Category: Marine invertebrates

Some ins and outs of raising larvae

Posted on by Allison J. Gong

Today my most recent batches of urchin larvae are six days old. Yesterday being Monday, I changed their water and looked at them under the scopes. I was pleased to be able to split each batch into two jars, as the larvae have already grown quite a bit; I now have a total of four jars to take care of. This makes me inordinately happy. Having only two jars is risky, as it wouldn’t take much for both of them to crash, but for some reason I feel more confident of success with four jars. It’s probably one of those all-your-eggs-in-one-basket things.

In any case, this is what they look like now:

Pluteus larvae of the sea urchin Strongylocentrotus purpuratus, age 5 days. 9 November 2015 © Allison J. Gong
Pluteus larvae of the sea urchin Strongylocentrotus purpuratus, age 5 days.
9 November 2015
© Allison J. Gong

These larvae are perfectly formed. At this point they are shaped essentially like squared-off goblets, with four arms sticking up at the corners of the goblet. They will continue to grow arms in pairs until they have a total of eight (four pairs). The stomachs (the round-ish pale red structures in the middle of the body) are big and round; the color of the stomachs is due to the food that the larvae are eating. And can you see the skeletal rods extending into each of the arms? Each of the eventual larval arms will be supported by one of these rods, and additional rods will serve as cross-braces going horizontally across the body.

Ever wondered what these animals eat? In the wild they would be feeding on whatever phytoplankton they can catch. In the lab we have several types of phytoplankton growing in pure culture, but trial and error has taught us that urchin larvae do best on a diet of the cryptophyte Rhodomonas sp.

The cryptophyte Rhodomonas sp., growing in pure culture. 9 November 2015 © Allison J. Gong
The cryptophyte Rhodomonas sp., growing in pure culture.
9 November 2015
© Allison J. Gong

The red color of the cultures is due to the color of the cells. When the larvae eat this food their stomachs turn pinkish. Rhodomonas cells are about 25 µm long and have two flagella that they use to zip around. Here’s a short video of a drop of Rhodomonas culture on a slide:

They sort of look like sperms, but the cells are much larger than sperms, the flagella are much shorter than the single flagellum of a sperm, and their swimming isn’t quite right to be sperms, either.

The larvae themselves live in glass jars in one of the seawater tables that I converted into a paddle table. The larvae are negatively buoyant and would sink to the bottoms of the jars if left unstirred, and the gentle back-and-forth motion of the paddles keeps them, and their food, suspended in the water column.

See my four jars? They are a sign of short-term success. There’s still a lot of time for things to go south with these larvae, and I certainly don’t take for granted that I’ll be able to keep them alive for the duration. But today, as my students were dissecting urchins in lab, I was able to show them the offspring of said urchins. I hope to keep the larvae alive through the end of the semester, to show the students as much as I can of larval development in one of my favorite animals.

Boy meets girl, urchin style

Posted on by Allison J. Gong

Having obtained decent-ish amounts of gametes from sea urchins, the next step is to get eggs and sperm together. The first thing I did was examine the spawned eggs to make sure they were round and all the same size. Lumpy eggs or a variety of sizes of eggs indicates that they are probably not fertilizable. These eggs from F1 looked just about perfect:

Freshly spawned eggs of Strongylocentrotus purpuratus. 4 November 2015 © Allison J. Gong
Freshly spawned eggs of Strongylocentrotus purpuratus.
4 November 2015
© Allison J. Gong

Note that the eggs are all similarly sized (80 µm in diameter) and round. These look good to go.

The next step is to dilute the sperm in filtered seawater and introduce a small amount to the eggs. The sperm need to be diluted because, believe it or not, in this case too much of a good thing is bad. There’s a phenomenon called “polyspermy” which is pretty much exactly what it sounds like: an egg being penetrated by more than one sperm. Polyspermy leads to wonky development down the road, and while it probably rarely happens in the field, where sperm would be diluted immediately upon being spawned, it definitely does occur in the lab. However, eggs are smart and have evolved a couple of mechanisms to prevent polyspermy.

The fast block to polyspermy occurs within a few seconds of the fusion of the sperm and egg plasma membranes. As the sperm nucleus begins to enter the cytoplasm of the egg, Na+ ion channels in the egg membrane open and cause a depolarization of the egg membrane; this depolarization makes the egg impenetrable to other sperm. However, the egg membrane cannot remain depolarized indefinitely, so after about a minute the slow block to polyspermy takes effect.

The slow block is the rising of the egg’s vitelline layer above the surface of the egg, creating what we call the fertilization membrane. This envelope acts as a physical barrier against additional sperm. The really cool thing about studying fertilization in sea urchins is that you can watch it happen in real time. I mean, how often do you get to observe the formation of a brand new life at the moment that is is being formed?

In this video there are 2.5 eggs in the field of view. Concentrate on the two whole eggs. The one on the top has already been fertilized, which you know because you can see the fertilization membrane surrounding it. You can also see a lot of sperm zooming around. Keep an eye on the lower of the whole eggs; can you see the rising of its fertilization membrane?

Of the two female urchins that spawned for me this morning, F2 had only a few eggs to give but her fertilization rate was 100%. F1, on the other hand, spawned a lot of eggs but only about 50% of them were fertilized. I have no explanation for this. Sometimes (quite a lot of times, actually) things simply don’t work.

That said, at our local ambient temperature the first cleavage division occurs about two hours post-fertilization. That’s when I saw this:

Two-cell embryo of Strongylocentrotus purpuratus, approx. two hours post-fertilization. 4 November 2015 © Allison J. Gong
Two-cell embryo of the sea urchin Strongylocentrotus purpuratus, approx. two hours post-fertilization.
4 November 2015
© Allison J. Gong

A few hours later the embryos had progressed to what I think is the 16-cell stage. At this point it starts getting difficult to distinguish the different cells without focusing up and down through the embryo. But if you know what you’re looking at, the three-dimensional structure does make some sense. In the embryo below I can talk myself into seeing two rings of eight cells each, one ring lying on top of the other.

16-cell embryo of the sea urchin Strongylocentrotus purpuratus. 4 November 2015 © Allison J. Gong
16-cell embryo of the sea urchin Strongylocentrotus purpuratus, approx. five hours post-fertilization.
4 November 2015
© Allison J. Gong

If the embryo is at the 16-cell stage, then it has undergone four cleavage divisions. The early divisions of an embryo are called “cleavages” because the cells divide in half to form equal-sized daughter cells. In other words, the cell cleaves. During cleavage the embryo doesn’t grow, which means that the average cell size necessarily decreases. Cleavage divisions will continue for a total of about 24 hours, resulting in a stage called a blastula.

UP NEXT (hopefully): hatching and swimming

Playing matchmaker

Posted on by Allison J. Gong

We are finally heading into the time of the year that our local intertidal sea urchin, Strongylocentrotus purpuratus, spawns. Usually I would wait until December or January to try to spawn urchins in the lab, but next week my students will be dissecting urchins in lab and I thought I might as well evaluate gonad development in the animals that are going to be sacrificed anyway. In early December I’m going to loan several urchins to a colleague who will be spawning them to show the earliest stages of development to students in one of the lower-division classes at the end of the semester. If I have any luck today, I’ll be able to: (1) start my own cultures of urchin larvae so that I can show the later larval stages to students in my upper-division class; and (2) let my colleague know how likely it is that the urchins I loan to her will be spawnable.

4 November 2015 © Allison J. Gong
4 November 2015
© Allison J. Gong

I know, it ain’t as romantic as the Ritz-Carlton but this is where I hope to make the sea urchins have sex. We have our victims lucky individuals in their “live only” tub, two beakers for eggs, two sperm dishes on ice, a box of glass pipets, a bottle of magic juice, and a syringe with needle to get the magic juice into the animals. Ready to go!

What is the magic juice, you ask? It’s a solution of KCl in filtered seawater. I’m not sure exactly how it works, but here’s what I think happens. We use a solution of MgCl2, a similar salt, to narcotize animals before dissecting them. Sea urchins sitting in a bath of  MgCl2 isotonic with seawater get sleepy pretty quickly, becoming entirely nonresponsive after about 30 minutes. I suspect that KCl has a similar effect. We inject KCl into the main body cavity of the urchin (I call this “shooting them up”) and I think it relaxes the muscles surrounding the gonopores. If the gonads are ripe, then gametes are released as the gonopores open. If gonads are immature, then nothing happens.

A sea urchin is a well-armored beast. Its endoskeleton, or test, is a solid structure composed of calcareous ossicles that are perforated only where tube feet extend. Getting a needle through the test without damaging the animal is pretty much impossible, so we go through the peristomial membrane instead. This membrane surrounds the mouth on the oral (bottom) side of the urchin. It’s the only way to get into an urchin without breaking the test.

The urchins don’t seem to like being injected with KCl–they wave their tube feet and spines all around and generally appear somewhat agitated–but they don’t suffer any lasting effects.

If the urchins are ripe, they should start spawning shortly after being injected with KCl. Sometimes the response is immediate, with urchins pouring out gametes through all five gonopores at an astounding rate. Today it was much slower. It took about 5 minutes for the first female to spawn:

Spawning female sea urchin (Strongylocentrotus purpuratus). 4 November 2015 © Allison J. Gong
Spawning female sea urchin (Strongylocentrotus purpuratus).
4 November 2015
© Allison J. Gong

That little blotch of pale orange is is the mass of eggs that she is spawning. At this point you can pipet off the eggs into a beaker of filtered seawater, but I decided to go the less-invasive route and simply invert the spawning animal onto a beaker filled with water and let the eggs drop to the bottom as they flowed out of her.

The only difficulty with this method is that the animal doesn’t like being upside down and immediately tries to right herself. I kept having to remove her from the beaker and replace her in the orientation we wanted. I designated this urchin as F1. She gave us a decent number of eggs. A second, smaller female (F2) spawned just a few eggs but we kept them all.

Sperm get a different treatment. I had only one male spawn this morning and he wasn’t exactly a gusher. I pipetted off the concentrated sperm into a cold dish on ice, and didn’t dilute the sperm until the eggs were ready for fertilization.

UP NEXT: Fertilization and subsequent events.

You are what you eat, part the fourth

Posted on by Allison J. Gong

The juvenile sea urchins I’ve been raising this year are now nine months old. Back in June I put them on three different macroalgal diets and have been measuring their test diameters monthly. I do the measuring in the first week of every month, and today was the day for November. Over the past few weeks I lost a lot of my Ulva urchins, for no reason that I could discern. Judging from the poop production they were definitely eating, but on some days there would be a handful of corpses in the bowl when I changed the water. They all seemed healthy and happy today, including this beautiful creature:

Juvenile sea urchin (Strongylocentrotus purpuratus) that has been eating Ulva, age 9 months. 2 November 2015 © Allison J. Gong
Juvenile sea urchin (Strongylocentrotus purpuratus) that has been eating the green alga Ulva sp., age 9 months.
2 November 2015
© Allison J. Gong

Seriously, this has to be the most gorgeous photo of a sea urchin I’ve ever taken. This individual is the largest of my Ulva urchins, with a test diameter of 12.7 mm. I love the coloration of this animal: the younger spines are green, the older spines are pale purple, and the tube feet are beautifully transparent and tipped with purple suckers.

By contrast, the urchins eating Macrocystis continue to be a more uniformly golden color:

Juvenile sea urchin (Strongylocentrotus purpuratus) that has been eating the kelp Macrocystis pyrifera, age 9 months. 2 November 2015 © Allison J. Gong
Juvenile sea urchin (Strongylocentrotus purpuratus) that has been eating the kelp Macrocystis pyrifera, age 9 months.
2 November 2015
© Allison J. Gong

This Macrocystis urchin is actually a tad bigger than the Ulva urchin and has a test diameter of 13.0 mm. It looks smaller because its tube feet are fully extended, so I had to zoom out a bit to get the entire body in the frame. It was also crawling around very fast and I had to hold it down to get it centered, then remove the forceps and take the picture quickly before it walked out of the picture. Every photo of this individual that I managed to get is a little blurry because of the movement.

Last but not least, the urchins eating coralline algae are hanging in there. None of them died in the past month and they are growing. Their color patterns are qualitatively different from the those of urchins eating Ulva or Macrocystis. To my eye there is more contrast in the coralline urchins; they all seem to have prominent dark coloration in the lines that radiate outward from the apical region. The other urchins have it too, but in the coralline urchins this dark pigmentation is concentrated into more clearly defined streaks and contrasts more strongly with the paler background color.

Juvenile sea urchin (Strongylocentrotus purpuratus) that has been eating coralline algae, age 9 months. 2 November 2015 © Allison J. Gong
Juvenile sea urchin (Strongylocentrotus purpuratus) that has been eating coralline algae, age 9 months.
2 November 2015
© Allison J. Gong

This animal, with a test diameter of 6.08 mm, is about half the diameter of the largest of its full siblings in each of the other food treatments. Food quality definitely has an effect on size, as these data indicate:

Test diameters of juvenile sea urchins (Strongylocentrotus purpuratus) on three food treatments. 2 November 2015 © Allison J. Gong
Test diameters of juvenile sea urchins (Strongylocentrotus purpuratus) on three food treatments.
2 November 2015
© Allison J. Gong

It remains to be seen whether or not I’ll be able to provide Ulva and Macrocystis to these animals throughout the winter. If we get the strong El Niño storms that are predicted, the nearshore algae could be wiped out for a while. I’ll make sure that if I run out of one food then urchins in the other treatment will also fast until I can feed both of them again. In the meantime, because the coralline urchins are so far behind in their growth, I’ll continue to give them access to food. I don’t want any of them to die of starvation, and the coralline eaters are the most vulnerable, I think.

Happiness is . . .

Posted on by Allison J. Gong

. . . taking a small group of highly motivated students into the field!

My invertebrate zoology class this semester has only 10 students, which allows me a lot more freedom to improvise on the fly and actually participate in the course instead of having to stand back and supervise 30 of them at the same time.

Most of my class getting started on their investigative journalist assignment at Point Pinos. 27 October 2015 © Allison J. Gong
Most of my class getting started on their investigative journalist assignment at Point Pinos.
27 October 2015
© Allison J. Gong

Their job was to interview at least six marine invertebrates and suss out answers to the Big 6 questions: Who? What? When? Where? Why? and How? In other words, to do a small bit of preliminary ecological investigation into animals they don’t already know much about. Some of the students also used the time to scope out the site for their independent research projects, which they will be starting soon.

. . . serendipity!

This past couple of classes I lectured on Platyhelminthes and Nemertea, and we saw both on the field trip.

The flatworm, Eurylepta californica, was spotted by a keen-eyed student, who thought at first it was a nudibranch but then noticed the ruffling edge and decided it must be something else.

Eurylepta californica, the "chocolate drizzle" polyclad flatworm, at Point Pinos. 27 October 2015 © Allison J. Gong
Eurylepta californica, the “chocolate drizzle” polyclad flatworm, at Point Pinos.
27 October 2015
© Allison J. Gong

This individual was a bit less than 2 cm long. I’ve only seen it at Point Pinos. Such a cool animal!

Some day I want to find one of these at a site where I can collect, and bring it back to the lab for closer observation.

On each of these class field trips to the intertidal there’s at least one conversation that goes something like this:

  • Student: Allison! I found this thing! What do you think it is?
  • Me, from several rocks over: Well, what does it look like?
  • Student gives a vague description, which usually isn’t very helpful.
  • Me: Is it alive?
  • Student: I think so.
  • Me: Color?
  • Student: Sort of orange. (or brown or purple or whatever)
  • Me: Shape? Size?
  • Student: This big (holds up fingers or hands to indicate size, then describes shape).
  • Me: Is it hard or squishy?
  • Student: I don’t want to touch it! Is it going to hurt me?
  • Me: Not unless it’s a big crab. Just touch it and tell me what it feels like!
  • <pause>
  • Student: Hey, it didn’t hurt me!

This conversation occurs as I make my way over to see what it is. Eventually I can take a look at the whatever-it-is and explain as best I can. The nemertean that we saw yesterday resulted in a conversation similar to this, but the student had pretty much decided on her own that she had found a nemertean. By the time I made it over to where she was pointing the worm had just about disappeared into a mussel bed, which is where they hang out. I could see enough to determine that it was Paranemertes peregrina.

Paranemertes peregrina, a nemertean worm, at Pistachio Beach. 31 January 2015 © Allisoin J. Gong
Paranemertes peregrina, a nemertean worm, at Pistachio Beach.
31 January 2015
© Allison J. Gong

Nemerteans are unsegmented, slimy, predatory worms that feed by shooting out a sticky proboscis and wrapping it around prey. Some have a stylet at the end of the proboscis with which they can repeatedly stab the prey and inject toxins. They may not be much to look at, but watching them in action should make you glad that you’re not a small animal.

. . . being in the right place at the right time!

Yesterday we saw octopuses! Three of them, I think. And one of the most glorious sea anemones I have ever seen.

Octopus rubescens crawling around at Point Pinos. 27 October 2015 © Allison J. Gong
Octopus rubescens crawling around at Point Pinos.
27 October 2015
© Allison J. Gong
A beautiful Anthopleura xanthogrammica anemone at Point Pinos. 27 October 2015 © Allison J. Gong
A beautiful Anthopleura xanthogrammica anemone at Point Pinos.
27 October 2015
© Allison J. Gong

The octopuses that were out of the water were duly rescued by my students. The red one that I photographed turned out to about the length of my hand when it swam away into the depths of a tidepool. Watching the students release this little animal back into the water was a fitting way to close out what had been a fantastic field trip.

Feeding the ‘droids

Posted on by Allison J. Gong

So, those bits of Ectopleura crocea that I grabbed from the harbor on Monday are voracious eaters. I didn’t feed them yesterday because I didn’t have time and the students spent the afternoon looking at them in lab, and I hoped that they’d be alive today. Some of the stalks had dropped their hydranths (the distal part that bears the feeding tentacles, mouth, and reproductive gonophores) but most of them were alive and lovely. I had a nice fresh batch of brine shrimp all hatched out and ready to go and thought I’d see if the hydranths could eat them. Little did I know that this simple exercise would occupy most of my day.

I started by squirting some of the brine shrimp onto the hydranths, which for the most were pretty lackadaisical about catching them. Then I decided to feed them the mashed up brine shrimp that I’m still feeding the tiny Melibe and WOW! that did the trick! Maybe it was the scent of the macerated brine shrimp that triggered the feeding response. I was fascinated.

They are beautifully and unexpectedly animated animals.

After I watched them feed for a while, it seemed to me that the outer ring of tentacles catches and holds onto prey, while the prehensile manubrium swings around and brings the mouth into contact with the food. In the meantime, while all this brine shrimp catching is going on there are other larger crustaceans crawling all over the hydranths, even onto the tentacles, without getting stung. I think their exoskeletons must be thick enough not to be penetrated by the hydroid’s cnidocytes (stinging cells).

Having discovered the trick to making the hydranths eat, I squirted brine shrimp mush on them and left them alone for about 20 minutes. When I came back they had eaten and I could see brine shrimp in their guts, so I gave them more. The feeding response was pretty much as vigorous as the first one had been. So I kept feeding them throughout the morning and early afternoon.

If I didn’t have other things to do, I could watch these all day. I hope that if I can keep feeding them this much they will regrow their dropped hydranths. Although I’m not sure how realistic it is to think that I can go through this routine every day. And do I really need a few dozen more mouths to feed on a regular basis? I seem to accumulate animals like other women accumulate shoes. On the other hand, I don’t expect the Ectopleura colonies to last long in the lab so even my “forever” relationship with these particular animals will likely be over in a week or so. I can probably keep up this level of effort for that long.

These ARE the droids I’m looking for

Posted on by Allison J. Gong

Tomorrow my students will be examining cnidarian diversity in lab, so early this morning I went to the harbor to collect hydroids. Or ‘droids, as I refer to them. These are not the droids of Star Wars fame, such as C-3PO and R2D2, but rather colonial cnidarians. As such, they are made up of many iterated units (called zooids) connected by a shared gastrovascular cavity (GVC), or gut. Despite how weird it seems to most people, this sort of colonial lifestyle is not uncommon among marine invertebrates; it occurs in several other taxa as well, most notably the Anthozoa (sea amenones, corals, and others), Bryozoa (bryozoans such as Membranipora), and Urochordata (sea squirts).

On my various trips to the harbor over the past few months I’ve been keeping an eye out for ‘droids, as I knew I’d need them. The one species I was glad to see getting established this summer is called Ectopleura crocea; it is one of the non-native members of the fouling community that shows up in harbors all along the California coast. It is a most beautiful animal, and quite conspicuous when it is present. This year I’ve seen it growing lustily on the docks, mussels, and any manmade object that has been marinating in the water for a while.

In situ it looks like this:

The hydroid Ectopleura crocea, at the Santa Cruz Yacht Harbor 19 October 2015 © Allison J. Gong
The hydroid Ectopleura crocea, at the Santa Cruz Yacht Harbor
19 October 2015
© Allison J. Gong

The stalks in this particular colony are 3.5-4 cm long. Each one of those tufts at the end of a stalk is a hydranth, the part of the zooid that bears the feeding tentacles and mouth. Hydroids are cnidarians and thus have stinging cells along their tentacles, which form a ring surrounding the mouth.

Ectopleura hydranths actually have two concentric rings of tentacles, with the mouth in the middle of the smaller ring. Between the tentacle rings there is a sort of empty space that is filled with reproductive structures called gonophores when the colony is preparing for sexual reproduction. In some hydroids gonophores release medusae, but in Ectopleura they release gametes. A given colony is either male or female, and any one of the hydranths can become reproductive and develop gonophores.

Hydroids are definitely animals whose beauty is better appreciated when observed under a microscope:

Hydranth of Ectopleura crocea 19 October 2015 © Allison J. Gong
Hydranth of Ectopleura crocea
19 October 2015
© Allison J. Gong

In the colonies of E. crocea that I’ve observed before, mature male gonophores are a solid white and female gonophores are pinkish. I collected three clumps of Ectopleura today, and none of the gonophores are mature. You can see why the common name for this animal is “pink mouth hydroid,” as the mouth is borne on a pink tubular structure called a hypostome.

I’ve tried multiple times to grow this animal in the lab. There are some experiments on resource sharing in hydroids that I’ve been wanting to do for years but haven’t yet found the right species to work with. In captivity Ectopleura eats well, then after several days all the hydranths drop off and the colonies die. I’ve never had success getting them to regrow their hydranths, either. So I bring them in for short periods and observe them up close while I can.

Off with the old, Case B

Posted on by Allison J. Gong

ORGANISM OF THE MONTH: Pugettia producta, the kelp crab

For a few months now, I’ve had a pet kelp crab running around in one of my seawater tables. I don’t remember where I collected it, or even whether or not I collected it at all; quite often crabs and other animals arrive as hitch-hikers on kelp that we bring into the lab to feed urchins, and I end up with many cool critters in my care that way. However she got here, this crab has been rather a pain in the butt during her stay with me. For at least a couple of weeks she got stuck in the drain of the table and would not come out despite three experienced marine biologists (including yours truly) trying to persuade her by altering water flow and offering food bribes. Then she disappeared from the table drain and I assumed that she had gone all the way through to the floor drain, where she could live quite happily for all eternity. Then she suddenly showed up again in one of my urchin baskets. When she came back up from the drain and how long she’d been hiding, I’ll never know.

Wondering why I keep referring to this crab as “she”? It’s because I know for certain that she’s a female. Here’s the secret to how you can determine the sex of brachyuran crabs (most of the common crabs: kelp crabs, shore crabs, rock crabs, even Dungeness crabs): You look at the shape of the abdomen, which is curved forward on the underside of the body. See here:

Abdomen of female Pugettia producta. 16 October 2015. © Allison J. Gong
Abdomen of female kelp crab (Pugettia producta)
16 October 2015
© Allison J. Gong

The abdomen is the broad flat upside-down-U-shaped panel that covers about half the width of the ventral surface. Female crabs brood their embryos under the abdomen, hence the broad shape. Male crabs of the same species have a much narrower, pointed abdomen.

Since her escapade with the drain the crab has been more, shall we say, co-operative. She’s still free to scurry around at will in the table, but I haven’t found her doing anything objectionable such as tormenting urchins or trying to get down the drain again. She has also been eating well.

Until this past week, that is. On Monday she accepted a piece of food but then abandoned it without even tasting it. On Wednesday she fled from the food, which I took to mean that she was getting ready to molt. Like all arthropods, crustaceans molt their exoskeletons every so often. The decapod crustaceans I’m most familiar with tend to off their feed for a few days before molting, and usually the actual shedding of the exoskeleton occurs at night. Then we show up the next day and voilà! like magic there’s a new, bigger crab in the table.

Ms. Kelp Crab stopped eating on Monday of this week. Today (Friday) I didn’t get to the lab until about noon, and one thing I noticed in the table was an empty carapace. Sure enough, she had molted. It took a little hunting to find the crab herself, but she wasn’t really hiding and her new exoskeleton had already hardened. I’m pretty sure she’ll eat on Monday.

Kelp crab (Pugettia producta) and carapace of its molted exoskeleton. 16 October 2015 © Allison J. Gong
Kelp crab (Pugettia producta) and carapace of her molted exoskeleton
16 October 2015
© Allison J. Gong

Living in a rigid exoskeleton means that a crustacean can increase in body size only in the time period between when an old exoskeleton is shed and the new one hardens up. I’m always curious about exactly how much crabs grow when they molt. So today I measured the crab and her old carapace at the same place, halfway between the two points on the lateral edges of the carapace. Huzzah for empirical data! The old carapace measured 27.6mm across, and the new one 33.8mm, for an increase in width of 6.2mm or 22.5%. Mind you, this is simply the increase in one linear dimension of the crab’s body. To obtain a more accurate measurement of body size increase, I’d have to have weighed the crab immediately before her molt and after it. Still, it does give an estimation of how much bigger a body part can get when a crab molts.

All good things. . .

Posted on by Allison J. Gong

. . . must come to an end, so they say. And Scott’s and my little experiment growing Pisaster ochraceus came to its end when the last of our teensy stars gave up the ghost a week ago. We aren’t entirely surprised, as nobody before us had succeeded in growing these guys in the post-larval stage, but it’s still sad to see the empty paddle table and disappointing to know that we haven’t really added to the body of knowledge about how to grow them.

But we did make a small bit of progress, at least to further our own understanding of exactly how difficult it is to do what we attempted. To summarize, here’s a timeline of what we did and what happened:

  • 18 and 20 May 2015 — Collected adult stars from local intertidal sites. Made up the solution of “magic juice” (100 µM 1-methyladenine).
  • 2 June 2015 — Shot up stars with 1-MA. Got usable amounts of gametes from a total of three stars: 2 Purple (1 female + 1 male) and 1 Orange (female). After examining gametes to make sure they were okay, set up two matings: Purple x Purple; and Orange x Purple. The Purple x Purple embryos went through the earliest developmental stages just fine. The Orange x Purple embryos got off on the wrong foot and never recovered.
  • 5 June 2015 — Purple x Purple embryos began undergoing gastrulation. Began feeding them. Orange x Purple embryos all dead.
  • 20 July 2015 (age 48 days) — Larvae began settling.
  • 27 July 2015 (age 55 days) — Counted a total of ~22 tiny stars in the jars. Removed a few to measure, and they were all 500 µm or smaller in diameter. It was very difficult keeping track of things this tiny in our 1-gallon jars.
  • 13 August 2015 (age 73 days) — Paintbrushed out all of the little stars into a bowl and divvied them up into six food treatments. Replaced bowls in the paddle table to provide very gentle stirring.
  • 21 August – 7 September 2015 — Stars died off in all but one of the food treatment bowls. By 7 September (age 96 days) the only surviving stars (N=4) were the ones we kept in a bowl with a small piece of mussel shell.
  • 11 September 2015 (age 100 days) — And then there were three.
  • 28 September 2015 (age 117 days) — Two survivors + 1 corpse.
  • 2 October 2015 (age 121 days) — And then there were none.

In a nutshell, the larval development went fairly well, as we expected, and the post-larval survival sucked, also as we expected. We did manage to get those last two stars to survive 48 days post-metamorphosis, which is something. I’m not sure how much credit we can take for that, though, as I suspect that the reason the other juveniles died had to do with poor water quality as much as lack of food.

Here’s what I think might have been going on: We are in our second consecutive year of elevated seawater temperature, and coupled with the massive El Nino that yesterday was proclaimed to be among the strongest ever this means that coastal animals are being subject to higher-than-normal temperatures. In ectothermic poikilotherms such as marine invertebrates, metabolic rate is directly related to environmental temperature. Thus, higher ambient seawater temperature should result, all else being equal, in a faster growth rate.

This sounds like it might be a good thing for the Pisaster larvae, especially if predation and other risks are higher in the planktonic larval stages than as benthic juveniles. However, I think there’s more to the problem than simple growth rate. What if success as a juvenile depends not only on how quickly an animal progresses through all of its developmental stages, but also on how much time it spends in the different stages? For some larvae, notably the nauplius larva of barnacles, the primary job is to eat as much as possible and deposit energy reserves in the form of oil droplets; these food reserves will be utilized by the second larval stage, the non-feeding cyprid, as it hunts around for a place to establish a permanent home in the benthos. Perhaps part of the job of the developing Pisaster brachiolaria larvae is also to sequester energy reserves. Although no oil droplets were visible in any of the larvae that Scott and I observed this summer, energy could have been stored in other tissues of the larval body.

Back to the problem of post-larval survival. Our larvae began metamorphosing after only 48 days in the plankton. One of our sources has Pisaster ochraceus undergoing metamorphosis at 76-228 days in culture, at temperatures of about 12°C (for the duration of our experiment this summer ambient seawater temps were 15-18.5°C). So, if the warmer temperatures caused the larvae to develop more quickly than normal, and the larvae spent ~25 fewer days in the plankton than they “should” have, they may simply not have had time to accumulate whatever energy reserves they’d need to draw on once they metamorphosed.

That’s just a guess on my part. I also imagine that poor water quality played a part in our juvenile stars’ demise. It proved to be impossible to make potential food available to such tiny animals while keeping their water clean. We thought that stirring on the paddle table might help, and who knows, maybe it did.

In any case, RIP, little guys. Thanks for what you taught us, and I’m sorry we weren’t able to help you succeed.

Juvenile Pisaster ochraceus, age 75 days. 16 August 2015 © Allison J. Gong
Juvenile Pisaster ochraceus, age 75 days.
16 August 2015
© Allison J. Gong

Cuteness strikes again!

Posted on by Allison J. Gong

That cute little Melibe I found last week is still alive, and still super cute. It lost one of the two large cerata on its back the second day I had it, and I wasn’t sure it would be able to survive long without it, but it has hung in there and started growing a replacement. This afternoon it was crawling on the underside of the surface tension in the bowl:

Melibe leonina crawling on underside of surface tension. 2 October 2015 © Allison J. Gong
Melibe leonina crawling on underside of surface tension.
2 October 2015
© Allison J. Gong

It is extremely difficult photographing transparent animals; this is the best shot I got. You are looking at the animal’s ventral surfaces. It is using its elongate foot to stick to the surface, and the rest of the body is suspended from the foot. The oral hood is wide open and you can see the little blue spots at the base of each tentacle.

The best news is that the tiny Melibe has learned how to eat! The first couple of days I offered it live brine shrimp nauplii, and the Melibe didn’t seem to like the thrashing of the nauplii. It cowered and shrank instead of trying to eat them. Then it occurred to me to mush up the nauplii first, so they wouldn’t be so active. I also thought that the Melibe might be able to eat the mush itself. Aha, success! Except that I wasn’t able to capture any video or photos then.

Today, though, the Melibe did this, while I had the camera all set up and ready to go:

Instead of cringing from the nauplii, today the Melibe was actively going after them. In this video it encloses its oral hood around a handful of nauplii and collapses the hood, forcing the nauplii into its mouth. You can actually see the nauplii stop struggling as they are ingested.

I think the Melibe is growing, too. I’ll have time to measure it on Monday.