OTOH, none of those individual points were truly new). The Nosenko paper came out in January 2013, and the Mnemiopsis genome paper was received by Science in July of the same year – I imagine most of the data had been generated way before then, and you can’t just redo all your data analysis and rewrite a paper on short notice.

I’m still going to view Ryan et al. (2013) in the light of Nosenko, because regardless of the genome team’s ability to answer them, some of Nosenko et al.’s points are very relevant to the claims they make. Their biggest claim, of course, being that ctenophores are the sister group to all other animals.

In Nosenko et al.’s experiments, this placement showed up in trees where faster-evolving genes, poorer models or more distant outgroups were used, but not when the slowest-evolving gene set was analysed with the best models and the closest outgroup.

Ryan et al. acknowledge that “supermatrix analyses of the publicly available data are sensitive to gene selection, taxon sampling, model selection, and other factors [cite Nosenko].” Their data are obviously sensitive to such factors. In fact, they behave rather similarly to what I saw in the Nosenko study.

Ryan et al. used two method/model combinations – one of the models was the preferred CAT model of Nosenko et al., and the other was the OK but not great GTR model that CAT beat by miles in terms of actually fitting Nosenko et al.’s data. (Caveat: in the genome paper, the CAT and GTR models were used with different treebuilding methods, so we can’t blame the models for different results with any certainty.) Also, they analysed the data with three different outgroups.

And guess what – the ctenophores-outside-everything tree was best supported with (1) the GTR model, (2) the more distant outgroups. There is not much testing of the effect of gene choice – there were two different data sets, but they were both these massive amalgamations of everything useable, and they also included totally different samples of species.

However, here comes another nod to Nosenko et al. and all the other people who advocated trying things other than “conventional” sequence comparisons through the years. Provided you can securely identify genes across different organisms, you can also try to deduce evolutionary history based on their presences and absences rather than their precise sequences. This is not a foolproof approach because genes can be (commonly) lost or (occasionally) picked up from other organisms, but it is often regarded as less artefact-prone than sequence-based trees.

But does it help with ctenophores? Like the GTR model-based sequence trees, the tree based on gene presence/absence (you obviously need complete genomes for this!) supports ctenophores being the outsider among animals:

Ryan_etal2014-RGCtree

My problem with this? Note what else it supports. The white circles indicate groupings that this method had absolutely no doubt about. And these groupings include things that frankly sound like abject nonsense. Here’s one annelid worm (the leech Helobdella) sitting next to a flatworm, while another annelid worm (Capitella) teams up with a limpet right next to a chordate. If anything, that is more controversial than the placement of ctenophores, because we thought we had it settled!

So if we’re concluding that ctenophores are basal to all other animals, why aren’t we also making a fuss about the explosion of phylum Annelida? Surely, if this method gives us strong enough conclusions to arbitrate between different sequence-based hypotheses about ctenophores, it’s strong enough to make those claims too. The cake can’t quite decide if it’s being eaten, I think.

I’m not sure what to think about the sequence trees. I’m far more confident about the presence/absence one. Maybe I’m just demonstrating the Dunning-Kruger effect here, but I’m not buying that tree for a second.

Overall verdict?

Not convinced. Not by a long shot.

Round 2: Nosenko vs. Moroz

The Pleurobrachia genome took me completely by surprise. I’d known Mnemiopsis was sequenced since Ryan et al. (2010). (Three years. Can you imagine the twitching?) I had no idea this other project was happening, so I nearly fell off my chair when Nature dropped it into my RSS reader yesterday. Another ctenophore genome – and another one that supports ctenophore separatism? (This hypothesis is becoming strangely popular…)

Bonus: it’s not just a genome paper, it also describes the transcriptomes of ten different ctenophores. Transcriptomes, the set of all active genes, are a little bit easier to sequence and assemble than genomes, and if you’re thorough they’ll catch most of the genes the organism has, so they can be almost as good for the analysis of gene content.

Which they kind of don’t do properly. There is a discussion of specific gene families that ctenophores lack – including many immune- and nervous system-related genes – but that’s not exactly saying much given that we know even “important” genes can be lost (case in point: the disappearing (Para)Hox genes of Trichoplax). The fact that ctenophores seem to completely lack microRNAs is interesting, but again, it doesn’t mean they never had them. Sponges do have microRNAs but don’t seem to be nearly as big on them as other animals.

As for the global analysis of gene content – I had to chase down a reference (Ptitsyn and Moroz, 2012) to understand what they actually did. As far as I can tell, there is no phylogenetic analysis involved – they just took a tree they already had, and used this method to map gene gains and losses onto that tree. Which is cool if you’re fairly sure about your tree, but pretty much meaningless when the tree is precisely the question. The Mammal is disappointed.

One of the problems with listing genes that aren’t there or don’t work in the “expected” way in ctenophores is that even if they’re not outside everything else, it’s still a distinct possibility that these guys branched off from our lineage before cnidarians did. For example, the Pleurobrachia paper spends a lot of time on “nervous system-specific” genes like elav missing or not being expressed in neurons, and common neurotransmitters like serotonin not being used by ctenophores.

But, assuming that the tree of animals looks something like (sponges + (ctenophores + (cnidarians + bilaterians))), we wouldn’t expect ctenophore nervous systems to share every property that cnidarians and bilaterians share. Remember: (1) sponges don’t have nervous systems, so they’re not much use as a comparison, (2) cnidarians + bilaterians had a longer common ancestry than either did with ctenophores. Genes possessed by sponges PLUS cnidarians and/or bilaterians but missing from ctenophores are more suggestive, but only if you can demonstrate that they weren’t lost. (We’re kind of going in circles here…)

The other problem is that pesky last common ctenophore ancestor. If it really is very recent, then taking even all living ctenophores to represent ctenophore diversity is like taking my close family to represent human diversity. Just like my family contains pale-skinned, lactose tolerant people, it is entirely possible that this lone surviving ctenophore lineage possesses (or lacks) important traits that aren’t at all typical of ctenophores as a whole. Ryan et al.’s supplementary data are clear that at least the Mnemiopsis genome is horribly scrambled, all trace of conserved gene neighbourhoods erased from it. That’s not exactly promising if you’re hoping for “trustworthy” animals.

The actual phylogenetic trees in Moroz et al. (2014) seem to follow an approach of throwing AAAALLL the genes at the problem. The biggest dataset contains 586 genes, compared to 122 in Nosenko et al.’s largest collection, and there is not much filtering by gene properties other than “we can tell what it is”. I have no idea how the CAT + WAG model they used compares to CAT or WAG or GTR on their own; unfortunately, the Nosenko paper doesn’t test that particular setup and this one doesn’t do any model testing. Moroz et al.’s supplementary methods claim it’s pretty good, cite something, and I’m not gonna chase down that reference. (Sorry, I’ve been poring over this for four hours at this point).

Interestingly, the support for ctenophores being apart from other animals increases when they start excluding distant outgroups. The only time it’s low is when they add all ten ctenophores and use fewer genes. Hmm. This is where I would like to hear some real experts’ opinions, because on the face of it, I can’t pinpoint anything obviously wrong. (Other than saying that chucking more genes at a problem tree is perfectly capable of making the problem worse)

TL;DR version: While I’m generally underwhelmed by the gene content stuff, I literally have no idea what to think about the trees.

I’m banking on the hope that someone will do.

***

And… I think that is all the opinion I’m going to have about ctenophores for a long time. Lunch was a long time ago, my brain is completely fried, and I’m not sure how much of the above actually makes sense. To be clear, I don’t really have a horse in this race, though I’d really like to know the truth. (Fat chance of that, by the looks of it…) I think I’m going to need a bit more convincing before I stop looking sideways at this idea that ctenophores are further from us than sponges. If anything is clear from recent phylogenomics papers, it’s that what data you analyse and how you analyse them makes a huge difference to the result you get, and this is happening with data and methods where it’s not necessarily easy to dismiss an approach as clearly inferior.

It’s a mess, damn it, and I’m not qualified to untangle it. Urgh.

***

References

Moroz LL et al. (2014) The ctenophore genome and the evolutionary origin of neural systems. Nature advance online publication, 21/05/2014; doi: 10.1038/nature13400

Nosenko T et al. (2013) Deep metazoan phylogeny: When different genes tell different stories. Molecular Phylogenetics and Evolution 67:223-233

Ptitsyn A & Moroz LL (2012) Computational workflow for analysis of gain and loss of genes in distantly related genomes. BMC Bioinformatics 13:S5

Ryan JF et al. (2010) The homeodomain complement of the ctenophore Mnemiopsis leidyi suggests that Ctenophora and Porifera diverged prior to the ParaHoxozoa. EvoDevo 1:9

Ryan JF et al. (2013) The genome of the ctenophore Mnemiopsis leidyi and its implications for cell type evolution. Science 342:1242592

OH MY GOD A FILTER-FEEDING ANOMALOCARIDID!!!

ETA: OK, technically it should be “suspension-feeding”, because there’s a good chance that its feeding mechanics involved more than simple filtering (see comments). I hate retconning, so I’ll leave the post as it is aside from this addendum. Thanks for the heads-up, Dave Bapst 🙂

This is when I put everything resembling work aside to squee madly over a fossil.

(Imagine me grinning like crazy and probably bouncing up and down a bit in my seat)

Tamisiocaris is a newly “updated” beast from the Cambrian, and the coolest thing I’ve seen since that helicoplacoid on a stalk (most cool things come from the Cambrian, right?). It is the Cambrian equivalent of a baleen whale.

Anomalocaridids were close relatives of arthropods and are among the most iconic creatures of the Cambrian. Most anomalocaridids we know of were large, swimming predators with large head appendages bearing sturdy spines to grab prey and bring it to that trilobite-crunching pineapple slice mouth. Going with the whale analogy, they were more like the killer whales of their time (although they would be easy snacks for an actual killer whale). In fact, when the putative head appendage of Tamisiocaris was originally described by Daley and Peel (2010), the only odd thing they noted about it was that it was not hardened or obviously segmented the way those of Anomalocaris were.*

Tamisiocaris was already cool back then, because it was the first animal of its kind found at Sirius Passet in Northern Greenland, one of the lesser known treasure troves of fabulous Cambrian fossils. However, since then, more appendages have been found, and it turns out that those long spines had been hiding a fascinating secret.

They were… kind of hairy.

Vinther_etal2014-sf3_crop

Closer examination of the appendages shows that their long, slender spines bore closely spaced bristles, making each spine look like a fine comb (whole appendage and close-up of a spine above from Vinther et al. [2014]). With all the spines next to each other, the bristles would have formed a fine mesh suitable for catching prey smaller than a millimetre. Compared with modern filter-feeding animals, Tamisiocaris fits right in – it would have “fished” in a similar size range as a greater flamingo. Vinther et al. (2014) suggest that Tamisiocaris would have brought its appendages to its mouth (which isn’t among the known fossils) one at a time to suck all the yummies off.

These guys are tremendously interesting for more than one reason, as the new study points out. First, HOLY SHIT FILTER FEEDING ANOMALOCARIDIDS! (Sorry. I’m kind of excited about this.) Second, the mere existence of large**  filter feeders implies a richness of plankton people hadn’t thought existed at the time. Third, there is some remarkable convergent evolution going on here.

Often, really big plankton eaters evolve from really big predators – see baleen whales, basking sharks, and these humongous fish for example. It’s not an already filter-feeding animal growing bigger and bigger, it’s an already big animal taking up filter-feeding. The interrelationships of anomalocaridids suggest the same story played out among them – ferocious hunters begetting “gentle giants” in a group with a totally different body plan from big vertebrates. For all the dazzling variety evolution can produce, sometimes, it really rhymes.

And finally, Vinther et al. did something really cool that tickles my geeky side in a most pleasant way. In their phylogenetic analysis that they did to find out where in anomalocaridid evolution this plankton-eating habit came along, they found that Tamisiocaris was closely related to another anomalocaridid with (on a second look) not dissimilar appendages. They named the group formed by the two the cetiocarids – after an imaginary filter-feeding anomalocaridid created by artist John Meszaros for the awesome All Your Yesterdays project.

Man. That’s definitely worth some squee.

***

*Disclaimer: I’m basing this on the abstract only, since palaeontological journals are one of the unfortunate holes in my university library’s otherwise extensive subscriptions.

**For Cambrian values of “large” – based on the size of the appendages, this creature would have been something like two feet long.

***

References:

Daley AC & Peel JS (2010) A possible anomalocaridid from the Cambrian Sirius Passet Lagerstätte, North Greenland. Journal of Palaeontology 84:352-355

Vinther J et al. (2014) A suspension-feeding anomalocarid from the Early Cambrian. Nature 507:496-499

Protocells YAY!

I’m briefly surfacing from the stress ocean that is paper writing to do a little dance of joy about the latest mind-blowing development in origin-of-life research.

(With my ability to go on endlessly about random scientific subjects, you’d think I’d love writing papers. No, no, no, hell no. I wish I could just upload my methods and figures to some database and be done with it. >.<)

My latest great squeal about abiogenesis research was due to an RNA enzyme that could copy long RNA strands. Well, that’s still bloody amazing, but maybe massive RNA enzymes are not how the thing we call life started. Jack Szostak’s group works witn a model of early life in which enzymes aren’t needed at all.

They’ve been working for years and years on their protocells (illustration above by Janet Iwasa via exploringorigins.org). These are basically little fat bubbles floating around in a watery solution, with a bit of nucleic acid inside. The fatty membranes of protocells are made of much simpler materials than modern cell membranes. Protocells haven’t got any proteins, and contain just a tiny “genome” that doesn’t encode anything meaningful. Yet they can, under the right circumstances, grow and divide and pass on that genome to their descendants through ordinary physical forces.

And now, they can also copy it.

The problem so far was magnesium. RNA can be replicated by an enzyme, or it can, to an extent, copy itself using base pairing. Magnesium is necessary for both kinds of replication. However, the Szostak group’s fatty protocells quickly fall apart in the presence of magnesium, spilling all their RNA content.

Adamala and Szostak (2013) tested a bunch of small molecules that bind magnesium to see if they could help. Many of them could protect the protocells, but only one, citrate, could do this without also stopping RNA replication. As a bonus, citrate prevented the degradation of RNA that, under normal circumstances, eventually happens at high magnesium levels.

Like other research toward RNA replication, this study isn’t quite there yet. For one thing, the “genomes” of these protocells are very limited – they are tiny, and they are just runs of a single RNA building block, so it’s hard to imagine how they could be precursors to more “meaningful” genomes. Also, although a lot of organic molecules just spontaneously show up when someone tries to recreate early Earth chemistry, citrate is not one of them.

Nonetheless, little by little we’re edging closer to a living RNA world. We may never know how life actually started, but the research on how it could have started looks more exciting by the day…

***

Reference:

Adamala K & Szostak JW (2013) Nonenzymatic template-directed RNA synthesis inside model protocells. Science 342:1098-1100

Fifty thousand generations, still improving

I take all my hats off to Richard Lenski and his team. If you’ve never heard of them, they are the group that has been running an evolution experiment with E. coli bacteria non-stop for the last 25 years. That’s over 50 000 generations of the little creatures; in human generations, that translates to ~1.5 million years. This experiment has to be one of the most amazing things that ever happened in evolutionary biology.

(Below: photograph of flasks containing the twelve experimental populations on 25 June 2008. The flask labelled A-3 is cloudier than the others: this is a very special population. Photo by Brian Baer and Neerja Hajela, via Wikimedia Commons.)

It doesn’t necessarily take many generations to see some mind-blowing things in evolution. An irreducibly complex new protein interaction (Meyer et al., 2012), the beginnings of new species and a simple form of multicellularity (Boraas et al., 1998) are only a few examples. However, a few generations only show tiny snapshots of the evolutionary process. Letting a population evolve for thousands of generations allows you to directly witness processes that you’d normally have to glean from the fossil record or from studies of their end products.

Fifty thousand generations, for example, can tell you that they aren’t nearly enough time to reach the limit of adaptation. The newest fruit of the Long-Term Evolution Experiment is a short paper examining the improvement in fitness the bacteria experienced over its 25 years (Wiser et al., 2013). “Fitness” is measured here as growth rate relative to the ancestral strain; the faster the bacteria are able to grow in the environment of the LTEE (which has a limited amount of glucose, E. coli‘s favourite food), the fitter they are. The LTEE follows twelve populations, all from the same ancestor, evolving in parallel, so it can also determine whether something that happens to one population is a chance occurrence or a general feature of evolution.

You can draw up a plot of fitness over time for one or more populations, and then fit mathematical models to this plot. Earlier in the experiment, the group found that a simple model in which adaptation slows down over time and eventually grinds to a halt fits the data well. However, that isn’t the only promising model. Another one predicts that adaptation only slows, never stops. Now, the experiment has been running long enough to distinguish between the two, and the second one wins hands down. Thus far, even though they’ve had plenty of time to adapt to their unchanging environment, the Lenski group’s E. coli just keep getting better at living there.

Although the simple mathematical function that describes the behaviour of these populations doesn’t really explain what’s happening behind the scenes, the team was also able to reproduce the same behaviour by building a model from known evolutionary phenomena. For example, they incorporated the idea that two bacteria with two different beneficial mutations in the same bottle are going to compete and slow down overall adaptation. (This is a problem of asexual organisms. If the creatures were, say, animals, they might have sex and spread both mutations at the same time.) So the original model doesn’t just describe the data well, it also follows from sensible theory. So did the observation that the populations which evolved higher mutation rates adapted faster.

Now, one of the first things you learn about interpreting models is that extrapolating beyond your data is dangerous. Trends can’t go on forever. In this case, you’d eventually end up with bacteria that reproduced infinitely fast, which is clearly ridiculous. However, Wiser et al. suggest that the point were their trend gets ridiculous is very, very far in the future. “The 50,000 generations studied here occurred in one scientist’s laboratory in ~21 years,” they remind us, then continue: “Now imagine that the experiment continues for 50,000 generations of scientists, each overseeing 50,000 bacterial generations, for 2.5 billion generations total.”

If the current trend continues unchanged, they estimate that the bugs at that faraway time point will be able to divide roughly every 23 minutes, compared to 55 minutes for the ancestral strain. That is still a totally realistic growth rate for a happy bacterium!

I know none of us will live to see it, but I really want to know what would happen to these little guys in 2.5 billion generations…

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References:

Boraas ME et al. (1998) Phagotrophy by a flagellate selects for colonial prey: a possible origin of multicellularity. Evolutionary Ecology 12:153-164

Meyer JR et al. (2012) Repeatability and contingency in the evolution of a key innovation in phage lambda. Science 335:428-432

Wiser MJ et al. (2013) Long-term dynamics of adaptation in asexual populations. Science, published online 14/11/2013, doi: 10.1126/science.1243357

Thumbs down, what?

Bird fingers confuse me, but the explanations confuse me more, it seems.

I didn’t mean to post today, but I’ve just read a new review/hypothesis paper about the identities of the stunted little things that pass for fingers in the wings of modern birds. The review part is fine, but I’m not sure I get the difference between the hypothesis Čapek et al. (2013) are proposing and the hypothesis they are trying to replace/improve.

To recap: the basic problem with bird fingers is that fossil, genetic and developmental evidence seem to say different things about them.

1. Fossils: birds pretty clearly come from dinosaurs, and the early dinosaurs we have fossils of have five fingers on their hands with the last two being reduced. Somewhat closer to birds, you get four fingers with #4 vestigial. And the most bird-like theropods have only three fingers, which look most like digits 1, 2 and 3 of your ordinary archosaur. (Although Limusaurus messes with this scheme a bit.)

2. Embryology: in developing limb buds, digits start out as little condensations of tissue, which develop into bits of cartilage and then finger bones. Wing buds develop a short-lived condensation in front of the first digit that actually forms, and another one behind the last “surviving” digit. Taking this at face value, then, the fingers are equivalent to digits 2, 3 and 4.

3. Genetics: In five-fingered limbs, each digit has a characteristic identity in terms of the genes expressed during its formation. The first finger of birds is most like an ordinary thumb, both when you focus on individual genes like members of the HoxD cluster and when you take the entire transcriptome. However, the other two digits have ambiguous transcriptomic identities. That is, bird wings have digit 1 and two weirdos.

Add to this the fact that in other cases of digit loss, number one is normally the first to go and number four stubbornly sticks around to the end, and you can see the headache birds have caused.

So those are the basic facts. The “old” hypothesis that causes the first part of my confusion is called the frame shift hypothesis, which suggests that the ancestors of birds did indeed lose digit 1, as in the digit that came from condensation 1 – but the next three digits adopted the identities of 1-2-3 rather than 2-3-4. (This idea, IMO, can easily leave room for mixed identities – just make it a partial frame shift.)

Čapek et al.’s new one, which they call the thumbs down hypothesis, is supposedly different from this. This is how the paper states the difference:

The FSH postulates an evolutionary event in which a dissociation occurs between the developmental formation of repeated elements (digits) and their subsequent individualization.

versus

According to the TDH no change of identity of a homeotic nature occurs, but only the phenotypic realization of the developmental process is altered due to redirected growth induced by altered tissue topology. Digit identity stays the same. Also the TDH assumes that the patterning of the limb bud, by which the digit primordia are laid down, and their developmental realization, are different developmental modules in the first place.

(Before this, they spent quite a lot of words explaining how the loss of the original thumb could trigger developmental changes that make digit 2 more thumb-like.)

I…. struggle to see the difference. If you’ve (1) moved a structure to a different position, (2) subjected it to the influence of different genes, (3) and turned its morphology into that of another structure, how exactly is that not a change in identity?

Maybe you could say that “an evolutionary event” dissociating digit formation and identity is different from formation and identity being kind of independent from the start, but I checked Wagner and Gauthier’s (1999) original frame shift paper, and I think what they propose is closer to the second idea than the first:

Building on Tabin’s (43) insight, we suggest causal independence between the morphogenetic processes that create successive condensations in the limb bud and the ensuing developmental individualization of those repeated elements as they become the functional fingers in the mature hand, thus permitting an opportunity for some degree of independent evolutionary change.

Am I missing something? I feel a little bit stupid now.

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References:

Čapek D et al. (2013) Thumbs down: a molecular-morphogenetic approach to avian digit homology. Journal of Experimental Zoology B, published online 29/10/2013, doi: 10.1002/jez.b.22545

Wagner GP and Gauthier JA (1999) 1,2,3 = 2,3,4: A solution to the problem of the homology of the digits in the avian hand. PNAS 96:5111-5116