Biologists from AZaquaculture will be attending the annual meeting of the Desert Fishes Council in Cuatrociénegas, Coahuila, México.  Only a skeleton staff will remain in Tucson.  As such, we will be closed during the period of November 10-20.  Phones will not be answered, but you can leave a message and we will do our best to get back to you in a timely fashion.  Emails should be answered, but this will be limited by the availability of wi-fi or other internet accessability in Cuatrociénegas, and we won’t know that for sure till we arrive.

Thanks for your patience and understanding, and we look forward to serving your aquaculture needs upon our return!

It seems only fitting that our first creature feature focuses on one of the first marine ornamentals to have been raised successfully in captivity, the Neon Goby, Elacatinus oceanops, (formerly Gobiosoma oceanops). First reared in the 1970’s the neon goby is a popular aquarium fish owing to its general hardiness, attractive appearance, and abilities in picking ectoparasites off of other fishes.

Size and Appearance

Neon gobies are small and typical gobiioid in shape. Most Elacatinus spp. are less than 5cm (2 inches) total length. Neon gobies are black overall with a neon blue stripe extending from front of eye to the base of caudal fin.

Broodstock Care

Omnivorous and hardy they will do well in nearly any species-only or reef aquarium situation, but due to their size should not be kept with larger predatory fishes. Neon gobies do best in water temperatures below 26.5 decrees C (80 degrees F). Foods should include a variety of grated frozen shrimp, squid and fish, as well as commercial gelatin or pellet diets. Multiple feedings daily will condition neon gobies for spawning.

Pair Formation

Hermaphroditic sexual patterns are common in the family Gobiidae. I am not aware of a definitive classification of Elacatinus oceanops, but experience in our lab suggests that they are sequential hermaphrodites rather than simultaneous hermaphrodites.
Males are often larger and more slender. Females will possess a swollen abdomen, particularly when ripe with eggs.

Neon Gobies may be kept as groups of 6 or more individuals when provided with sufficient hiding spots, as these gobies can be quite quarrelsome. If they are to be kept as a pair, they should be observed closely during the first week after introduction. If fighting is excessive, one member of the pair should be swapped until marital harmony ensues. Groups of fewer than 6 individuals are not suggested, as pairs will begin to try to “evict” other gobies in their territory. Larger groups dampen and disperse these aggressive tendencies.

Spawning and Hatching

Spawning can occur as often as every fourteen days with plenty of feeding and warm water conditions. In their natural environment, demersal eggs are laid in small holes and crevices in the reef and under discarded bivalve shells. In captivity, small Tridacna sp shells serve well, as do halved clay flowerpots and short sections of half inch PVC pipe. Both parents tend eggs. Depending on temperature, hatching will commence in 6-8 days. Hatching occurs after dark.

Larval Rearing

Neon goby larvae are slightly shorter and substantially slimmer than clownfish larvae. The larval period ranges from 18-25 days depending on temperature and food type. The first diet is rotifers, followed by Artemia nauplii. The transition period is variable between these foods. Elacatinus larvae can be transitioned to Artemia as early as day 6, and while growth is more rapid, mortality is often high. Waiting until day 12-15 to begin Artemia feedings will delay metamorphosis by a few days, but will also significantly increase survivorship. We have successfully reared batches of neon goby hatchlings through metamorphosis only on rotifers, but metamorphosis took 30-35 days.

A critical factor in the success of a plankton culture system is the proper sterilization of both culture vessels and any solutions that go into them. Such sterilization prevents the overgrowth of the target species with microbial contaminates. Such contaminates may include undesirable bacterial, fungal, and protozoan species. Microbial contaminates may exert deleterious effects on the target species via predation, release of toxins, or through secretion of harmful metabolic byproducts and competition for nutrients and space. Contamination of rotifer and microalgae cultures by certain species of dinoflagellates has been shown to be a major factor in the mortality of clownfish larvae in hatchery settings. At the very least, maintaining a clean and sterile culture system will go a long way in producing reliable production levels of planktonic or larval organisms and will help speed the troubleshooting process when things go wrong.

Below are descriptions of various methods used in aquaculture hatcheries, as well as an explanation of their typical applications, limitations, and in some cases, links to protocols for implementation.

STERILIZATION BY MEMBRANE FILTRATION:

Primarily utilized for small culture volumes, membrane filtration provides a high level of sterility while being extremely gentle to water chemistry. Sterilization is accomplished by forcing liquid through a filter that has a defined pore size, typically either 0.45 or 0.22 microns. This allows for the elimination of bacteria and fungi, which are too large to fit through the pores, without modification of the chemical constituents of the culture media. Effective for bacterial, fungal, and protozoan species, these filters are not generally effective against viruses as these are small enough to pass easily through the pores.

Membrane filters typically come in two flavors:
Syringe filters are handy for sterilizing very small volumes of liquid, less than 100ml. These filters come in convenient disposable cassettes that attach to the tip of syringes. Liquid is forced from the syringe through the filter and the exiting solution is sterilized.
Vacuum filtration setups are effective for larger volumes, up to a liter. These allow the culture media to be pulled through a larger filter membrane via a vacuum pump. The sterilized culture media is collected in a receiver vessel, often a side-arm Erlenmeyer flask. It is important to note that for either of these methods, all downstream vessels and apparatus that contact the sterilized media must be sterile themselves. Thus, it is common to use these methods only for making small stocks of solutions such as f/2 that are sensitive to other forms of sterilization, and may be stored in pre-sterilized, disposable containers.

HEAT STERILIZATION:

Heat sterilization, when properly performed, can be among the best methods of sterilization. However, many desirable constituents of culture media may be temperature sensitive and can be destroyed by heat. Most notable are vitamins, fertilizers, and antibiotic solutions, which are typically filter sterilized and added to heat sterilized media after it cools. In addition to being damaging to additives, high temperatures can cause undesirable precipitation of a variety of constituents of seawater, especially as temperatures approach boiling. Such precipitation may or may not have adverse effects on the culture, depending on the species and conditions that are utilized.

Autoclaving is the most common and effective method for sterilizing moderate amounts of material, especially in a laboratory setting. Requiring specialized equipment, material is heated under pressure in the presence of steam. Given an adequately long exposure time, this is an effective method of destroying bacteria, fungi, spores, and viruses. The most common exposure conditions are 121 degrees C at 15psi. Similar levels of sterility can be attained in a pressure cooker without the expense of an autoclave.
Boiling is a moderately effective method of sterilization. It does a good job of killing most bacteria, viruses, and fungi. However, it often is not successful at destroying the environmentally resistant spores produced by some species. To ensure complete killing of spores, it may be necessary to boil the medium on 2 or 3 consecutive days, allowing the medium to cool between treatments.
Pasteurization is not quite as assured a method for complete destruction of microorganisms as autoclaving, but properly executed can reach nearly the same kill rates without the problems of precipitation that may occur with boiling or autoclaving. Pasteurization can be accomplished by heating the solution to 80 degrees Celsius, allowing the solution to cool naturally, then heating again, generally 24 hours later. This may be repeated a third time for extra safety.

CHEMICAL STERILIZATION

The most economical and convenient method for sterilizing moderate to large volumes of water is chemical sterilization. This is most often accomplished through the addition of strong oxidizing agents such as chlorine, or by dropping the pH below 4 through the use of hydrochloric or muriatic acid. Highly effective, these methods are affected by factors such as temperature, contact time, dissolved organics etc., therefore, sterilization parameters will need to be adjusted in response to these conditions.
It is important to return the pH to normal before using the media if acid-sterilization was employed, and chlorine solutions must be neutralized, generally through additions of sodium thiosulfate. Chemical sterilization may destroy additives such as vitamins, antibiotics, and fertilizers, so sterile stocks of these should be added only after neutralization of the chlorine or acid.