A few days ago I told my friend Brenna that I’d hunt around in the marine lab for a bit of a green alga that she wants to press. I had a pretty good idea of where to look, only the animals I’d seen it on had been removed from the exhibit hall. I asked for and got permission to examine the animals behind the scenes. And fortunately I had remembered correctly, and I was able to pick off some nice clumps of dark green stuff.
Bryopsis corticulans is a filamentous green alga. It grows to about 10 cm in length and is a dark olive color. When emersed it sometimes looks almost black. I’ve seen it in the intertidal in a few places, where at low tide it resembles nothing so much as a shapeless slime. It’s very difficult to see the beauty of organisms when they’re out of their natural element, which in this case is water.
One of the reasons I love the algae is their very inscrutability. I enjoy discovering the beauty of organisms that, at first glance, don’t look like much. Many of the filamentous algae, both the greens and the reds, have a delicate structure that requires close examination to be appreciated. Fortunately, I have access to microscopes, so close examination is very easy.
The thallus of B. corticulans is relatively simple, consisting of a bipectinate arrangement of filaments.
This is a shot of the main axis and side filaments. The small green blobs are chloroplasts. One thing to notice is that there are no crosswalls separating any of the filaments. That’s because the thallus is coenocytic, essentially one large cell with a continuous cytoplasm. Coenocytic cells are common in fungi, the red and green filamentous algae, and a few animals. In animals, coenocytic cells are often referred to as syncytial. They can arise in one of two ways: (1) adjacent cells fuse together; or (2) nuclear replication occurs as usual during normal mitosis but cytokinesis (division of the cytoplasm) does not. However the syncytium arises, it can result in very large cells. Even though B. corticulans itself is a small organism, some algae in the Bryopsidales consist of single cells that can be over 1 meter long!
Sometimes things that appear simple at first glance conceal a deeper complexity when you look more closely.
It has been almost three and a half years since I first documented seastar wasting syndrome (SSWS) in the lab. Since then many stars have died, in the field and in the lab, and more recently some species seem to be making a comeback in the intertidal. This circumstantial evidence may not be reason enough to conclude that the epidemic is over, but I think there is reason to be hopeful. Any disease outbreak eventually runs its course, and despite its death toll there are always at least some survivors. And I have an individual star that was very sick but seems to be recovering.
In September of 2015 one of my bat stars (Patiria miniata) developed the first tell-tale lesion of SSWS on its aboral surface. At the time the lesion was small (less than 10 mm in diameter) and superficial. Knowing that SSWS starts with minor symptoms and rapidly progresses to something horrific within a day or so, I wanted to keep an eye on this star. It held the same morbid fascination as a car accident or any other impending catastrophe.
By November 2015 the main lesion hadn’t grown much but a few others had developed. The star still wasn’t acting sick and was eating every once in a while, although it occasionally ignored the food that I offered.
So far, so good. I was thinking that the star doesn’t look too much worse, so maybe it wouldn’t keep getting sicker. I checked on it regularly, offered food a few times a week, and left it alone.
4 May 2016
Several months later I noticed that the first lesion had gotten much deeper. The outer dermal layers had been completely compromised, exposing the animal’s internal organs (gonad and digestive caecum) to the external environment. This was bad, very bad. Even in stars, internal organs are supposed to be internal, except when stars extrude their stomachs to feed.
This was the point in time when things started going south. The star lost the ability to maintain its internal turgor pressure and became lethargic and floppy. It stopped eating, or even responding to food. It spent most of its time in a corner of the seawater table where it lives, although a few times I saw it wrapped around one of the hoses that feeds the table. However, its body never started disintegrating the way I’d seen with other SSWS victims.
19 January 2017
Fast-forward another several months. About a month ago the sick bat star began perking up a bit when I placed food near the tip of one of the arms. A week later it actually wrapped its arm around the food, and I assume ate it. It has since been eating about once a week, after fasting for at least eight months. I began to think it would recover.
Today I had some time to photograph the star again, and it really appears to be doing much better!
The lesions are apparently healing over; at any rate, the internal organs are no longer exposed to the outside. The body margin between the arms has a few small divots, but they look superficial. Lately the star has been more active, too, cruising around the table instead of hunkering down in a corner. I’m going to keep feeding it to see if it continues to improve.
One of the most remarkable things about many animals with a decentralized nervous system, such as echinoderms and cnidarians, is their ability to regenerate lost parts and repair damage to their bodies. This bat star is a prime example. It has been sick for almost a year and a half now, and for at least half that time it hasn’t eaten. Yet it somehow had the metabolic reserves to heal a major wound to its body wall. That’s some astounding resilience there. I am very impressed, and you should be, too.
Although at this stage it’s a close race. Two and a half weeks ago I spawned sea urchins in the lab, setting up several purple urchin crosses with the hope of re-doing the feeding experiment that I lost this past summer when I was on the DL (that’s Disabled List, for those of you who don’t speak baseball). I was also fortunate enough to set up a hybrid cross, fertilizing purple urchin (Strongylocentrotus purpuratus, or “Purp”) eggs with red urchin (Mesocentrotus franciscanus, or “Red”) sperm. I would have done the reciprocal hybrid cross (red eggs by purp sperm) as well if I’d gotten any of female red urchins to spawn. However it wasn’t really spawning season for the reds, and I consider myself lucky to have persuaded that one male to release some sperm for me.
This is the first time that I’ve tried to raise the hybrid larvae, although I know it can be done because my colleagues Betsy and John did it many years ago, before I came to the marine lab. All of my larvae are the exact same age and are being raised side-by-side, so I can make direct comparisons between the Purp by Purp crosses and the Purp by Red hybrids. Incidentally, when speaking or writing about a hybrid cross the convention I’ve adopted is to reference the female parent first, so when I say Purp by Red I mean a Purple eggs fertilized by Red sperm. A Red by Purp hybrid would logically result from red urchin eggs fertilized by purple urchin sperm.
My experience raising sea urchin larvae is that things almost always go well for the first 48 hours or so; most (but not all) of the fertilized eggs develop into embryos and undergo the crucial processes of gastrulation and hatching. In some cultures the hatching rate is close to 100%. After that there’s a window of 3-4 days when cultures can crash for no apparent reason, although food availability or quality may be a factor. If the larvae make it past their first week of post-hatching life they generally cruise along until the next danger period which occurs at about 24 days. I change the water in the culture jars and observe the larvae under the microscope twice a week.
Today the larvae are 18 days old. It’s a little early for that second mortality period, but some of the Purp by Purp cultures never really took off. The larvae don’t seem to be growing or developing as quickly as I’m used to. Perhaps this has to do with lower water temperatures, especially after the prolonged period of high temps in 2014-2015. In any case, two of the four Purp by Purp crosses are doing well and the other two are just hanging in there.
There are two things I can see with the naked eye that give me a heads-up when cultures are crashing: the first sign is an accumulation of debris at the bottom of the jar and the second is an absence of larvae in the water column. The debris can be due to excess food, a build-up of fecal matter (not usually the case, as I’m pretty good at doing the water changes on time), the disintegration of larval bodies, or some combination thereof. If the water column is clear then the culture has already crashed and everybody is dead.
Today one of my jars had crashed. The water column was very clear and there was a lot of fluff at the bottom of the jar. I’d been wondering if I could figure out what the fluff was made of, so I sucked up a bit in a pipet and examined it under the microscope. I thought I’d see dead algal cells or pieces that look like defecated algal cells. This is what I saw:
Silly me. I had forgotten that the corpses of pluteus larvae would disintegrate pretty quickly, leaving behind only the skeletal rods. The rods get all tangled together and trap the organic stuff, which is probably a mixture of uneaten and defecated algal cells and the soft tissues of the larval bodies. This explains the clear water column in the jar.
While the Purp by Purp larvae have had mixed success so far, the Purp by Red hybrids have been doing well. From the outset they appeared to be more robust than the Purps, and even though the fertilization rate was only about 50% the post-hatching mortality seems low. The hybrid larvae are also larger than the Purps, and are developing more quickly. In the two photos below the scale bar indicates 100 µm.
The hybrid larva is about 10% larger than the Purp larva. Other than that they look similar, but to me the hybrid larva seems farther along the developmental process: its arms are proportionally longer and have a more mature look (although I don’t have any way to describe that to a naive observer). There’s something about the gestalt of the animal that makes me think it’s more robust than the Purp individual.
We’ll see how the pure Purps and the hybrids do from here on. I actually have the Purp larvae divided up into different feeding treatments, which I may discuss in a future blog post. In the meantime I’m trying to baby the hybrid larvae as much as possible, to maximize their probability of successful metamorphosis in six weeks or so.
Sea urchins have long been among my favorite animals. From a purely aesthetic perspective I love them for their spiky exterior that hides a soft squishy interior. I also admire their uncanny and exasperating knack for getting into trouble despite the absence of a brain or centralized nervous system. Have you ever been outsmarted by an animal without a brain? I have. It’s rather humbling.
Red sea urchins (Mesocentrotus franciscanus) and purple sea urchins (Strongylocentrotus purpuratus) share a common geographic range along the northeastern Pacific but generally live in different habitats. S. purpuratus is the common urchin in tidepools, while reds are almost always subtidal (although I have seen them in the intertidal on very low minus tides). The two species’ habitats do overlap a bit, as the purple urchin can live in subtidal kelp forests alongside the reds. There is a commercial fishery for the gonads of red urchins, which are prized as uni by sushi aficionados. I’ve tried uni once, and it tasted exactly the way I imagined the gonads of a sea urchin would taste. Not a fan. I’d much rather make a different use of urchin gonads.
The other week I collected some urchins from the field, hoping that they’d have nice full gonads. Gametogenesis in many marine invertebrates, including sea urchins, is governed at least partly by annual light cycles. Provided they have sufficient food, purple urchins have ripe gonads and spawn in the winter, from December through March. Reds spawn in the spring, from March through June. In my experience the best time to induce spawning of purps in the lab is December or January, when the urchins have developed gonads but likely haven’t spawned yet. There is no way of knowing the sex of any given urchin or the condition of its gonads, so this exercise is somewhat of a crap shoot even with the best of planning.
Today I shot up my eight field-collected purps, hoping to get at least one male and one female out of the deal. I got lucky with the timing, as one of the smallest urchins was a female and began spewing out eggs. This little female gave a lot of eggs! She was followed by three males and two more females. So out of my eight purps I ended up with three of each sex, and a spawning rate of 75% ain’t bad.
I set up some mating crosses and fertilized all of the eggs. I divided the little female’s eggs into two batches and fertilized them with the sperm of two different males (M1 and M2). Each of the other females’ eggs was fertilized by M1, who gave huge amounts of sperm. When I checked on the eggs about two hours post-fertilization most of them had gone through the first cleavage division and seemed to be developing normally and on schedule.
Just for the hell of it I decided to shoot up some of the red urchins we have in the lab. I didn’t really think they’d spawn, as it’s not the season for them to be gravid. Red urchins are large, heavy animals with long and sharp spines and they are much more difficult to handle. Four of the five that I shot up did nothing, as expected. It took a long time, but just as I was about to give up on them the biggest red began dribbling out a couple thin streams of sperm. I examined the sperm under the microscope and they were very active and healthy. Fortunately I hadn’t returned the purps to their tanks, and two of the female were still putting out some eggs. I rinsed the purp eggs into a clean beaker, pipetted up some of the red sperm, and added it to the eggs.
Sea urchin eggs are covered by a thick jelly coat. In the video you can see many of the red urchin sperm embedded in the jelly coat of the egg. Despite the frantic activity of the sperm, fertilization (as evidenced by the rising of the fertilization envelope off the surface of the egg) took much longer than it does when eggs and sperm come from the same species.
Look at that beautiful zygote! Fertilization success in this hybrid cross was low, only about 50%. The eggs that did get fertilized went through the first cleavage division after about two hours later, which is right on time.
It remains to be seen whether or not the few hybrid embryos I have continue to develop. I have a colleague who has hybridized red and purple urchins successfully in the past, and has raised the offspring to adulthood. I don’t have any expectations of great success with this little experiment, but it would be very informative to raise known hybrid urchins. I’ve seen animals in the field that look like hybrids and there’s no reason to assume that hybridization between these two free-spawning species never occurs. The adults can be found living side-by-side subtidally, and there’s enough overlap in their reproductive seasons that some individuals of each species could very well spawn at the same time. On the other hand, hybridization that can be forced in the lab doesn’t necessarily occur in the field. I dumped a lot of red urchin sperm on those purple urchin eggs, and such high sperm concentration may overcome any mechanisms of reproductive isolation that exist under real-life conditions.
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.
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).
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.
About three weeks ago I collected some mussels from the intertidal, to use both in the lab and in the classroom. A mussel can itself be an entire habitat for many other organisms. Many of the mussels I brought into the lab this last time were heavily encrusted with barnacles and anemones. I wanted to look more closely at one of the anemones so I took the mussel to the microscope. And, as often happens when I look at stuff under the microscope, I got totally distracted by things other than what I intended to study.
But for the record, this is the anemone that started the whole chain of events:
Below the anemone there’s a thick mat of small acorn barnacles (Balanus glandula) and a couple of leaf barnacles (Pollicipes polymerus). They were all alive when I brought the mussel into the lab, and over the weeks a few of them have died. But many of them are still kicking, both figuratively and literally.
Barnacles are most strange animals. Believe it or not, they are crustacean arthropods, somewhat closely related to crabs and lobsters. They live encased within a shelter of calcareous plates, which they can close seal up against predators and desiccation. I’ve never figured out why they are called “acorn barnacles,” as they don’t look anything like acorns to me, but in Balanus and such the base of the shelter is glued directly to a rock or some other hard surface. Leaf barnacles are shaped very differently, and have a fleshy stalk between the shelter that houses the main body of the animal and the rock surface.
To picture what’s going on with a barnacle, imagine a shrimp lying on its back, then curl it up and stick the whole thing inside some calcareous plates. The thoracic appendages would be facing up. In barnacles the thoracic appendages are modified to be clawlike feeding structures called cirri. Barnacles are filter-feeders, collecting particles from the water by maneuvering the cirri in a sort of grasping fashion. So in a nutshell, or more precisely a test, a barnacle lies on its back and kicks its legs out to catch food.
Here’s what B. glandula looks like when feeding. Note the clearly jointed cirri, with fine hairs that help catch particles. The cirri can be controlled independently, as you can see when they flick towards the center, and the entire apparatus can be rotated quite a bit.
Same deal with Pollicipes.
So that’s the feeding part. A little strange, but not as interesting as the barnacles’ sex lives. Let’s start with some background about sexual function. And get your mind out of the gutter; this is real science stuff! Most of the animals that you’re familiar with are described as dioecious (Gk: “two houses”). This means that female and male sexual functions are segregated; in other words, there are male bodies and female bodies. Other animals are described as monoecious (Gk: “one house”), so that a single body has both female and male sexual function. Monoecious animals could also be described as hermaphroditic. Some monoecious animals have male and female function in a single body at the same time; we call these simultaneous hermaphrodites. If a body first functions as one sex and then either acquires or switches to the other sex, we say the animal is a sequential hermaphrodite. Many fishes, including the California sheephead and the anemone fishes of coral reefs, are sequential hermaphrodites. Make sense?
Barnacles are simultaneous hermaphrodites. If you dissect an adult barnacle you will find mature ovaries and testes. This means that every barnacle can be both a mother and a father. The logical assumption is that monoecious animals should just fertilize their eggs with their own sperm. . . however, this generally isn’t the case. The whole point of sexual reproduction is to combine the genomes of two individuals, and self-fertilization obviously doesn’t accomplish this. So even though there are many hermaphroditic animals, very few of them are self-fertile.
One other weird thing about barnacles, and crustaceans in general, is their sperm. Arthropods have non-flagellated sperm, which means they don’t swim (although some of them have amoeboid sperm that can ooze around a little bit). Many marine animals reproduce by broadcast spawning; that is, by throwing their gametes into the water, where fertilization takes place. Fertilization is facilitated by the sperms’ ability to swim towards conspecific eggs.
Barnacles, with their non-swimming sperm, generally cannot rely on broadcast spawning to get sperm to egg. They must copulate. How do you suppose they do this? The same way that other animals (e.g., Homo sapiens) copulate, by using a penis or some other structure to transfer sperm from one individual to the body of another. In barnacles the penis’s technical name is intromittent organ. The penis is inserted into the test of a neighboring barnacle and sperm is delivered. The receiving barnacle uses the sperm to fertilize its eggs. Unlike the cirri, the penis is unjointed and flexible, the better to seek out and slip into potential mates. You can see the intromittent organ unrolled and poking around.
Now think about the ramifications of these constraints. Barnacles live their entire post-larval lives permanently cemented to a rock. They also have non-motile sperm so sperm transfer can occur only by copulation. If the key to reproductive success is to mate with as many other individuals as possible, what do you suppose natural selection has done? That’s right: barnacle anatomists, including the great Charles Darwin himself, have noticed that barnacles have incredibly long penises. In fact, compared to overall body size, barnacles have the longest penises in the animal kingdom, up to 15 times the length of the body! That’s what you call bragging rights. Not all barnacle species are so amply endowed, however. The same leaf barnacle that I observed today (P. polymerus) has recently been reported to be a spermcaster; their penises are shorter than body length, and they release sperm that are captured by their downstream neighbors.
My friend Peter Macht is the aquarium curator at the Seymour Marine Discovery Center. He is responsible for all of the live (i.e., wet) exhibits and has a team of student and volunteer aquarists who help him care for the animals in the hall and behind the scenes. Peter and I go way back together, to years before the Seymour Center opened in 2000. Back then the only public space at Long Marine Lab was called the Shed Aquarium because it was, literally, in a wooden shed. I do miss the marine lab the way it was then, when I knew everybody who was there and it was a quieter and more peaceful place to work. However, we’ve come a long way, baby, and the Seymour Center is in just about every way imaginable, a huge improvement over the Shed Aquarium.
For one thing, there are two large exhibits in the Seymour Center, each of which would occupy about half the volume of the old Shed Aquarium. One of these tanks, the Sandy Seafloor tank, has housed many different animals over the years: surf perches, sand dabs, sharks, rays, señoritas, and various invertebrates. My personal favorite continues to be the burrowing sea star Astropecten, although she hasn’t been on exhibit for several years now. The current inhabitants are a close second favorite, even though when they first arrived I didn’t expect them to be nearly as fascinating as I’ve found them.
Pleuroncodes planipes is a little red crab commonly called the pelagic crab or the tuna crab. For once the common names reveal something about the biology of the animal–these crabs spend their lives in the water column over the continental shelf, at least as youngsters, and are one of the favored food items of tunas. They are usually found in the waters of southern California and Mexico, but during the El Niño event of 2015 they washed onto the beaches around Monterey Bay in humongous numbers; they also did so during the ENSO event of 1982-1983.
Although they resemble crayfish, Pleuroncodes is a crab. They are anomuran crabs more closely related to hermit crabs and porcelain crabs than to “regular” brachyuran crabs such as shore crabs and rock crabs. The way you tell the difference between anomuran and brachyuran crabs is to count the number of thoracic walking legs, keeping in mind that the claws are included as walking legs: anomurans have four pairs while brachyurans have five pairs. You can see in the picture of the lateral view that this crab has three pairs of stick-like legs and one pair of chelipeds (claws).
Being arthropods, red crabs molt periodically. Peter has been collecting data on frequency of molts for individual crabs since the spring of 2016. Doing so requires isolating crabs in separate containers, to keep track of which crab molts when and also to prevent the crabs from ripping apart a freshly molted compadre, which they do with great enthusiasm. It is not unusual to see one or more of the inhabitants of the Sandy Seafloor tank missing a leg.
Here’s one of Peter’s tables containing crabs in baskets:
It’s just as well that these guys have extraordinary regenerative capabilities, as they are eager to rip each other’s legs off. With most crabs that I’ve observed in the lab limb regeneration is a gradual process, with the new leg growing a bit with each successive molt. Chelipeds, even with their increased size and complexity, seem to regrow faster than the other walking legs, likely reflecting their importance to the animal’s lifestyle.
Peter told me last week that he’d seen one of his isolated crabs regenerate an entire cheliped with a single molt, going from nothing to an almost-full-size functional limb essentially overnight. This seemed very unlikely to me, but Peter said he’d seen the before (the empty molt) and after (the actual crab) together in the same container. Unfortunately the crabs end up demolishing and eating their molts within a couple of days, so the evidence doesn’t stick around very long.
Sometimes, though, you get lucky. When I was at the lab yesterday morning Peter told me that he’d seen another of his crabs molt, and that it had grown a missing cheliped since the previous day. And this time he could show me the proof. Voilà!
Note that the molt has only one cheliped, the left, while the crab itself has two. How cool is that? The crab’s right cheliped is a bit smaller than the left, as might be expected of a regenerating limb, but it’s definitely intact and functional. It was pretty exciting to see evidence of wholescale limb regrowth taking place in such a short period of time, which must be incredibly energetically expensive. On the other hand, chelipeds are extremely important for defense, and there is obvious selective pressure to regrow them as quickly as possible should a crab be unfortunate enough to lose one.
Peter gave me permission to examine the molt more closely, so I took it back to the lab where the lighting is better. And surprise! The right cheliped apparently didn’t grow from nothing overnight. If you look really hard at the photo above, you can just barely see a ghostly transparent sheath where the missing arm would be. Hmm. This was not at all what I expected. Did I really see that?
It turns out that, yes, that is exactly what I saw.
See that translucent tiny limb up front? That’s a little cheliped! And it had been there at least six months, as this crab’s last recorded molt was in April. Why hadn’t anyone seen it before? I think because this limb was so small that the crab kept it tucked underneath the carapace, where it wouldn’t be seen from the dorsal (top) side.
In the course of one morning I got taken for quite a roller coaster ride. Peter reminded me that he’d seen a crab apparently regrow a missing appendage in a single molt cycle . . . and had just found a crab whose molt showed exactly that . . . and then that molt ended up including a claw after all. What fun!
Now, why is that little claw so transparent? An arthropod’s exoskeleton is made of a material called chitin, with varying degrees of calcification depending on species. The large marine crustaceans (e.g., crabs and lobsters) have heavily calcified exoskeletons, while insects have much more lightweight, less-calcified exoskeletons. As a crab prepares to molt, one of the things its body does is resorb some of the minerals that it had deposited in the soon-to-be-discarded exoskeleton, so they can be re-used in the new one. If you find a discarded molt on the beach, pick it up and note how little it weighs; you’d be surprised at how flimsy it is.
Here’s my hypothesis. I think that this little cheliped, because it was newly regenerated before this most recent molt, was only lightly calcified. The crab may have used it, but it wouldn’t have been much use for defense. Then, the next time the crab molted the claw was shed along with the rest of the exoskeleton, and the limb was significantly larger. This crab now possesses a complete pair of chelipeds again. After examining the molt I returned it to the crab, which has probably torn it to pieces and eaten already. It’s a way for the animal to recover some of the nutrients it allocated into building the exoskeleton in the first place.
This past weekend I was trying to manage some concussion headache issues and stayed away from the marine lab for four days. Usually that’s not a big deal. Since I’ve been absent so much of the summer due to the head injury, the lab assistants whose job it is to make sure that everybody has air and water and food have been told to check my stuff and change water daily. They’ve been keeping things alive when my headache wouldn’t tolerate my being at the lab, and I’ve gone in when I could (usually on weekends) to take care of the big chores. And so far, under normal conditions at the lab, this has worked.
But every so often conditions stray from the norm, and we are in one of those situations now. It isn’t uncommon at this time of year for us to experience an algal bloom in Monterey Bay. This isn’t the sort of spring phytoplankton bloom we get in the upwelling season, but a massive population explosion of a single species, usually a dinoflagellate. This kind of algal bloom is referred to as a “red tide,” even though the organism that causes it isn’t so much red as golden.
I went to San Francisco yesterday afternoon, and the water was brownish like this all the way up the coast. The bloom wasn’t evenly distributed; there were large patches of brown water interspersed with areas of clear blue water. At Scott Creek and Waddell Creek the breaking waves were distinctly tea-colored, which did not keep the kite surfers out of the water.
It might be easier to see the discoloration when the water is moving:
The seawater intake for the entire marine lab is straight off the point here in the surf zone, so this mucky water is the exact same stuff that’s trickling through our labs. When I returned to the lab on Monday after a 4-day absence the first thing I noticed when I opened the door was the smell, which I recognized immediately because we get red tides like this every year or so. It’s not really a horrible smell, like the smell of dead sea things, but it gets classified in my mind as ‘bad’ because of what it connotes. And it can get really bad, if the gunk accumulates and begins to rot.
When the cell concentration is this high, filter apparatuses get clogged up fast. This applies to both mechanical and biological filters. Unlike, say, small sediment particles that get suspended in water but act more or less independently of each other, the cells of these blooming dinoflagellates are sticky. They glom together in stringy mucilaginous masses, and tend to settle out in little eddies and areas with less water movement. When this muck settles on animals’ bodies, it can clog up gills or other respiratory surfaces, making gas exchange difficult or impossible. So while the red tide persists we siphon out tanks and flush tables at least once daily.
I guess when you see the color of these masses of cells, it makes sense to call this phenomenon a red tide. Under the microscope, however, the cells are golden. Based on the guilty party of the last big red tide event we had and some sampling data from Santa Cruz and Monterey dated 7 September, I’m pretty sure the cells are Akashiwo sanguinea. The cells are fairly large by dinoflagellate standards, ~100 µm long, and have the usual pair of flagella (1 wrapped around the middle and the other trailing free) that propel the cells through the water.
The groove around the middle of the cell is called the cingulum; one of the cell’s flagella sits in this groove like a belt going around your waist. The other indentation that runs from the cingulum to the posterior end is the sulcus, and houses the other flagellum that trails free like a very skinny tail. The beating of this pair of flagella causes the cell to swim in a spiral fashion:
People always want to know if a red tide is toxic, and if they need to stay out of the water. Akashiwo sanguinea, as far as anybody knows, does not produce toxins like some other dinoflagellates do. However, it does secrete surfactants that produce foam in agitated water, and a report from 2007 correlates a mass stranding of seabirds in Monterey Bay with a large bloom of A. sanguinea. The authors hypothesize that the foam from the surfactants of A. sanguinea coated the feathers of seabirds and hindered their ability to thermoregulate.
This afternoon I am heading out to the intertidal. One of the things I’ll be looking for is signs of the bloom. I do want to take some pictures in the tidepools, so I hope the discoloration isn’t too bad. Fingers crossed!
Seeing as today is the third anniversary of the first blog post I wrote about sea star wasting syndrome (SSWS), I thought it would be appropriate to take inventory of my remaining stars and see how they’re doing. Right now I have custody of ~10 bat stars (Patiria miniata), 7 ochre stars (Pisaster ochraceus–collected last year for the juvenile survival experiment I did with Scott), and 1 Mediaster aequalis. For whatever reason the M. aequalis hasn’t been affected by SSWS so I’m going to disregard it for now. Of the 10 or so bat stars, four live in one of my seawater tables, roaming free-range in quite a large volume of water. The other half-dozen or so live in a tank in a different building. The Pisasters live in 1s and 2s in tanks distributed in two rooms in the same lab.
After the initial horror and shock of the spectacular onset of SSWS, in which we watched stars rip themselves into pieces right before our eyes, what we’ve seen has followed the standard epidemiology pattern. Any time a novel pathogen enters a population, the individuals that have no immunity or resistance are the first to die. The disease spreads rapidly through the population, wiping out all of these weaker individuals. However, not everyone dies. Even during the Black Death of the 14th century, the very fact that 1/3-1/2 of the human population died of bubonic plague means that 1/2-2/3 survived. Those survivors presumably had some degree of resistance to the disease.
At the same time three years ago that all of my forcipulate stars died, divers were noticing similar phenomena happening subtidally. It didn’t take long for us to realize that Something Big was going on, which was eventually dubbed SSWS. Fast-forward three years and now I’m seeing healthy, hand-sized P. ochraceus in the intertidal again. These individuals are certainly survivors from the SSWS outbreak; they were likely small juveniles during the plague, and were able to come out of hiding and expand into open niches after so many of the adults died. Whether or not natural populations will recover completely remains to be seen, but as of right now things look promising.
About a year ago, having gone two years without showing any signs of being sick, one of my bat stars developed lesions on its aboral surface. It’s the red star in the middle of that blog post. This star is one of the four that live in my shallow table. It has now been sick for a year. See how it has changed since then:
The lesions have all gotten worse–the largest is about 2 cm long now–and the body margin has some ripples that it didn’t have before, but the star is still alive. For a while it wasn’t eating, as far as I could tell, but two days ago I watched it eat a piece of fish. Perhaps the return of cooler water is helping this animal survive.
One of its tablemates, however, hasn’t been so lucky. I first noticed apparent SSWS damage in a second star several months ago. Today was the first chance I had to look closely at it.
The most noticeable injury to this star is that big interradial divot. It looks like someone took a bite out of the body at that spot. The margins of the wound are white and fluffy, similar in appearance to the lesions caused by SSWS.
For years now this star has had an abnormal spot on its aboral surface. I’ve been calling it a bubble, for lack of a better word. The bubble may be an over-inflated papulla (skin gill) and it didn’t seem to be causing any problems for the star. I’d touch it and it would deflate, then re-inflate almost immediately. When I touched it today, it shrank back a little but didn’t really deflate.
If you look really closely at the above photo, you can make out clusters of small, clear, clublike projections. These are papullae, extensions of the internal body lining that project through the skeletal ossicles to the outside and act as gas exchange surfaces. The bubble is many times larger than the normal papullae. Because it has been there for so long, years before the divot in the interradial margin, I don’t think the bubble is due to SSWS. I don’t even know if it’s a wound, or merely an overinflated papulla. The largest star in this table has also had a bubble for many years, but no lesions or wounds indicating SSWS or other disease.
So. Three years after the outbreak of SSWS I still have stars that are sick. They’ve been sick for a long time and aren’t getting worse very quickly, from which I conclude they may eventually recover. At the very least they must have some resistance to the SSWS pathogen because they’ve managed to survive so far. One more thing. Way back in 2013 when all of the forcipulates were tearing themselves into pieces and melting into piles of goo, these bat stars were among them, scavenging on the dead and decaying tissue. For a while I feared that eating contaminated tissue might cause the disease, but that doesn’t seem to be the case, as these two didn’t get sick until two years after the initial exposure.
I hope these two stars make it. Cooler water temperatures should help. When they’re really sick they stop eating (they haven’t eaten much in the past year) but if they’re going to eat now I’ll keep feeding them. Fingers crossed!
Spending the summer trying to heal a concussion brain injury means that not much science has been happening in my life lately. Now three months post-accident, I’m finally able to do a little bit of thinking and am not quite as exhausted as I was, although extended periods of concentration are still taxing and usually result in what I’ve come to call the concussion headache. I’m very disappointed to have been on the DL (disabled list) for most of the summer intertidal season, and hope that when the afternoon minus tides return this fall I’ll be able to take advantage of them. Fortunately my condition has progressed to the point that I can drive myself out to the wharf to collect a plankton sample and spend a couple of hours looking through microscopes at what I’ve caught. That’s about the limit of what I can do these days; it’s not much, but at least it’s something.
As we approach the autumn equinox I would expect to see signs that the summer growing season is winding down. Days are noticeably shorter than they were a month ago, and the major upwelling season has passed. In terms of plankton, this should mean a reduction in phytoplankton abundance and diversity, with an overall shift in population makeup away from the strictly photosynthetic diatoms and favoring dinoflagellates, many of which are at least sometimes heterotrophic.
The water at the wharf is remarkably clear right now. Visibility would be fantastic for anyone who wanted to dive under the wharf. September and October tend to be the best months for SCUBA diving in Monterey Bay because the natural cessation of coastal upwelling results in clearer and warmer surface water. I didn’t have a Secchi disk or any other way to measure turbidity, but judging by how far below the surface I could see the plankton net as it sank I’d guesstimate that visibility was about 7.5 meters. For people used to diving in the oligotrophic waters of the tropics this level of visibility is downright awful, but for those who dive in productive areas this is not bad.
As expected, when I pulled up the net there wasn’t much phytoplankton in the net, and none of the diatom smell I get in spring plankton tows. The net came up pretty clear and rinsed easily into my jar. There was, however, a lot of zooplankton. When I got back to the lab I started looking through small aliquots to see what was there.
The usual suspects were quite plentiful. These included:
copepods, in both larval and adult stages
polychaete worms
veliger larvae, of both gastropod and bivalve types
medusae from the hydroid Obelia sp.
tintinnids, a type of protozoan that lives in a goblet-shaped glass shell
echinopluteus larvae, probably of the sand dollar Dendraster excentricus
Especially beautiful in today’s sample were the acantharians:
Acantharians are large single-celled protozoans; I’ve seen some that are 3 mm in diameter. They build spines of strontium sulfate, which are arranged in precise geometric formations. The protoplasm of the cell extends partway along the spines, which are thought both to deter predation and provide buoyancy. Acantharians are predatory, feeding on smaller unicellular organisms, but also form symbiotic relationships with unicellular algae. The algae are given safe harbor within the cell of the acantharian, and in return provide fixed carbon to the protozoan. Although the players are different, this is pretty much the exact same symbiosis as occurs between reef-building corals and zooxanthellae in the tropics (and also between some of our temperate sea anemones and zooxanthellae).
Here’s a puzzle for you. Take a look at this pair of animals:
Both consist of a roundish body and a tail. The one on the left is much larger, about 5 mm long, and more opaque. The one on the right is about 2 mm long and is very transparent.
Question: Do you think these animals are the same thing?
Answer: It can often be a mistake to assume any close evolutionary relationship between animals that appear to share a morphological similarity, but in this case shape does result from genetic relatedness. Both of these animals are chordates, my (and your!) closest invertebrate relatives. Yes, we share a closer kinship to these critters than we do to any other invertebrates. We also share with them the following morphological characteristics: pharyngeal gill slits, a dorsal hollow nerve cord, a notochord, and a post-anal tail. Of course, for us the gill slits, notochord, and tail are gone long before we are born, but if you look at pictures of human embryos you can see them. Once we are born the only chordate characteristic remaining to us is the dorsal hollow nerve cord, which runs up through our vertebral column.
The animal on the left is called a tadpole larva, probably of one of the benthic solitary or colonial tunicates. Tadpole larvae are short-lived and lecithotrophic (i.e., non-feeding); the opacity of the body is an indicator of energy reserves stored in body tissues. Tadpole larvae have a short larval life. They typically don’t disperse far from the parent, and within a few hours metamorphose into new tunicates.
The animal on the right is a larvacean. It bears a superficial resemblance to the tadpole larva, but is an adult. Larvaceans are entirely planktonic and have one of the most interesting lifestyles imaginable. They live in a house of snot. The house is secreted from an area on the back of the animal, and is inflated as the animal pumps its tail up and down in a rhythmic sinusoidal fashion. The mucus house actually consists of two distinct meshes: the outer mesh is coarse and serves to keep large particles from clogging up the finer feeding mesh. The feeding mesh collects very small particles, which are transported in a mucus thread to the animal’s mouth.
Larvacean in its mucus house.
Larvaceans are prodigious mucus makers. As any filter does, the house eventually clogs up. Instead of trying to backflush and clean out its house, the larvacean wiggles out of it and secretes a new one. They can build up to three houses a day when the water is full of plankton! The discarded houses of countless larvaceans slowly sink from the surface and are a major source of food to animals in the deep sea.
Larvaceans caught in a plankton net are almost always dislodged from their houses. In a dish or a drop of water on a microscope slide, they thrash about in a characteristic larvacean sort of way. Only once have I caught a larvacean and then been able to watch it build a new house in my dish of water. What I saw today is much more typical.
This poor animal was trapped under a cover slip so it can’t move freely, but the tail still thrashes about. You can also see its little heart beating like mad.
The tadpole larva, on the other hand, is a much more sedentary creature. It doesn’t disperse far so its tail remains still, and its heart rate is much slower than that of its pelagic cousin:
To shift to a completely different taxon there were, as usual, many crustaceans. In addition to the larval and adult copepods, today I saw several examples of Podon, a type of crustacean called a cladoceran. The most familiar cladocerans are the freshwater Daphnia species, but in Monterey Bay we see Podon on a fairly regular basis. Cladocerans reproduce via parthenogenesis, in which unfertilized eggs develop into daughters, and in the springtime most of the Podon I catch are gravid. At this time of year, however, they are not reproducing, at least not parthenogenetically.
The most striking feature of Podon is its large compound eye, which causes problems. For many creatures living up in the water column, the only way to hide is to be transparent. This invisibility would be interrupted by any pigment in or on the body. Unfortunately for Podon and other animals that try to hide in plain view, eyes are, at bare minimum, a collection of pigmented cells that detect light. For them, eyes are both a useful sensory structure and a big “Here I am!” signal for predators.
The best thing I saw in today’s sample, aside from the acantharians, was a small ciliated blob with little ciliated flaps. This cute little creature is the Müller’s larva of a polyclad flatworm. It’s hard to appreciate the cuteness of Müller’s larva in a 2-dimensional still shot, so here’s a video:
Okay, so maybe it’s not the cutest larva in the plankton. It was swimming really fast and I had to squash it a bit under a cover slip to slow it down enough that I could keep up with it. But I don’t come across them very often, so it’s always a pleasure when they show up. You’ll have to take my word that they’re cute.
Oh, and by the way, I kept the tadpole larva and a couple of other shmoo-like larvae in a dish of seawater to see what they will turn into. Tomorrow I may have new things to look at. I dumped the rest of the plankton into a tank of filter-feeders, where they will resume their place in the food chain.
