Sea urchin embryology lab

This document was prepared for a school-teachers' workshop. Others may find it useful too. This advice is specific to species available at Friday Harbor Labs.

Note: everything, including the spawning, has to take place within the temperature comfort zone for the species you're using. For the local urchins, if shallow seatables are not available, then a refrigerator set to about 10 C is ideal. Materials to be used, including seawater and culture-ware, should be chilled to this temperature in the fridge in advance. Sand dollars are much more temperature-tolerant (they live in the intertidal zone), and you may be able to do everything at room temperature.

How to get gametes from urchins

The most common method is to inject potassium chloride (KCl) into the body cavity. The injected solution is 0.55 M KCl in distilled water. You need 1-2 ml per urchin, less than 1 ml per sand dollar. There is no need to sterilize it.

KCl stimulates the gonad wall to contract; ripe gametes emerge from the gonopores surrounding the anus on the aboral (up) side of the animal. Use a syringe and 18 gauge or smaller needle (22 gauge is ideal for both urchins and sand dollars). Insert the needle, angled radially away from the mouth, into the edge of the disc of soft tissue surrounding the mouth. Don't force it. Go straight in about 1 cm or so and inject slowly, no more than 2 ml (1 ml is usually sufficient), making sure not to inject any bubbles. Sand dollars initially seem trickier because there is so little space in the body and it's hard to find the soft spot around the mouth, but with a little practice they are just as easy. Insert the needle near the mouth, then tip it to a steep angle, and insert further until you feel the firm test on the opposite side. You should inject less: 0.5 ml is usually adequate for a medium-sized sand dollar.

Injected animals may not begin shedding immediately. Give them a bit of a shake, and then set them down dry (sprinkle them with seawater occasionally) until you see something coming out of the gonopores. Be patient; some animals take up to 10 min. to spawn, and some just aren't ripe. Often not all the gonopores emit gametes. If it's milky-white, it's sperm. Either use a glass pipette to transfer dry semen from the top of the spawning male to a test tube (sand dollars usually don't make enough semen to allow this), or invert him over a shallow dish with just enough water to cover the gonopores. If it's yellow, it's eggs. Invert the spawning female over a small beaker (that is, smaller than the urchin!) filled to the brim with chilled seawater. She will release eggs in streams and clumps which accumulate on the bottom. When the animals are done spawning, or when they've provided enough gametes, return them to a tank — BUT: not to the tank with all your other animals, because they may be induced to spawn by the presence of gametes in the water.

Eggs don't usually remain viable for more than 6-8 hours unless drastic measures are taken (like using antibiotics, etc.). Semen that has been collected dry or nearly dry may last until the next day if kept at 4 C

Once you have some urchin gametes...

Leave the unfertilized eggs in a settled mass until you need them. Usually you will get vastly more eggs than you can handle, if the animals are ripe. Using a glass Pasteur pipette, and without disturbing the settled mass, transfer 1 ml of settled eggs to a new beaker with 100 ml chilled seawater. For urchin eggs, stir them a bit to rinse clumps apart, and if time allows let them settle and give them a rinse by pouring off and replacing most of the seawater. DON'T rinse sand dollar eggs; just transfer them over with as little swirling as possible.

Meanwhile, you need to dilute the sperm. A good target is to prepare a 1:10,000 dilution – e.g., 10 microliters in 100 ml chilled seawater. Stir it around a bit. Before fertilizing the eggs, make sure the sperm are motile. Place a drop of sperm suspension on a slide without a coverslip, and examine it with a 20x lens. You may need to close the condenser aperture to see the sperm. Don't be alarmed if most of the sperm are just sitting there; sperm often stick to glass, and furthermore peptides released by eggs stimulate motility. You should see that at least 10% are swimming vigorously. The diluted sperm will only be good for maybe a half hour or so.

Note: keep pipettes for sperm and unfertilized eggs separate! Color-coding with tape helps.

Add 1 ml (or one Pasteur pipette-full) of dilute sperm to the settled eggs, and stir them up. You can immediately begin checking for fertilization either by making clay-feet preps (see below) or by pouring a few mls into a dish to examine on a dissecting scope (you will need a pretty good dissecting scope, though). What you are looking for is the elevation of the fertilization envelope. If the sperm and eggs are both in good condition, you should see that 90% of the eggs have an elevated envelope within 5 minutes.

Once the eggs are fertilized, let them settle. Give them an occasional rinse by pouring off most of the seawater and replacing it. If your cultures are too dense they won't develop normally. For early development, aim for a monolayer of embryos on the bottom of a beaker. That is plenty to observe development and raise larvae. Once the larvae hatch out of the fertilization envelope, they must be diluted still further. Once they are actively feeding (see below for feeding instructions), they should be kept at approximately one larva per ml.

Clay-feet preps

To observe embryos on a compound microscope, make a chamber on a slide by supporting a glass coverslip at its corners with little feet of modeling clay. The best modeling clay in the world is the "Roma Plastilina" No. 1. Simply place a drop of your culture (not too big a drop, not too small a drop) on a clean slide, scratch each corner of the coverslip over a ball of modeling clay, and place it over the drop. Press down gently until the drop meets the coverslip. The feet should be less than a millimeter thick, and you can press them down (if you use a small enough drop) to trap swimming larvae or to improve the optics. Use plain slides and 18 mm square #1 coverslips; these are the most for the money and you can fit three preps on one slide. You can also prepare the chambers in advance; just line up some clean slides, and put the coverslips down in with clay feet in a row, and then fill them from the edge with a Pasteur pipette.

Some interesting activities

Observing fertilization and early cleavage

Fertilization can be observed in clay-feet preps. Put some unfertilized eggs under a coverslip, and then add a drop of sperm suspension, more concentrated than one would normally use, to the edge of the prep. Sperm may gradually make it across the entire prep, although of course the nearest eggs are the ones to watch first.

Immediately upon fertilization the eggs release the contents of "cortical granules" which include enzymes that release a membrane tied to the surface. This membrane initially looks like bubbles on the surface, but rapidly balloons out and rounds up and soon hardens, preventing more sperm from entering. This fertilization membrane is part of the "slow block to polyspermy" (the "fast block" involves very rapid ionic changes in the egg membrane). As the fertilization membrane elevates, the eggs often become very irregular in shape, but round up again within minutes. Sand dollar eggs are especially rewarding because the unfertilized eggs are often rather flabby-looking, but they round up quite nicely upon fertilization.

Soon after fertilization (10-30 min.), one can observe the "sperm aster" and the female pronucleus in the center of the zygote. The sperm aster brings the tiny sperm pronucleus together with the egg pronucleus. Although you cannot see the events of pronuclear fusion with a standard compound microscope, you will be able to see the subsequent formation of two asters – radiating lines coming from centers on either side of the zygote nucleus. The radiating lines reflect the creation of microtubule bundles initiated from the centrosomes. As the eggs prepare to divide, the asters become more prominent, the nucleus breaks down, and in especially clear eggs (e.g. white urchins or sand dollars), with good optics one can occasionally see condensed chromosomes in the mitotic spindle.

Unfortunately, none of the local species of urchin will go through first cleavage within an hour. At room temperature, sand dollars will make it in about 90 minutes. Therefore, to observe cleavage stages, plan to fertilize several small batches of eggs a few hours ahead of time, and space them an hour apart or so. See below for rough developmental time-tables.


Urchin eggs have several mechanisms to prevent more than one sperm from entering each egg. Textbooks refer to the "fast block", which involves an influx of sodium ions, and the "slow block", which involves the secretion of cortical granules. Cortical granule release causes, among other things, the removal of sperm receptors from the egg plasma membrane, severing of the bonds holding the vitelline envelope to the egg surface (once it rises up we call it the "ferilization envelope" instead), and hardening the fertilization envelope.

Of course, one way to induce polyspermy is to add way too much sperm. Polyspermic eggs will do odd things at first cleavage. Often, dispermic eggs will divide from one to four cells at first cleavage. More-than-dispermic eggs often fail to exhibit any cleavage furrows at all because mitosis gets held up before anaphase.

Another way to induce polyspermy is to conduct fertilization in artificial seawater that contains very little sodium. Various recipes exist, and here is one which I found on the internet in which choline replaces sodium:

Low sodium artificial sea water (50 mM sodium ion)
NaHCO30.2 g
Choline Cl28.3 g
KCl0.75 g
CaCl2•2 H2O1.6 g
MgCl2•6 H2O5.5 g
MgSO4•7 H2O7.1 g
Distilled water to 1 liter, adjust pH to 7.8

Add a small amount of settled eggs to 100 ml or so of low-sodium seawater, let them settle a bit, and add diluted sperm.

Other agents that are supposed to interfere with the fast block include nicotine (at ~1 mM) or barium chloride (also ~1 mM).


It's usually obvious what distinguishes phyla and classes of animals; for example, the many ways in which sea stars are different from snails and couldn't conceivably interbreed. However it's trickier at the level of genus and species. Why do we have green, red, white, and purple urchins, all living together in more or less the same geographical area, all belonging to the same genus, all evolutionary separated by millions of years only. Biological species are individuated from one another by the existence of barriers to hybridization (more literally, gene flow) between populations. Barriers might be developmental or physiological (e.g., hybrids develop abnormally), genetic (e.g., chromosomes don't match and meiosis fails in hybrids), or may involve mechanisms to inhibit creation of hybrid zygotes. The latter seems to be a major impetus for speciation in echinoderms, with other incompatibilities following.

Sperm and egg recognize each other as compatible gametes through at least two major mechanisms: first, the activation of the sperm acrosome reaction upon contact with the egg's jelly coat, and second, the binding of sperm to the egg membrane through the sperm protein bindin (displayed on the acrosome) and its receptor on the egg surface. These mechanisms are evidently subject to high rates of evolutionary change.

Among the local species of urchin, it is possible to make hybrids by adding a vast excess of sperm from one species to eggs of another. There are polarities to this interaction:

S. purp. spermS. drob. spermS. pallidus spermS. franciscanus sperm
S. purp. eggs10-50%; reach early pluteus40%; survive cleavage
S. drob. eggs50-98%; reach pluteus stage40-98%; can make hybrid urchin
S. pallidus eggs<1%; can make hybrid urchin
S. franciscanus eggs80%; reach gastrulation

These observations were copied from M. Strathmann's book "Reproduction and Development of Marine Invertebrates of the Northern Pacific Coast", which, incidentally, contains a vast treasury of information, descriptions, methods, and references on the title subject.

One curiosity is that, since one of the barriers to hybridization takes place at the level of the triggering of the acrosome reaction upon contact of the sperm with egg jelly, hybridization between some pairs of species may be potentiated by mixing eggs from the target species and eggs of the sperm donor, and then fertilizing both together.

Dissociating embryos

Urchin embryos can be dissociated using artificial seawater lacking calcium and magnesium ions (Ca/Mg-free ASW). After return to normal seawater, cells will adhere and continue to develop. In order to dissociate cells, one must first remove the fertilization envelope. There are several ways, including careful use of calcium-free seawater, but the easiest is to use an agent that prevents hardening of the fertilization envelope. Embryos which have been fertilized in the presence of 5 mM para-amino-benzoic acid (PABA) can be stripped of their envelopes by running them through either a drawn-out Pasteur pipette (broken to the right size) or by sieving them through Nitex mesh of the proper mesh size. For purple urchins, the proper mesh to use is somewhere between 60 and 70 microns; for green urchins, use 150 micron mesh. Sand dollar fertilization membranes remain soft enough to strip with Nitex alone (no inhibitor present) for as long as 30 min. after fertilization.

Stripped embryos still possess a hyaline layer, a sticky mat of glycoproteins that holds the embryo together. Ca/Mg-free ASW will cause the hyaline layer to loosen or, with washing or agitation, dissolve altogether. Embryos cultured in Ca/Mg-free ASW will not usually cleave normally, but one can dissociate them, either en masse or with a fine tool like a pulled glass needle, and then return them to normal seawater. After dissociation embryonic cells are often very sticky both to each other and to glass, and they also shear easily. Therefore it is a good idea to keep them in a Petri dish coated with a thin layer of 2% agar in seawater. Dissociation usually takes no more than a few minutes, with agitation, after which embryos should be rinsed extensively with normal seawater.

One interesting experiment to try is to dissociate green urchin embryos at the 4-cell stage, and then allow the dissociated blastomeres to continue development. The result, which once upon a time was so surprising that it drove many embryologist to mysticism, drink, or both, is that each fourth will develop into a small, but normal, larva. Actually, these quarter-larvae resemble the larvae of purple urchins, whose eggs are in fact much smaller than green urchin eggs.

Another interesting experiment, although it requires a delicate touch and a lot of patience, is to use a fine tool (e.g., a pulled glass needle, maybe an eyelash glued to a toothpick) to separate the animal half of the eight-cell embryo from the vegetal half, and culture them apart. The vegetal half should develop into a more or less normal larva, but the animal half will not gastrulate or make a skeleton.

Recipe for Ca/Mg-free ASW, copied from M. Strathmann's book
NaCl26.2 g/l
KCl0.67 g/l
Na2SO44.62 g/l
NaHCO30.21 g/l
Na2•EDTA *0.37 g/l
* that is, disodium ethylene-diamine tetra-acetic acid)
One may want to add Tris buffer (10 mM) as well. After bringing near to final volume with distilled water, adjust pH to 8.0 with NaOH. This solution should be stored in plastic bottles, not glass (which is rumored to leach calcium).
Disrupting the unequal fourth cleavage with SDS treatment

The first cleavage in urchins is as geometrically simple as cell division gets: a sphere turns itself into two spheres. Same for the second, and, more or less, the third division, and all the divisions in the other echinoderms (sea stars, sea cucumbers, and brittle stars). But the fourth division in urchins is highly unequal in some cells: the vegetal ("south pole") cells in the eight-cell embryo, when they divide, make one large daughter (a "macromere") and one small daughter (a "micromere") each. This unequal division is important for the subsequent differentiation of cells into skeletogenic, mesodermal, and endodermal populations.

Unequal cleavage requires re-orientation of the mitotic spindle, and eccentric positioning of the spindle with respect to the cell cortex. These events can be disrupted with various agents, including sodium dodecyl sulfate (SDS; a.k.a "sodium lauryl sulfate"). SDS is a strong ionic surfactant (look for it in the ingredients list for your shampoo or toothpaste), and at too high concentration disrupts cell membranes and denatures proteins. However, treatment of embryos with 20-25 micrograms per ml SDS during the interphase between second and third cleavage will equalize fourth cleavage.

Immediately following second cleavage, add a thousand or so embryos to 5-10 ml of seawater containing SDS (make fresh that day). Disposable plastic Petri dishes are convenient for this treatment because they provide an easy means to examine the embryos on a dissecting microscope and concentrate them by swirling. After 30-45 minutes, swirl the dish so that settled embryos concentrate in the center, and pipette them (taking as little water as possible) into another dish of normal seawater to rinse out the SDS.

This treatment works reliably on purple urchin and sand dollar embryos. For reasons we don't understand, fourth cleavage in green urchins is extremely sensitive, and even untreated embryos often fail to form micromeres. If all goes well, you should find that all cells of the eight-cell embryo cleave equally. Furthermore, if the embryos are cultured carefully, they will continue to develop, and even recover, replacing the cell types that normally would have differentiated as a consequence of micromere formation. This reflects the famous regulative capacity of urchin development – the ability to recover from drastic perturbation of early development and pattern formation, regenerating missing structures and cell populations.

It might therefore be interesting to compare gastrulation and early larval development in SDS-treated and control embryos. The SDS-treated embryos, missing the micromeres, should be unable to form primary mesenchyme (cells which invade the blastocoel cavity prior to gut invagination). These cells are fated to form the larval skeleton. SDS-treated embryos will nevertheless form such a skeleton by recruiting cells from other fates to do so. Does primary mesenchyme form? Does the skeleton begin to form late? Are the arms as long as in controls?


Sea urchins beautifully exhibit the stereotype for which the deuterostomes are named: the vegetal-most region of the blastula invaginates as a single epithelial layer to form a pit, which then elongates until it crosses the blastocoel cavity (extending itself into a thin tube, the archenteron). The archenteron joins a slight invagination on the opposite side of the embryo from where it started; when the two fuse, the alimentary canal is complete. The site at which gastrulation began forms the anus – hence, the term "deuterostomes" to describe the echinoderms and related phyla (including vertebrates).

Before archenteron invagination, a population of cells descended from the micromeres appear in the blastocoel. These cells are the "primary mesenchyme" and they will form the larval skeleton. Embryos in which these cells have ingressed, but which have yet to begin archenteron invagination, are called "mesenchyme blastulas". The primary mesenchyme cells migrate within the blastocoel into a ring surrounding the site of where the archenteron will form.

As the embryo prepares to gastrulate, the vegetal region thickens noticeably, then flattens. As the vegetal portion of the blastula wall invaginates, it thins out. Once the invagination has progressed a bit, one will see that the cells at the tip of the archenteron no longer have an exclusively epithelial character; cells at the tip express protrusions, including fine filopodia, and begin to migrate out of the epithelium. These are the "secondary mesenchyme", and they form the coelomic sacs. By the end of archenteron elongation these should be apparent as pouchs of mesenchymal cells on the left and right sides of the archenteron, near the mouth.

During archenteron elongation, primary mesenchyme cells continue to rearrange, and they begin to form the skeletal rods that will define the larval form. The first skeletal rudiments appear on the vegetal side of the embryo, one each on left and right sides. These have a distinctive refractility, and are always tri-radiate in normal embryos. They consist of calcite crystals secreted intracellularly within a syncytium made by fusion of the primary mesenchyme cells. Because they are refractile, one can visualize them either with darkfield illumination or, better, polarized light. For the latter technique, use two squares of Polaroid film; place one on top of the illuminator in the base of the compound microscope, and place the other in the light path between the objectives and the microscope head. Rotate the lower one until the field of view is dark, and the skeletal spicules should appear (with sufficiently bright light) as bright objects within the embryo.

Various treatments perturb gastrulation. Raising embryos in sulfate-free artificial seawater is reported to prevent gastrulation because the invagination of the archenteron requires production of sulfate-containing proteoglycans. Treating embryos with lithium ions (e.g., up to 50 mM LiCl in seawater) during early cleavage stages (e.g., from the sixteen-cell to the mesenchyme blastula stage) will cause more cells to contribute to the endoderm, and consequently causes drastic changes in gastrulation movements. In contrast, nickel ions (e.g. up to 10 mM NiCl2 added to seawater) will inhibit formation of skeletal structures. Many protocols for such treatments and many others can be found in the primary scientific literature, and even on websites.

Raising larvae

Urchin larvae are remarkably easy to raise as long as you have plenty of clean, filtered seawater, a supply of small algal cells for them to eat, and enough space keep them at a healthy density – less than one larva per ml is ideal for normal development. Indeed, this is the main challenge: don't be greedy.

Once embryos have finished gastrulation, they begin to grow arms supported by the larval skeleton, their mouths open, and they begin to feed by capturing suspended particles using the band of narrow, tightly-packed ciliated cells that run up and down the arms and around the mouth. Without food they will grow a little further, enough to illustrate what a pluteus larva looks like. However they will become truly spectacular little organisms within weeks with just a small culture of Rhodomonas (preferred) or Dunaliella on hand.

Both algae are easy to culture; use f/2 medium in seawater, which is available cheaply in stock form. Aseptic technique is not required for short-term cultures, although one should autoclave or otherwise sterilize the culture flasks in which algae are to be grown, and one should sterile filter or autoclave (or at least boil) the medium before use. Autoclaved f/2 medium should be cooled overnight before adding cells. Cultures do not need grow lights (a north-facing window suffices), but they will benefit from bubbling if an aquarium pump is available. The cultures should not be allowed to become too dense; dense algal cultures seem to sicken larvae. If the Rhodomonas turns orange instead of red, or if the Dunaliella goes muddy-green instead of fluorescent green, throw them out (although you might want to inoculate a new culture from the overgrown one).

Ideally, one would assess the algal culture density using a hemocytometer; in practice, it's usually easier to guess. A couple milliliters of a reasonably-dense algal culture per liter of larval culture, per day or two days, is about right. If the larvae have big round stomachs with lots of color and lots of cells in them, they're well-fed. Starved larvae will look, well, starved – skinny arms, small stomachs, little color.

To keep larval cultures healthy one must change the water at least every couple of days. To do so, use a plastic beaker in which the bottom has been replace by Nitex mesh of a size smaller than the larvae. Dip the beaker bottom down into the culture, and use a turkey baster to draw off the water, concentrating the larvae in what remains. It is enough to replace 90% of the water every other day, as long as the cultures aren't too dense and you aren't feeding them too many algae.

Developmental time-tables for some local urchins (copied from M. Strathmann's book)

Please take these with a grain of salt; they were compiled from different individuals' observations in different years and seasons, etc., and batches of embryos may differ slightly in development rate at a particular temperature anyway.

Dendraster excentricus (sand dollar)
Stage10-11 C11-12 C13-14 C17 C22 C
Two cells3 hr2 hr
Four cells4 hr3 hr2 hr
Eight cells5.5 hr
Sixteen cells7 hr3 hr
Hatching30-32 hr19-23 hr17 hr15 hr10.5 hr
Mesenchyme blastula36 hr27-29 hr
Gastrula38-42 hr32-35 hr20 hr
Prism55-60 hr40-46 hr32 hr24 hr17 hr
Early pluteus (2 arms)65 hr57 hr35.5 hr24 hr
4-arm pluteus4 day2.5 day2 day
6-arm pluteus8 day6 day4 day2.5 day
Pluteus with juvenile rudiment11 day

S. purpuratus (purple urchin)
Stage10 C12 C15 C
Two cells3.5 hr2.5 hr
Four cells5 hr4 hr
Eight cells6 hr5.5 hr
Sixteen cells8.5 hr6.5 hr5 hr
Hatching27-28 hr24 hr18 hr
Gastrula - halfway2 day30 hr
Prism3 day3 day2 day
Early pluteus4-5 day4 day3 day

S. droebachiensis (green urchin)
Stage4 C *8-9 C9-10 C
Two cells5 hr3 hr3 hr
Four cells8 hr5 hr4 hr
Eight cells10.5 hr6.5 hr6 hr
Sixteen cells14 hr8.5 hr8.5 hr
Hatching50 hr32 hr28 hr
Prism6.5 day4 day66 hr
4-arm pluteus11 day7 day5 day
* Unless you obtain Arctic urchins they are unlikely to develop at 4 C

Page written by George von Dassow; last updated Sept. 12 2003