Promoting Ethics and Sustainability

Within The Marine Aquarium Trade

Help The Reefs

Propagate Corals

Propagate all your corals and swap or sell these frags with other hobbyists

Breed Marine Fish

Attempt to breed the marine fish you keep and pass on your knowledge and experiences to others in the hobby

Buy Frags

Buy frags/propagated corals from other hobbyists rather than buying corals removed from the reefs

Make A Pledge

Commit to purchase only those marine animals which have been propagated or are MAC certified

Offset Your Impact

Donate a percentage of the purchase price for each marine animal you buy to a reef conservation organisation.

Promote MAC

Ask your aquarium store to become MAC certified

Marine Aquarium Home Breeders FAQ

Introduction

I guess this is where I'm supposed to put all the typical stupid legal stuff about copyrights and such. That will follow after giving my rationale for starting this project in the first place. I have decided to write this guide for a number of reasons:

I have been asked to write something by a number of fellow inhabitants of the *.aquaria.* newsgroups.
I agree with the current thread on the ethics of our hobby being primarily in the education of others about the natural functions of marine ecosystems; a reef tank is indeed probably the best example of a microcosm possible for a home setting. Thus, given that we have these great microcosms in many of our living rooms, the next step is learning to breed these animals and make them more generally available to our fellow hobbyists from captive bred rather than wild collections;
The greatest challenge of most aquarists (whether marine or freshwater) is not simply to keep an animal alive and happy - that is considered the basics by most - but to actually breed that rare and treasured critter.

I hope that I can help people to become more successful at doing just that. I also wanted to say that although this guide is written with marine invertebrates in mind, these techniques will also work perfectly well for the culture of marine fishes with planktonic larvae. It may be that you substitute rotifers (which are cultured in basically the same way as described for larvae herein) for the algal cultures for feeding fish larvae, but the techniques I describe in this document can easily be adapted to marine fishes with pelagic larvae as well.
So the legal rubbish is that this text is copyright Robert Toonen, but may be freely reproduced for non-profit purposes as long as it is not altered in any way and the authors name is not removed (that's what every FAQ seems to say, is not it?). Any commercial use of this text (i.e., in publications) is only permitted with the written consent of the author. I must make a disclaimer that this text is as accurate as I could make it, to the best of my knowledge, for the size of this document. Despite the many omissions and probable inaccuracies herein, I covered what I deemed important for home applications. I did the best I could (or was willing to do), so if you notice blatant errors, please contact me (Email at "rob@biogeek.ucdavis.edu" - preferably with the text that you think should be included to "fix" that error) and I will update the document. To quote Mike Noreen's bristleworm FAQ, "I am not responsible for any actions or losses or altered mental states resulting from the reading of this text or following advice given in it." I'll do my best to give you my honest opinion on this stuff and tell you how I've had success with larval culture, but from there, you're on your own.
Now, the last thing you need to know before getting into this is that in this document, I make the assumption that reef keepers are sophisticated enough to know the basic levels of classification of marine invertebrates (e.g., shrimps, crabs, copepods, amphipods, etc. are all crustaceans). If you are not familiar with the taxonomic classifications or terms mentioned herein, there are a couple of sources for you to find out what I'm talking about: 1) any invertebrate zoology textbook from a local library (see OTHER RESOURCES); 2) Martin Moe's handbook (see OTHER RESOURCES); 3) Megumi Strathmann's handbook of Pacific Marine Invertebrate Development is considered the Marine Biologist's Bible by many - it contains a lot of information and culture techniques for most common temperate Pacific Northwest marine invertebrates (see OTHER RESOURCES). I also use terms like Molar (remember basic chemistry?), ml and micrometer - if you have trouble with these, please look them up before you contact me to explain how to make a 0.5 M solution of KCl - I have enough Email as it is, thanks.

About the Author

I am a Canadian, born and raised in Edmonton, Alberta. If I had to pigeon-hole myself into a specific field, I would say that I am an Invertebrate Larval Biologist, but my interests are fairly diverse, and my current research is more along the lines of marine invertebrate population genetics than larval biology. I got my undergraduate degree working in the laboratory of Fu-Shiang Chia, at the University of Alberta in Edmonton. While in Chia's lab, I worked on several projects in progress, inclucing developmental variability in the peocilogonous opisthobranch, Haminoea callidiginata (the graduate research of Glenys Gibson), and the tubiculous polychaete, Phragmatopoma lapidosa (the post-graduate research of Joe Pawlik). While working for Chia in Friday Harbor, I completed a project on the feeding behaviour and prey preferences of the hydrozoan jellyfish Proboscidactyla flavicirrata under the guidance of several people to whom I am eternally grateful, including Chia himself, Glenys Gibson, Jon Havenhand, Joe Pawlik, and Claudia Mills (Toonen and Chia 1993). I subsequently went on to a Masters degree in Marine Biology at the University of North Carolina, where I studied larval settlement cues and the foundations of gregarious settlement in the serpulid polychaete Hydroides dianthus under Joe Pawlik (Toonen and Pawlik 1993, 1994, in press, submitted a,b, in prep). While at UNC, I also was involved in a survey of the chemical defenses of Caribbean sponges- we dove every island in the Bahamas for each of two summers, in order to complete a comprehensive survey of sponge diversity and chemical defenses (Pawlik et al. 1995, Chanas and Pawlik 1995). I am currently a Ph.D. candidate in the Population Biology program at the University of California in Davis working under Rick Grosberg, Gary Vermeij, Dennis Hedgecock, Brad Shaffer and John Gillespie. I am interested in the role of larval dispersal mode on the population genetic structure of sessile marine invertebrates (I work on over twenty species distributed among several phyla), and the subsequent effects of genetic structure on geographic range, species longevity and speciation/extinction rates.
Although I'm a marine biologist, I would have to say that aquaria are my hobby rather than my work. My dad owned a pet shop when I was a child; I've had freshwater tanks as long as I can remember, and my dad got me into breeding guppies on my own about the age of 5. I've been keeping fish and trying to breed everything I can get my hands on ever since. Aside from maintaining over 30 tanks in my home, I gained much of my experience with various aquarium-related jobs. I worked part-time as a dolphin trainer, and also helped to maintain the numerous reef, salt and freshwater tanks in West Edmonton Mall for about a year or so (before the cut-backs-when there were still some decent tanks set up in the Mall). I also worked as a marine mammal trainer and volunteer zoo keeper at the Edmonton Valley Zoo, where I helped maintain the aquaria and a lot of other animals. Most of my early experience came when I started working at the Koi Petshop (IMHO the best pet shop in Edmonton) when I was in high school, and continued to work there through my undergraduate time at the UofA. I had plenty of opportunity to experiment with new setups, various filtration methods and such things in the pet shop, and I accumulated a lot of my practical experience that way. I've now had a little over 10 years of experience with marine/reef aquaria, and about 6 years of experience breeding and raising marine invertebrates for both my research and at home.
If you want to contact me about any comments, questions, suggestions, complaints, death threats, etc., etc., please send me Email at: rjtoonen@ucdavis.edu or rob@biogeek.ucdavis.edu.

Background Information (aka the basics)

There are two ways in which animals can reproduce: sexually and asexually. If the animals are asexual, then a single individual can split itself into multiple copies and develop into a genetically identical colony. In this case, there is obviously little care required for propagating such animals at home - good conditions and plenty of food usually suffice. Asexual reproduction broadly covers both the addition of units (such as cnidarian polyps or bryozoan zooids) to an existing colonial growth form, and the fission of individuals to produce two (or more) identical individuals that are not attached in any way (such as some echinoderms and polychaetes). For sexual reproduction, at least two individuals are obviously required. If sexes are separate, the species is said to be gonochoric (=dioecious in plants), but if a single animal possesses both male and female reproductive organs at any point in it's life, the species is said to be hermaphroditic (=monoecious in plants). SIMULTANEOUS hermaphrodites are those that possess functional male and female gonads during the same reproductive season, SEQUENTIAL (or protandrous) hermaphrodites are those that never possess functional gonads of both sexes at the same time, but rather "change sex" at some point in their life (sometimes repeatedly). The "typical" pattern of protandrous hermaphroditism is that younger/smaller individuals will be the male, while older/larger individuals will be the female (this is the case with many shrimps, the Coral Banded Shrimp or CBS, Stenopus hispidus, for example). Within the broad category of sexual reproduction there are a variety of developmental modes common among marine invertebrates, some of which are simple to raise and require no additional care, and others of which require both special care and facilities.
The three primary sexual developmental modes of sessile marine invertebrates are: 1) DIRECT DEVELOPMENT, in which larvae are either held by the parent or within an egg case, or the larval stage is bypassed entirely; 2) LECITHOTROPHY, in which pelagic larvae are released fully provisioned and do not feed in the water column; and 3) PLANKTOTROPHY, in which pelagic larvae must feed in the water column before becoming competent to settle. These developmental modes can vary dramatically in the length of the developmental period; planktotrophic larvae typically spend months in the plankton; lecithotrophic larvae usually spend from less than an hour to weeks in the plankton; direct developers typically spend no time in the water column. Of course, the real patterns of larval development are more complex than this gross simplification (See Levin and Bridges 1995 for a more detailed account), but this simplification is accurate enough that it is widely used, even within the scientific community. In the home aquarium, direct and even some lecithotrophic developers can survive and reproduce without any special care or attention of the aquarist (i.e., those species with very short-term planktonic or nonplanktonic nonfeeding larvae). Species with long-term planktonic development, and even species with very short-term planktotrophic larval development, however, will require special care and equipment to raise at home. It is the latter group (species with feeding larvae or those with relatively long-term planktonic larvae regardless of feeding mode) that will be the primary focus of this document.

Setting Up For Larval Culture

Before we even start this section, let me warn you that larvae are VERY sensitive creatures, and need very clean culture conditions. You must not EVER wash the culture containers with any detergent - if you're worried about the cleanliness of your containers, rinse them with a strong acid (pool "pH down" is typically Muriatic Acid) or soak them in full strength, unscented bleach, followed by several boiling water rinses and finally a soak with concentrated sodium thiosulphate (dechlorinator) in cold water. Rinse the dishes several times with fresh water before standing them upside down on cotton towels and allowing them to air dry (and if you have access to DI/RO water, it is best to rinse a final time with that). You must not use hand cream or any other lotions on your hands, and should not use soap to wash your hands any time prior to working with the cultures (make sure you rinse very well with lots of warm water every time you plan to work with your cultures). You should also not use paper towels for drying anything in your culture equipment - they contain enough residual formalin to alter the development of most larvae. You cannot use any sprays (like disinfectant, deodorant or pest aerosols) in the room where your larvae are to be kept. You must prevent any contaminants (like heavy metals, fixatives, detergents, solvents, etc.) coming into contact with the dishes to be used for culture, and they should be of an inert substance, like glass or acrylic, that are resistant to corrosion by seawater.
OK, I've made my point: enough of that. So there is a basic set of equipment that you'll NEED, some that you'd LIKE, and some that is just REALLY nice if you can get your hands on it. The equipment in these lists will vary to a large degree on the scale of larval culture in which you're interested. This is something that you may have to decide in advance of spawning your animals. For many marine invertebrates, it is not uncommon to produce tens of thousands, or even hundreds of thousands of viable eggs in a single spawn. If you plan to raise hundreds of larvae, your technique will likely be very different than if you plan to raise hundreds of thousands of larvae! If you have no idea how many eggs your animal(s) may produce, you may want to try a "learning" spawn on a small scale before moving up to large scale culture techniques. In this case, you would attempt the normal spawning and simply keep a few hundred fertilized embryos to raise. Of course, if you find yourself in this situation for the first time, it is very hard when you have hundreds of thousands of fertilized eggs to just throw most of them out, but believe me, if you are not set up for it, and try to keep too many larvae, you WILL fail. It is much better to successfully raise a few hundred animals than try desperately to keep up with a lot of cultures and eventually have them all die because you cannot keep them properly. If you have to discard a lot of eggs the first time, you can then prepare for a large-scale culture the next time around.
For small-scale cultures, you'll need a series of deep glass bowls (like custard bowls or wide-mouth highball glasses or something), something to loosely cover the bowls (primarily to decrease evaporation and prevent anything from falling into the cultures), and a good quality pipette (=fancy eye dropper, the reason you want a good one is so that you can reasonably remove larvae from the culture without having to suck them up one at a time - if you haven't ever seen a typical research pipette, the glass part is finer and more tapered than your average eye dropper, and the rubber bulb is about 3-4 times the size of an eye dropper). That is the primary equipment you'll NEED to culture relatively few larvae at home. It would be NICE to also have a dissecting microscope to examine the larvae from time to time as they grow, both for interest's sake, but also to watch for clumping (caused by high bacterial growth) and deformities or illness. Depending on the species you are culturing, you might also LIKE to have some Acetyl Alcohol (looks and feels like soap flakes) around to sprinkle lightly on the surface of the culture water. Many species (especially gastropods) have larvae notorious for getting stuck in the surface tension of the culture vessel. Adding the acetyl alcohol decreases the surface tension (without any toxicity effects) and inhibits the trapping of larvae at the surface of your culture vessel. Another way to prevent larvae from getting stuck in the surface tension is to build a "lift-drop stirrer" to turn over the culture and constantly disturb the surface to prevent larvae becoming trapped at the air-water interface. See Strathmann's book (1987 - see OTHER RESOURCES) for instructions on how to build an "airlifted-droplet stirrer" (pgs 15-16 in Strathmann) if you are interested in trying this (it will also be discussed further below in the BASIC LARVAL CULTURE TECHNIQUES section).
There have evolved a wide variety of methods for the large-scale culture of marine invertebrates, and virtually every larval biologist you speak to will explain their method is the best, because they have tried everything else, and this is the method on which they have settled because it works the best (I'm sure this logic will sound familiar to anyone who has keep aquaria for very long). I think that the best method is the one that gives you the best results, and that will vary from person to person to some degree - that is why we have such diverse "best" techniques from which to choose. I will only describe a couple herein, and I will (of course) endorse my own method as most effective with the least amount of work and special equipment. If you are REALLY interested in finding out more about different larval culture techniques (which is a good idea if you are serious about pursuing this beyond a hobby - you should probably take every piece of information available to you and distil it down to what will work best for your specific application) check the culture texts in the OTHER RESOURCES section.
Large-scale larval culture varies from typical research applications (thousands to hundred of thousands of larvae in tens of gallons at a time) to typical aquaculture applications (millions of larvae in hundreds of gallons at a time). I am assuming that no one reading a "home breeders FAQ" is interested in running an aquaculture size or style facility (if I am incorrect in this assumption, I'd suggest you move on to the OTHER RESOURCES section - check out Spotte's writings), and so will concentrate on the research application scale of culture (which would probably even be appropriate for the home breeder who wanted to produce many juveniles for local distribution). In order to do a large-scale culture of larvae you'll NEED several wide-mouthed large glass jars (I use well-cleaned 1 gallon pickle jars from restaurants), a good quality air pump and a VERY sensitive, good quality gang valve (i.e., you don't want to get the $0.49 special plastic 5-way gang valves from "QuickieMart" - use whatever department store name you want here - for this purpose!). The number of jars you'll need depends on the number of larvae you'll ultimately want to culture. Plan on an absolute maximum of 10,000 larvae per gallon jar (ideally you should aim for about 1000 larvae per gallon, but it become very space consuming if you want to culture a lot of animals at this density!) and you'll need 2 jars for each culture. Thus, if you want to raise 100,000 larvae, you'll actually need a minimum of 20 jars for that culture effort (the reason for the 2 jars is to keep cultures cleaner - every time you feed a culture, you transfer the larvae to a new jar and scrub out the old jar with boiling water before allowing it to air dry). You'll also need some very fine mesh screen (this should ideally be Nitex 56 micrometer mesh, but I couldn't find it in any easy to access catalogs. I did find 75 micron Nitex available from several supply companies like Fritz Aquaculture - now called Aquacenter - 1-800-748-8921, so I'll use that as my base size herein. Although Nitex is the best thing to use, even really fine cheesecloth should probably do the trick - but realize that 75 micrometers is 75/1000 of a millimeter - that is VERY fine mesh), some coarser mesh screen (again, this should ideally be Nitex on the order of 300-500 micrometer mesh - depending on the size of the larvae you are culturing - but even fine fiberglass mosquito mesh would probably work), and a couple of stackable plastic beakers. Now, I realize this is starting to sound strange, but after having experimented with various techniques, I assure you this is by far the easiest method (even if not the most successful for all species - for example most nudibranch larvae are too delicate to be cultured in the following method, but most polychaetes handle it quite well). See, already my method is the best - just ask me! The purpose of the beakers and mesh is to build filters with which to sieve the larvae when transferring them from one culture vessel to the next. Use a hacksaw to cut the base out of the stackable plastic beaker, and hot glue (use the `soft' clear hot melt glue - yes this is the stuff from Home Depot or whatever - rather than the `hard' white stuff) the mesh across the bottom of beaker. If you put down a bead along the plastic, lay the mesh over that and press it down into the glue and let it cool. Then, trim around the edge with a sharp razor or such, and place another generous bead of glue over the top of that, you'll have a very strong bond. You use the large mesh size beaker as a prefilter in which to catch flocculant debris that has built up in the culture (dead and aggregated larvae, excess food, feces, etc), while the healthy larvae pass directly through the large mesh screen and into the lower screen. The reason that you use stackable beakers is so that you can keep the lower screen submerged in a dish the whole time, without trying to hold layers of beakers and pour simultaneously. This is easier if you take a third beaker, cut the bottom off, and cut large notches around that edge of the beaker. Then when you stack the three together, you'll have the large mesh on top, the small mesh beneath, and the open beaker with notches cut into it on the bottom. This stack can be placed into a casserole dish or other such container that is tall enough that the small mesh beaker remains submerged, but the large mesh beaker is kept out of the water. The culture jars can then be picked up, swirled (to lift everything into suspension) and easily poured through the stacked filters to isolate the larvae. Once completed, you'll need some sort of plastic spray bottle (like a cyclists spray bottle or such) to wash the larvae out of the filter and back into a culture dish. Again, it would be NICE to also have a dissecting microscope to examine the larvae from time to time as they grow, both for interest's sake, but also to watch for clumping (caused by high bacterial growth) and deformities or illness.
The party line among larval biologists is that artificial seawater is inadequate to raise most species of marine invertebrates, but my experience is that reef tank water SOMETIMES works just fine. Newly-mixed seawater does not seem to be nearly as successful, however. My guess for the reasons behind this is that there are many dissolved organics in our reef tank water that are lacking in freshly-made artificial seawater. You may be able to bypass this problem by adding the invertebrate feeding formulae used for reef tanks (those claiming to be a rich food source with all the essential amino acids, vitamins and such but look clear or possibly just a bright artificial colour when you add them - they contain a lot of dissolved organic material (DOM) so that may help a lot - but I have yet to try this). The other option is to take natural seawater and either Millepore filter, pasteurize or microwave sterilize it. These will be discussed further in the ALGAL CULTURE section below. Also, be warned that many buffers commonly added to artificial seawater mixes (for example TRIS - this is a chemical stabilizer commonly used to buffer fast dissolving seawater salt mixes) are toxic to some marine species, and tend to uniformly kill larvae, regardless of adult tolerance to the chemical. This may or may not be a problem to home breeders, depending on the species which you are trying to culture.

Algal Culture

If you hope to raise any species which has feeding larvae, you'll need to setup algal cultures to feed those larvae. This sounds easy, but is generally much more difficult than it sounds. Many people have more trouble getting the algal cultures running well enough to start raising larvae than they do raising the larvae themselves. This should be your first step if you want to culture any species with planktotrophic larvae! The first step is getting both the nutrient supplement and the pure strain algal inoculants for your cultures. The supplement can come from a number of sources. First, you can buy sterile, premixed and measured aliquots of f/2 algal growth supplement from companies such as Carolina Biological Supply . This is probably the easiest and also the most expensive way to feed your algal cultures. Second you can buy bulk f/2 supplement from many aquaculture supply companies like Aquatic Ecosystems, Inc (1-800-422-3939), Fritz Aquaculture - now called Aquacenter (1-800-748-8921), etc. who provide premixed supplement which can be measured and added to culture vessels as needed. This second alternative is probably the best for people who want things easy, but still relatively inexpensive. The final method is the cheapest, but also the most work: you can mix your own nutrient supplement. Strathmann (1987) has a recipe, and Guillard (1975) is considered the authoritative source for all subsequent instructions and nutrient supplement recipes, if you're really interested. If you want a copy of the recipe I use, please get in touch with me (rob@biogeek.ucdavis.edu) and I'll FAX it to you (I'm far too lazy to type the damn thing in here - that may change, depending on interest, in future versions - if any - of this document).
Once you have the nutrient supplement, you'll need an inoculant to start your cultures. Again, you have a number of options. First, you can simply buy a pure strain culture from a company such as Carolina Biological Supply or Florida Aqua Farms. Again this is the easiest and most expensive method of obtaining cultures, but in this case also by far the most reliable. Second, you can try to get inoculant from some local source (check with a University if there is one in your area, or try to contact other breeders in your area). Third, you can try to isolate algal cells from natural seawater, or even possibly from well established reef tanks. This third option will again be the cheapest, but also the most work. In order to isolate algae to inoculate a culture, you can add a few drops (and I mean a FEW - like 2 or 3 drops) of natural seawater (or water from a well-established reef tank) to about 250 ml of sterile sea water (see sterilization techniques below) with culture supplement added (hereafter called simply medium). Split this plankton medium into a series of petri dishes (just enough water to cover the bottom of the dish) kept somewhere with constant temperature and subdued lighting. They should be in a place where they can be viewed with a magnifying glass without moving or disturbing them in any way. In the course of a few days, colonies of different species of diatoms will be seen growing at different spots on the bottom of the dishes. A drop from the center of this spot may be removed and placed into fresh culture media. This isolation procedure must be completed as early as possible (i.e., when the colonies first become visible), before the entire water is `infected' with organisms and no "pure" strain can be isolated. This procedure may need to be repeated a number of times to ensure that the culture is actually "pure" (I use "pure" in quotations because technically pure cultures are bacteria-free, which these will most certainly not be - but for all intents and purposes at home these cultures can be considered pure). There are a number of fancier and more complex methods, but this is the simplest and least expensive method for home isolation (actually, another very simple one - if you have a good microscope handy - is to examine a droplet of water under a microscope and remove a single cell of an appropriate diatom for isolation).
The species of unicellular algae typically used for the culture of marine invertebrate larvae are as follows (this information is largely lifted from Strathmann 1987):

Cricosphaera carterae - Gold-brown flagellate with cells in the 10-18 micrometer range. Particularly good food for small prosobranch veligers.
Dunaliella primolecta - Green alga with cells in the 4-10 micrometer range. Particularly poor food for most species with the exception of asteroid larvae, possibly because it lacks certain polyunsaturated fatty acids (Peirson 1983).
Dunaliella tertiolecta - Green alga with cells in the range of 10-15 micrometers. Easy to culture and nutritious, this alga makes a good choice for the culture of many echinoderm larvae, but apparently poor for prosobranchs.
Isochrysis galbana - Gold-brown alga on the order of 5-8 micrometers which contain large lipid stores. An excellent food for most small planktotrophic species, especially in mixture with Dunaliella, Pavlova, or Thalassiosira. Recommended for mollusc larvae, copepods and rotifers.
Pavlova (=Monochrysis) lutheri - Gold-brown flagellate approximately 7-10 micrometers. Tends to be a poor food source alone, and may be harmful in dense culture due to the accumulation of algal byproducts. There are better species available, so it is best to avoid this one, unless you are willing to rinse the algae before feeding.
Procentrum micans - Large dinoflagellate good when mixed with Isochrysis for advanced larval stages or large larvae.
Skeletonema costatum - Chain diatom, each cell being about 6 micrometers. Fair to good food for crustacean larvae such as barnacles.
Tetraselmis (=Paltymonas) suecica - Tiny green alga, about 10-12 micrometers, recommended for rotifer culture, or when mixed with Isochrysis for molluscs.
Thalassiosira (=Cyclotella) pseudonana - Small diatom averaging about 3 micrometers in diameter. A good food for molluscs and early crustaceans. When mixed with Dunaliella also for echinoderms.
Thalassiosira weissflogii (=fluviatilis) - Large diatom, averaging about 20 or so micrometers in length. An excellent food for juvenile and adult bivalves, and a good food for echinoderms and crustaceans.

It should be obvious from my flippant and superficial treatment of nutrition (good vs poor doesn't tell you anything, really, other than my opinion) that relatively little is known about either the nutritive requirements of marine larvae or the nutritive content of marine phytoplankton. It is always best, whenever possible to feed a mixture of algal sources to larvae.
In order to be at all successful in culturing "pure" strain algal cultures, you will first need to sterilize the seawater in which algae are to be introduced. The best way to do this is, of course to autoclave it, but this is not always easy or desirable for the hobbyist at home. That leaves a couple of options. The first, if you have the equipment available or money and desire to buy it, is to Millepore filter your seawater to one micrometer (there aren't any cheap alternative filters I know of to get 1 micrometer filtered seawater - you have to bite the bullet and go for the Millepore stuff). This could be made cheaper (actually only by making each filter last longer before clogging and becoming useless) if you drip the water from your tank through a coffee filter (I HOPE coffee filters are not exposed to formalin in processing the way paper towels are!! Does anyone know this for sure?). The second (pasteurization) and third (microwaving) methods are much cheaper and more likely to be used at home. Pasteurization involves heating the seawater to about 65 C (I think this is about 145-150 F) for at least 30 minutes (keep it covered to prevent significant evaporation - this would change the salinity; in fact, some people add 50ml if DI/RO water per l of seawater prior to heating to offset this effect), followed by rapid cooling to about 4 C (put it in your fridge), and store it in a covered container (a large saucepan with a good lid will probably do the trick) until you are ready to use the "sterilized" water (I put sterilized in quotes because many bacteria survive this process, but it's better than nothing, and Stan Brown tells me it works well for him - he says that calcium tends to precipitate slightly with the elevated temperature, but otherwise the treated medium seems fine). The final method is microwaves sterilization. Microwaves have been found to kill most everything but fungal spores, so if you live in a relatively dry part of the country, and fungal spores are not a source of significant contamination, you need only microwave your culture medium. Bob Guillard (the phytoplankton expert I cited earlier) found that the medium need not even boil for the microwaves to kill most things in seawater. Generally if you place the medium in the microwave in the flask in which you intend to culture the algae for about 2 minutes on high, you'll have pretty sterile culture medium. If, however, you happen to live somewhere like Florida or the Carolinas, where fungal spores will definitely be a major problem, you'll need to get a special microwave pressure cooker. I have heard of them, and know people who use them, but have never tried it myself. The major concern for a pressure cooker is that you add some tapwater to the inside of the cooker to prevent your algal growth medium from boiling over and making not only a mess, but wasting the medium. When either pressure cooking or autoclaving, you must aerate the "sterilized" medium for a while prior to adding the algal inoculant to remove excess carbon dioxide from the medium. Once the medium is sterilized (or nearly so), make sure that the stopper stays in the beaker, and that you never let anything that has lain around on counters or such come into contact with the medium or the edge of the stopper to go into the flask (that ESPECIALLY includes your hands - never touch the flask or stopper on the "inside" parts once sterilized - the only part of our body more filthy than our hands is our mouth). You can open the flask to add algal inoculant water and/or nutrient supplement, but make sure that you only lift the stopper enough to get whatever you're trying to add into the flask, and keep it directly over the culture flask to prevent anything settling into the flask while you're doing that.
If you plan to get into this seriously, I would suggest that you find a few glass 1 or 2L Erlenmeyer or Fernbach flasks (large flat-bottom, narrow-necked flasks) and get a good quality rubber stopper with 3 holes drilled through it to fit each flask. You can drill the holes yourself by cutting a piece of copper pipe or some such thing of the appropriate diameter and sticking it into your home drill. The sharp edges of the cut pipe (you can even get the hardware store to do this for you if you don't have a pipe cutter at home) will cut through the rubber and remove a core from the stopper. Stan Brown tells me that he's found it much easier to drill the stopper if you put it in the freezer until it gets good and hard. You'll also want some glass tubing that will (snugly) fit through those holes (and also into airline tubing, so think about this before you drill the holes in your stopper). Two pieces of glass should be tall enough to reach all the way to the bottom of the flask and still stick up half an inch through the stopper so it can be attached to a good airpump (if you have a very hard time with contamination of your cultures, you can add a bacterial filter from Carolina Biological Supply inline, otherwise a typical aquarium floss-filled plastic inline filter should suffice) to turn over the culture continually. One of these tubes should be bent slightly (you can do this by heating it over a small flame from a bunsen burner or BBQ or such, or even the burner of your stove at home - be careful! Glass is a good conductor of heat!!) near the middle, so that it will end up releasing the stream of air bubbles at the front edge of the bottom of the flask. This tube should have the NON-BENT end inserted through the stopper (be very careful inserting glass tubing though a stopper - you don't realize the amount of force you're using, and it is very easy to break the tubing and stab yourself, as the scars all over my hands will attest!! - always hold the glass as near to the stopper as possible and twist back and forth while pushing down, using a copious amount of spit on the hole to lubricate it while inserting the glass tubing) far enough to attach an airline to it, but not so far that you're likely to break it when moving the thing around. The other tube should be straight inserted through the stopper so that it almost touches the bottom, and again have an airline attached to it. This airline will be your tube from which algae are removed from the culture. It should hang down at least to the bottom (outside of course) of the culture flask when not in use. The final tube should be much shorter (just enough to stick through the stopper and then a couple of inches outside), and also be slightly bent (make sure that the bending of the tube does not close it off - if that happens you're being impatient and not warming enough of the tube before trying to bend it) and then inserted (once cooled, of course!) through the stopper with the bent side OUT of the flask. This is an air outlet, and by bending the tube you simply decrease the chance that any spores (algal, fungal or bacterial) can get into the culture (and constant air flow out also helps). Then set the flask on an eraser or similar nonslippery thing to raise the back side (away from the air outlet inside the flask) and turn up the air. That should prevent your algal culture from settling and decreasing growth rate or building up bacteria. Once your culture is growing well, you won't want to open the thing and risk contaminating it, so you can simply cover the air outlet with your finger, and place a flask under the piece of aquarium tubing that you have attached to the straight piece of glass. Culture will be forced out of the flask by the increased air pressure as your pump fills the flask with air, and your finger prevents it from escaping. Because this tube will frequently contain seawater, rich with growth medium and dead or dying algae it will be a haven for bacterial and fungal growth, so should NEVER be lifted above the level of the culture when algae are growing in the flask, or you will surely contaminate the culture.
Once you have the thing set up, and are ready to add the algae, you can set the flasks on a shelf in front of a window that gets lots of sunlight, or set up a light specifically for it - which ever works best for your needs. Both GE and Phillips make cheap full-spectrum lights that are good alternatives to spending a mint on petshop aquarium bulbs (e.g., GE Chroma 50, Philips colortone 50). I'd use at least 80 W of light directly above (or better yet beside) the flasks if you want decent growth and final culture densities. I use 160 W of full spectrum light in the door of a light box (sealed and with reflective Mylar all around the inside) made specifically for this purpose, and I don't start to feed the cultures until they reach about 10,000,000 cells per ml but home use probably doesn't require anything so drastic, unless you plan to run a major breeding facility. The recommendation Guillard makes is about 4000-6000 lux for about 14 hours per day. This is about the output I get at 10-14" away from the 4 Hagen 48" Powerglow lights in my lightbox door, just to make this lux measurement into something tangible for most people. Many people grow their algae under regular cool white fluorescent bulbs, but in my experience, the cultures SEEM to grow faster (reach density sooner) and to have better color under the full spectrum lights than under the cool whites. I say SEEM because that is my impression - I have not done any experiments to see whether or not there is any significant difference between the two light sources, or whether my subjective impression of "algal growth/quality" is, in fact, real. Either way you decide to go, you need to have a lot of light on your algal cultures, so pick what you think works best for you.

Fertilization and Artificial Spawning

Many marine invertebrates have yet to be spawned under "natural" conditions in the lab, and in such cases they are induced to spawn, or are "artificially spawned" (a lovely euphemism for being killed to strip the animals of ripe gametes). Although this is common, and sometimes desirable in research settings, it is probably far from desirable in a home breeding situation. There are a number of methods to induce "natural" spawning in ripe specimens, including: 1) strong mechanical agitation (e.g., vigorous shaking or tapping causes some echinoderms and bivalves to spawn); 2) pouring cooled seawater over the animals in a small room-temperature container induces spawning in many intertidal species (e.g., some anthozoans chitons, gastropods and bivalves); 3) light changes may induce spawning in many species, but it varies as to how - keeping the animals in complete darkness for about 48 hours prior to intense illumination often causes spawning (sometimes up to several hours later) in species that normally spawn in response to light changes (e.g., some hydrozoans, bryozoans, tunicates and crustaceans); 4) immersion in water densely populated with phytoplankton causes spawning in some species (e.g., some chiton, bivalves and echinoderms); 5) dramatically increased current flow has been reported to stimulate spawning in some species kept in low current conditions (e.g., some ophiuroids, gastropods and bivalves); 6) although I would NEVER recommend this at home (electricity and seawater are NOT something people should play with in the basement!!), it has been found that running a low level DC current through seawater excites many echinoderms and some molluscs to spawn (e.g., Suquira, 1962); and finally, 7) Chemical treatments have been found to induce spawning in a variety of species (e.g., some echinoderms, bivalves and gastropods). The chemical effective for treatment varies by group: for example, 0.55 Molar potassium chloride (KCl) or 0.1 Molar acetylcholine works best for echinoids; 0.1 N ammonium hydroxide or seratonin work best for bivalves; 1-methyladenine works best for asteroids; and hydrogen peroxide works best for prosobranch gastropods. I am not going to go into detail on any of these chemical techniques for spawning with the exception of the hypodermic injection of 0.1-5.0 ml (depending on the size of the animal) of 0.55 Molar KCl directly into the body cavity. Once an animal is injected, place it into a separate container with a small volume of seawater (just enough to cover the animal) and allow it to release all it's gametes (you don't want to stick it back into your tank until it has finished spawning, or it may trigger other animals to spawn). However, you also don't want to leave the animal for too long while spawning, because most gametes are only viable for a short period (maybe an hour or two) once released (and it takes about 30 minutes to complete the fertilization sequence). Although techniques to induce spawning (such as injection of KCl), may still be useful for home aquaria, there is a significant risk to the animals anytime spawning is forced by some mechanical or chemical shock method. Assuming that you don't OD the animal on KCl, and that it does not develop a secondary infection from the treatment, it should recover after being placed back into the tank immediately following spawning. The other techniques are similarly effective if done in a careful manner so that the animal is dosed to the point of inducing spawning but not death. The amount required and sensitivity of animals to these chemical stimulants will vary by individual, and is something that cannot be safely predicted for long-term survival of the animals. Of course, it will therefore be far better for your pets if they spawn on their own and you to collect the gametes.
Regardless of how you obtain the gametes, once you have both sperm and eggs in isolated containers, you need to fertilize the eggs. The first step is to dilute the sperm. The amount of sperm released by a single male is enough to ensure polyspermy (egg penetration by multiple sperm, resulting in abnormal development and rapid death), therefore, you must dilute the sperm with seawater to a suspension that appears nothing more than a slightly milk haze to the water. Once diluted, add a reasonable amount (like 10 drops of sperm suspension into 50 - 250ml of seawater containing eggs, for example) to the dish with the eggs in it. Very gently swirl the suspension and eggs for about 10 minutes, and then add a doubled volume of sperm suspension to the dish (in our example, we'd now add 20 drops into the 50 - 250ml). Again, very gently swirl the mixture for about 10 minutes, and then add something like a tripled or quadrupled volume of sperm suspension to the dish (50-75 drops now in our example) and swirl very gently one last time for about 10 minutes. The eggs should now be fertilized, and the embryos are ready to begin culturing.

Basic Larval Culture Techniques

A lot of the information important here has already been given in the EQUIPMENT SECTION above, so I will not repeat it. I will assume that you have already read the sections on both ALGAL CULTURE and EQUIPMENT before starting in on this, because there is a fair amount of overlap. I may refer to techniques or simply skip discussion of topics covered in those earlier sections, so be forewarned - this is a minimalist guide to culture techniques.

Small Scale Cultures

OK, so this is probably where you want to start out with your culture attempts. Once you have raised larvae for a while, and have some experience with it, you'll realize that this is the more labour intensive way to raise larvae. BUT, until you have experience raising larvae, you probably won't be very successful with large-scale culture attempts (but you're welcome to try). If you are planning on raising larvae at home for your own use and amusement, the best way is probably to stick to small scale cultures, but if you want to raise larvae to sell or to supply juveniles to the local petshops or such, you're better off with stepping up to large-scale culture pretty quick, or you'll spend all your time working to raise relatively few larvae.
So you start off with about twice as many larvae as you'd like to eventually raise (with the assumption that you'll lose a bunch along the way). Set these larvae up in small culture vessels (like large custard dishes or wide-mouthed highball glasses or something similar) at a concentration NO HIGHER than 1 larva per ml in each vessel (1 larva per 10 ml is probably ideal, but very space intensive). To the top of each larval culture vessel, add a few flakes of Acetyl Alcohol (if you have access to it) to keep the surface tension low and prevent trapping of larvae in the surface film. The importance of keeping the surface tension lowered depends on the species which you are trying to raise. In some groups, such as opisthobranch molluscs, the larvae have hydrophobic shells, and will be frequently trapped in the surface tension without the Acetyl Alcohol, in other groups, such as polychaetes, there is little need for decreasing the surface tension. Larvae should be fed algal cultures (uni-algal if you have to, a combination selected from the list provided earlier if you can) at a concentration of approximately 100,000 cells per ml every second day. A feeding concentration of about 100,000 cells per ml is around 5 ml of very dense (we start to use an algal culture when it hits about 50,000,000 cells per ml - at this density, algal cultures look sorta like coffee) algal culture added per 1 l of larval culture (you should adjust the feeding amount according to your algal culture density). There are a variety of methods to determine algal density. Most researchers use either a Coulter-counter (an electronic counter to determine the cell density in an aliquot of the stock culture), or a hatched microscope slide on which a known volume of stock culture is added (you add a drop of culture to the slide - if I remember my basic chemistry, there are about 20 drops per ml from a research pipette - count the number of algal cells in the drop and multiple that by the total culture volume to get cell density). Some people use Secchi-disks, but I am not sure how that works or how accurate it is. Anyhow, once you have added the algae, the culture water should look only SLIGHTLY cloudy - if the water is dark, you have added a lot more algae than needed. SLIGHTLY cloudy means the water should look basically clear until you hold it up to the light, when it should be obvious that there is something in it. Even if you can't figure out (or don't care) what your algal culture density is, you can use adding algae until your culture just starts to look slightly cloudy as a gauge for the amount to feed - keep in mind, though, if you overfeed, it will lead to more bacteria and debris in your cultures, your larvae will grow more slowly and you will lose more. It is better to underfeed your cultures (it just takes them longer to reach competency with less food - and this experiment I have done!) than to overfeed them. Every second day the cultures should be fed and cleaned. Like I said earlier, you'll want 2 culture `jars' for each larval culture that you intend to keep, one to keep the culture in and another for changing the culture. Culture dishes that larvae are to be moved into should be set up with the appropriate amount of seawater (treated/prepared as for the algal cultures), and larvae should be transferred individually by pipette from the old culture to the new one. Depending on your eyes, and the size of the larvae, this may be made much easier with a good hand lens or a dissecting scope. This is likely the stage during which most larvae will be lost, so be careful to try to pick out every healthy larva you can find. You will undoubtably lose some, some will die, etc., and that is why you want to start this with twice the number of larvae that you eventually want to end up with of juveniles. The idea is to transfer as little water as possible while still getting all the larvae out of the old culture vessel into the new one. This is to keep the bacterial numbers down to a reasonable level through out the culture duration. Once all the larvae are transferred, the old culture jar should be rinsed out with water as hot as possible (boiling is best, but use what ever seems reasonable to you) and then allow the jar to air dry (again, if you have access to RO or DI water, give the vessel a rinse with that before drying) on cotton towels. If you keep the cultures clean and transfer the larvae with a minimal amount of the old culture water, this culture technique should work the best for any species that you want to culture in small quantities.
Depending on where you live, you may need to regulate the temperature of your cultures. If you need to watch the temperature in your tank (i.e., do you have a heater in your tank?) then you'll also need to maintain the temperature of your larval cultures. This is easily done by setting your cultures into a large, flat-bottomed, low-sided plastic tub (you can get them from WalMart or any of a million other places) with enough freshwater in the tub to cover about halfway up the side of the culture dishes (there should be enough water to cover a submersible heater - ones like the Hagen thermal compact are very small - but still be low enough that the culture jars seat firmly on the floor of the tub: you don't want them tipping over accidentally). This apparatus can have a simple shop light set up above it with a couple of cool white or full spectrum fluorescents in it and set on a timer to simulate day/night cycles if you are concerned about larvae and/or the algae they are fed behaving `normally.' As far as I can tell, the development of larvae through metamorphosis does not require regular light/dark cycles, however, so the addition of lights is, IMHO optional.
After the larvae have reached competency (this is a whole other issue again), you have to figure out how to get them to settle and metamorphose into adults. This is the subject of another section (see below - Section VIII: Larval Settlement).

Large Scale Cultures

So, once you've figured out how to culture larvae, you may want to step up the scale of culture to expand your production. The market for reef-tank inverts is huge, and a garage business of raising tube worms, nudibranchs, sea apples, and other marine invertebrates certainly never hurt my income. Besides the income, the enjoyment and satisfaction of raising the animals is worth the effort and time involved. For large-scale cultures, your best bet is to use 1 gallon wide-mouth pickle jars (or some such thing) that are either new, or have been used only for food (human consumption is probably a safe indication that nothing permanently damaging has ever been used in the vessel, although you'll still want to acid wash or bleach your new culture vessels). These jars should be set up with seawater (again, prepared/treated as for algal cultures) and larvae should be added at a concentration around 1 larva per ml of seawater (you can go as low - 1 larva per 10 ml is probably best - or as high - 10 larvae per ml is probably as high as you want to reasonably go - as you want, though. This is a judgement call and will come with experience and depend critically on the species which you are trying to raise). Again, larvae should be fed at approximately 100,000 algal cells per ml for a larval culture of 1 larva per ml, and that amount should be adjusted according to your culture protocol. Adding too much algae is not only wasteful but it increases the likelihood of your algal cultures becoming fouled. Use the least algae you can to ensure that all larvae have and adequate food supply and will grow/develop normally. Again, if you think that larval cultures are likely to experience temperature swings, set them into a constant-temperature bath to maintain the culture temp (as described in Small-Scale Cultures above). Large-scale culturing is basically the same as the small-scale culture technique: you want to maintain the cleanest culture conditions possible with the least amount of bacterial growth. Again, you should have 2 culture jars per culture, and the new one should be practically set up (i.e., be about half-full of prepared seawater and have algae ready to add) before you start to clean the old cultures (the reason you only fill the new cultures half-way is that you want to leave room to wash larvae into the culture jar, and once you add the larvae and algae, you can top the culture jar up to the appropriate level). If you need to regulate the temperature of your culture vessels, you may want to set up water in advance and let it sit in the culture `bath' beside the cultures which you are going to change to ensure the temperature remains constant while changing the larvae. Some larval biologists add antibiotics to their long-term (i.e., several months) cultures to inhibit bacterial growth, but I do not think this is necessary at home unless your vessels are dirty or your technique is sloppy. If you find that your larvae are clumping together into lumps of debris, but still look healthy and active (aside from being stuck in a pile of flotsam), you likely have a problem with bacteria. This is why the cleaning of cultures is so important. Another alternative to stringent cleaning, which I do NOT recommend, is to treat your cultures with antibiotics during each cleaning, prior to the addition of larvae to the cleaned culture jar. (I am STRONGLY against the superficial and ignorant use of antibiotics at home, because with improper disposal - like down your sink or such - new bacteria are constantly exposed to the antibiotic, and eventually some individual that is resistant to that compound will have a huge selective advantage and rapidly spread. Chalk up another useless antibiotic to carelessness...)
For large-scale cultures, you want to gently pour the entire culture through the filters you made (see EQUIPMENT) to filter out any large particulate debris, retain the larvae and drain the water and bacteria out. It is VERY important when doing this that you keep the filters constantly submerged. What I mean is that you want to have your large mesh filter on top, your small mesh filter underneath (remember I said these should be made from stackable beakers, so they should only fit about halfway down into the one below it once you add the filter mesh and hot glue), and both of these should sit about halfway down (you don't want water to back up and overflow any of your larvae out the top of the filter). If you have made the beakers and found an appropriate height casserole dish (as described in EQUIPMENT), you should practice before you start culturing larvae so that you don't lose any of your work from inexperience once you start. Try pouring water through this a few times with a gallon jug of tapwater (DO NOT try this for the first time with your larvae, or you will likely be very upset when you lose them) to ensure that the first filter (large mesh) sits just below the water level of the water as you pour (although it is not essential that this pre-filter remain submerged, it is easier on the larvae if they pass through this filter while submerged, and it will decrease your losses with sensitive larvae); the second filter (small mesh) should ALWAYS remain submerged (this is the filter on which larvae will be retained). Practice swirling the culture jar with one hand while steadying the beakers (if they fall over, you've probably lost a LOT of work) and pouring the "culture" through the filters. When you're finished, you should have the waste debris on the large mesh pre-filter (which may or may not be underwater), the larvae on the small mesh filter (which must be underwater), and a casserole dish full of old culture water. At this point, you want to take new culture water and fill your spray bottle. Lift the pre-filter from the stack and discard it (to be washed later in hot water as well, although boiling is out, because it will melt the hot-melt glue). Swirl the small mesh filter in the casserole dish, trying to concentrate any particulate matter that made it through the prefilter towards the center of the filter. If you have good eyes or a hand lens, you should be able to carefully remove any particulates with an eye dropper or pipette, while leaving the larvae behind. This will become more important the longer the culture runs. Once you have checked the culture and/or removed excess organic wastes from the filter, lift the filter from the casserole dish, and VERY GENTLY spray the contents with the spray bottle (even pouring water from a glass or some such thing will probably work fine - you just want to rinse the larvae off in clean water, but need to do this as gently as possible), and then invert the filter and using the spray bottle, wash all the larvae into the new culture vessel. Once the larvae are added to the new culture jar, you should add the algae to the culture (again you want the culture jar just slightly cloudy - something on the order of 100,000 cells per ml to feed 1 larva per ml, adjusted accordingly). This procedure should be repeated every second day to ensure the larvae are well-fed and the bacterial populations are keep to an acceptable (minimum) level. Once you have added the larvae and algae, and have topped the cultures up to the appropriate level with seawater, you need to stir the cultures to prevent both the larvae and the algae from settling out of the culture water. There are a number of ways that larval biologists typically keep their cultures in suspension: 1) paddles, 2) aeration, and plankton wheels (in about that order). The construction of paddles involves setting up a frame with a long stemmed paddle hanging down into each jar, and setting a small motor with a wheel on it to drive the paddles. The motor should be a very slow motor (about 5-15 rpm) and should be mounted to a wheel with a shaft on one edge. To that shaft you attach a rope such that when the wheel turns the shaft directly away from the cultures, the paddles are pulled toward the motor and when directly towards the cultures, the paddles relax back to their original position. The paddles are typically made from some flexible material such as thin acrylic, or are more rigid but are suspended from strings or wires above the culture bath. Jars are then placed such that the `resting' position of the paddles is near the "back" wall of the culture jar, and motion of the motor pulls them towards the "front" wall, stirring the culture and keeping everything in suspension. This technique is probably the most commonly used in larval biology for research, but I personally use the second option for my research, and think it a far better choice for home culture. For aeration, you need a good quality gang-valve, and a length of silicone tubing for each culture you intend to keep - these should all be run off a good quality (preferably variable output) airpump. To each culture jar, you run a length of airline tubing, and simply slide a glass pipette (I use the 23 cm long-tip Pasteur pipettes because they work best, but any eyedropper tube will probably work) over the airline tubing (it should the exact size to slip the airline tubing snugly into the back end of the pipette) and place the tip of the pipette into one corner of the culture jar. Then adjust the air output to that line such that you get no more than 2 bubbles in the water at any time (that is about 2-3 bubbles per second from the pipette). This gentle airflow will be enough to keep the culture water turning over and both the larvae and the algae in suspension. The final method of keeping the culture in suspension is to use a plankton wheel, in which a slow motor (like that for the paddles) is used to turn a large diameter disc to which tightly sealed culture jars are attached directly. The wheel is usually made of wood with straps of rubber tubing used to affix the cultures to the wooden frame. This frame is usually set at about a 45 degree angle and the cultures are turned over as the wheel continually turns the entire culture vessel over (upside-down to upside-right as the wheel turns). This is probably the least common and most difficult method used to culture larvae and is surely the most impractical to use at home.

Larval Settlement

Now here is a subject scientists know very little about. Larval settlement has been a strong field of interest since Gunnar Thorson arguably made all the important discoveries in the field in the 1950's (this ought to cause some controversy with my colleagues!). In general, I don't think there is a single species for which the basis of settlement is known unequivocally - there are several (still not that many, though), for which a chemical settlement inducer has been identified (and every one is a different class of molecule altogether), but someone somewhere contests virtually every settlement inducer currently known. The problem with larval settlement, and why I even bother to mention it herein, is that there are still a number of species of marine invertebrates, for which there is no hope of culture, because we know nothing about the settlement requirements or inducers. I have raised several species of nudibranchs only to have the larvae remain in culture until they begin to die, as I try desperately to give them everything I could think of to settle on. If you can't get the larvae to settle (fortunately, there are a few chemical ways to cheat here), there is no point in going through the time and effort of raising the larvae in the first place. Determining whether or not larvae are competent to settle is a whole other issue. Many lecithotrophic larvae spend as little as minutes to hours in the water column, do not feed and need very little culture effort (however, even some lecithotrophic larvae spend in excess of a year swimming in the water column); planktotrophic larvae feed and typically spend weeks, months or even over a year in the plankton, and some need a huge culture effort to raise any juveniles. Larvae are said to be competent when they become capable of settling (and metamorphosing) if provided with the appropriate cue. If you have good culture techniques, and you see that larvae are not growing despite appearing healthy (competent larvae typically reach a given size and then keep the status quo), it is likely that your larvae are competent but have not yet been provided with the appropriate cue to metamorphose.
In general, there are many species which need only reach a certain developmental stage and be presented with a hard substrate from your tank on which to settle and metamorphose (virtually all species need a surface which has become covered with bacteria and other organics - called "biofilm" - before they will settle) into the adult body form, but there are also many species which need some specific (and unknown) cue to settle and metamorphose. It is these latter species which are of concern. If you are confident that the larvae are competent to settle, but refuse to do so for whatever reason, you can attempt to induce settlement and metamorphosis artificially in many taxa (but apparently not in most anthozoans, bivalves or tunicates) by raising the K+ ionic concentration by about 5-10 milliMoles over standard seawater (e.g., Pechenik and Gee 1993, Pearce and Scheibling 1994, Wendt and Woollacott 1995) with KCl. If dosing with KCl in various concentrations does not work (the required dose varies and settlement is typically inhibited by high doses of KCl, so try a series of concentrations from very low to VERY VERY low), you are on your own for experimenting to figure out what the larvae require to settle and metamorphose. Larval biologists typically try prey species, certain algae common in the same habitats or conspecifics (other living adults of the same species) as good choices to induce settlement. If these don't work, your guess will likely be as good as mine as to what to try next....

Summing Up

Well, I hope that this has given you some idea of the difficulties associated with raising marine invertebrates, but also has shown you how to be successful at it, should you decide to try it yourself. If you have any questions, comments, suggestions, improvements, etc., please feel free to contact me - I don't guarantee you that I'll answer right away (Sorry, but playing on *.aquaria is my hobby, and it has to come a distant second to my responsibilities at work! Besides, I am often in the field for weeks or even months at a time), but I will try to get back to you eventually. If not re-send the post some time later to flog my feeble memory.
I also wanted to thank the members of the breeders registry, and in particular Stan Brown, for comments, feedback and suggestions regarding this document.

Literature Cited

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Guillard, R.R.L. 1975. Culture of phytoplankton for feeding marine invertebrates. In: Culture of Marine Invertebrate Animals. W.L. Smith and M.H. Chanley (eds.), Plenum Publishing Corp., New York, NY. pp. 29-60.

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By Rob Toonen

Section of Evolution and Ecology
University of California
Davis, CA