Things I Did Right
(for a change!)
I outgrew my first reef tank, a 30-gallon acrylic aquarium with a built-in trickle filter, in eight months. The 90-gallon glass tank that followed was running for two years before it, too, was outgrown. This article is about the upgrade to my new 220-gallon glass tank. With each of my first two tanks I learned invaluable lessons that became more firmly engrained in a direct relationship to the amount of time it took to clean up the mess. (I wish I had a dime for every quart of water I've mopped off the floor.) Now, sitting in front of my gorgeous new tank, I'm finally able to say "I think I did it right this time." This is not to say that everything is perfect; it simply means I think I've done more things right than wrong. The tank has been up and running for nine weeks now, and I'm very happy with both the setup of the system and the improvements in the overall environment for my animals.
From a maintenance standpoint, the design of the stand and canopy was critical. My husband, while never claiming to be a carpenter, nonetheless designed and built both the stand and canopy using my 90-gallon stand and a set of Do-It-Yourself plans from Reef Central as guides. He was determined to correct some of the shortcomings of my previous stand and he did a wonderful job, if I do say so myself. One of these improvements was the interior finishing. The sides and bottom of the new stand are lined with Formica (the material that is laminated to particle board to make kitchen countertops) and all the inside seams are caulked to make it waterproof. The interior of the canopy was primed and painted white. As is often the case, my previous stand was lined with unfinished particleboard that would swell at the mere mention of the word "water". (I believe a little time spent waterproofing the inside of any stand and canopy prior to installation is well worth the effort.) The design of the canopy is simply ingenious. It was built with a removable front panel secured by wooden dowels.
With the front panel removed, the canopy slides backwards, or forwards, up to ten inches, allowing complete access to the front or rear walls of the tank. I can now do almost any type of maintenance, including removing or placing large rocks, without having to work through the doors in the front of the canopy or needing assistance to remove the canopy.
The second important maintenance improvement was the placement of the tank twelve inches from the wall. Some of you may be thinking, "So what? Everyone knows that." Go ahead, snicker if you must, but my 90-gallon tank was set so close to the wall that I could not get behind it. (Well, you don't want to be able to see all the ugly equipment BEHIND the tank, now do you?) This poor planning was never so apparent as when the main circulation pump failed and had to be replaced. Removing it required contortions that no one, save a yoga master, should ever attempt.
I solved my aesthetics issue of the large gap behind the tank with a well-placed corner shelf that nicely hides the equipment in back. (Also ingeniously built, the shelf is attached to the wall on the left side and swings open like a door to allow easy access to the back of the tank.)
Prior to its purchase, I put some serious thought into the dimensions of my new tank. I chose a tank that is 72" long by 30" wide by 24" deep for several reasons. The large footprint allows maximum space for creative aquascaping and an extensive sand bed area. All of the rockwork is far enough away from the glass to allow me to use a magnet cleaner on all the viewable surfaces without catching the retrieval string on any delicate coral branches. With the 24" depth I can reach the floor of the tank with my hand by standing on a stepstool. Yes, I know there are numerous long-handled tools available for deeper tanks, but I still find it easier to do tasks, such as moving a coral, by hand.
The design of the tank's circulation and plumbing scheme was also carefully considered. The tank was predrilled for two drains in the built-in overflow box, as well as three holes across the back wall of the tank for the circulation pump return lines. I use two pumps whose output is split into two lines each. Three of the four return lines are attached directly to the tank through bulkheads fitted with flexible ball-socket tubing that allow me to direct the water flow to cover the whole tank. The fourth line is connected to a SeaSwirl oscillator mounted to the edge of the tank. This means I am not using any power heads inside the tank. I see this as a major coup, since not only does this eliminate the chance of animals being injured or killed by being sucked into a power head, but it also eliminates the extreme irritation of suction cups that don't… suck. (Or perhaps they do and that's the point!). In my previous tank, I had quite the sand storm when the power head came loose and aimed directly at the bottom. It could have been worse, though; I was lucky the power head didn't aim straight up and pump untold gallons of water all over my floor!
The two drains in the overflow box are fitted with PVC standpipes drilled with holes and capped with U-shaped fittings. The fittings were made from two 90-degree PVC elbows glued together fitted with a short piece of PVC pipe at the end. The standpipes keep the overflow box full of water, so there is no sound of water crashing into the box. The U-fittings put the intake just below the waterline, so there is no sound from the water being sucked into the standpipes. In fact, my new tank is much quieter than my 90-gallon tank that always had that slightly irritating bathtub-draining sound. Other plumbing improvements include using ball valves in front of each pump and check valves on the return lines into the tank. The ball valves allow easy removal of the main circulation pumps for maintenance or repair, and the check valves prevent the sump from overflowing in the event of a power outage. I opted for two 1200 gallon per hour return pumps rather than one larger pump to allow for a pump failure without compromising the whole system.
Electrical improvements were not overlooked in designing the new system. First, my husband installed a shop-grade five-foot power strip on the inside back wall of the stand. This keeps all electrical plugs off the floor and away from any water spills. Additionally, the spacing of the power strip is ideal for the X-10 appliance modules I use with my Neptune Aquacontroller. The controller itself was put on an uninterruptible power supply. Clean, consistent power to my controller means less chance of failure.
A water spill on the ballasts with my first tank taught me the futility of placing them on the floor. Mounting the ballasts to the sidewall of my previous 90-gallon stand protected them from wetting, but resulted in an 'octopus' of wiring and timers that was impossible to keep neat and took up too much valuable space inside the stand. All lighting ballasts are now securely mounted on top of the canopy. While my husband and I still have to build some sort of fence to hide the ballasts from the view of anyone taller than five foot three, I think this is a small price to pay.
I've also learned a thing or two from my previous endeavors about setting up the environment inside the tank. I believe my best discovery to date is the Orange County silicate sand (size 60) I bought from a local sand and gravel supplier.1 It comes in hundred pound bags and is used, for one thing, in ashtrays in hotel lobbies. The advantages to this sand are: it is inexpensive ($6.50 per hundred pounds); it does not need to be washed and does not cloud the tank; and, best of all, the sand bed critters seem to thrive in it. (I had used the same sand in my 90-gallon tank for over a year with great results. No clumping, good denitrification and healthy critter counts.) I poured five hundred pounds of sand in my tank, which made a six-inch sand bed, added my pre-mixed saltwater, and in less than an hour the water had cleared sufficiently to allow me to work with the live rock.
Concerning live rock, I've conquered my fear of the drill and masonry bit and found that it is really quite easy to drill holes in Fiji rock. Cable ties, pieces of solid plastic coat hangers (used as dowels) and underwater epoxy, work wonders to secure reef structures. I was able to build much more interesting structures with caves and arches using far less rock, secure in the knowledge that it will withstand the bulldozing of turbo snails or abalone.
Last, but certainly not least, I have learned to give all the animals plenty of growing space. It is truly amazing how, in less than a year, a coral fragment the size of a nickel became a colony the size of a softball.
I have vowed to resist the urge to buy too many new animals to fill up all the empty spaces in my new tank. I know my current animals will continue to grow and will expand into the extra space in their own good time.
Basically, I have to say I couldn't be happier with my new tank. I know that I owe its success to the experiences, hard lessons and spilt water from my first two tanks. Whatever doesn't drown you makes you better, right?
If you have any questions about this article, please visit the Notes from the Trenches forum on Reef Central.
from: reefkeeping.com
Senin, 30 Juni 2008
Additional Commentary on Dyed Corals
Additional Commentary on Dyed Corals
I apologize for not immediately continuing the series on coral nutrition, but Anthony Calfo's article in this issue prompts me to write on another subject this month. I'd like to thank Anthony for writing an informative article on this most disturbing recent trend to artificially dye corals. I feel it an important enough issue to warrant a second commentary on the subject. It amazes me that anyone would feel that the naturally glorious colors already present in corals and coral reef organisms should need to be enhanced.
The majority of the dyed corals entering the market began appearing more prolifically late in 2001 and originated in Indonesia. However, soon after the first reports began trickling in of dyed Sinularia and Turbinaria, in early 2002, I was unfortunate enough to have first hand (and unintentional) experience with a dyed coral. This time, its origin was Fiji, and many other reports of dyed Fijian corals, including ones not commonly exported from Indonesia, are now confirmed. Perhaps more disturbing is what I have ascertained may be an ulterior motivation for using dye in Fiji. The corals appear to be totally or partially bleached, and would otherwise be unsellable.
Indonesian dyed corals seemed to be enhanced to command either higher prices or to make them more desirable or attractive. The Indonesians are, as a culture, fond of bright colors, and it shows in their clothes and even their food! Being surrounded by the bright colors of nature both on land and sea, it is hardly surprising, and I rather suspect that deception is not always the primary motivation for this practice. It may very well be that those involved feel they are doing a service in the practice.
In Fiji, however, I feel the practice is clearly deceptive. Once again, Fiji has undergone significant bleaching events and many corals being cultured and collected have also bleached. This is truly an unfortunate predicament for those involved in the trade. However, the introduction of artificial colorants to mask a bleached coral and make it marketable is a practice that must not continue.
I ordered an orange morph of a Lobophyllia - an unusual, but by no means impossible, color morph. Lobophyllia generally are very hardy and transport well. However, this specimen arrived with tissue that appeared grossly swollen and abnormal. I suspected significant damage that appeared necrotic, and foresaw an impending jelly-type infection. I opted to treat the coral, and placed it in a quarantine tank with newly made seawater.
I changed the water several times over several days, and once the dye was eliminated from the tissues, it was apparent that the coral was clearly severely bleached. It is possible that the dye caused the bleaching prior to obtaining the coral, but logic would dictate otherwise. The coral is still alive, but its health has remained compromised and has been very sensitive to even slight aquarium water perturbations.
As to dyed corals, I personally collected threads and documented cases of dyed corals being sold from aquarists and stores across the country. I wrote a letter to the US Coral Reef Task Force, AMDA, Marine Aquarium Council, and seven separate persons in the Indonesian government and the coral collecting group AKKII about this issue. I have received responses from USFWS, NMFS, AKKII and other organizations in response, all saying that they would take steps to correct the problem. I have included some of the letters of response at the end of this article.
In terms of buying livestock from any source, certain corals are going to be wild collected. There are no alternatives. Similarly, there is no one source of dyed corals, and Indonesia recognizes that certain wholesalers, though only a few, may be participating in this practice. However, the operators of the facilities are apparently not doing the dying, and may not be knowledgeable enough to even know or recognize corals from collectors or middlemen that are dyed. Thus, orders are placed and corals are sent. No store ordering corals from Indonesia, and perhaps now even Fiji, will necessarily be assured of not receiving dyed corals. Unless a coral is aquacultured or collected from a known source, every store that sells wild collected corals will potentially be a source of dyed corals.
Given that the vast majority of corals collected for the aquarium trade are from Fiji and Indonesia, both now known to be sources of dyed corals, should these countries be economically boycotted? I don't believe so, and I am not sure that not supporting stores that may have dyed corals is a proper response. However, it is unquestionably the case that at least some suppliers, from exporter to retailer, are knowingly and intentionally purchasing and supplying dyed corals. For those cases, non-cooperative non-support may be a good choice. For others, making sure to inform the facility that the coral is dyed, and explaining the reasons why you don't approve is a good way to voice concern and dissatisfaction. Urging the facility to do the same to the supplier "upstream" is also a good idea. Education is the key to eliminating this practice, especially when coupled with ethical and economically compelling reasons.
As to the survival and health of dyed corals, I cannot say what the true effects the dyes have on coral tissue. To be able to investigate their effect on animal survival and health, it would be very helpful to know what are the dyes being used. Some may be non-toxic, at least some are clearly water soluble, and may be relatively innocuous - just gaudy and annoying! However, there is also a very real possibility that the dyes are directly toxic or indirectly causing stress, distress, and compromising the health of the treated corals. Calfo's thoughts and proposed effects of dye are possible, if not likely, to be true.
It appears that some corals tolerate being dyed better than others. For some, it appears that the dye is actively removed from the tissues and transported out of the coral through solenia or gastrovascular canals (Figure 3). For others, signs indicative of stress exist, including tissue recession, poor feeding response, poor polyp expansion, and increased secretion of mucus. In some cases, death appears to occur as result of the dye. However, studies would need to be done to be sure that mortality could indeed be attributed to the dye, either directly or indirectly.
In conclusion, I urge anyone seeing such corals to be pro-active and vocal in expressing their concern and dissatisfaction with the practice. Obviously, no one should intentionally purchase a dyed coral, as also mentioned and supported by Calfo (this issue). I would also welcome all reports and documentation of this practice wherever seen so that I can perhaps continue to use channels of inquiry and within trade groups and interested parties to help stop the practice. If you are able to help in this regard, the following information will be helpful, if not required, to be useful in supportive evidence of the practice:
1. Date
2. Coral type - genus level, species if possible
3. Color of dye
4. How many present
5. Name, address, phone number and contact at the facility
6. Photos, if possible
7. Supplier of facility
8. Country of origin for the specimen, if known
Furthermore, I would also request any reports and documentation of dyed corals that have been unintentionally purchased and are being cared for by an aquarist be sent to me. Particularly useful, in addition to the information above, would be:
1. Length of time since purchase
2. Length of time until dye is lost from the tissue (if ever)
3. General tank information
4. Obvious changes in behavior, health, or survival of the dyed coral or others in the tank
5. If the tissue is bleached or normally pigmented once the dye is lost
6. If there are any measured, observed, or anecdotal long-term consequences from the dye
7. Any other notable remarks, measurements, or photos
from:reefkeeping.com
I apologize for not immediately continuing the series on coral nutrition, but Anthony Calfo's article in this issue prompts me to write on another subject this month. I'd like to thank Anthony for writing an informative article on this most disturbing recent trend to artificially dye corals. I feel it an important enough issue to warrant a second commentary on the subject. It amazes me that anyone would feel that the naturally glorious colors already present in corals and coral reef organisms should need to be enhanced.
The majority of the dyed corals entering the market began appearing more prolifically late in 2001 and originated in Indonesia. However, soon after the first reports began trickling in of dyed Sinularia and Turbinaria, in early 2002, I was unfortunate enough to have first hand (and unintentional) experience with a dyed coral. This time, its origin was Fiji, and many other reports of dyed Fijian corals, including ones not commonly exported from Indonesia, are now confirmed. Perhaps more disturbing is what I have ascertained may be an ulterior motivation for using dye in Fiji. The corals appear to be totally or partially bleached, and would otherwise be unsellable.
Indonesian dyed corals seemed to be enhanced to command either higher prices or to make them more desirable or attractive. The Indonesians are, as a culture, fond of bright colors, and it shows in their clothes and even their food! Being surrounded by the bright colors of nature both on land and sea, it is hardly surprising, and I rather suspect that deception is not always the primary motivation for this practice. It may very well be that those involved feel they are doing a service in the practice.
In Fiji, however, I feel the practice is clearly deceptive. Once again, Fiji has undergone significant bleaching events and many corals being cultured and collected have also bleached. This is truly an unfortunate predicament for those involved in the trade. However, the introduction of artificial colorants to mask a bleached coral and make it marketable is a practice that must not continue.
I ordered an orange morph of a Lobophyllia - an unusual, but by no means impossible, color morph. Lobophyllia generally are very hardy and transport well. However, this specimen arrived with tissue that appeared grossly swollen and abnormal. I suspected significant damage that appeared necrotic, and foresaw an impending jelly-type infection. I opted to treat the coral, and placed it in a quarantine tank with newly made seawater.
I changed the water several times over several days, and once the dye was eliminated from the tissues, it was apparent that the coral was clearly severely bleached. It is possible that the dye caused the bleaching prior to obtaining the coral, but logic would dictate otherwise. The coral is still alive, but its health has remained compromised and has been very sensitive to even slight aquarium water perturbations.
As to dyed corals, I personally collected threads and documented cases of dyed corals being sold from aquarists and stores across the country. I wrote a letter to the US Coral Reef Task Force, AMDA, Marine Aquarium Council, and seven separate persons in the Indonesian government and the coral collecting group AKKII about this issue. I have received responses from USFWS, NMFS, AKKII and other organizations in response, all saying that they would take steps to correct the problem. I have included some of the letters of response at the end of this article.
In terms of buying livestock from any source, certain corals are going to be wild collected. There are no alternatives. Similarly, there is no one source of dyed corals, and Indonesia recognizes that certain wholesalers, though only a few, may be participating in this practice. However, the operators of the facilities are apparently not doing the dying, and may not be knowledgeable enough to even know or recognize corals from collectors or middlemen that are dyed. Thus, orders are placed and corals are sent. No store ordering corals from Indonesia, and perhaps now even Fiji, will necessarily be assured of not receiving dyed corals. Unless a coral is aquacultured or collected from a known source, every store that sells wild collected corals will potentially be a source of dyed corals.
Given that the vast majority of corals collected for the aquarium trade are from Fiji and Indonesia, both now known to be sources of dyed corals, should these countries be economically boycotted? I don't believe so, and I am not sure that not supporting stores that may have dyed corals is a proper response. However, it is unquestionably the case that at least some suppliers, from exporter to retailer, are knowingly and intentionally purchasing and supplying dyed corals. For those cases, non-cooperative non-support may be a good choice. For others, making sure to inform the facility that the coral is dyed, and explaining the reasons why you don't approve is a good way to voice concern and dissatisfaction. Urging the facility to do the same to the supplier "upstream" is also a good idea. Education is the key to eliminating this practice, especially when coupled with ethical and economically compelling reasons.
As to the survival and health of dyed corals, I cannot say what the true effects the dyes have on coral tissue. To be able to investigate their effect on animal survival and health, it would be very helpful to know what are the dyes being used. Some may be non-toxic, at least some are clearly water soluble, and may be relatively innocuous - just gaudy and annoying! However, there is also a very real possibility that the dyes are directly toxic or indirectly causing stress, distress, and compromising the health of the treated corals. Calfo's thoughts and proposed effects of dye are possible, if not likely, to be true.
It appears that some corals tolerate being dyed better than others. For some, it appears that the dye is actively removed from the tissues and transported out of the coral through solenia or gastrovascular canals (Figure 3). For others, signs indicative of stress exist, including tissue recession, poor feeding response, poor polyp expansion, and increased secretion of mucus. In some cases, death appears to occur as result of the dye. However, studies would need to be done to be sure that mortality could indeed be attributed to the dye, either directly or indirectly.
In conclusion, I urge anyone seeing such corals to be pro-active and vocal in expressing their concern and dissatisfaction with the practice. Obviously, no one should intentionally purchase a dyed coral, as also mentioned and supported by Calfo (this issue). I would also welcome all reports and documentation of this practice wherever seen so that I can perhaps continue to use channels of inquiry and within trade groups and interested parties to help stop the practice. If you are able to help in this regard, the following information will be helpful, if not required, to be useful in supportive evidence of the practice:
1. Date
2. Coral type - genus level, species if possible
3. Color of dye
4. How many present
5. Name, address, phone number and contact at the facility
6. Photos, if possible
7. Supplier of facility
8. Country of origin for the specimen, if known
Furthermore, I would also request any reports and documentation of dyed corals that have been unintentionally purchased and are being cared for by an aquarist be sent to me. Particularly useful, in addition to the information above, would be:
1. Length of time since purchase
2. Length of time until dye is lost from the tissue (if ever)
3. General tank information
4. Obvious changes in behavior, health, or survival of the dyed coral or others in the tank
5. If the tissue is bleached or normally pigmented once the dye is lost
6. If there are any measured, observed, or anecdotal long-term consequences from the dye
7. Any other notable remarks, measurements, or photos
from:reefkeeping.com
Naked ...Gills on Snails
Naked ...Gills on Snails
Well, it got your attention, didn't it? Nudibranch means "naked gills" and this month I thought I would briefly review these gorgeous and wonderful animals; animals that, almost without exception, have no place in aquaria, but which find their way there anyway.
Now, I will freely admit, right up front, I don't have many pictures of beautiful tropical nudibranchs, and so I won't put any pictures of them in this column. However, I will refer you to a site that has a lot of good images; tropical eye candy, if you will. Nudibranchs are exceptionally interesting creatures, and I will concentrate my efforts in this column discussing how to recognize them. In doing so, I hope to give you enough information that you can recognize a nudibranch when you see one, which is not a difficult proposition, and decipher a bit of its biology simply from its appearance, which is a bit more difficult.
For much information and a lot of good images, and references, go to The Sea Slug Forum.
Nudibranchs are mollusks. Following from last month's column on molluscan diversity, they therefore must have all those typical molluscan characters such as a radula, a foot, and specific types of gills, guts, and nervous system. This, they do, and possibly more importantly for the topic at hand, they reproduce in a standard molluscan manner. All nudibranchs are hermaphrodites. This means they have the sexual organs and plumbing of both genders, and they are frequently simultaneously active. During copulation, they both give and receive sperm. They cannot, however, fertilize themselves.
After mating, they lay eggs, often by the millions, all encased in some sort of jellylike mass. All of these develop into embryos, and these embryos develop into small larvae, which hatch from the egg mass. These larvae are called veligers, and each of them has the beginnings of small snail shell. The shell is transparent, because at this stage of the animal's life it can't secrete the calcareous shell of an adult snail; nevertheless, it is a perfectly good little snail shell. Nudibranch larvae look like any other good planktonic snail larvae, and live much the same way. They swim in the plankton, and feed on phytoplankton.
If they get enough food, and if it is the correct food, they will grow, and develop further into recognizable snails, which do have a calcareous shell. They are normally still planktonic when this occurs. During this period they are getting larger, and their internal structure is developing as they get more complex. Eventually, they become able to live on the bottom in the benthic environment. Biologists, who study larvae, refer to them at this stage as being "competent" to settle.
Competent larvae don't just rain out of the plankton down to the ocean floor and change into small snails. Instead, they swim down until they encounter the substrate and then they chemically analyze it. Or, to throw in a highly technical, jargon term from biology, they "taste" it. These larvae will not settle from the plankton until they find a suitable substrate, and if they don't find one, they perish. What constitutes an appropriate substrate varies with each nudibranch species. Often, it is something that is correlated with their adult prey. So, for example, a coral-eating nudibranch may regard a species of coralline algae typically found near the corals on which the adults feed as the appropriate substrate, as it provides a cue for future food.
If they find the substrate, they settle or touch down on the substrate. Following this they metamorphose, or change their body, into a small juvenile nudibranch. In this process, they discard the shell; it is simply shed into the water. They also generally reabsorb the larval feeding/swimming organ called a velum. Typically, the gut reorganizes; the cells capable of digesting phytoplankton die or change, and from this point forward the animal can only digest animal prey. All nudibranchs are carnivores. This process takes from a few hours to a few days, and during this time, the little nudibranch lives on fat reserves it accumulated during its larval planktonic period; it can't begin to feed until the gut has completed its changes. Additionally, during this time in the life of most nudibranchs, gills develop from the upper surface, which in other snails is covered by a shell. Consequently, as the shells have been shed, these animals have "naked gills" exposed to the environment.
See The Slugs, Er, Nudibranchs, Er, Sea Slugs…. What Are These Things, Anyway?
Humans look at the world and from this looking, they make decisions. We are predominantly visual animals and we share, with the other higher primates, a visual system that is probably the best in the natural world. So, it stands to reason that we often make sweeping generalizations based on exterior appearances. For example, politicians look like humans, close enough to fool some experts, but examination of their internal morphology will disclose they belong to a different sort of species. Snails without shells all really look alike. Basically, there is this blobby body which may or may not have any permanent appendages on it, and underneath is a muscular foot. And they secrete mucus. Always. Lots of it.
We call such animals, slugs, and recognize they all basically look alike. Unfortunately, the appearances of similarity are only skin deep, and actually in many cases, not even that, as their skins may be different, too. If all snails are in one group, the group we call the Class Gastropoda of the Phylum Mollusca, nudibranchs and all marine slugs, belong to a subdivision of that large group. They are put in the sub-grouping called the opisthobranchs. Branch, pronounced "brank" which rhymes with "bank," is a derivative of Branchia, a Greek word for the gills of fishes. We ignore the niceties that snails are not fishes and use the term anyway. Opistho- is a derivative of opisthen, meaning "at the back" or "behind," so this group of marine snails, the Opisthobranchia, are those snails with the gills behind, or at the back. Actually, they are the snails with their gills behind or to the rear of their hearts. Terrestrial slugs, by the by, don't belong to this group. They are all air breathers and have a lung. However, in their favor, even if they aren't as colorful as the nudibranchs, some terrestrial slug species are close to being the champion mucus producers of the animal kingdom.
Not all opisthobranchs are slugs; many of them have shells. As an aquarium example, the Pyramidellid snails that parasitize Tridacna, are also opisthobranchs, as are the "bubble shells." The groups of snails characterized by the pyrams and bubble shells have representatives that may have good shells. However, in the bubble shell group, the "head-shield snails" or Cephalaspidiceans, there appears to be a trend toward the loss of the shell, with some species having only a small internal shell, and looking quite sluggish.
So, what is a nudibranch, and what isn't, and how can you tell? More importantly, why should you care?
I will address the second question first. Many sea slugs (notice I was careful not to use the term "nudibranchs") are quite beneficial, as they may be predators on some of the algae that become problems in aquaria. An example of one such animal is the lettuce slug, Elysia cristata, which may eat various green algal pests. A similar situation is found with the sea hares. These beneficial animals all look "nudibranchish," but they are not, belonging to different groups, the Sacoglossans and the Aplysiids, respectively.
Nudibranchs, fortunately, are grouped together in what taxonomists call, the "Order Nudibranchia." Among some other things, this means that all of them have some characteristics in common that will help in the determination of whether or not a newly found slug becomes a pest or a pet. Unfortunately, the Order Nudibranchia is huge, with several thousand species having been scientifically described, and more being found regularly that are not yet described. Some of these undescribed species may be quite uncommon or very cryptic. I have personally found an example of one undescribed species, which has been subsequently described and named by a nudibranch taxonomist using the specimens I sent to her. I found this species in a very well-studied locality near a marine laboratory, and it has subsequently been found elsewhere, but all of these other collections have only been through environmental grab samples. As far as I know, I am still the only person to have seen a living example in its natural environment. It is a small, very cryptic, species, and looks like a small piece of organic leaf litter. It may well be common, but if so, it is commonly overlooked. Other scientifically undescribed or "new" species may be quite large and often very strikingly colored. During my stay, in the early 1970's, as a graduate student at the University of Washington Laboratories, several large and obvious species were found and named from localities around that marine station. That is the largest marine biological laboratory on the west coast of North America, and has been operational since early in the twentieth century, and yet new species of nudibranchs are still being found in that locality.
So, what can an aquarium hobbyist do when confronted with a sluggish blob? How can such a person identify it? Well, identification to species is likely impossible, even for the experts, as in many cases you have to know where it was found in nature to be able to separate it from dozens of other look-alike species. However, to identify the unknown critter as a nudibranch or some other snail, and then (if it is a nudibranch) to be able to identify it to a major group is certainly something that an observant aquarist should be able to do.
Probably the first thing you should do if you find an animal like this in your aquarium is to remove it from the system and put it in a clean container with some tank water. Once it relaxes enough to move around, use a good digital camera to take some pictures. Don't bother taking pictures with a camera that has less than about 3 Mpixels if the slug is small, as the enlarged image will be too blurry to decipher. You can often send good pictures to an expert and get a valid name for the animal.
Once the animal is moving around, examine it. Does it look like it has a hard lump in the middle or does the middle of the body appear swollen? If so, gently touch it. If you feel any hardness or resistance to your touch, it is likely a shell, and the animal isn't a nudibranch, and you probably have a bubble shell. Check references on the Sea Slug forum for more precise Cephalaspidacea identifications.
Figure 1. A diagram of a bubble shell or cephalaspidean.
Figure 2. A small bubble shell with an exposed shell. In most bubble shells, the shell is completely internal.
Figure 3. A small, 1/8th inch long, bubble shell found on a reef in Palau.
If the animal lacks an internal shell, or you don't feel one, examine the head as it is moving. There will be two large tentacles or projections pointing up and, often, forward from the front end. These tentacles are typically larger than other tentacles in the "head" region, if there are any. These are the main sensory organs for the animals, and are called rhinophores (in nudibranchs) or cephalic tentacles (in most other groups). Use a magnifying glass to examine them, and note their structure. If they appear to consist of a fold of tissue rolled into a cylindrical shape, the animal is a sacoglossan, such as Elysia, the lettuce slug, or a sea hare, such as Aplysia. Check other references for these animals.
Figure 4. Most sacoglossans are green and eat algae. I wanted to show you something different. This tiny slug, Olea hansinensis, about 1/50th of an inch long, finds its way to bubble shell egg masses and eats the eggs. I took this photo in nature using a 35 mm camera, and the image on the slide is in crisp focus, but even scanned at 4500 dpi. I couldn't get a crisp image here. The faint dots on the back of the slug are individual cells in the epidermis.
If the back of the animal is covered by a more-or-less smooth surface (there may be small bumps on it), and if the two sensory tentacles arise from under the front of this surface, not through it, you may have either a side-gill slug, or a nudibranch. Use a medicine dropper tip and gently lift up the dorsal surface on the right side of the animal. If there is an elongate structure covered under the flap of the dorsal surface, the animal is a side-gill slug, or Notaspidean. Remove the animal to another aquarium and consult other references for the biology of these animals. Most of them are predatory, and if bothered some will secrete sulfuric acid as a defensive agent. If there is no elongate structure along the right side, although there may be warts, bumps or circular depressions present there, you have an arminid nudibranch. Arminids eat soft corals, and will not be reef safe.
Figure 5. Berthella californica, a side-gill slug, or Notaspidean. The gill, peeking out from under the dorsal mantle, is normally invisible, being covered by the flap of tissue on the animal's right side.
Figure 6. An arminid nudibranch, Armina californica. Animals essentially identical to this individual, except for slight color differences, are found in the tropics.
If your specimen is unlike the examples covered so far, gently touch the tentacles with the dropper tip or some other glass probe. They should retract into sheaths of one sort or another. If so, you have one of the several other types of nudibranchs. None of them are generally reef safe, although a few are useful as they may prey on undesirable reef-aquarium animals such as Aiptasia.
Examine your critter again, this time concentrating at the general shape of the beast, and the number and kind of projections off of the top of the animal. If the animal looks rather like the slug version of a small brick, more-or-less rectangular in cross-section and with defined sharp right-angled corners at the front end, it is likely a "Dendronotacean." These often have two rows of projections called "cerata" on the back. Each row is found on the edge of the back where it meets the side of the animal, and these projections generally have small branches off of them. Dendronotaceans eat soft corals, sea pens, sea anemones, and jellyfish polyps. They are definitely not reef safe, and they appear in aquaria with some frequency. The largest dendronotaceans are found off the Pacific Coast of North America, where Tokuina tetraquetra may reach wet weights exceeding 20 pounds, and Dendronotus iris is commonly seen at lengths exceeding 16 inches. The Tritonia festiva in the slide show in this issue of Reefkeeping Magazine is also a dendronotacean.
Figure 7. Tokuina tetraquetra, the largest nudibranch in the world, is a dendronotacean. Note the small frilly gills along the sides of the flattened dorsal surface. This animal, a small individual, was about 15 inches long, and what is not readily apparent is that the top of the animal is about three inches above the substrate.
Figure 8. An unidentified dendronotacean, possibly a species of Doto, eating an aquarium coral. Photo courtesy of Skip Attix.
Upon examination, if the body is not rectangular in cross-section, and if any projections along the back are not found in linear rows along the back, then you have one of the other kinds of nudibranchs, most likely either a dorid, or an aeolid. Examples of each of these kinds of nudibranchs are found rather frequently, actually, in marine reef aquaria.
Aeolid nudibranchs are a bane for some aquarists, particularly those involved with stony coral culture, as they eat the corals. Identifying them is, fortunately, an easy process. The top surface of aeolids is covered by unbranched projections, also, as in the dendronotaceans, called cerata. These cerata may be arranged in linear rows, running from the front to the back of the animal, or in paired groups or clusters, one of each cluster on each side of the midline on the back of the animal. The smallest aeolids may have only one or two cerata, and are found living on, or between, sand grains. The largest ones cruise tropical and temperate seas in search of their prey. Aeolid cerata, or gills, are distinctive. If you have a "furry" nudibranch, you almost certainly have an aeolid. Examine one of the cerata, and you will truly see one of the wonders of nature.
The cerata have at their center a brown, tan, or beige colored tubular core. Occasionally, in some species a different color is found, but only rarely. This is an extension of the digestive gland in the gut running up through the gill to its tip. The digestive gland is the part of the molluscan gut which actually digests and assimilates food. In some ways it is an analog of the human small intestine. Aeolids are generally predators on corals, anemones, hydroids, and other cnidarians. Each nudibranch species is probably specialized to eat only one or a few species of prey. When they eat their prey, they typically bite off rather large chunks; large relative to the nudibranch, that is. In their digestive process, all the flesh is moved to the digestive gland where it is digested and assimilated. That is, with the exception of the stinging capsules or nematocysts, which are moved up the digestive gland tube in the center of each gill and come to reside in the tissue under the bright white tip, in what is called the cnidosac. The tips of the gills actually are usually the first thing one sees, when an aeolid is noticed.
Figure 9. Flabellina rufobranchialis, a common aeolid from the Northeastern Pacific. Note the brown digestive gland cores in the cerata, which are also tipped with the bright white cnidosacs.
The nematocysts in the cnidosac remain fully capable of discharging and are oriented so that their threads will fire through the epidermis covering the cerata. The nematocysts are provided with a nutritive environment, and immature nematocysts will mature in the cnidosac. Old nematocysts, those incapable of extending firing, are extruded from the cnidosac tips. Nematocysts in the cnidosac remain capable of firing for several days to a couple of weeks after the animal that secreted them has been eaten. If the aeolid is threatened by some potential predator, the nematocysts can be discharged, stinging the potential predator, and presumably deterring predation. As a result of this, relatively few animals will eat aeolids. Consequently, the prey of an aeolid not only nourishes it, but also protects its killer by providing, in the form of nematocysts, protection against predation.
Over the past couple of years, aeolid nudibranchs have been increasingly found in aquaria on Montipora colonies and other small-mouthed stony corals. These tiny aeolids are white or translucent gray, small, about one millimeter long to maybe about one centimeter, and are easily overlooked. They appear to rapidly pass totally through their larval stages in the egg mass, and hatch as crawl-away slugs. As such, once they are found infesting a tank, it may be very difficult to get rid of them, and if they become established in some dealer's tanks, they may be spread far and wide very rapidly. Others have been found just "roaming" tanks. Other aeolids, such as species of Berghia, have been used with varying success in aquaria to eradicate pest anemones such as Aiptasia. These species die when individuals of their prey become so rare that the slugs starve before finding more food. If this starvation occurs when they have eaten all of the pests, then they have succeeded. More often, it simply means they can't find the last of the pests, and with the last nudibranch gone, the pests return.
The remaining nudibranchs typically found in aquaria are in the group called the Doridacea, or the dorid nudibranchs. These animals are characterized by having their anus on the top midline of the body, about two-thirds to three-fourths of the distance from the front to the back. The anus is surrounded by a tuft of gills, which when extended look rather like a feather duster (the implement, not the worm). The rest of the top surface of the dorid may be either smooth, often to the point of "glossiness," or bumpy like the surface of a raspberry, or with appendages that look like the cerata of an aeolid. In these latter species, the cerata are not typically arranged in rows or clusters, but rather are scattered randomly over the back of the dorid. In all cases, however, the anus will be on the mid-line of the dorsal surface, and if the animal is "relaxed" will be surrounded by its tuft of gills.
Figure 10. Archidoris montereyensis, a typical dorid nudibranch. The front of the animal is at the lower right, and the two rhinophore tentacles are visible. The tuft of gills at the upper left surrounds the anus. All dorids have this basic gill and tentacle structure.
Dorids are often the most colorful of the nudibranchs and are often purchased by well-meaning aquarists who bring them home, only to watch them die. Dorids generally eat one of three different types of sessile or non-moving animals: sponge, tunicates, or bryozoans. Often the nudibranch species can only eat one or two closely related species of prey, so it is effectively impossible to keep dorid nudibranchs alive for extended periods. Not because aquarists can't provide the physical conditions necessary, but simply because the food is impossible to get. Few tropical dorids have been well studied, and in most cases we don't even know what they eat, so it would impossible to import food for them, even if we wished to.
Once you have a dorid in a tank, however, you really often want to keep it alive, if for no other reason, that they are often so filled with toxic chemicals that they can kill a tank upon their death. This is especially true for some of the beautiful brightly patterned blue, black and gold Phyllidia. (Check these out on the Sea Slug Forum). The bright, beautiful colors of nudibranchs are some of the best-known examples of aposematic or warning coloration. Warning coloration patterns are found on animals that in some way have a good defense against predators that hunt using vision, such as fishes or crabs. These bright colors and striking patterns are present to be visually obvious and thus to warn the predators away.
The system of dorid nudibranchs and their prey forms an interesting "double warning system." Sponges, tunicates, and bryozoans, the foods of dorid nudibranchs, are also often quite toxic, and brightly colored, providing a signal to fend off their own visually oriented predators. This is the first warning component, and in most cases, these poisons and bright colors work. Very few predators will eat these animals. However, one group of predators that has species that do eat them is the Doridacea, which specializes on these prey. Coincidently, the nudibranchs are blind and can't see the warning color on their prey. But that doesn't matter, as the poisons don't have any effect upon them.
When nudibranchs eat their toxic prey, they often modify the prey's own internal toxins. In many cases this makes them even more toxic. Dorid nudibranchs are typically animals that marine predators such as fish sample only once in their life. If they survive, they never eat or attempt to eat a nudibranch again, and the nudibranch's bright colors are there to tell them what to avoid eating, so this is the second warning in this particular system of predators and prey. As many marine fish live several years to several decades, this is a lesson that, once learned, is of significant benefit to the snails.
Aquarists often see some beautiful nudibranchs in their local fish stores, where they have been starving, and bring them home as an addition to the tank. Generally, the slugs continue to starve in the aquarists' tanks and within a few weeks they die. The toxins that they have accumulated during the life of the nudibranch may be released into the tank water and cause serious problems. These are animals truly best left in the wild, or if inadvertently collected, they are best left for someone else to deal with.
Conclusion:
I have tried here to give you a little bit of background on many nudibranchs and how to identify them to their major group. Once identified, you may follow links online or do some research at a library about them and try to determine if you really want to tackle the problems of trying to maintain them. They are truly beautiful and easy to maintain, once you have accepted the cost of that maintenance, which is, of course, the cost of their foods. Although often attractive, they are destructive of other decorative animals such as corals, soft corals, anemones, sponges, tunicates, and bryozoans. They are not reef safe unless your reef is very big indeed.
from: reefkeeping.com
Well, it got your attention, didn't it? Nudibranch means "naked gills" and this month I thought I would briefly review these gorgeous and wonderful animals; animals that, almost without exception, have no place in aquaria, but which find their way there anyway.
Now, I will freely admit, right up front, I don't have many pictures of beautiful tropical nudibranchs, and so I won't put any pictures of them in this column. However, I will refer you to a site that has a lot of good images; tropical eye candy, if you will. Nudibranchs are exceptionally interesting creatures, and I will concentrate my efforts in this column discussing how to recognize them. In doing so, I hope to give you enough information that you can recognize a nudibranch when you see one, which is not a difficult proposition, and decipher a bit of its biology simply from its appearance, which is a bit more difficult.
For much information and a lot of good images, and references, go to The Sea Slug Forum.
Nudibranchs are mollusks. Following from last month's column on molluscan diversity, they therefore must have all those typical molluscan characters such as a radula, a foot, and specific types of gills, guts, and nervous system. This, they do, and possibly more importantly for the topic at hand, they reproduce in a standard molluscan manner. All nudibranchs are hermaphrodites. This means they have the sexual organs and plumbing of both genders, and they are frequently simultaneously active. During copulation, they both give and receive sperm. They cannot, however, fertilize themselves.
After mating, they lay eggs, often by the millions, all encased in some sort of jellylike mass. All of these develop into embryos, and these embryos develop into small larvae, which hatch from the egg mass. These larvae are called veligers, and each of them has the beginnings of small snail shell. The shell is transparent, because at this stage of the animal's life it can't secrete the calcareous shell of an adult snail; nevertheless, it is a perfectly good little snail shell. Nudibranch larvae look like any other good planktonic snail larvae, and live much the same way. They swim in the plankton, and feed on phytoplankton.
If they get enough food, and if it is the correct food, they will grow, and develop further into recognizable snails, which do have a calcareous shell. They are normally still planktonic when this occurs. During this period they are getting larger, and their internal structure is developing as they get more complex. Eventually, they become able to live on the bottom in the benthic environment. Biologists, who study larvae, refer to them at this stage as being "competent" to settle.
Competent larvae don't just rain out of the plankton down to the ocean floor and change into small snails. Instead, they swim down until they encounter the substrate and then they chemically analyze it. Or, to throw in a highly technical, jargon term from biology, they "taste" it. These larvae will not settle from the plankton until they find a suitable substrate, and if they don't find one, they perish. What constitutes an appropriate substrate varies with each nudibranch species. Often, it is something that is correlated with their adult prey. So, for example, a coral-eating nudibranch may regard a species of coralline algae typically found near the corals on which the adults feed as the appropriate substrate, as it provides a cue for future food.
If they find the substrate, they settle or touch down on the substrate. Following this they metamorphose, or change their body, into a small juvenile nudibranch. In this process, they discard the shell; it is simply shed into the water. They also generally reabsorb the larval feeding/swimming organ called a velum. Typically, the gut reorganizes; the cells capable of digesting phytoplankton die or change, and from this point forward the animal can only digest animal prey. All nudibranchs are carnivores. This process takes from a few hours to a few days, and during this time, the little nudibranch lives on fat reserves it accumulated during its larval planktonic period; it can't begin to feed until the gut has completed its changes. Additionally, during this time in the life of most nudibranchs, gills develop from the upper surface, which in other snails is covered by a shell. Consequently, as the shells have been shed, these animals have "naked gills" exposed to the environment.
See The Slugs, Er, Nudibranchs, Er, Sea Slugs…. What Are These Things, Anyway?
Humans look at the world and from this looking, they make decisions. We are predominantly visual animals and we share, with the other higher primates, a visual system that is probably the best in the natural world. So, it stands to reason that we often make sweeping generalizations based on exterior appearances. For example, politicians look like humans, close enough to fool some experts, but examination of their internal morphology will disclose they belong to a different sort of species. Snails without shells all really look alike. Basically, there is this blobby body which may or may not have any permanent appendages on it, and underneath is a muscular foot. And they secrete mucus. Always. Lots of it.
We call such animals, slugs, and recognize they all basically look alike. Unfortunately, the appearances of similarity are only skin deep, and actually in many cases, not even that, as their skins may be different, too. If all snails are in one group, the group we call the Class Gastropoda of the Phylum Mollusca, nudibranchs and all marine slugs, belong to a subdivision of that large group. They are put in the sub-grouping called the opisthobranchs. Branch, pronounced "brank" which rhymes with "bank," is a derivative of Branchia, a Greek word for the gills of fishes. We ignore the niceties that snails are not fishes and use the term anyway. Opistho- is a derivative of opisthen, meaning "at the back" or "behind," so this group of marine snails, the Opisthobranchia, are those snails with the gills behind, or at the back. Actually, they are the snails with their gills behind or to the rear of their hearts. Terrestrial slugs, by the by, don't belong to this group. They are all air breathers and have a lung. However, in their favor, even if they aren't as colorful as the nudibranchs, some terrestrial slug species are close to being the champion mucus producers of the animal kingdom.
Not all opisthobranchs are slugs; many of them have shells. As an aquarium example, the Pyramidellid snails that parasitize Tridacna, are also opisthobranchs, as are the "bubble shells." The groups of snails characterized by the pyrams and bubble shells have representatives that may have good shells. However, in the bubble shell group, the "head-shield snails" or Cephalaspidiceans, there appears to be a trend toward the loss of the shell, with some species having only a small internal shell, and looking quite sluggish.
So, what is a nudibranch, and what isn't, and how can you tell? More importantly, why should you care?
I will address the second question first. Many sea slugs (notice I was careful not to use the term "nudibranchs") are quite beneficial, as they may be predators on some of the algae that become problems in aquaria. An example of one such animal is the lettuce slug, Elysia cristata, which may eat various green algal pests. A similar situation is found with the sea hares. These beneficial animals all look "nudibranchish," but they are not, belonging to different groups, the Sacoglossans and the Aplysiids, respectively.
Nudibranchs, fortunately, are grouped together in what taxonomists call, the "Order Nudibranchia." Among some other things, this means that all of them have some characteristics in common that will help in the determination of whether or not a newly found slug becomes a pest or a pet. Unfortunately, the Order Nudibranchia is huge, with several thousand species having been scientifically described, and more being found regularly that are not yet described. Some of these undescribed species may be quite uncommon or very cryptic. I have personally found an example of one undescribed species, which has been subsequently described and named by a nudibranch taxonomist using the specimens I sent to her. I found this species in a very well-studied locality near a marine laboratory, and it has subsequently been found elsewhere, but all of these other collections have only been through environmental grab samples. As far as I know, I am still the only person to have seen a living example in its natural environment. It is a small, very cryptic, species, and looks like a small piece of organic leaf litter. It may well be common, but if so, it is commonly overlooked. Other scientifically undescribed or "new" species may be quite large and often very strikingly colored. During my stay, in the early 1970's, as a graduate student at the University of Washington Laboratories, several large and obvious species were found and named from localities around that marine station. That is the largest marine biological laboratory on the west coast of North America, and has been operational since early in the twentieth century, and yet new species of nudibranchs are still being found in that locality.
So, what can an aquarium hobbyist do when confronted with a sluggish blob? How can such a person identify it? Well, identification to species is likely impossible, even for the experts, as in many cases you have to know where it was found in nature to be able to separate it from dozens of other look-alike species. However, to identify the unknown critter as a nudibranch or some other snail, and then (if it is a nudibranch) to be able to identify it to a major group is certainly something that an observant aquarist should be able to do.
Probably the first thing you should do if you find an animal like this in your aquarium is to remove it from the system and put it in a clean container with some tank water. Once it relaxes enough to move around, use a good digital camera to take some pictures. Don't bother taking pictures with a camera that has less than about 3 Mpixels if the slug is small, as the enlarged image will be too blurry to decipher. You can often send good pictures to an expert and get a valid name for the animal.
Once the animal is moving around, examine it. Does it look like it has a hard lump in the middle or does the middle of the body appear swollen? If so, gently touch it. If you feel any hardness or resistance to your touch, it is likely a shell, and the animal isn't a nudibranch, and you probably have a bubble shell. Check references on the Sea Slug forum for more precise Cephalaspidacea identifications.
Figure 1. A diagram of a bubble shell or cephalaspidean.
Figure 2. A small bubble shell with an exposed shell. In most bubble shells, the shell is completely internal.
Figure 3. A small, 1/8th inch long, bubble shell found on a reef in Palau.
If the animal lacks an internal shell, or you don't feel one, examine the head as it is moving. There will be two large tentacles or projections pointing up and, often, forward from the front end. These tentacles are typically larger than other tentacles in the "head" region, if there are any. These are the main sensory organs for the animals, and are called rhinophores (in nudibranchs) or cephalic tentacles (in most other groups). Use a magnifying glass to examine them, and note their structure. If they appear to consist of a fold of tissue rolled into a cylindrical shape, the animal is a sacoglossan, such as Elysia, the lettuce slug, or a sea hare, such as Aplysia. Check other references for these animals.
Figure 4. Most sacoglossans are green and eat algae. I wanted to show you something different. This tiny slug, Olea hansinensis, about 1/50th of an inch long, finds its way to bubble shell egg masses and eats the eggs. I took this photo in nature using a 35 mm camera, and the image on the slide is in crisp focus, but even scanned at 4500 dpi. I couldn't get a crisp image here. The faint dots on the back of the slug are individual cells in the epidermis.
If the back of the animal is covered by a more-or-less smooth surface (there may be small bumps on it), and if the two sensory tentacles arise from under the front of this surface, not through it, you may have either a side-gill slug, or a nudibranch. Use a medicine dropper tip and gently lift up the dorsal surface on the right side of the animal. If there is an elongate structure covered under the flap of the dorsal surface, the animal is a side-gill slug, or Notaspidean. Remove the animal to another aquarium and consult other references for the biology of these animals. Most of them are predatory, and if bothered some will secrete sulfuric acid as a defensive agent. If there is no elongate structure along the right side, although there may be warts, bumps or circular depressions present there, you have an arminid nudibranch. Arminids eat soft corals, and will not be reef safe.
Figure 5. Berthella californica, a side-gill slug, or Notaspidean. The gill, peeking out from under the dorsal mantle, is normally invisible, being covered by the flap of tissue on the animal's right side.
Figure 6. An arminid nudibranch, Armina californica. Animals essentially identical to this individual, except for slight color differences, are found in the tropics.
If your specimen is unlike the examples covered so far, gently touch the tentacles with the dropper tip or some other glass probe. They should retract into sheaths of one sort or another. If so, you have one of the several other types of nudibranchs. None of them are generally reef safe, although a few are useful as they may prey on undesirable reef-aquarium animals such as Aiptasia.
Examine your critter again, this time concentrating at the general shape of the beast, and the number and kind of projections off of the top of the animal. If the animal looks rather like the slug version of a small brick, more-or-less rectangular in cross-section and with defined sharp right-angled corners at the front end, it is likely a "Dendronotacean." These often have two rows of projections called "cerata" on the back. Each row is found on the edge of the back where it meets the side of the animal, and these projections generally have small branches off of them. Dendronotaceans eat soft corals, sea pens, sea anemones, and jellyfish polyps. They are definitely not reef safe, and they appear in aquaria with some frequency. The largest dendronotaceans are found off the Pacific Coast of North America, where Tokuina tetraquetra may reach wet weights exceeding 20 pounds, and Dendronotus iris is commonly seen at lengths exceeding 16 inches. The Tritonia festiva in the slide show in this issue of Reefkeeping Magazine is also a dendronotacean.
Figure 7. Tokuina tetraquetra, the largest nudibranch in the world, is a dendronotacean. Note the small frilly gills along the sides of the flattened dorsal surface. This animal, a small individual, was about 15 inches long, and what is not readily apparent is that the top of the animal is about three inches above the substrate.
Figure 8. An unidentified dendronotacean, possibly a species of Doto, eating an aquarium coral. Photo courtesy of Skip Attix.
Upon examination, if the body is not rectangular in cross-section, and if any projections along the back are not found in linear rows along the back, then you have one of the other kinds of nudibranchs, most likely either a dorid, or an aeolid. Examples of each of these kinds of nudibranchs are found rather frequently, actually, in marine reef aquaria.
Aeolid nudibranchs are a bane for some aquarists, particularly those involved with stony coral culture, as they eat the corals. Identifying them is, fortunately, an easy process. The top surface of aeolids is covered by unbranched projections, also, as in the dendronotaceans, called cerata. These cerata may be arranged in linear rows, running from the front to the back of the animal, or in paired groups or clusters, one of each cluster on each side of the midline on the back of the animal. The smallest aeolids may have only one or two cerata, and are found living on, or between, sand grains. The largest ones cruise tropical and temperate seas in search of their prey. Aeolid cerata, or gills, are distinctive. If you have a "furry" nudibranch, you almost certainly have an aeolid. Examine one of the cerata, and you will truly see one of the wonders of nature.
The cerata have at their center a brown, tan, or beige colored tubular core. Occasionally, in some species a different color is found, but only rarely. This is an extension of the digestive gland in the gut running up through the gill to its tip. The digestive gland is the part of the molluscan gut which actually digests and assimilates food. In some ways it is an analog of the human small intestine. Aeolids are generally predators on corals, anemones, hydroids, and other cnidarians. Each nudibranch species is probably specialized to eat only one or a few species of prey. When they eat their prey, they typically bite off rather large chunks; large relative to the nudibranch, that is. In their digestive process, all the flesh is moved to the digestive gland where it is digested and assimilated. That is, with the exception of the stinging capsules or nematocysts, which are moved up the digestive gland tube in the center of each gill and come to reside in the tissue under the bright white tip, in what is called the cnidosac. The tips of the gills actually are usually the first thing one sees, when an aeolid is noticed.
Figure 9. Flabellina rufobranchialis, a common aeolid from the Northeastern Pacific. Note the brown digestive gland cores in the cerata, which are also tipped with the bright white cnidosacs.
The nematocysts in the cnidosac remain fully capable of discharging and are oriented so that their threads will fire through the epidermis covering the cerata. The nematocysts are provided with a nutritive environment, and immature nematocysts will mature in the cnidosac. Old nematocysts, those incapable of extending firing, are extruded from the cnidosac tips. Nematocysts in the cnidosac remain capable of firing for several days to a couple of weeks after the animal that secreted them has been eaten. If the aeolid is threatened by some potential predator, the nematocysts can be discharged, stinging the potential predator, and presumably deterring predation. As a result of this, relatively few animals will eat aeolids. Consequently, the prey of an aeolid not only nourishes it, but also protects its killer by providing, in the form of nematocysts, protection against predation.
Over the past couple of years, aeolid nudibranchs have been increasingly found in aquaria on Montipora colonies and other small-mouthed stony corals. These tiny aeolids are white or translucent gray, small, about one millimeter long to maybe about one centimeter, and are easily overlooked. They appear to rapidly pass totally through their larval stages in the egg mass, and hatch as crawl-away slugs. As such, once they are found infesting a tank, it may be very difficult to get rid of them, and if they become established in some dealer's tanks, they may be spread far and wide very rapidly. Others have been found just "roaming" tanks. Other aeolids, such as species of Berghia, have been used with varying success in aquaria to eradicate pest anemones such as Aiptasia. These species die when individuals of their prey become so rare that the slugs starve before finding more food. If this starvation occurs when they have eaten all of the pests, then they have succeeded. More often, it simply means they can't find the last of the pests, and with the last nudibranch gone, the pests return.
The remaining nudibranchs typically found in aquaria are in the group called the Doridacea, or the dorid nudibranchs. These animals are characterized by having their anus on the top midline of the body, about two-thirds to three-fourths of the distance from the front to the back. The anus is surrounded by a tuft of gills, which when extended look rather like a feather duster (the implement, not the worm). The rest of the top surface of the dorid may be either smooth, often to the point of "glossiness," or bumpy like the surface of a raspberry, or with appendages that look like the cerata of an aeolid. In these latter species, the cerata are not typically arranged in rows or clusters, but rather are scattered randomly over the back of the dorid. In all cases, however, the anus will be on the mid-line of the dorsal surface, and if the animal is "relaxed" will be surrounded by its tuft of gills.
Figure 10. Archidoris montereyensis, a typical dorid nudibranch. The front of the animal is at the lower right, and the two rhinophore tentacles are visible. The tuft of gills at the upper left surrounds the anus. All dorids have this basic gill and tentacle structure.
Dorids are often the most colorful of the nudibranchs and are often purchased by well-meaning aquarists who bring them home, only to watch them die. Dorids generally eat one of three different types of sessile or non-moving animals: sponge, tunicates, or bryozoans. Often the nudibranch species can only eat one or two closely related species of prey, so it is effectively impossible to keep dorid nudibranchs alive for extended periods. Not because aquarists can't provide the physical conditions necessary, but simply because the food is impossible to get. Few tropical dorids have been well studied, and in most cases we don't even know what they eat, so it would impossible to import food for them, even if we wished to.
Once you have a dorid in a tank, however, you really often want to keep it alive, if for no other reason, that they are often so filled with toxic chemicals that they can kill a tank upon their death. This is especially true for some of the beautiful brightly patterned blue, black and gold Phyllidia. (Check these out on the Sea Slug Forum). The bright, beautiful colors of nudibranchs are some of the best-known examples of aposematic or warning coloration. Warning coloration patterns are found on animals that in some way have a good defense against predators that hunt using vision, such as fishes or crabs. These bright colors and striking patterns are present to be visually obvious and thus to warn the predators away.
The system of dorid nudibranchs and their prey forms an interesting "double warning system." Sponges, tunicates, and bryozoans, the foods of dorid nudibranchs, are also often quite toxic, and brightly colored, providing a signal to fend off their own visually oriented predators. This is the first warning component, and in most cases, these poisons and bright colors work. Very few predators will eat these animals. However, one group of predators that has species that do eat them is the Doridacea, which specializes on these prey. Coincidently, the nudibranchs are blind and can't see the warning color on their prey. But that doesn't matter, as the poisons don't have any effect upon them.
When nudibranchs eat their toxic prey, they often modify the prey's own internal toxins. In many cases this makes them even more toxic. Dorid nudibranchs are typically animals that marine predators such as fish sample only once in their life. If they survive, they never eat or attempt to eat a nudibranch again, and the nudibranch's bright colors are there to tell them what to avoid eating, so this is the second warning in this particular system of predators and prey. As many marine fish live several years to several decades, this is a lesson that, once learned, is of significant benefit to the snails.
Aquarists often see some beautiful nudibranchs in their local fish stores, where they have been starving, and bring them home as an addition to the tank. Generally, the slugs continue to starve in the aquarists' tanks and within a few weeks they die. The toxins that they have accumulated during the life of the nudibranch may be released into the tank water and cause serious problems. These are animals truly best left in the wild, or if inadvertently collected, they are best left for someone else to deal with.
Conclusion:
I have tried here to give you a little bit of background on many nudibranchs and how to identify them to their major group. Once identified, you may follow links online or do some research at a library about them and try to determine if you really want to tackle the problems of trying to maintain them. They are truly beautiful and easy to maintain, once you have accepted the cost of that maintenance, which is, of course, the cost of their foods. Although often attractive, they are destructive of other decorative animals such as corals, soft corals, anemones, sponges, tunicates, and bryozoans. They are not reef safe unless your reef is very big indeed.
from: reefkeeping.com
Captive Rearing of Peppermint Shrimp (Lysmata wurdemanni):
Captive Rearing of Peppermint Shrimp (Lysmata wurdemanni):
A Hobbyist's Tale
April Kirkendoll writes in the conclusion of her entertaining book entitled "How to Raise & Train Your Peppermint Shrimp" to, 'tell the world what you discover and don't skimp on the details. Our hobby may depend on it." I sincerely believe these words, and have tried to detail as much of my experiences attempting to raise Peppermint shrimp, so that you might not only observe many of the aspects of shrimp growth, but also may learn and develop novel ways of increasing the post-larval settlement survival rate.
For the first three years of trying to spawn and raise Peppermint shrimp, I could never get the shrimp larvae to survive more than a week or so before they died en masse. Recently, however, I was able to rear them to 5 weeks before they died by feeding copepods, in addition to baby brine shrimp (bbs) and plankton flake food. Here's my hobbyist's tale.
Figure 1. Breeding Adult
The Breeding Adults
Approximately three years ago, I obtained two small Peppermint shrimp, Lysmata wurdemanni, from a local fish store. Since then I've been able to capture roughly half of the larvae (about 50 larvae captured) almost every month. No special attention is given to the adult shrimp. Generally, frozen Mysis and brine shrimp are fed to the tank every day, as well as Tahitian Blend algae paste (see note 1). The adult shrimp will generally eat anything that they can catch and tear apart with their pincers. The shrimp are full sized adults (3") ,and regularly produce free swimming larvae every month (about 100 larvae at each event).
Capturing Larvae
Spawning inevitably occurs late at night, usually around midnight. If the larvae are not removed immediately from a fully stocked reef tank, they stand very little chance of survival because of predation by fish. In my 100 gallon main tank, it is obvious when the larvae are present as the presence of a swarm of shrimp larvae induces the Anthias to go into a plankton feeding frenzy. At that point, I start removing the larvae using a 10 ml plastic syringe with 5 inches of airline host attached to the end. Do not attempt to use a fine mesh net to collect the fragile larvae as the physical contact tends to break appendages. I've observed larvae with missing appendages circle and spin in the grow-out tank and usually die off very soon after. Additionally, I suspect the larvae don't feed as well as those with all appendages. The pumps are all turned off during this process to prevent larvae from going over the overflow, or being distributed around the tank, making it harder to siphon them out. In dark and still water, the larvae naturally group together on the bottom of the tank; which makes it easier to siphon them out. If there is light, the larvae tend to swim toward the light but don't group together well. Even grouped together, it is quite laborious to get them out, and I usually only manage to retrieve about half before tiring. The fish consume the rest.
Initial Grow-Out Tank
Water from the main tank (see Table 1) is transferred to a tiny 1/2 gallon tank containing a single airstone that generates very fine bubbles. Fresh artificial salt water (ASW) is not used because of very high mortality rates (personal observation, Toonen 1999, Strathmann 1987) possibly due to the use of Tris-EDTA or like chelator. One workaround maybe to first run the ASW through granular activated carbon then age under heavy aeration. If the bubbles are too large, the larvae can be damaged from the turbulence. Again, larvae with broken appendages appear to feed less effectively and die sooner than larvae with all appendages. Using an extra small tank for the initial grow-out of the larvae increases the density of newly hatched Artemia, and provides a higher probability that weakly swimming larvae will make contact with the food. Brine shrimp nauplii stocking concentration was roughly 25 per milliliter. A single 9-watt normal output fluorescent light is used to attract both the larvae and brine shrimp to the water surface and away from the airstone. My next version of the rearing tank will be a tank within a tank so I can isolate the larvae from dangerous filtration equipment. The water temperature is kept between 74 and 78 degrees, and no water changes are made for the two weeks of the initial rearing period. Only RO/DI water is added, very slowly, to keep the specific gravity roughly constant at 1.025. The chance of changing water and inadvertently transferring larvae out of the tank is great, and fishing larvae out of water with a plastic 10ml suringe is not an easy task. Larval mortality during the first two weeks is high; I estimate it to be about 50 percent. When the tank bottom begins accumulating detritus, the larvae are transferred to the final rearing tank via a ½ inch inner diameter hose to reduce the chance of larvae becoming trapped in the bottom debris (Castro, 1983).
Table 1: Main Tank Water Chemistry
Temp
79.5 - 82.5 deg F
pH
7.95 to 8.22
ORP
278 - 300
Specific Gravity
1.025
NH3
Not detectable
NO2
Not detectable
NO3
1.0 ppm
Alk
8.0kH/2.86 meq
Ca
450 ppm
Mg
1250 ppm
PO4
<0.05 ppm
BART-HAB
12 hour incubation
Larval Rearing Tank
A 10 gallon rearing tank is set up in parallel to the initial grow-out tank using water from the main tank, and any copepods are allowed to grow under 40 watts of normal output fluorescent light. Additionally, copepod growth is encouraged by additions of cryopreserved algae (see note 1). After two weeks, the surviving larvae from the initial grow-out tank are transferred via hose to the rearing tank. Freshly hatched Artemia nauplii are added every other day at a density of about one fifth of what it was in the mini-tank (about 5 per milliliter). However, since the larvae are bigger and are better swimmers with more appendages, finely smashed dried plankton flake is added to the tank as well (Kirkendoll 2001). It is believed that 2-4 week old larvae eat bbs, phytoplankton (Toonen, personal communication 2002; Jaime 2000; Ronquillo 1997), flake food (Kirkendoll 2001) and baby copepods (personal observation; Shishehchian 1999). I've observed 5 week old larvae catch a copepod and eat it, look quite fat afterwards and pigmentation increases afterwards. (See photos 1 & 2 of 4-week old larvae ). Toonen believes (personal communication, 2002) shrimp larvae consume some amount of phytoplankton as part of their diet since shrimp larvae have phyllopodous legs that are typically an adaptation for filter-feeding in marine invertebrate larvae. Toonen has also found Lysmata larvae guts loaded with phytoplankton (Tahitian Isochrysis ) fed hours before. Toonen (pers. comm., 2002) believes only live phytoplankton (e.g. DT's Phytoplankton) should be used since the chemical used to preserve algae paste contributes to sticky surface films and hence a greater chance of trapping larvae.
Figure 2: 4 week old [side view]
Figure 3: 4 week old [top view]
Copepods
I do not know what species of copepod grow abundantly on the glass surfaces of the 10 gallon tank, but I believe them to be harpacticoids (Shimek, personal communication, 2002). The young white copepods (1-2 mm) often dart out into the water column to move from place to place or so make small forays into the water to catch food and are occasionally caught by larvae. The copepods are clearly benthic, spending most of their time in the bottom debris or on the tank sides. The copepods in my tank under 40x magnification have long twin first antennae in the front and very long twin 'tails' in the end (see Figure 4 for an approximate sketch). The females carry sacks of eggs in the tails. These copepods appeared to reproduce rapidly under 40 watts of normal output fluorescent light, daily algae paste and brine shrimp nauplii additions.
Figure 4.
The adults (5 mm) do not exhibit darting behavior and spend most of their time near the bottom. My theory is that the shrimp larvae are able to catch the free-swimming copepod juveniles as they dart out, even though the copepods are much faster swimmers. Occasionally, I would see a shrimp larvae jerk in response to being hit by a young copepod 'missile,' and then retreat. The shrimp larvae intestines contained colored contents (probably flake food), as well as white/gray matter (probably bbs and, I believe, the small copepods). Copepods are believed to be a good larvae food source because copepods are high in EPA/DHA (Toonen, personal communication, 2002) and waxy esters and marine oils (Hoff 1999). Strathmann (1987) reports some epibenthic harpacticoid species are heavily preyed upon by juvenile salmon.
Larval Settlement
According to Riley (1994), Lysmata settlement can occur anywhere between 40 and 65 days, and it is not clear exactly what settlement cue triggers the larvae to become post larvae shrimp. The earliest settlement of 42 days by L. amboinensis was observed at the Waikiki Aquarium in a special 1000 gallon flow-through system using natural seawater, and the larvae were fed heavily on Tetraselmis and rotifers, switching to enriched Artemia at a later time (Toonen, personal communication with Waikiki Aquarium staff, 2002).
Conclusion
I was not able to get larvae past 5 weeks even though the larvae were large (1cm), colorful, and were freely using their pleiopods to swim towards food. I believe my high mortality rate at week 2 then again at week 5 were do to the following four issues that will be the focus of follow-up experiments:
* Entrapment of larvae on bottom debris and on sticky slime/film on the tank surface do to use of preserved algae paste. I've already redesigned the larvae growout tank so that larvae are contained in a inside tank with mesh bottom and side to encourage detritus to fall into the outside water column and processed by mechanical and biological filters (Strathmann 1987)
* Poor water quality. I did not actively measure nitrogenous waste, but there was tremendous algae growth on the bottom and sides of tank. I plan to reduce the amount of light in the growout tank from 40 to 20 watts of fluorescent light, replace 50 percent of the tank water per day from my main tank and routinely test for ammonium and nitrite ions.
* Poor food quality. The majority of brine shrimp napulii were not the characteristic orange color they should be for newly hatched San Francisco Bay Brand brine shrimp but were pale brown. I don't have the means to measure fatty acid content of the brine shrimp but they didn't look as good as I've seen them. To ensure the nauplii are as nutritious as possible, I've purchased a new lot of premium grade San Francisco Bay Brand from Brine Shrimp Direct and will feed a mixture of nauplii and brine shrimp gut loaded with algae paste and SELCO.
* Physical damage/stress due to collection and aeration of water. Even using a large 10 ml syringe and gentle aeration I personally observed several larvae with missing appendages. There may be other stress indicators but physical damage is the most obvious. I'll probably try using the overflow and capture method mentioned by Strathmann (1987) where Zoeae are isolated in a baffle system. The missing appendages weren't simply the result of a molt, but a single missing appendage that caused the larvae to spin uncontrollably.
Riley (1994) and Kirkendoll (2001) report maximum survival rates of 22 and 35 percent to post larval settlement, respectively.
…Till the next collection of free-floating Zoea!
If you have any questions about this article, please visit the Notes from the Trenches forum on Reef Central.
Notes
1. Algae products used:
Tahitian Blend algae paste - an cryopreserved algae product from Brine Shrimp Direct that is a combination of Nannochloropsis, Tahitian Isochrysis, Tetraselmis, and Pavlova plus NatuRose astaxanthin (Haematococcus sp.)
Acknowledgments
The author wishes to thank Dr. Ron Shimek, Eric Borneman and the Saltwater Enthusiasts Association of the Bay Area (SEABay) board members for reading and criticizing the paper. Special thanks to Rob Toonen, larval biologist, for advice and references on phytoplankton, marine invertebrate larvae diets and settlement.
References
Castro, Alceu de, Darryl E. Jory, 1983. Preliminary experiments on the culture of the banded coral shrimp Stenopus hispidus Oliver. Journal of Aquaculture & Aquatic Sciences Volume 3, Number 4.
Hoff, Frank 1999. Plankton Culture Manual Fifth Edition. Florida Aqua Farms. Florida. P106-111.
Jaime, Barbaro; Artiles, Miguel; Fraga, Iliana; Galindo, Jose. Substitution of Chaetoceros muelleri for spray-dried Chlorella vulgaris in feeding protozoae of Litopenaeus schmitti. In: Boletin del Centro de Investigaciones Biologicas Universidad del Zulia Agosto, 2000. 34 (2): 127-142.
Kirkendoll, April 2001. How To Raise & Train Your Peppermint Shrimp. Lysmata Publishing, Miami, Florida. 117 pages.
Riley, Cecilia 1994. Captive Spawning and Rearing of the Peppermint Shrimp (Lysmata wurdemanni). SeaScope Vol 11, Summer 1994. 3 pages.
Ronquillo, Jesse D.; Matias, Jonathan R.; Saisho, Toshio; Yamasaki, Shigehisa. Culture of Tetraselmis tetrathele and its utilization in the hatchery production of different penaeid shrimps in Asia. In: Hydrobiologia Dec. 22, 1997. 358 (0): 237-244.
Shishehchian, F.; Yusoff, F. M.; Omar, H.; Kamarudin, M. S.. Nitrogenous excretion of Penaeus monodon postlarvae fed with different diets. Marine Pollution Bulletin 1999. 39 (1-12): 224-227
Strathmann, M. F., 1987, Reproduction and Development of Marine Invertebrates of the Northern Pacific Coast, University of Washington Press, Seattle and London, 670 pages.
Toonen, Rob 1999. Culturing Shrimp. www.reefs.org 2 pages.
Toonen, Rob 1999. Larvae Settlement Cues. www.reefs.org 3 pages.
from: reefkeeping.com
A Hobbyist's Tale
April Kirkendoll writes in the conclusion of her entertaining book entitled "How to Raise & Train Your Peppermint Shrimp" to, 'tell the world what you discover and don't skimp on the details. Our hobby may depend on it." I sincerely believe these words, and have tried to detail as much of my experiences attempting to raise Peppermint shrimp, so that you might not only observe many of the aspects of shrimp growth, but also may learn and develop novel ways of increasing the post-larval settlement survival rate.
For the first three years of trying to spawn and raise Peppermint shrimp, I could never get the shrimp larvae to survive more than a week or so before they died en masse. Recently, however, I was able to rear them to 5 weeks before they died by feeding copepods, in addition to baby brine shrimp (bbs) and plankton flake food. Here's my hobbyist's tale.
Figure 1. Breeding Adult
The Breeding Adults
Approximately three years ago, I obtained two small Peppermint shrimp, Lysmata wurdemanni, from a local fish store. Since then I've been able to capture roughly half of the larvae (about 50 larvae captured) almost every month. No special attention is given to the adult shrimp. Generally, frozen Mysis and brine shrimp are fed to the tank every day, as well as Tahitian Blend algae paste (see note 1). The adult shrimp will generally eat anything that they can catch and tear apart with their pincers. The shrimp are full sized adults (3") ,and regularly produce free swimming larvae every month (about 100 larvae at each event).
Capturing Larvae
Spawning inevitably occurs late at night, usually around midnight. If the larvae are not removed immediately from a fully stocked reef tank, they stand very little chance of survival because of predation by fish. In my 100 gallon main tank, it is obvious when the larvae are present as the presence of a swarm of shrimp larvae induces the Anthias to go into a plankton feeding frenzy. At that point, I start removing the larvae using a 10 ml plastic syringe with 5 inches of airline host attached to the end. Do not attempt to use a fine mesh net to collect the fragile larvae as the physical contact tends to break appendages. I've observed larvae with missing appendages circle and spin in the grow-out tank and usually die off very soon after. Additionally, I suspect the larvae don't feed as well as those with all appendages. The pumps are all turned off during this process to prevent larvae from going over the overflow, or being distributed around the tank, making it harder to siphon them out. In dark and still water, the larvae naturally group together on the bottom of the tank; which makes it easier to siphon them out. If there is light, the larvae tend to swim toward the light but don't group together well. Even grouped together, it is quite laborious to get them out, and I usually only manage to retrieve about half before tiring. The fish consume the rest.
Initial Grow-Out Tank
Water from the main tank (see Table 1) is transferred to a tiny 1/2 gallon tank containing a single airstone that generates very fine bubbles. Fresh artificial salt water (ASW) is not used because of very high mortality rates (personal observation, Toonen 1999, Strathmann 1987) possibly due to the use of Tris-EDTA or like chelator. One workaround maybe to first run the ASW through granular activated carbon then age under heavy aeration. If the bubbles are too large, the larvae can be damaged from the turbulence. Again, larvae with broken appendages appear to feed less effectively and die sooner than larvae with all appendages. Using an extra small tank for the initial grow-out of the larvae increases the density of newly hatched Artemia, and provides a higher probability that weakly swimming larvae will make contact with the food. Brine shrimp nauplii stocking concentration was roughly 25 per milliliter. A single 9-watt normal output fluorescent light is used to attract both the larvae and brine shrimp to the water surface and away from the airstone. My next version of the rearing tank will be a tank within a tank so I can isolate the larvae from dangerous filtration equipment. The water temperature is kept between 74 and 78 degrees, and no water changes are made for the two weeks of the initial rearing period. Only RO/DI water is added, very slowly, to keep the specific gravity roughly constant at 1.025. The chance of changing water and inadvertently transferring larvae out of the tank is great, and fishing larvae out of water with a plastic 10ml suringe is not an easy task. Larval mortality during the first two weeks is high; I estimate it to be about 50 percent. When the tank bottom begins accumulating detritus, the larvae are transferred to the final rearing tank via a ½ inch inner diameter hose to reduce the chance of larvae becoming trapped in the bottom debris (Castro, 1983).
Table 1: Main Tank Water Chemistry
Temp
79.5 - 82.5 deg F
pH
7.95 to 8.22
ORP
278 - 300
Specific Gravity
1.025
NH3
Not detectable
NO2
Not detectable
NO3
1.0 ppm
Alk
8.0kH/2.86 meq
Ca
450 ppm
Mg
1250 ppm
PO4
<0.05 ppm
BART-HAB
12 hour incubation
Larval Rearing Tank
A 10 gallon rearing tank is set up in parallel to the initial grow-out tank using water from the main tank, and any copepods are allowed to grow under 40 watts of normal output fluorescent light. Additionally, copepod growth is encouraged by additions of cryopreserved algae (see note 1). After two weeks, the surviving larvae from the initial grow-out tank are transferred via hose to the rearing tank. Freshly hatched Artemia nauplii are added every other day at a density of about one fifth of what it was in the mini-tank (about 5 per milliliter). However, since the larvae are bigger and are better swimmers with more appendages, finely smashed dried plankton flake is added to the tank as well (Kirkendoll 2001). It is believed that 2-4 week old larvae eat bbs, phytoplankton (Toonen, personal communication 2002; Jaime 2000; Ronquillo 1997), flake food (Kirkendoll 2001) and baby copepods (personal observation; Shishehchian 1999). I've observed 5 week old larvae catch a copepod and eat it, look quite fat afterwards and pigmentation increases afterwards. (See photos 1 & 2 of 4-week old larvae ). Toonen believes (personal communication, 2002) shrimp larvae consume some amount of phytoplankton as part of their diet since shrimp larvae have phyllopodous legs that are typically an adaptation for filter-feeding in marine invertebrate larvae. Toonen has also found Lysmata larvae guts loaded with phytoplankton (Tahitian Isochrysis ) fed hours before. Toonen (pers. comm., 2002) believes only live phytoplankton (e.g. DT's Phytoplankton) should be used since the chemical used to preserve algae paste contributes to sticky surface films and hence a greater chance of trapping larvae.
Figure 2: 4 week old [side view]
Figure 3: 4 week old [top view]
Copepods
I do not know what species of copepod grow abundantly on the glass surfaces of the 10 gallon tank, but I believe them to be harpacticoids (Shimek, personal communication, 2002). The young white copepods (1-2 mm) often dart out into the water column to move from place to place or so make small forays into the water to catch food and are occasionally caught by larvae. The copepods are clearly benthic, spending most of their time in the bottom debris or on the tank sides. The copepods in my tank under 40x magnification have long twin first antennae in the front and very long twin 'tails' in the end (see Figure 4 for an approximate sketch). The females carry sacks of eggs in the tails. These copepods appeared to reproduce rapidly under 40 watts of normal output fluorescent light, daily algae paste and brine shrimp nauplii additions.
Figure 4.
The adults (5 mm) do not exhibit darting behavior and spend most of their time near the bottom. My theory is that the shrimp larvae are able to catch the free-swimming copepod juveniles as they dart out, even though the copepods are much faster swimmers. Occasionally, I would see a shrimp larvae jerk in response to being hit by a young copepod 'missile,' and then retreat. The shrimp larvae intestines contained colored contents (probably flake food), as well as white/gray matter (probably bbs and, I believe, the small copepods). Copepods are believed to be a good larvae food source because copepods are high in EPA/DHA (Toonen, personal communication, 2002) and waxy esters and marine oils (Hoff 1999). Strathmann (1987) reports some epibenthic harpacticoid species are heavily preyed upon by juvenile salmon.
Larval Settlement
According to Riley (1994), Lysmata settlement can occur anywhere between 40 and 65 days, and it is not clear exactly what settlement cue triggers the larvae to become post larvae shrimp. The earliest settlement of 42 days by L. amboinensis was observed at the Waikiki Aquarium in a special 1000 gallon flow-through system using natural seawater, and the larvae were fed heavily on Tetraselmis and rotifers, switching to enriched Artemia at a later time (Toonen, personal communication with Waikiki Aquarium staff, 2002).
Conclusion
I was not able to get larvae past 5 weeks even though the larvae were large (1cm), colorful, and were freely using their pleiopods to swim towards food. I believe my high mortality rate at week 2 then again at week 5 were do to the following four issues that will be the focus of follow-up experiments:
* Entrapment of larvae on bottom debris and on sticky slime/film on the tank surface do to use of preserved algae paste. I've already redesigned the larvae growout tank so that larvae are contained in a inside tank with mesh bottom and side to encourage detritus to fall into the outside water column and processed by mechanical and biological filters (Strathmann 1987)
* Poor water quality. I did not actively measure nitrogenous waste, but there was tremendous algae growth on the bottom and sides of tank. I plan to reduce the amount of light in the growout tank from 40 to 20 watts of fluorescent light, replace 50 percent of the tank water per day from my main tank and routinely test for ammonium and nitrite ions.
* Poor food quality. The majority of brine shrimp napulii were not the characteristic orange color they should be for newly hatched San Francisco Bay Brand brine shrimp but were pale brown. I don't have the means to measure fatty acid content of the brine shrimp but they didn't look as good as I've seen them. To ensure the nauplii are as nutritious as possible, I've purchased a new lot of premium grade San Francisco Bay Brand from Brine Shrimp Direct and will feed a mixture of nauplii and brine shrimp gut loaded with algae paste and SELCO.
* Physical damage/stress due to collection and aeration of water. Even using a large 10 ml syringe and gentle aeration I personally observed several larvae with missing appendages. There may be other stress indicators but physical damage is the most obvious. I'll probably try using the overflow and capture method mentioned by Strathmann (1987) where Zoeae are isolated in a baffle system. The missing appendages weren't simply the result of a molt, but a single missing appendage that caused the larvae to spin uncontrollably.
Riley (1994) and Kirkendoll (2001) report maximum survival rates of 22 and 35 percent to post larval settlement, respectively.
…Till the next collection of free-floating Zoea!
If you have any questions about this article, please visit the Notes from the Trenches forum on Reef Central.
Notes
1. Algae products used:
Tahitian Blend algae paste - an cryopreserved algae product from Brine Shrimp Direct that is a combination of Nannochloropsis, Tahitian Isochrysis, Tetraselmis, and Pavlova plus NatuRose astaxanthin (Haematococcus sp.)
Acknowledgments
The author wishes to thank Dr. Ron Shimek, Eric Borneman and the Saltwater Enthusiasts Association of the Bay Area (SEABay) board members for reading and criticizing the paper. Special thanks to Rob Toonen, larval biologist, for advice and references on phytoplankton, marine invertebrate larvae diets and settlement.
References
Castro, Alceu de, Darryl E. Jory, 1983. Preliminary experiments on the culture of the banded coral shrimp Stenopus hispidus Oliver. Journal of Aquaculture & Aquatic Sciences Volume 3, Number 4.
Hoff, Frank 1999. Plankton Culture Manual Fifth Edition. Florida Aqua Farms. Florida. P106-111.
Jaime, Barbaro; Artiles, Miguel; Fraga, Iliana; Galindo, Jose. Substitution of Chaetoceros muelleri for spray-dried Chlorella vulgaris in feeding protozoae of Litopenaeus schmitti. In: Boletin del Centro de Investigaciones Biologicas Universidad del Zulia Agosto, 2000. 34 (2): 127-142.
Kirkendoll, April 2001. How To Raise & Train Your Peppermint Shrimp. Lysmata Publishing, Miami, Florida. 117 pages.
Riley, Cecilia 1994. Captive Spawning and Rearing of the Peppermint Shrimp (Lysmata wurdemanni). SeaScope Vol 11, Summer 1994. 3 pages.
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Let's Clown Around With More Gobies: The Gobiodon Species
Let's Clown Around With More Gobies: The Gobiodon Species
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Last month I introduced you to a small genus in the Gobiidae family; a huge family that is found most anywhere bodies of saltwater are present. Due to the overall size and diversity of Gobiidae, I decided to continue exploring this family by discussing another small genus from the vast assortment of reef tank-suitable genera. The gobies of the genus Gobiodon, often called "clown gobies" or "gum drop gobies," are my fish of choice for October. Meet the Family Gobiidae is the largest family of marine fish with over 2,000 members and still growing. Gobiodon is a small genus within Gobiidae, comprised of only 15 recognized species (Harold & Winterbottom, 1995) (see below); though more than 30 nominal species have been described (Munday et al., 1999) including the recently classified Gobiodon brochus (Harold & Winterbottom, 1999).
The reader may note the lack of G. brochus in the list above. Gobiodon brochus was not recognized during the most recent revision of Gobiodon (Harold & Winterbottom, 1995). A new revision is in preparation by the same authors, however, and it will include G. brochus as a valid species (Munday et al., 1999).
Individuals of Gobiodon species are small fish, rarely growing larger than 60mm. Though some species are predominantly scale-less, the majority of them are totally scale-less. Instead of scales, their smooth-sided skin has a thick, toxic mucus covering (Hasimoto et al., 1974). The jaws contain predominately small teeth, though two pairs of well-developed canine teeth are present. At least one species, Gobiodon brochus, has a modified jaw. The exact function of this modified, protruding, deflected lower lip is not known, although it has been suggested that it utilizes its extra "toothy pad" to rasp tissue from coral polyps (Winterbottom, pers. comm.). Gobiodon species are also equipped with modified ventral fins which have joined as one and developed small suction cups on the end. These aid the gobies in grasping onto corals in high current areas.
In the Wild Gobiodon species are found throughout the Indo-Pacific, and some species extend to the Red Sea and to the western Pacific Ocean. They are abundant in these waters, and range from depths of 5 feet to as deep as 60 feet. Their depth distribution is directly dependant upon available corals within which they can find shelter. Generally speaking, though, the corals these fish inhabit are found in shallow waters. Like many other gobies, species of Gobiodon partake in a special symbiotic relationship of their own. Fish of this genus are obligate coral dwellers, usually exhibiting a relationship known as "commensalism" with corals of the genus Acropora (Tyler, 1971), though some species relate with a few other corals, namely Echinopora spp., Hydnophora spp., and Stylophora spp. High upon the reefs, within the branches of these corals, the clown gobies will take refuge and wait for a passing morsel. Examination of the gut contents by Harold and Winterbottom (1999), found that these "passing morsels" were copepods, foraminifera, and unidentifiable flocculent material. Another interesting and uncommon trait has been uncovered and described. Nakashima et al., (1996), have described a two-way sex change, known as bi-directional sex change, within two Gobiodon species, G. micropus and G. oculolineatus. This led to the discovery that other Gobiodon also have this ability, and it is now believed that all Gobiodon species can change sex. In doing so, Gobiodon deviates from the size-advantage model (Ghiselin 1969) that states if an individual could significantly increase its chance of reproduction success after a certain size was reached, it would change to that sex. Instead, bi-directional protogynous hermaphrodites are the ultimate in sex-changing species, as it guarantees a heterosexual pair at any given time. In most cases, the smaller fish of the heterosexual pair is the female. The only time this is not true is when a small male is placed in the same coral head as a large female. It was found that in pairs which started as two females, the larger of the two became male. In pairs that started as two males, the smaller of the two changed to female (Munday et al. 1998). In the Home Aquarium Gobiodon sp. have a tough time adjusting to captive care, since it is all too often that these fish arrive at our local fish stores emaciated. This is most likely due to the stressful transit period, which results in the fish not eating. Being a smaller fish, and having what appears to be a fairly quick metabolism, not many of these fish make it to the hobbyist tank without having been starved to some degree or another. To compound this problem, they regularly require live foods and special attention until accustomed to aquarium life. Frozen/thawed foods can be offered first, but if they are not accepted, be prepared to offer live food. Upon arrival of a new goby, special care should be taken to ensure large quantities of food are offered to the new arrivals without fouling the aquarium water. Live brine shrimp is the most available type of live food. Try to "gut-load" these live Artemia with phytoplankton if the possibility exists. Once the goby has accepted live food, they may slowly be weaned off the live food until they eventually are accepting frozen/thawed or prepared foods. Any of the commercially available foods suitable for a carnivore should be sufficient. Be sure to provide a varied diet, and that the food is small enough to fit into their tiny mouths.
Tank mates for Gobiodon must be carefully chosen. Though these fish do have a toxic mucus on their skin for protection, it doesn't mean you can mix and match haphazardly. Fish that are aggressive feeders should be avoided as tankmates, at least until after the Gobiodon has been fattened up and is readily accustomed to the aquarium. A refugium can be helpful during the transition period, hopefully offering an abundance of their favorite natural foods, copepods. An established tank stocked heavily with Acropora spp. will assist in mixing these fish with more aggressive swimmers. The Gobiodon species will take refuge within the branches of these corals. The more threatening the tank mates, the deeper into the branches the Gobiodon will retreat. In a peaceful aquarium Gobiodon spp. will normally remain on the tips of the corals within full view of the hobbyist. Compatibility chart for members of the genus Gobiodon:
Note: While many of the fish listed are good tank mates for members of the genus Gobiodon, you should research each fish individually before adding it to your aquarium. Some of the fish mentioned are better left in the ocean, or for advanced aquarists. Although fish of the genus Gobiodon are obligate coral dwellers, usually utilizing the genus Acropora, it is not absolutely necessary that Acropora be present in their tank. In the home aquarium Gobiodon sp. will often co-exist with other stony corals, or even soft corals, lacking more preferable options. In all instances, if their preferred coral is available, it will be utilized. In aquariums without sufficient coral growth, it is likely that the Gobiodon sp. will either hide within rockwork, or seek out any other hiding places it can find. Given this type of environment, it will not be comfortable, and most likely will not adapt well to captivity. Due to their obligate coral dwelling nature, they can easily be kept in small or nano reef aquariums. Captive Reproduction Although captive spawning is a regular occurrence in Gobiodon species, raising the small fry seems to be less than easy. Obtaining a pair is simply a matter of obtaining two of the same species, since they will change sex to become a mated pair. Once mated, the female will attach circular bands of eggs around the branches of their preferred coral. The male immediately fertilizes and guards the eggs. The egg mass, that can contain up to 1000 eggs, hatches on the evening of the fourth or fifth day. Rotifers should be the first food offered to fry, with a possible transition to newly hatched Artemia nauplii around day 25. Around day 33, the fry go through a metamorphosis, settle, and begin to perch on the sides of the aquarium's glass. Their first coloration has been noted to occur on day 40 (Breeder's Registry).
Meet the Species Two species of Gobiodon are regularly regarded as being the most popular. The first is Gobiodon okinawae, or the yellow clown goby. Luckily for aquarists, this bright yellow fish is one of the most outgoing of Gobiodon. It will frequently perch on the glass of the aquarium, and regularly sits on the tips of the corals, rather than deep inside the colony. Gobiodon okinawae can be found on the largest variety of acroporid corals (Myers, 1991).
The other favorite Gobiodon among hobbyists is the Green Clown Goby, or Gobiodon histrio. One look at the clown-like paintwork on the face and body is usually enough for the aquarist to fall in love. Two color varieties, possibly separate species, exist for G. histrio. The first variety of G. histrio has a black spot on the upper margin of the operculum. The pectoral fins are generally pale green to light brown. The second coloration pattern has not yet been definitively published, largely due to inadequate analysis, but is commonly referred to as G. histrio "erythropilus." The coloration of this variation is nearly identical to G. histrio, except the black spot on the operculum is missing and the pectoral fins are generally yellowish with a fine black margin (Suzuki et al., 1995). In the wild both species colonize Acropora nasuta most frequently, but can also be found on A. valida, A. millepora, and sometimes A. tenuis. It is rare to find more than a single pair of G. histrio per coral colony (Patton, 1994)
Although a large percentage of the clown gobies available in the trade will be one of the two previously mentioned species, a couple more show up less frequently. Gobiodon citrinus, or the Citron goby, is the largest of clown gobies, measuring up to 60 mm. Four light blue bands are located on the head as well as single blue lines along the base of the dorsal and anal fins. A single black spot is located on the operculum. Overall, the fish have a yellow to brownish yellow coloration. They are most likely to be found residing within the branches of A. nobilis (Kailola, 1991, Munro, 1967, Randall et al., 1990).
The Black clown goby, or Gobiodon ceramensis, is an intriguing Gobiodon due to its intense black coloration. The entire fish is midnight black. Unlike its congeners, G. ceramensis is most likely found on corals from the family Pocilloporidae, most notably Stylophora pistillata (Tyler, 1971). Don't confuse G. ceramensis with its nearly identical cousin, Gobiodon acicularis. Only one physical difference exists between the two; G. acicularis has an extra long first dorsal spine. However, G. acicularis can also be found on different corals than G. ceramensis. G. acicularis can only be found on Echinopora horrida, E. mammiformis and Hydnophora rigida (Munday et al. 1999).
Gobiodon rivulatus has occasionally shown up in the trade in recent times. This fish lacks a common name, though it is usually sold under the common name of "Citron Goby." Gobiodon rivulatus is similar in appearance to G. citrinus, but is missing the blue lines on the base of the dorsal and anal fins. The blue lines it does have are narrow and wavy, and extend down the body of the fish rather than just appearing on the head as with G. citrinus. The overall color of these fish are highly variable, from a dark brown to light brown (Winterbottom & Emery, 1996). This species is most frequently found associated with Acropora gemmifera and A. secale, but will inhabit a large variety of acroporid species (Munday et al., 1997).
Up until 1999, Gobiodon brochus was known as Gobiodon micropus. It was then that Harold and Winterbottom (1999) discovered the protruding lip containing tiny teeth, which set it apart from G. micropus. Also, G. brochus has 10 - 12 dorsal fin rays and 9 - 10 branched anal fin rays, whereas G. micropus has 12 - 13 dorsal fin rays and 11 anal fin rays.
In Conclusion Gobiodon gobies make adorable additions to the reef aquarium. If the hobbyist has an aquarium with suitable coral growth, and is willing to take on the challenge of fattening one up upon purchase, these gobies can fare extremely well in captivity. Never before would you have believed you could find so much personality in a fish that barely moves! |
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References:
Burgess, W.E., et al. 1991. Dr. Burgess's Mini-Atlas of Marine Aquarium Fishes Mini Edition. T.F.H. Publications. Neptune City. pp. 579 - 591.
Ghiselin, M.T. 1969. The evolution of hermaphroditism among animals. Q. Rev. Biol. 44: 189-208.
Harold, A. S., Winterbottom, R. 1995. Gobiodon acicularis, a new species of gobiid fish from Belau, Micronesia. Proc. Biol. Soc. Wash., 108: 687-694
Harold, A. S., Winterbottom, R. 1999. Gobiodon brochus, a new species of gobiid from the western South Pacific, with a description of jaw morphology. Copeia, 1999, 1 : 49-57.
Hashimoto, Y., Shiomi, K., Aida, K. 1974. Occurrence of a skin toxin in coral gobies Gobiodon spp. Toxicon, 12: 523-528.
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Michael, S. W. 1999. Gobies. Marine Fishes: 500 + Essential-To-Know Aquarium Species. Microcosm. Shelburne. p. 350.
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Munday, P.L., Caley, M.J., Jones, G.P. 1998. Bi-directional sex change in a coral-dwelling goby. Behav. Ecol. Sociobiol., 43: 371-377.
Munro, I.S.R. 1967. The fishes of New Guinea. Department of Agriculture Stock and Fisheries, Port Moresby.
Myers, R.F. 1991. Micronesian reef fishes. Coral Graphics, Guam.
Patton, W.K. 1994. Distribution and ecology of animals associated with the branching corals (Acropora spp.) from the Great Barrier Reef, Australia. Bull. Mar. Sci., 55: 193-211.
Randall, J.E., Allen, G.R., Steene, R.C. 1990. Fishes of the Great Barrier Reef and Coral Sea. Crawford House Press. Bathurst.
Suzuki, T.M., Aizawa, M., Senou, H. 1995. A preliminary review of three species of the Gobiodon rivulatus complex from Japan. I.O.P. Diving News, 6: 2-7.
Tyler, J.C. 1971. Habitat preferences of the fishes that dwell in shrub corals on the Great Barrier Reef. Proc. Acad. Nat. Sci. Phila., 123: 1-26.
Winterbottom, R., Emery, A.R. 1986. Review of the gobiod fishes of the Chagos Archipelago, central Indian Ocean. Life Sciences Contribution 142. Royal Ontario Museum, Ontario.
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