Tag: sea stars

Return of the natives

Posted on by Allison J. Gong

When the most recent epidemic of seastar wasting syndrome (SSWS) began back in 2013, the forcipulate stars were the first to succumb. This group includes conspicuous members of intertidal and subtidal habitats, such as:

  • Pisaster ochraceus — the intertidal ochre star
  • Pisaster giganteus — the giant spined star, which lives in the low intertidal and subtidal
  • Pycnopodia helianthoides — the sunflower star, a huge monster of the low intertidal and subtidal.

In the past year or so, I’ve noticed P. ochraceus making a comeback at local intertidal sites. At first I was seeing stars in the 2-3 cm size range, and now I’m regularly seeing hand-sized ones clinging to the rocks.

4 mm juvenile Pisaster ochraceus star at Pescadero State Beach.
11 May 2016
© Allison J. Gong

You read that right. 4 mm in diameter. This is the tiniest forcipulate star that I’ve ever been able to ID in the field with any certainty.

Pair of Pisaster ochraceus stars in the low-mid intertidal at Natural Bridges.
22 July 2016
© Allison J. Gong
A hand-sized (dark orange) and much smaller (dark purple, tucked far back in the little cave) Pisaster ochraceus at Mitchell’s Cove.
28 November 2016
© Allison J. Gong

It seems pretty clear that the ochre stars, at least, are making a comeback. It’s likely that the larger ones are survivors of the SSWS plague. That little tiny one, though, may well be a post-SSWS recruit. Unfortunately we don’t know how fast they grow once they recruit to the benthos. We do know that when they recruit they’re about 500 µm in diameter, so even that little guy has grown a lot in however long it has been since it settled.

The really exciting news is that yesterday I saw my first P. giganteus since the SSWS outbreak began! I was up at Davenport Landing collecting sea urchins and saw this star in an urchin hole. The rock around here is a soft mudstone that is easily eroded. Urchins excavate holes by twisting their spines against the rock, and then live in them. Holes that are urchinless, for whatever reason, are quickly colonized by other organisms (including baby urchins).

A not-so-gigantic Pisaster giganteus star in an urchin hole at Davenport Landing.
13 December 2016
© Allison J. Gong

For a sense of size, this urchin hole is about 8 cm in diameter. The star is sharing it with a small anemone, most likely Anthopleura elegantissima.

Pisaster giganteus generally occurs lower in the intertidal than P. ochraceus, and I wouldn’t expect to see it on a tide that isn’t at least as low as -0.8 ft. It isn’t as closely associated with mussel beds as P. ochraceus, either, because it lives lower in the intertidal. Fortunately, this week’s low tide series includes a few days with tides below -1.0 ft, and I’m going back out today. I’ll be keeping my eyes open for not only Pisaster stars, but also the Pycnopodia that disappeared a few years ago. Although Pycnopodia gets very large, I don’t expect to see any really big ones running across the intertidal. However, Pycnopodia juveniles would indicate at least the beginning of a possible population recovery  from the SSWS plague.

So, wish me luck and keep your fingers crossed!

Seeing (wannabe) stars

Posted on by Allison J. Gong

So. I have a batch of larvae from a spontaneous spawning of the leather star, Dermasterias imbricata, that occurred four weeks ago tonight. Until now I’ve never had an opportunity to work with this species, even though we have quite a few of them at the marine lab. I had my own for several years, until they became casualties of the plague about a year into the current sea star wasting syndrome event. In any case, this is the first time I’ve been able to spend time with larvae of this species. At the very least I wanted to see how big they would get and how quickly they would develop, compared to the species I’m more familiar with, Patiria miniata (bat star) and Pisaster ochraceus (ochre star).

When the Dermasterias spawned, the first thing I noticed was that the eggs are huge. I measured them at 220 µm in diameter, which is big even compared to what I’ve seen in other stars. Hatch rates were pretty good, and four days later the larvae were already in the 400-430 µm range. Since I have no experience culturing this species, I thought I’d divvy up my larvae and put them into three feeding treatments to see which larval diet resulted in the best overall success. According to the literature, Dermasterias larvae can be raised on a mixture of the unicellular algae Dunaliella tertiolecta (green) and Isochrysis galbana (golden). My three feeding treatments are: Dun only, a Dun/Iso mix, and Iso only.

A week into the experiment there was a clear difference between the larvae eating only the green food, and those eating either a mixture of green and golden or only the golden. Larvae from all food treatments were about the same size, but the ones eating only Dunaliella had noticeably green guts.

Bipinnaria larva of Dermasterias imbricata. 29 February 2016 © Allison J. Gong
Bipinnaria larva of Dermasterias imbricata, fed on Dunaliella tertiolecta, age 7 days.
29 February 2016
© Allison J. Gong
Bipinnaria larva of Dermasterias imbricata, fed on a mixture of D. tertiolecta and I. galbana. 29 February 2016 © Allison J. Gong
Bipinnaria larva of Dermasterias imbricata, fed on a mixture of Dunaliella tertiolecta and Isochrysis galbana, age 7 days.
29 February 2016
© Allison J. Gong
Bipinnaria larva of Dermasterias imbricata, fed on I. galbana. 29 February 2016 © Allison J. Gong
Bipinnaria larva of Dermasterias imbricata, fed on Isochrysis galbana, age 7 days.
29 February 2016
© Allison J. Gong

Fast forward two weeks, and the larvae were 20 days old. By this time they had progressed from the bipinnaria stage to the brachiolaria stage. The interesting thing was the absence of green pigment in any of the guts, even those that were eating only green food. The D. tertiolecta larvae looked good, actually. They were a little smaller than the other larvae but were perfectly formed.

Brachiolaria larva of Dermasterias imbricata, fed D. tertiolecta, age 20 days. 14 March 2016 © Allison J. Gong
Brachiolaria larva of Dermasterias imbricata, fed D. tertiolecta, age 20 days.
14 March 2016
© Allison J. Gong
Brachiolaria larva of Dermasterias imbricata, fed D. tertiolecta and I. galbana, age 20 days. 14 March 2016 © Allison J. Gong
Brachiolaria larva of Dermasterias imbricata, fed D. tertiolecta and I. galbana, age 20 days.
14 March 2016
© Allison J. Gong
Brachiolaria larvae of Dermasterias imbricata, fed I. galbana, age 20 days. 14 March 2016 © Allison J. Gong
Brachiolaria larvae of Dermasterias imbricata, fed I. galbana, age 20 days.
14 March 2016
© Allison J. Gong

Obviously all of the larvae are assimilating enough of their food to grow and develop normally. I looked at them today but didn’t have time to take pictures. Qualitatively there is no difference between the Dun larvae and the Dun/Iso larvae. In the Iso jars, however, there are many larvae at earlier stages; some are still at the “jellybean” stage. I don’t know if this is because these larvae are developing more slowly, or because of some nonrandom distribution of earlier stages into those jars when I was setting up the feeding treatments.

Next week I’ll measure the larvae again, and will have three data points to track growth trajectories.

Swimming jellybeans

Posted on by Allison J. Gong

When serendipity strikes, I try to go with the flow and ride it as long as I can. The latest wave is my batch of Dermasterias larvae, which are developing nicely for the first four days of life. And now they look just like jellybeans!

They have complete guts now and have already grown a bit, measuring 400-430 µm long. It’s not always easy to catch these guys in the right orientation to take a photo, as they are spinning and swimming through three-dimensional space, but I got lucky:

4-day-old dipleurula larva of Dermasterias imbricata. 26 February 2016 © Allison J. Gong
4-day-old dipleurula larva of Dermasterias imbricata.
26 February 2016
© Allison J. Gong

For now I’ve got the larvae divvied up into different feeding treatments. More on that later.

Serendipity strikes again

Posted on by Allison J. Gong

Actually, it was a fortunately placed phone call from an aquarium curator that struck the other night. I was at home, having eaten dinner and reviewed my lecture for the following morning, when my phone rang. It was the curator, saying that he was making his last rounds of the evening and had noticed that some of his sea stars were spawning. Echinoderm sex–more specifically, the opportunity to collect gametes and observe larval development–always grabs my attention, so I told him I’d throw on some shoes and meet him at the marine lab in five minutes.

Lo and behold, there were leather stars (Dermasterias imbricata) spawning in several of the tanks and seawater tables. Many of the tables were cloudy with sperm, but I found only one female, which seems strange but isn’t so unusual. These spawning events occur in response to some environmental cue, such as day-length, a chemical of some sort, or the phase of the moon. When a sea star (or sea urchin) spawns it also releases chemicals that trigger spawning in nearby conspecifics, as to spawn by oneself is an enormous waste of energy. A single spawning animal can result in all the others of its kind spewing out huge numbers of gametes in an orgy of passive sex. However, an animal can be induced to spawn only if its gonads are ripe. Ripeness depends on the overall health of the animal and requires adequate food; animals that don’t receive enough food don’t have energy to allocate towards gamete production. As eggs are energetically expensive to produce, compared to sperm, it is not unusual for males of a species to mature earlier in the reproductive season than the females. In Washington the spawning season for D. imbricata is April-August. Here in California the reproductive season hasn’t been clearly defined, but I do remember a springtime spontaneous spawning event in the lab several years ago.

Spawning female leather star (Dermasterias imbricata) at Long Marine Lab. 22 February 2016 © Allison J. Gong
Spawning female leather star (Dermasterias imbricata) at Long Marine Lab.
22 February 2016
© Allison J. Gong

That creamy looking mass of goo on the star’s aboral surface is a pile of eggs. Sea star eggs are fairly large, compared to the urchin eggs I’m used to, and sticky. They tend to clump together in stringy globs until they are dispersed by water currents. The star whose arm is photobombing in the lower right corner is a male. He was also spawning copiously and is probably the individual who fertilized most of this female’s eggs.

Given the lateness of the hour and the fact that I had to get up early the next morning I didn’t take many pictures of the eggs, although I did look at them to make sure they were fertilized. They were, so I put them into a 1000-mL beaker of seawater and let them do their thing.

Fast forward to today, about a day and a half after fertilization. About two-thirds of the embryos had hatched and were swimming in the water column. Here’s what they look like under the dissecting scope:

I poured off the swimmers into jars and set them up on the paddle table. I gave them a little bit of food, in case their mouths break through before I can get back to the lab tomorrow afternoon. In the meantime, I took a sample of embryos and examined them under the microscope. They look really cool!

Almost-two-day-old embryo of Dermasterias imbricata. 24 February 2016 © Allison J. Gong
Almost-two-day-old embryo of Dermasterias imbricata.
24 February 2016
© Allison J. Gong

The embryos are almost spherical, measuring 290 µm long and 270 µm wide. They are ciliated all over and swim with the rounded end forward. The flattened end is where the process of gastrulation started. That visible invagination begins at a section of the embryo called the blastopore; the channel is the archenteron, the first gut of the larva. In echinoderms, as in chordates (including us humans), the blastopore will end up being the larva’s anus; the mouth breaks through later at the other end of the archenteron. This is why I don’t need to start feeding the larvae right away even though their gut has begun forming.

Tomorrow afternoon I’ll have a brief window of time when I can check on the larvae and see how they’re doing. I think they may have complete guts by then!

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

Constellations

Posted on by Allison J. Gong

I did a quick search, and there doesn’t seem to be a collective noun for sea stars. I’m going to remedy that by declaring “constellation” to be the official term for a group of sea stars. And by “official” I mean that’s the term I’m going to use. Who knows, maybe it’ll take.

In any case, I certainly have a constellation of sea star larvae in each of my jars. Today I pipetted a lot of them into a bowl, and they look pretty cool all swimming together, like strange alien spaceships. What do you think?

The largest of the larvae are over 2 mm long now, and the brachiolar arms have grown much longer. They have three adhesive papillae on the ventral side of the anterior projection and well-formed juvenile rudiments, where the water vascular system is forming. They’re much too big to fit under the compound scope, so the only way to get pictures of the entire body is through the dissecting scope:

Brachiolaria larva of Pisaster ochraceus, age 31 days. 3 July 2015. © Allison J. Gong
Brachiolaria larva of Pisaster ochraceus, age 31 days
3 July 2015
© Allison J. Gong

In the above photo you are looking at the larva’s ventral surface, so the animal’s left side on the right side of the photo, and vice versa. If you squint you might be able to convince yourself that you see a small whitish bleb on the left side of the stomach; that’s the rudiment. Since it doesn’t make much sense under this magnification, I removed this individual to a slide and put it under the compound scope. It doesn’t fit in the field of view, so I took pictures of each half of the body. If I were clever with photo editing software I’d be able to mesh these photos into a single image. Alas….

Ventral view of the anterior end of a brachiolaria larva of Pisaster ochraceus, age 31 days. 3 July 2015. © Allison J. Gong
Ventral view of the anterior end of a brachiolaria larva of Pisaster ochraceus, age 31 days
3 July 2015
© Allison J. Gong
Ventral view of the posterior end of a brachiolaria larva of Pisaster ochraceus, age 31 days. 3 July 2015. © Allison J. Gong
Ventral view of the posterior end of a brachiolaria larva of Pisaster ochraceus, age 31 days
3 July 2015
© Allison J. Gong

This gives you a better view of the juvenile rudiment on the animal’s left. Those three roundish blobs are tube feet! I think it’s likely that at some point in the not-too-distant future the larvae will be competent, which means they’d be physiologically and anatomically capable of metamorphosis. It seems to me that they are still developing very quickly, and with seawater temperatures consistent at 15-16°C I don’t expect that to change. So far, so good!

Edit 4 July 2015:  Look at what my online friend Becca can do! She was able to merge my photos into a single image. Now you can see the entire body! Thanks, Becca!

Composite image of brachiolaria larva of Pisaster ochraceus, age 31 days. 3 July 2015.
Composite image of brachiolaria larva of Pisaster ochraceus, age 31 days
3 July 2015

Growing fast

Posted on by Allison J. Gong

Today my Pisaster ochraceus larvae are 10 days old. Although they seemed to be developing slowly, compared to the urchins that I’m more used to, in the past several days they have changed quite a bit. They’ve also been growing quickly, which makes me think that they’re off to a strong start. Of course, there’s still a lot of time for things to go wrong, as they have another couple of months in the plankton. However, at this point in time I feel optimistic about their chances.

In the dish under the dissecting scope they swim around like bizarre space ships. All the bits of detritus in the water add to the effect. The only thing missing is the sound effects.

The magnification of my dissecting scope goes up to 40X. To see any details of anatomy I have to use the compound microscope, through which I can see this, under 100X magnification:

10-day-old bipinnaria larva of Pisaster ochraceus, 12 June 2015. © Allison J. Gong
Ventral view of 10-day-old bipinnaria larva of Pisaster ochraceus
12 June 2015
© Allison J. Gong

Aside from the dramatic increase in overall size (almost 1 mm long now!), the larva’s body has gotten a lot more complicated. For one thing, the animal’s marginal ciliated band, which propels the larva through the water, has started becoming more elongate and elaborate. In this view the larva is lying on its back, and I have focused on the plane of its ventral surface. The left and right coeloms are in the plane of the dorsal surface, and thus are not really in focus. You should still be able to see how long they have gotten, though. Eventually they will fuse anteriorly to form a single cavity. The stomach of the larva has a nice green-golden color due to the food it has been eating. It makes perfect sense, as we are feeding them a cocktail of green algae and a diatom-like golden alga.

The larvae are very flexible and can be quite animated when they’re swimming around. They bend, scrunch up, and swallow food cells. They have already gotten so big that they take up much of the field of view under the microscope, even at the lowest magnification. Watch some larval gymnastics:

Part of the reason that I wanted to spawn Pisaster and raise the larvae this summer is that I want to put together a series of pen-and-ink drawings of the developmental stages. I did the same for the bat star Patiria miniata several years ago, but the Pisaster larvae will have longer and more elaborate arms when they mature; capturing these in drawings will be a challenge for me. I also hope to include the juveniles in this set of drawings. With that goal in mind, I’ve been sketching the larvae every few days, just to get some practice under my hand and remind myself what it feels like to draw. I’ve missed it!

10-day-old bipinnaria larva of Pisaster ochraceus, drawn from life. 12 June 2015. © Allison J. Gong
10-day-old bipinnaria larva of Pisaster ochraceus, drawn from life
12 June 2015
© Allison J. Gong

For whatever reason, I really like how this sketch turned out. It’s not pretty, but it does truly represent what I saw under the microscope. I’m going to have to work on depicting three-dimensional structures on a two-dimensional page, which will take some practice. Fortunately I have several weeks to brush up on my skills!

Slow and steady (I think)

Posted on by Allison J. Gong

Today the Pisaster larvae that Scott and I are following are a week old. Happy birthday, little dudes! Yesterday we did the twice-weekly water change and looked at them. They’re getting big fast since we started feeding them on Saturday when their mouths finally broke through. At this stage they are sort of jellybean-shaped and extremely flexible–they don’t have the calcified skeletal rods that sea urchin larvae have so they bend and flex quite a lot. They are also beautifully transparent, which allows us to see their guts in fine detail. We can even watch them swallow food cells!

Front view of Pisaster ochraceus bipinnaria larva, age 7 days, 8 June 2015. © Allison J. Gong
Front (ventral) view of Pisaster ochraceus bipinnaria larva, age 7 days
8 June 2015
© Allison J. Gong

In profile view you can see that the larvae are shaped sort of like fat C’s. Here’s a side view of a different individual:

Right side view of bipinnaria larva of Pisaster ochraceus, age 7 days. © Allison J. Gong
Right side view of Pisaster ochraceus bipinnaria larva, age 7 days.
© Allison J. Gong

In the short term (over the next couple of weeks or so) the larvae will continue to get longer. Their guts won’t change much, but their coelomic systems will develop and become more complex. I’ll try to capture that in photos and drawings to share with you.

Questions and answers

Posted on by Allison J. Gong

I’ve been fielding questions about my recent sea star spawning work from people I’ve shared this blog with, which is a lot of fun! To streamline things and make the info available to anybody who might be following, I decided to put together a very brief FAQ-like post to address the most recent questions.

Question:  Can you watch the eggs divide in real time?

In a time-lapse sense you can watch cleavage divisions occur, but not in real time. What I can do is set up a slide on the microscope and leave it there for a while. The gradually warming temperature speeds up development to the point that I can sort of see the division in real time. Of course, the danger is that the embryo will cook on the slide. I generally figure that once I’ve pipetted some embryos onto a slide and dropped a cover slip on top of them, they’re goners (it’s not really possible to remove the cover slip without damaging the cells underneath it) so I feel marginally less bad about sacrificing a few to the gods of observation.

Questions:  I’m fairly certain that the stars can go back to the sea, but are you able to keep their eggs with them, too? How difficult is that transport?

Actually, my scientific collecting permit specifically states that I’m not allowed to return animals to the wild. If I needed to, I could apply for additional permits but it has never been necessary for the work I do. Surplus eggs and larvae, therefore, are discharged into the seawater outflow at the lab and do return to the ocean but the parents remain in my care.

Question:  Are orange and purple stars usually able to cross with each other?

As far as anyone has been able to determine, the color of stars has zero effect on whether two individuals’ gametes are able to do the nasty together. The sea stars that I’m working with–Pisaster ochraceus, the ochre star–are broadcast spawners, meaning that each individual spews his/her gametes into the water, where fertilization and development occur. The stars are also synchronous spawners, meaning that if one individual in an area begins spawning other stars in the immediate vicinity will also spawn. After all, it does take two to tango, and to spawn while nobody else does is a tremendous waste of energy.

So yes, a purple star and an orange star should be able to mate without any problems… at least not any problems due to the parents’ colors.

Question:  If so, what color do they end up being, statstically?

This is a very interesting question. Two of my colleagues are going to spawn Patiria miniata (bat stars) next week to address this. Their plans are to cross a Blue female with an Orange male, an Orange female with a Blue male, and both pure-color matings. They did a preliminary version of this experiment a couple of years ago but didn’t end up with enough juveniles at a size that color could be ascertained; thus they couldn’t calculate any statistically meaningful color ratios.

Questions:  Do you suppose that the wasting disease could be now in the genetic makeup? Any thoughts (unofficial of course) about this?

My thought is sort of the opposite, actually. The animals that we brought in from the field are all survivors of SSWS; if anything, I’d expect them to be resistant to whatever causes the plague, and to (hopefully) pass on this resistance to their offspring. Of course, there’s no way of knowing if and how exposure to SSWS affects the quality of the gametes. It’s quite possible that these survivors are less fit after the SSWS outbreak than they were before.

Question:  Purple Male with Purple Female developed well and purple Male with Orange female didn’t…some sort of incompatibility?

Well, given what I saw today the Orange (female) x Purple (male) cross almost certainly did not work. Fertilization occurred, but almost none of the embryos had any indication of normal development. Since we know the Purple male was able to mate successfully with the Purple female, we can infer that his sperm were fine. It could be that there was something going on with the Orange female’s eggs; there were a lot of them, but maybe their quality just wasn’t very good. Or perhaps we somehow mistreated and wrecked them the other day.

Any other questions? Use the Comments section to ask them, and I’ll address them in a future post.

Strangeness abounds

Posted on by Allison J. Gong

Wow, they weren’t kidding about “early developmental asynchrony” in sea stars! This morning I looked at the embryos that I had started almost 24 hours earlier, and noticed two things right off the bat:

Thing #1:  Within the F1 x M1 (Purple female x Purple male) mating , developmental rates among full siblings were all over the map. Some embryos had progressed to the blastula stage, which is essentially a hollow ball of ciliated cells, while others were still in the early cleavage stages and rather a lot hadn’t divided at all. In fact, with 24 hours of hindsight I can see that several of these eggs had not even been fertilized.

Embryos of Pisaster ochraceus, age 24 hrs. 3 June 2015. © Allison J. Gong
Embryos of Pisaster ochraceus, age 24 hrs
3 June 2015
© Allison J. Gong

My first reaction upon looking into the microscope and seeing all these assorted blobs was, “Oh, crap.” But then I looked more closely at some of the embryos and realized that they had become blastulae!

Here’s a picture of a blastula. This embryo is freely swimming inside its fertilization envelope, although it doesn’t have a lot of space (remember that narrow perivitelline space from yesterday? that’s all the elbow room it has). The hollow space in the center of the embryo is the blastocoel ‘sprout cavity.’ Given that the embryo hasn’t grown (or even hatched) yet, it’s still ~165 µm in diameter, the size of the original egg.

Blastula of Pisaster ochraceus, 3 June 2015. © Allison J. Gong
Blastula of Pisaster ochraceus
3 June 2015
© Allison J. Gong

The stage that precedes the blastula (a hollow ball of cells) is called a morula (a solid ball of cells). The embryo that is partially visible in the bottom of the above photo may be a morula. Imagine the following sequence of events: (1) an egg is fertilized by a sperm, forming a zygote; (2) the zygote undergoes a number of cleavage divisions, with the cells becoming more numerous and smaller in size; (3) at some stage a solid ball of small cells, the morula, is formed; (4) as cell division continues, the cells migrate toward the outside of the sphere, forming a cavity (the blastocoel) in the middle.

The blastula is a ciliated stage, and in this video clip you can see the cilia moving. I shot this video at only 100X magnification to capture as much depth of field as possible, and suggest viewing at full-screen. This should enable you to see the three-dimensional structure of the embryo, and that it is indeed a sphere.

Thing #2:  The F2 x M1 mating (Orange female X Purple male) isn’t doing well at all. I looked at several slides and didn’t see any embryos that were developing normally. They had all been fertilized, as I could see the fertilization envelope surrounding each egg, but most had not even divided. The ones that had divided were all strange and just plain wrong. Here, see for yourself:

24-hr embryos of Pisaster ochraceus, 3 June 2015. © Allison J. Gong
24-hr embryos of Pisaster ochraceus
3 June 2015
© Allison J. Gong

Many of the eggs are blurry because they’re below the focal plane of the microscope. But see how many of them are undeveloped? And how, in the ones that have started dividing, the cells are disorganized and of different sizes? Typical echinoderm cleavage, as I see in echinoids (our local urchins and sand dollars) and in my other crossing of these ochre stars, results in a blastula made up of cells that are all approximately the same size. Most of these embryos, on the other hand, appear to consist of one large cell and a bunch of tiny ones.

I assume that these abnormal-looking-to-me embryos will not hatch, although I could be pleasantly surprised tomorrow. I don’t yet have much of an intuition about these Pisaster ochraceus embryos, so this is a huge learning experience for me. I do expect to see hatching in the F1 x M1 cross tomorrow. Fingers crossed!