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

The use of a larva?

Hi! Long time no see!

(I think we’ve reached the point where it’s weird to say happy new year. I could swear xkcd had a pertinent chart of funny, but I couldn’t find it.)

Once upon a time, I briefly mentioned the problematic relationships of hemichordates. Since a short paper bearing on the subject came out relatively recently (i.e. in December, yes, I’m far behind the times ;)), I thought I’d revisit it.

To begin, let’s orient ourselves on my trusty old animal phylogeny.

animalPhylogeny

Hemichordates are a phylum of deuterostomes, and their closest relatives appear to be echinoderms like starfish. The inside of Deuterostomia looks something like this:

deuterostomes

Hemichordates come in two flavours: the butt-ugly (but nevertheless intriguing) acorn worm, which even the artistic eye of 19th century zoologists couldn’t make appealing (a selection of them from Johann Wilhelm Spengel’s work below):

… and the slightly nicer-looking pterobranch. Well. They’re kind of fluffy. That counts as “nicer,” right? (A couple of Cephalodiscus from the Halanych lab below):

Acorn worms and pterobranchs have different bodies adapted to very different lifestyles. Pterobranchs are stalked, tentacled filter-feeders that often clone themselves into colonies that live together in a branching tube system. Acorn worms are solitary burrowers without tentacles, tubes or shells. Hemichordates possess features in common with vertebrates, such as gill slits, and they seem a lot less freakish than their sister phylum Echinodermata. So hemichordates are kind of the natural go-to group to look for properties of the deuterostome common ancestor.

The only problem is, to do that, you need a solid understanding of hemichordate phylogeny itself. Because there are two very different kinds of hemichordates, you have to first figure out which of those best represents their common ancestor: the sit-at-home plankton sifter or the roaming mud-eating worm. (Maybe neither. Wouldn’t that be funny.) And, as it happens, there’s some disagreement about that.

One view, espoused by the mighty zoological tome of Brusca and Brusca (2002) among others, puts acorn worms and pterobranchs as separate sister groups, and considers pterobranchs the more conservative of the two. The Bruscas write, on page 869, that “the enteropneusts [= acorn worms] have lost [their tentacles], no doubt in connection with their development of an infaunal lifestyle.” In this view, the deuterostome ancestor was a sessile filter feeder, and the long worm-like body and general moving-aboutiness of other deuterostomes is a new feature.

The other hypothesis, backed by DNA sequence data (Cannon et al., 2009)* and more recently the discovery of a tube-dwelling acorn worm from the Cambrian (Caron et al., 2013), is that pterobranchs are a weird subgroup of acorn worms and therefore unlikely to say much about our own distant ancestors.

One thing that AFAIK both camps agree on is that the ancestral acorn worm had a larva that looked nothing like an acorn worm. That’s something pretty common for marine invertebrates. Creatures as different as sea urchins and ragworms explore the seas by way of tiny, planktonic larvae that later metamorphose into a completely different animal**. (Tornaria larva of an unidentified hemichordate below by Alvaro E Migotto from the Cifonauta image database.)

However, the specific family of acorn worms that pterobranchs supposedly come from does not have such a larval stage. They develop more or less directly from fertilised eggs into mini-acorn worms.

Pterobranchs are poorly studied, so not much is known about their babies. Are they like the conventional acorn worm larva, with its distinctive body plan and curly rows of cilia? Or are they more straightforward precursors of the adult, like their presumed closest cousins? Stach (2013) describes a larva of the pterobranch Cephalodiscus gracilis that looks more like the latter. He found the minuscule creature crawling around in a colony of adult Cephalodiscus, and used thin sections and transmission electron microscopy to make a 3D reconstruction of it.

(His account of finding the baby makes me wonder how the hell he knew it did belong to Cephalodiscus. If my experience with tube-dwelling marine invertebrates is anything to go by, being found in a certain animal’s home is no guarantee that you’re related to said animal. I suppose, incomplete though they may be, older descriptions of pterobranch babies were good enough to identify the little guy?)

The image that emerges is of a rather featureless little sausage. According to Stach, it has a through gut, one full-fledged and one partially formed gill opening (asymmetry like that is not unheard of in deuterostome embryos/larvae), as well as a body cavity and a bunch of muscle cells. What it doesn’t have is any trace of the bands of cilia that “typical” acorn worm larvae use to swim and feed, nor some other structures (e.g. nerve centres) that characterise such larvae.

Taken at face value, this would suggest (assuming this is a typical pterobranch larva) that the pterobranchs-are-acorn worms people are right. I have my reservations, and not just because a sample size of one makes me statistically nervous. Using this description as evidence for evolutionary relationships assumes that traditional larvae with ciliary bands are hard to lose. But that’s quite possibly not the case.

Echinoderm larvae, for example, have changed a lot even in the last few million years. The changes occurred many times independently, and often involved a return from a full-fledged larval stage to more direct development (Raff and Byrne, 2006). I don’t know whether acorn worms display a similar sort of flexibility. How many have even been studied in terms of development?

So: detailed internal structure of a pterobranch larva? Cool. As to the worms first hypothesis… “consistent with” would be a better description than “supports”, I think.

***

Notes:

*Although microRNAs beg to differ (Peterson et al., 2013).

**The history of these larvae is a mighty can of worms, or trochophores and tornariae as the case may be. I shall say no more on the matter here. 🙂

***

References:

Brusca RC & Brusca GJ (2002) Invertebrates (second edition). Sinauer Associates.

Cannon JT et al. (2009) Molecular phylogeny of hemichordata, with updated status of deep-sea enteropneusts. Molecular Phylogenetics and Evolution 52:17-24

Caron J-B et al. (2013) Tubicolous enteropneusts from the Cambrian period. Nature 495:503-506

Peterson KJ et al. (2013) MicroRNAs support the monophyly of enteropneust hemichordates. Journal of Experimental Zoology B 320:368-374

Raff RA & Byrne M (2006) The active evolutionary lives of echinoderm larvae. Heredity 97:244-252

Stach T (2013) Larval anatomy of the pterobranch Cephalodiscus gracilis supports secondarily derived sessility concordant with molecular phylogenies. Naturwissenschaften 100:1187-1191

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…

***

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.

***

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

New genes, new tricks, part 2

In my previous post, I marvelled over the strange and unexpected way duplicated genes behave in fruit flies. The second study I wanted to discuss is also about new fruit fly genes gaining new functions, but unlike the other one, it’s about new genes that didn’t come from pre-existing genes.

Reinhardt et al. (2013) wasn’t the best written paper I’ve read, and I had some difficulty figuring out exactly what was going on in places, but there is some interesting stuff in there nonetheless.

The authors investigated six recently evolved new ?protein-coding genes in Drosophila. They wanted to know how they came about and managed to stick. For example, did they first originate as non-coding RNA genes? Did they gain a function through their RNA copies alone before they began to encode a protein? Or did they first awaken from the no man’s land between old genes with protein-coding potential already present?

This harkens back to one of the papers about new genes that I’d previously discussed. Xie et al. (2012) found that the genes for several human-specific proteins began life (and function?) as RNA genes expressed in particular tissues in ancestral primates. What about the six fly genes the new study investigated?

Reinhardt et al.‘s illustration of the two routes to protein-coding geneness is below. Starting with an inactive stretch of DNA (black line), you need two things: (1) an “on” switch or promoter (green box), which causes the transcription of RNA (blue) from the region, and (2) a sequence that can be translated into a decent length protein (an open reading frame or ORF, pink box). These two can theoretically appear in either order.

Before we get into the meat of the paper, let’s borrow the Drosophila family tree from the 12 genomes project page:

D. melanogaster, third from the top, is the species that has been used for every variety of biological investigation for over a hundred years, and also the focus of this study. However, the other species were also used for comparison, to see exactly where and how the genes originated.

Five of the six genes had a relatively long history, with similar sequences being found in D. yakuba and erecta or even further out in D. ananassae. Three of them were not only there in those species, but could also potentially make a nice protein. In two genes, the sequence or part of it was recognisable all the way to ananassae, but it only had long sensible ORFs in melanogaster itself.

In terms of activity… well, first of all I think they screwed up Figure 2. Supposedly, the names of the species in which transcription of these genes was detected are bolded, but actually, all the names are bolded in all the trees, which doesn’t agree with what they say (or with the green dots signifying the origin of transcription in the same figure). Anyway, assuming the bolding was a mistake and the green dots are in the right place, it sounds like four of the six genes were already active in the common ancestor of melanogaster and yakuba or earlier, while another two were only turned on in the melanogaster/sechellia/simulans lineage.

The order of events varies from gene to gene: four genes had good solid ORFs right from the start, while two were transcribed before they were suitable protein templates. The authors note that we can’t actually be sure whether or not the first four developed an ORF before they became active. To be certain of that, we would need more distantly related species with a matching ORF that isn’t transcribed, but in all four cases the species lacking expression of the gene also totally lack any trace of the sequence. So, while the remaining two genes provide positive evidence for the transcription-first scenario, the jury is still out on the ORF-first option.

In D. melanogaster, the presence of the protein product was confirmed for the four genes with the oldest ORFs. The two youngest may still be translated: the protein data came only from embryos, and in fact all six genes contain short signals that are normally associated with the transport of proteins to specific parts of the cell. You might reason that a gene that never makes a protein doesn’t need such signals, but nevertheless, the authors couldn’t positively confirm the existence of these proteins without data from other life stages.

Where these genes are active brings us back to a common theme we encountered in the previous post. In adult D. melanogaster, all six are most strongly expressed in the testicles, and the products of one of them are exclusive to those organs. Likewise, male larvae show more expression of all six genes than females do. The other species show basically the same pattern.

What do these genes do? Actually, do they do anything? Being expressed, even being translated to protein, doesn’t necessarily equate to having a function. Luckily, “function” is not terribly difficult to test for in fruit flies. There are lots of clever tricks that allow you to manipulate their genes and look at the consequences. In this case, Reinhardt et al. bred flies where these genes were turned off. If I understood them correctly, they managed to do this for five genes, four of which resulted in very dead flies. Weirdly, for all four, the affected flies died at the same life stage, just before hatching from the pupa.

With a different strategy that produced only partial knock-down of the genes, they got themselves some grown-up survivors, which allowed them to test the effect of the genes on male fertility (a sensible question given where these genes are most active). Out of three knock-downs with surviving adults of both sexes, only one showed a serious effect, and that was the one that produced generally crappy, short-lived weakling males anyway, so while these genes are active in the testicles and they might disproportionately affect males, they don’t seem to have much to do with fertility per se.

In general, the results sound like new genes that come from random bits of DNA can very quickly become essential to the organism, and it also sounds very much like an overabundance of transcripts in the testicles doesn’t mean that that’s where their function lies – it’s probably more that all kinds of things are expressed in testicles, and these genes are still expressed there because that’s how they started their lives.

Something big missing from the study is actually testing when these genes became functional – we’re told when they became expressed and when they started making a protein, but without manipulating them in relevant non-melanogaster species, it’s impossible to tell whether either of those means function. *disappointed pout*

And what’s up with those four genes that were necessary for the flies’ survival? The knock-downs all did their killing at the same stage. I don’t know what to think about that, and the authors don’t really offer an explanation beyond describing control experiments to make sure the deaths weren’t an unfortunate side-effect of the manipulation itself. Is there something about the development of adults that attracts new genes? Is the process of metamorphosis especially sensitive to even minor mess-ups? (More sensitive than early embryonic development?) Intuitively, I’d find the first possibility more likely, but gods know intuition is a poor guide to reality…

***

References:

Reinhardt JA et al. (2013) De novo ORFs in Drosophila are important to organismal fitness and evolved rapidly from previously non-coding sequences. PLoS Genetics 9:e1003860

Xie C et al. (2012) Hominoid-specific de novo protein-coding genes originating from long non-coding RNAs. PLoS Genetics 8:e1002942

New genes, new tricks

I’ve previously written about the birth of new genes. Since new genes are cool, and I just found two recent papers on them, you’re getting more of them.

Part 1: how to survive duplication

Technically, the first paper isn’t about new new genes: Assis and Bachtrog (2013) examined recently duplicated genes in fruit flies. But screw technicalities, what they’re saying makes my eyes pop.

When a gene is accidentally copied, a variety of possible fates can await it. Most of the time, the extra copy just dies. Some mechanisms of gene duplication just take the gene without the regulatory elements it needs to function properly. Even if the new copy works, it’s still redundant, so there’s nothing stopping mutations from destroying it over time. However, sometimes redundancy is removed before the new gene breaks irrevocably, and both copies are kept. This can, in theory, happen in a number of ways. Because I’m feeling lazy, let me just quote them from the paper (square brackets are mine, because I hate repeatedly typing out long ugly words :)):

Four processes can result in the evolutionary preservation of duplicate genes: conservation, neofunctionalization, subfunctionalization, and specialization. Under conservation, ancestral functions are maintained in both copies, likely because increased gene dosage is beneficial (1). Under neofunctionalization [NF], one copy retains its ancestral functions, and the other acquires a novel function (1). Under subfunctionalization [SF], mutations damage different functions of each copy, such that both copies are required to preserve all ancestral gene functions (9, 10). Finally, under specialization, subfunctionalization and neofunctionalization act in concert, producing two copies that are functionally distinct from each other and from the ancestral gene (11).

We might add a variation on NF, too: Proulx and Phillips (2006) theorised that differences in function that arise in different alleles (variants) of a single gene can turn duplication into an advantage, turning the conventional duplication-first, new function-next scenario on its head.

Either way, genomes contain lots of duplicated genes, there’s no question about that. What isn’t nearly as well understood is the relative importance of various mechanisms in producing all these duplicates. It’s much easier to theorise about mechanisms than to test the theories. Since evolution doesn’t stop once a new gene has earned its place in the genome, it can be hard to disentangle the mechanism(s) responsible for its preservation from the stuff that happened to it later. Also, to really assess the relative role of different mechanisms, you’ve got to look at whole genomes.

(Assis and Bachtrog say that this hasn’t been done before, and then go right on to cite He and Zhang [2005], which is a genome-wide study of SF and NF. I guess it doesn’t look at all the mechanisms…)

Assis and Bachtrog used the amazing resource that is the 12 Drosophila genomes project, focusing on D. melanogaster and D. pseudoobscura to find slightly under 300 pairs of genes that duplicated after the divergence of those two species. Since Drosophila genomes are very well-studied, they were able to identify the “parent” and “child” in each pair based on where they sit on their chromosomes. They then also extracted thousands of unduplicated genes from the melanogaster and pseudoobscura genomes, to use as a measure of background divergence between the two species.

To measure changes in gene function, they compared the expression of parent and child genes to each other and to the “ancestral” copy (i.e. the unduplicated gene in the other species) in different parts of the body (if a gene is suddenly turned on somewhere it wasn’t before, it’s probably doing something new!).

Long story short, it turned out that in the majority of cases (167/281) cases the child copy behaved much more differently from the “ancestor” than expected, while the parent copy stayed pretty close. These child copies also showed faster sequence evolution than their parents. This means that NF – and specifically that of the new copy – is the most common fate of newly duplicated genes in these animals. There’s also a fair number of gene pairs where both copies gained new functions or both stuck with the old ones, but only three where both copies lost functions. Pure SF, which very influential studies like Force et al. (1999) championed as the dominant mode of duplicate gene survival, appears to be an incredibly rare occurrence in fruit flies!

A few paragraphs ago I mentioned the caveat that duplicated genes don’t stop evolving just because they’ve managed to survive. Well, the advantage of having all these Drosophila genomes is that you can further break down “young” duplicates into narrower age groups, using the species that fall between melanogaster and pseudoobscura on the tree. However, looking at this breakdown doesn’t change the general pattern – NF of the child copy is the most common and SF is rare or nonexistent in even the youngest age groups, along both the melanogaster and the pseudoobscura lineages.

So what exactly is going on here?

Part of the difference in expression patterns between parent/ancestral and child copies is because these new genes are turned on in the testicles, which might give us a big clue. Testicles, you see, are a bit anarchical. Things that are normally kept silent in the genome, like various kinds of parasitic DNA, wake up and run wild during the making of sperm. If you remember my throwaway reference to duplication mechanisms that cut the gene off from its old regulatory elements – well, the balls are a place where even such lost and lonely genes get a second chance.

The genomic anarchy of testes is also one of the reasons these duplications happen in the first place; the aforementioned mechanism involves those bits of parasitic DNA that copy and paste themselves via an RNA intermediate. The enzymes they use to reverse transcribe this RNA into DNA and insert it back into the genome aren’t particularly discerning, and they’ll happily do their thing on a piece of RNA that isn’t the parasite. Indeed, slightly more NFed child genes than you’d expect originated via RNA, although it’s worth noting that more than half of them still didn’t. So while the testes look like a good place for new gene copies to find a use, they aren’t totally responsible for their origins.

Why is there so little SF among these genes?

This is the Obvious Question; my jaw nearly landed on my desk when I saw the numbers. The authors have two hypotheses, both of which may be true at the same time.

First, SF assumes that the two copies have the same functions to begin with. This is not necessarily true when just a small segment of DNA is duplicated – even when it’s not just a bare gene you’re copying, the new copy might lose part of its old regulatory elements and/or land next to new ones, not to mention Proulx and Phillips’s idea of new functions appearing before duplication. So maybe SF is more common after wholesale duplications of entire genomes, and Drosophila species didn’t have any of those recently.

Secondly, SF happens by genetic drift, which is a random process that works much better in small populations. Fruit flies aren’t known for their small populations, and therefore the dominant evolutionary force acting on their genomes will be selection.

This makes sense to me, but the degree to which NF dominates the picture is still pretty amazing. I wonder what you’d get if you applied the same methods to different species. Would species with smaller populations, or those that recently duplicated their whole genomes, show more evidence for SF as you’d expect if the above reasoning is correct? Or would the data slaughter all those seemingly reasonable explanations? What would you see in parthenogenetic species that have no males (and testicles)?

Part two, with really new genes, hopefully coming soon…

***

References:

Assis R & Bachtrog D (2013) Neofunctionalization of young genes in Drosophila. PNAS 110:17409-17414

He X & Zhang J (2005) Rapid subfunctionalization accompanied by prolonged and substantial neofunctionalization in duplicate gene evolution. Genetics 169:1157-1164

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

Proulx SR & Phillips PC (2006) Allelic divergence precedes and promotes gene duplication. Evolution 60:881-892

Lamprey Hox clusters and genome duplications, oh my!

What the hell is up with lamprey Hox clusters?

Lampreys are among the few living jawless vertebrates, creatures that parted evolutionary ways with our ancestors somewhere on the order of 500 million years ago. If you want to know where things like jaws, paired fins or our badass adaptive immune systems came from, a vertebrate that doesn’t possess some of these things and may have diverged from the rest of the vertebrates soon after others originated is just what you need for comparison.

The vertebrate fossil record is pretty rich thanks to us having hard tissues, so a lot can be inferred about these things from the wealth of extinct fishes we have at our disposal. However, there are times when comparisons of living creatures are just as useful, if not more, than examinations of fossils. (Fossils, for example, tend not to have immune systems. ;))

One of the things you absolutely need a living animal to study is, of course, genome evolution. Vertebrates – well, at least jawed vertebrates – are now generally accepted to have the remnants of four genomes. Our long-gone ancestors underwent two rounds of whole genome duplication. Afterwards, most of the extra genes were lost, but evidence for the duplications can still be found in the structure of our genomes, where entire recognisable gene neighbourhoods of our close invertebrate relatives often still exist in up to four copies (Putnam et al., 2008).

Among these neighbourhoods are the four clusters of Hox genes most groups of jawed vertebrates possess. A “normal” animal like a snail or a centipede only has one of these. Since Hox genes are involved in the making of body plans, you have to wonder how suddenly having four sets of them and other developmental “master genes” might have influenced the evolution of vertebrate bodies.

Of course, to guess that, you need to know precisely when these duplications happened. That’s where lampreys come in: their lineage branched off from our definitely quadruple-genomed one after the next closest, definitely single-genomed group. But was it before, between, or after, the two rounds of duplication?

A few years ago, a phylogenetic analysis of 55 gene families by Kuraku et al. (2009) suggested that the lamprey-jawed vertebrate split happened after the 2R. Just this year, the genome of the sea lamprey Petromyzon marinus was finally published (Smith et al., 2013), and its authors agreed that yes, lampreys probably split off from us post-2R. (I don’t entirely get all the things they did to arrive at this conclusion. Groups of linked genes show up again, among other approaches.)

However, that isn’t the whole story, the latest lamprey genomics paper argues (Mehta et al., 2013). The P. marinus genome assembly couldn’t stitch all the Hox clusters properly together. There were two that sat on nice big scaffolds with the whole row of Hox genes and a few of their neighbours, and then there were a bunch of “loose” Hox genes that they couldn’t link to anything (diagram comparing humans and P. marinus below from Smith et al., 2013; the really pale blue boxes under the numbers represent Hox genes):

Smith_etal2013-F4

Given that Hox9 genes exist in four copies in this species, it seems like there may be four clusters. However, in hagfish, the other kind of living jawless vertebrate, a study found Hox genes that seemed to have as many as seven copies (Stadler et al., 2004). Another round of duplication? It wouldn’t be unheard of. Most teleosts, which include most of the things we call “fish” in everyday parlance, have seven Hox clusters courtesy of an extra genome duplication and loss of one cluster*. Salmon and kin have thirteen, after yet another duplication. Maybe hagfish also had another one – but did lampreys? How many more clusters do those lonely Hox genes belong to?

Mehta et al. hunted down the Hox clusters of Japanese lampreys (Lethenteron japonicum), hoping to pin down exactly how many there were. They used large chunks of DNA derived partly from the testicles, where sperm cells and their precursors keep the full genome throughout the animal’s life (lampreys throw away large chunks of the genome in most non-reproductive cells [Smith et al., 2009]). They probed these for Hox genes and sequenced the ones that tested positive. Plus they also got about two-thirds of the full genome together in fairly big pieces. Together, these data allowed them to get a better idea of the mess that is lamprey Hox cluster genomics.

They assembled four whole clusters, including their neighbouring genes, and a partial fifth cluster. A bunch of other genes sat on smaller sequence fragments containing only a couple of Hoxes, or a Hox and a non-Hox, but they were tentatively assigned to a total of eight clusters, eight being the number of different Hox4 genes in the data (no known vertebrate Hox cluster contains more than one Hox4 gene). The L. japonicum equivalents of the 31 publicly available Hox sequences from P. marinus spread out over six of these, which indicates that both species have at least six clusters. Seems like lampreys had another round of genome duplication after 2R? (Summary of L. japonicum Hox clusters from Mehta et al. below.)

But wait, that’s not the end of it.

First of all, although there are undoubtedly four complete Hox clusters in there L. japonicum, the relationships of these clusters to our four are terribly confused. Whether you look at the phylogenetic trees of individual genes, or the arrangement of non-Hox genes on either side of the cluster, only a big pile of what the fuck emerges. Phylogenies are problematic because the unusual composition of lamprey genes and proteins (Smith et al., 2013) could easily throw them off. All the complete lamprey clusters have a patchwork of neighbours that look like a mashup of more than one of our Hox clusters. Might it mean that lampreys’ proliferation of Hox clusters occurred independently of ours? Did we split before 2R after all?

Hox genes are not the only interesting things in a Hox cluster. In the long gaps between them, there are all sorts of little DNA switches that regulate their behaviour. Some of these are conserved across the jawed vertebrates. Mehta et al. aligned complete Hox clusters of humans, elephant sharks and lampreys to look for such sequences – called conserved non-coding elements or CNEs – in the lamprey.

They only found a few, but that’s enough for a bit more head-scratching. Most CNEs in, say, the human HoxA cluster are only found in one elephant shark cluster, and vice versa. Humans have a HoxA cluster, elephant sharks have a HoxA cluster, they’re clearly the same thing, pretty straightforward. Not so for lampreys. Homologues of individual CNEs in the complete lamprey clusters are spread out over all four human/elephant shark clusters. More evidence for independent duplications?

Mehta et al. are cautious – they point out that the silly mix of Hox cluster neighbours in lampreys could just be due to independent post-2R losses, which is plausible if the split between lamprey and jawed vertebrate lineages happened not too long after 2R. There’s also the fact that the weird lamprey sequences are phylogenetic minefields – however, that’s a double-edged sword, since the same caveat applies to analyses that support a post-2R divergence. Then, perhaps the same argument that goes for Hox cluster neighbours could also apply to CNEs. And, of course, this is just Hox clusters. Smith et al.‘s (2013) findings about overall genome structure don’t go away just because lamprey Hox clusters are weird.

So, in summary, thanks, lampreys. Fat lot of help you are! 😛

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*Actually, two losses of two separate clusters in two different teleost lineages. Because Hox evolution wasn’t already complicated enough.

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References

Kuraku S et al. (2009) Timing of genome duplications relative to the origin of the vertebrates: did cyclostomes diverge before or after? Molecular Biology and Evolution 26:47-59

Mehta TK et al. (2013) Evidence for at least six Hox clusters in the Japanese lamprey (Lethenteron japonicum). PNAS 110:16044-16049

Putnam NH et al. (2008) The amphioxus genome and the evolution of the chordate karyotype. Nature 453:1064-1071

Smith JJ et al. (2009) Programmed loss of millions of base pairs from a vertebrate genome. PNAS 106:11212-11217

Smith JJ et al. (2013) Sequencing of the sea lamprey (Petromyzon marinus) genome provides insights into vertebrate evolution. Nature Genetics 45:415-421

Stadler PF et al. (2004) Evidence for independent Hox gene duplications in the hagfish lineage: a PCR-based gene inventory of Eptatretus stoutii. Molecular Phylogenetics and Evolution 32:686-694