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Archive for November, 2017

As 2017 approaches its end, there have been a few papers I’ve been involved in that I thought I’d point out here while I have time. Our DAWNDINOS project has been taking up much of that time and you’ll see much more of that project’s work in 2018, but we just published our first paper from it! And since the other two recent papers involve a similar theme of muscles, appendages and computer models of biomechanics, they’ll feature here too.

Stomach-Churning Rating: 0/10; computer models and other abstractions.

Mussaurus patagonicus was an early sauropodomorph dinosaur from Argentina, and is now widely accepted to be a very close relative of the true (giant, quadrupedal) sauropods. Here is John Conway’s great reconstruction of it:

We have been working with Alejandro Otero and Diego Pol on Mussaurus for many years now, starting with Royal Society International Exchange funds and now supported by my ERC grant “DAWNDINOS”. It features in our grant because it is a decent example of a large sauropodomorph that was probably still bipedal and lived near the Triassic-Jurassic transition (~215mya).

In our new study, we applied one of my team’s typical methods, 3D musculoskeletal modelling, to an adult Mussaurus’s forelimbs. This is a change of topic from the hindlimbs that I’ve myopically focused on before with Tyrannosaurus and Velociraptor [in an obscure paper that I should never have published in a book! pdf link], among other critters my team has tackled (mouse, elephant [still to be finished…], ostrich, horse, Ichthyostega… dozens more to come!). But we also modelled the forelimbs of Crocodylus johnstoni (Australian “freshie”) for a key comparison with a living animal whose anatomy we actually knew, rather than reconstructed.

Mussaurus above; Crocodylus below; forelimb models in various views; muscles are red lines.

The methods for this biomechanical modelling are now standard (I learned them from their creator Prof. Scott Delp during my 2001-2003 postdoc at Stanford): scan bones, connect them with joints, add muscle paths around them, and then use the models to estimate joint ranges of motion and muscle moment arms (leverage) around joints. I have some mixed feelings about developing this approach in our 2005 paper that is now widely used by the few teams that study appendicular function in extinct animals. As a recent review paper noted and I’ve always cautioned, it has a lot of assumptions and problems and one must exercise extreme caution in its design and interpretation. Our new Mussaurus paper continues those ruminations, but I think we made some progress, too.

On to the nuts and bolts of the science (it’s a 60 page paper so this summary will omit a lot!): first, we wanted to know how the forelimb joint ranges of motion in Mussaurus compared with those in Crocodylus and whether our model of Mussaurus might be able to be placed in a quadrupedal pose, with the palms at least somewhat flat (“pronated”) on the ground. Even considering missing joint cartilage, this didn’t seem very plausible in Mussaurus unless one allowed the whole forearm to rotate around its long axis from the elbow joint, which is very speculative—but not impossible in Crocodylus, either. Furthermore, the model didn’t seem to have forelimbs fully adapted yet for a more graviportal, columnar posture. Here’s what the model’s mobility was like:

So Mussaurus, like other early sauropodomorphs such as Plateosaurus, probably wasn’t quadrupedal, and thus quadrupedalism must have evolved very close to in the Sauropoda common ancestor.

Second, we compared the muscle moment arms (individual 3D “muscle actions” for short) in different poses for all of the main forelimb muscles that extend (in various ways and extents) from the pectoral girdle to the thumb, for both animals, to see how muscle actions might differ in Crocodylus (which would be closer to the ancestral state) and Mussaurus. Did muscles transform their actions in relation to bipedalism (or reversal to quadrupedalism) in the latter? Well, it’s complicated but there are a lot of similarities and differences in how the muscles might have functioned; probably reflecting evolutionary ancestry and specialization. What I found most surprising about our results was that the forelimbs didn’t have muscles well-positioned to pronate the forearm/hand, and thus musculoskeletal modelling of those muscles reinforced the conclusions from the joints that quadrupedal locomotion was unlikely. I think that result is fairly robust to the uncertainties, but we’ll see in future work.

You like moment arms? We got moment arms! 15 figures of them, like this! And tables and explanatory text and comparisons with human data and, well, lots!

If you’re really a myology geek, you might find our other conclusions about individual muscle actions to be interesting—e.g. the scapulohumeralis seems to have been a shoulder pronator in Crocodylus vs. supinator in Mussaurus, owing to differences in humeral shape (specialization present in Mussaurus; which maybe originated in early dinosaurs?). Contrastingly, the deltoid muscles acted in the same basic way in both species; presumed to reflect evolutionary conservation. And muuuuuuch more!

Do you want to know more? You can play with our models (it takes some work in OpenSim free software but it’s do-able) by downloading them (Crocodylus; Mussaurus; also available: Tyrannosaurus, Velociraptor!). And there will be MUCH more about Mussaurus coming soon. What is awesome about this dinosaur is that we have essentially complete skeletons from tiny hatchlings (the “mouse lizard” etymology) to ~1 year old juveniles to >1000kg adults. So we can do more than arm-wave about forelimbs!

But that’s not all. Last week we published our third paper on mouse hindlimb biomechanics, using musculoskeletal modelling as well. This one was a collaboration that arose from past PhD student James Charles’s thesis: his model has been in much demand from mouse researchers, and in this case we were invited by University of Virginia biomechanical engineers to join them in using this model to test how muscle fibres (the truly muscle-y, contractile parts of “muscle-tendon units”) change length in walking mice vs. humans. It was a pleasure to re-unite in coauthorship with Prof. Silvia Blemker, who was a coauthor on that 2005 T. rex hindlimb modelling paper which set me on my current dark path.

Mouse and human legs in right side view, going through walking cycles in simulations. Too small? Click to embiggen.

We found that, because mice move their hindlimb joints through smaller arcs than humans do during walking and because human muscles have large moment arms, the hindlimb muscles of humans change length more—mouse muscles change length only about 48% of the amount that typical leg muscles do in humans! This is cool not only from an evolutionary (mouse muscles are probably closer to the ancestral mammalian state) and scaling (smaller animals may use less muscle excursions, to a point, in comparable gaits?) perspective, but it also has clinical relevance.

Simulated stride for mouse and human; with muscles either almost inactive (Act=0.05) or fully active (Act=1). Red curve goes through much bigger excursions (along y-axis) than blue curve), so humans should use bigger % of their muscle fibre lengths in walking. Too small? Click to embiggen.

My coauthors study muscular dystrophy and similar diseases that can involve muscle stiffness and similar biomechanical or neural control problems. Mice are often used as “models” (both in the sense of analogues/study systems for animal trials in developing treatments, and in the sense of computational abstractions) for human diseases. But because mouse muscles don’t work the same as human muscles, especially in regards to length changes in walking, there are concerns that overreliance on mice as human models might cause erroneous conclusions about what treatments work best to reduce muscle stiffness (or response to muscle stretching that causes progressive damage), for example. Thus either mouse model studies need some rethinking sometimes, or other models such as canines might be more effective. Regardless, it was exciting to be involved in a study that seems to deliver the goods on translating basic science to clinical relevance.

Muscle-by-muscle data; most mouse muscles go through smaller excursions; a few go through greater; some are the same as humans’.

Finally, a third recent paper of ours was led by Julia Molnar and Stephanie Pierce (of prior RVC “Team Tetrapod” affiliation), with myself and Rui Diogo. This study tied together a bunch of disparate research strands of our different teams, including musculature and its homologies, the early tetrapod fossil record, muscle reconstruction in fossils, and biomechanics. And again the focus was on forelimbs, or front-appendages anyway; but turning back the clock to the very early history of fishes, especially lobe-finned forms, and trying to piece together how the few pectoral fin muscles of those fish evolved into the many forelimb muscles of true tetrapods from >400mya to much more recent times.

Humerus in ventral view, showing muscle attachments. Extent (green) is unknown in the fossil but the muscle position is clear (arrow).

We considered the homologies for those muscles in extant forms, hypothesized by Diogo, Molnar et al., in light of the fossil record that reveals where those muscles attach(ed), using that reciprocal illumination to reconstruct how forelimb musculature evolved. This parallels almost-as-ancient (well, year 2000) work that I’d done in my PhD on reconstructing hindlimb muscle evolution in early reptiles/archosaurs/dinosaurs/birds. Along the way, we could reconstruct estimates of pectoral muscles in various representative extinct tetrapod(omorph)s.

Disparity of skeletal pectoral appendages to work with from lobe-fins to tetrapods.

Again, it’s a lengthy, detailed study (31 pages) but designed as a review and meta-analysis that introduces readers to the data and ideas and then builds on them in new ways. I feel that this was a synthesis that was badly needed to tie together disparate observations and speculations on what the many, many obvious bumps, squiggles, crests and tuberosities on fossil tetrapods/cousins “mean” in terms of soft tissues. The figures here tell the basic story; Julia, as usual, rocked it with some lovely scientific illustration! Short message: the large number of pectoral limb muscles in living tetrapods probably didn’t evolve until limbs with digits evolved, but that number might go back to the common ancestor of all tetrapods, rather than more recently. BUT there are strong hints that earlier tetrapodomorph “fishapods” had some of those novel muscles already, so it was a more stepwise/gradual pattern of evolution than a simple punctuated event or two.

Colour maps of reconstructed right fin/limb muscles in tetrapodomorph sarcopterygian (~”fishapod”) and tetrapod most recent common ancestors. Some are less ambiguous than others.

That study opens the way to do proper biomechanical studies (like the Mussaurus study) of muscle actions, functions… even locomotor dynamics (like the mouse study)– and ooh, I’ve now tied all three studies together, tidily wrapped up with a scientific bow! There you have it. I’m looking forward to sharing more new science in 2018. We have some big, big plans!

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This week we conducted wallaby leg dissections for a study of the kneecaps of marsupials (pouched mammals). Placental (non-pouched) mammals like us almost all have bony kneecaps but many marsupials do not. Kneecaps do important things, acting like gears around the knee joints (e.g. this old post), and yet it is unclear why some marsupials have lost, kept or even re-evolved them as bones. So we’re investigating that and already noticed that one of our wallabies has bony kneecap(s) whereas the other doesn’t, so we’re checking out why and taking tissue samples to do histology (sectioning for microscopic imaging of tissue composition and structure) on so we can see what the knee tendon/kneecap tissues are made of. Some marsupials turn their kneecaps into fibrocartilage rather than bone or tendon and that can be impossible to identify without histology.

The wallabies are small, about 20lbs or so and just take a day or so. Like a turkey. And it’s Thanksgiving today, so here I am with a post about thawing specimens for science, rather than for food. Maybe the title will make sense now.

Stomach-Churning Rating: 7/10; thawed wallaby bits from the get-go.

Thawed lower leg and foot of wallaby. The stickers are for an old study that would take too long to explain…

This post was directly inspired by journalist Jason Bittel’s inquiry to me about my tweet on the wallaby thawing; he wondered if there might be a fun story linking thawing-for-science with thawing-for-Thanksgiving. Some highfalutin editors didn’t agree, so no printed/online story came of this, but I am not so highfalutin, hence this blog post.

Thawed wallaby forelimbs. I’m also looking into the “false thumbs” that some marsupials have (“sixth fingers”), much as elephants and other mammals may have.

Thawing is second nature for our lab’s team; we do it all the time. Avid readers will be unsurprised to learn that just about everything I’ve worked on has been frozen at some time, and thus has been thawed out at some time(s). Normally we don’t freeze if we need live tissue or undistorted tissue, e.g. to measure physiology or very fine microstructure– freezing disrupts all of that. We would instead use physiological saline solution or else a preservative like formalin. And you can only freeze and then thaw a specimen for two times or so before it becomes too useless even for anatomical study.

A small specimen like this salamander can be thawed out simply by running it under warm water for a little while or leaving it out for an hour.

We just leave specimens in a cart, or on a table or sometimes in a cold-room shelving area, for slower thawing. Space heaters tend to overdo things. We don’t do any rough calculation from some sort of thermodynamic first principles of time-to-thaw vs. specimen size (I wish we were that smart!); just seat-of-pants guessing and checking (yes, poking specimens to check their thawedness is a method of choice). Cutting things in half along the way, or skinning them, may be used to accelerate the thawing process. But it’s about as unscientific a method as we use.

The hardest specimens to thaw of course have been the largest specimens. Elephant legs can be >2 metres long and hundreds of kilograms (especially when frozen). A week at room temperature tends to work OK for getting them to a dissectable state. One has to balance the outer deterioration with the inner frigidness. We’re not so concerned about microbe growth in most cases, as one would be with a thawing turkey, and not at all about consumption. We just want to be able to dissect it and make observations, mostly via eyeballing the specimens as we dissect them,

Left hindfoot of an Asian elephant. Still frozen; this was bandthawed- I mean bandsawed- to see its internal anatomy nice and clearly. You may see this specimen again somewhere else– stay tuned! 🙂

Moisture and fluids can be a challenge: generally the rooms we thaw in are low humidity so moisture may not be an issue once the ice melts away, and we have drains nearby. We try to remove ice first or have towels to wipe/soak fluids up as thawing progresses. But if a specimen is sitting in a cart or storage bag with too much ice early on, that can thaw first and then turn the specimen into a nasty slurry of the stuff you’re interested in and the less desirable muck. So we try to avoid that.

De-thawing too early is bad. The smell gets progressively worse– and once the interior of the specimen is thawed enough, then bacteria get in there and the interior becomes a brewing ground for heat production (rather than remaining a cooler region), which accelerates decay, so we don’t want that. We have to check on thawing specimens regularly and move them to cooler storage areas, or begin dissection earlier, if the decay process is noticeably getting excessive.

Any insulation affects thawing time- so scales, feathers, thick skin, shells, fat (for a short while until it decays), and other layers will slow thawing—and may keep heat inside, if there begins to be thawing of the core. So sometimes you open up a specimen that seems dry and clean on the outside and the inside is unpleasant. But with experience that is not hard to avoid.

Thawed wallaby patella prepared for histology.

The foulest specimen I’ve thawed by far was a monitor lizard… it was shipped to me in California from Arizona when I was a PhD student. This was in August’s heat and the box of the big lizard sat thawing at the post office for 2 weeks before they contacted me and asked why a smelly box was bleeding. I came and got it and brought it back to our department but the smell was so bad it set off our building health & safety person’s alarm bells (sorry, David!) and they emailed around a “toxic alert” warning, until I bashfully made it clear that my lizard was the cause, not some toxic chemical. I got in some trouble and was very ashamed. But we put the specimen into a big tank of brine solution and the smell was reduced—the specimen may well still be preserved there 20 years later; I do wonder! Anyway, that experience was so horrendous – and I have a strong stomach—that I regularly recall it and seek to avoid a repeat. It was the most disgusting thing I’ve ever experienced. I do not recommend it.

What we tend to want to get from thawed specimens is: (1) descriptive anatomy (what connects where), and maybe (2) quantitative measurements (laborious metrics of “muscle architecture”– how much does each muscle weigh, how long is it, etc; over and over again for many muscles…). These data not only serve to tell us what makes animals different (and how this evolved) but also the data are used to test questions such as how animals work. In the case of things like wallabies, ultimately we’d love to know what their kneecaps do if they are bony or not; what difference does it make and why might there be differences? We’d spotted one wallaby already that seemed to have a bony kneecap on one leg, and a non-bony one on the other leg, so that asymmetry got us excited.

What’s surprising to learn about thawing animals for science? Well, my first thought is that it’s beautiful. I don’t tend to think of it as gross. I’ve rhapsodized about this before. Animals are wonderful inside and out, and I regularly pause during a dissection to marvel at how amazing the anatomical specializations of animals are. Simple details- shapes, colours, configurations- can be gorgeous. (Often the blood is minimal, drained out early, so that doesn’t detract from or hide the detailed imagery) The gentle yet complex path of a tendon around a joint can yield profound visual enchantment in its elegance. This is all the more true once one ponders how these complex structures evolved, and how much diversity of form and function is out there to study—and how little we know about it! We still don’t know well how to fix many problems humans have with their anatomy, and that’s orders of magnitude worst for most animals, because we don’t understand how anatomy works, or even what the anatomy is like in some cases. So that keeps me busy discovering things. Every specimen is different with surprising little variations, or big ones—sometimes there is one muscle, sometimes it is clearly divided into two muscles, in the same species or even the left vs. right legs. I love seeing those intricacies and wondering about them.

Thawed wallaby shank sliced open to show lovely digital flexors and gastrocnemius muscles. So many questions are raised by this!

If you’re thawing for Thanksgiving, or thawing for science, or thawing out family relations during a gathering, or thawing yourself out from the winter’s cold– my best wishes to you! May we all enjoy what we thaw.

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