— 
The Fishroom Library Archives —
Articles on Breeding Killifish, Incubating and
Hatching Eggs, and Raising Fry (1996-2000)
An archived collection of articles, tips, and information about tropical fishkeeping in general, and killi-keeping in particular, from past issues of the G.C.K.A. Newsletter. To read an article, please click on the title (in blue).
All material is copyright © G.C.K.A. or the authors, unless otherwise noted. Reproduction is permitted for non-commercial purposes only (i.e., club bulletins). We do request that you provide source credit, and send us a copy of the publication in which the article appears. Please forward to G.C.K.A., c/o Recktenwalt, 4337 Ridgepath Drive, Dayton, OH 45424.
Breeding the Diapteron Subgenus. By Mark DelRaso.
The Case for “Benign Neglect” – another point of view on breeding killies.
To Cull or Not to Cull … It’s an Important Question
Diapause in Annual Killifish Eggs … some observations
Breeding Tanks: The European Method – Worth a Try? – a more “natural” species tank setup.
A few misconceptions about handling annual eggs – one aquarist’s experiences.
A Few Techniques for Hatching Eggs
A Few Tricks to Induce Spawning – some things to try when the fish just won’t spawn.
Finding and Storing the Eggs – an overview on how to handle the eggs of annual species.
Fry the Easy Way – a “no fuss” method for raising plant-spawners.
A Guide to Raising Fry – some pointers to raising those precious hatchlings.
Handling Eggs – Now What? – when water incubated eggs are problematic.
How Big Are They, Anyway? – egg sizes of some representative species.
How I Do It – one breeder’s method.
Inbreeding in Killifish – An Overview
Incubating and Hatching Eggs – an overview.
Incubating and Hatching Eggs – How I Do It (Al Anderson)
Incubating Eggs … A Judgement Call – a few tips on development and hatching.
Incubating Eggs on Peat
I’ve got eggs – now what? What to do with that spawn once you have some.
Keep More Than One Pair! – as protection against “Murphy’s Law.”
Killie Soup – Yum! – a new “recipe” for handling those semi-annual eggs.
Lunar Effects on Egg Production?
More on Greensand – more on this alternate spawning medium for annuals.
Selecting the Spawning Media – spawning media for annual fishes.
Sex Ratios in Aphyosemion zygaima – one aquarist’s experience.
Skewed Sex Ratios – what happened? and possible solutions.
Skewed Sex Ratios – Again – more on this topic.
Skewed Sex Ratios – Not Just an Interesting Question – more on the various influences that influence the result
Small Protection – a few tips for saving fry.
Some Views on … Skewed Sex Ratios in Killifish – more on this topic.
Storing Eggs – various methods.
Vacation Care – A Trick for Fry
Breeding the Diapteron Subgenus
By Mark DelRaso, GCKA/AKA. This article first appeared in the G.C.K.A. Newsletter in March 1990.
Breeding killies of the subgenus Diapteron (Huber & Seegers, 1977) is not too difficult once certain conditions are met. The first problem that arises is finding good stock that is mature. Sexual maturity in the Diapteron subgenus is reached at nine months to one year of age. Even if eggs are picked or transferred without touching them, [eggs laid] at an early age will result in fungusing. Diapteron georgiae (Lambert & Gery, 1967) is the easy one to spawn at seven to nine months of age. However, viability of the eggs and the resulting fry is noticeably poor. With this particular species, viable fry and eggs are produced at an age reaching over one year. An additional point to be made is that the fish in this subgenus are rather long-lived. The oldest specimen kept by me currently is a healthy five and a half year old female D. abacinum (Huber 1976). D. abacinum is also the most challenging to breed beyond the first generation from the wild. According to some experts, D. abacinum may not be a valid species due to crossing. I am not sure about this because of the same problem with the Aphyosemions in the bualanum/dargei group. To breed these fishes, the aquarists must follow certain guidelines to care for their Diapterons. Once the conditions are met, you will be able to propagate these killies for several generations.
The first item on the checklist is suitable temperature. These fishes are from cool highland forest brooks in Gabon. In this biotope, heavy shade provided by the forest canopy prevents direct solar radiation from heating the water above about 70° F. A constant fishroom temperature of 68° F is the magic number. If you cannot give these fish a water temperature of less than 70° F, then you should not attempt these fish. At temperatures above 72° F, Diapteron do not keep their body weight.
The second requirement for these fish is very dim lighting. This is due to the effects of the very heavy forest canopy. The aquaria that you keep these fishes in may be sidelit from another tank to the point of recognition of the bottom of the tank. Some Diapteron specimens will do well in a tank bright enough to barely grow Microsorium pteropus and Anubias nana. The water should be quite soft and acid. You can use rainwater filtered through a high quality carbon to remove pollutants. To keep them comfortable, a pH of 5.5 to 6.2 is required. The hardness should be less than 50 ppm. More specifically, the carbonate hardness should be less than 2° DH and the general hardness less. The water must be very clean – no metabolites.
The third requirement is the proper foods. While white worms and grindal worms can be fed occasionally, crustaceans are the primary diet in the wild. Rinsed artemia nauplii are excellent foods for both the fry and adults. Daphnia is the best food, however. The Diapterons do not accept prepared or frozen foods. This is another requirement: LIVE FOOD ONLY. I feed all my killies live foods most of the time. So I always feed the Diapterons first if I am running low on live food. It is neat to watch a male D. fulgens stalk a baby brine shrimp.
After you have met the above conditions, you can pick the eggs very carefully or scoop the fry. I suggest you scoop the fry or transfer the mops, floating and sunken, to a container with water from the adults’ tank. The fry can be very nasty and cannibalistic. Remove the largest fry if it becomes a problem. Usually, if you keep a lot of mop material in the rearing tank, you will end up with a colony that will get along together.
There is a sex ratio problem with fry from young pairs. The first batch is very long in males. After the pair has been producing for one and a half to two years, the problem is much less pronounced. My last batch of D. fulgens was actually long on females. This works out well for young males of the next group of fry paired up with older females.
Remember: breeding these fish is a project taking at least one and one half years. Feed live food. Keep the tanks dim. Maintain soft, acid, and cool water. Use floating and sunken mops. Try not to touch the eggs. Enjoy the following Diapterons in the order of increasing difficulty:
A. (Diapteron) georgiae (Lambert & Gery 1967)
A. cyanostictum (Lambert & Gery 1967)
A. fulgens (Radda 1975)
A. abacinum (Huber 1976)Note: A. (Diapteron) seegersi (Huber 1980) has not been in the hobby for quite some time. There is confusion as to its validity as a species with some people. Also, there are some new locations of undescribed Diapterons making their way into the country currently: A. (Diapteron) sp. GBN 88/29 and one other species I am currently breeding.
– G.C.K.A. Newsletter, June 1999 Return to top of page
Raisin’ Killies
The Case for “Benign Neglect”
Most of us who try to raise killifish have our own preferred methods for producing fry. But some killikeepers have found that there are simpler methods that work very well. The following are a few examples.
“An approach of ‘benign neglect’ has worked wonderfully well” with Aphyosemion australe, reports Frank Louden. He breeds them in a 10 gallon tank with an undergravel filter and lots of watersprite, and collects fry on a regular basis.
“I get mostly males using this method,” he admits. “I have been told that young AUS will produce more males than females. As they grow older the ratio begins to even out and then as the breeders reach an older age, they produce many more females. This makes some sense in the ‘Darwinian’ mode of thinking, as the males come along first, their natural aggressiveness reduces the genetic pool to only the strongest, and then there are more females for the fittest males to breed with.”
Brad Higgins also does very well with killies such as AUS by putting them in 10 gallon tanks with a peat moss bottom and lots of Water Sprite (or India Fern in the UK). When fry appear, he pulls the breeders and begins to judiciously feed baby brine shrimp.
For substrate spawners, many use spawning containers (margarine tubs, small ceramic or glass bowls), but some use the divider method. A 4 inch high divider is placed in the center of a 10 gallon tank, with one side of the tank left bare and the rest filled with peat moss. The fish use the side with peat for spawning, while food, filters, and other equipment can be placed in the other. This method has proven useful for a number of the substrate spanwers, both annual and non-annual species.must be the best possible mate you can find.
Sexual selection is an aspect of killikeeping whose details have yet to be worked out for most species. However, it appears to involved with inherited male and female colors, and inherited preferences. Those individuals who readily recognize species recognition markings enjoy more reproductive success, thus perpetuating the species and their own selection criteria.
— G.C.K.A. Newsletter, November 1998 Return to top of page
To Cull or Not to Cull … It’s an Important Question
By Donna M. Recktenwalt
Cull: Verb: to examine carefully in order to make a selection from; pick over.
Noun: something picked out, especially something rejected as not up to standard.
Culling is a necessary evil, one of those things that we often don’t want to think about, but that’s essential if we’re trying to breed strong, healthy fish. If you’re only a person who keeps a few fish, culling is probably a subject of little importance and of less interest. But if you’re actively breeding fish, it’s absolutely essential for the continuing improvement of the fish you are breeding, and for the overall good of the species.
"The future health of any strain of fish depends upon just how handy we are with the culling net," says Greg Niedzielski, discussing this subject. "Culling refers to the process of selecting the best individuals of a strain of fish for future reproduction, and the disposal of inferior specimens. [It]… is an important technique for maintaining the quality of a strain of fish and preventing the circulation and distribution of inferior specimens. Not rigorously culling … may, to some, seem to be a more ‘humane’ approach, but … over time [it] can do great harm to … a species." Culling eliminates the weak and the deformed, removes weaknesses from the breeding strain, and allows selection of the best possible individuals as breeding stock.
"It goes without saying that only the best individuals should be used for breeding purposes, but what many people don’t realize is that the corollary to this is that inferior specimens should be destroyed," Greg continues. "Selling [culls] to local pet stores is not indicated, as this merely distributes inferior fish to unwary customers."Why Should We Cull?
In our artificial aquarium environments, we often are able to hatch a high percentage of the eggs produced in a spawn, and often raise a high percentage of the fry. This is not the norm in nature, where successful reproduction by a pair of fish may result in the survival to adulthood of only a few of the strongest offspring; the weaker, slower, less healthy or less well adapted young ultimately fall prey to predators or disease.
Culling mimics this natural selection process, but other factors may influence our decisions with the culling net as well: size and growth rate, coloration, and personal preference. For the continuing vigor of a species or a strain, we must cull for deformity (missing fins, bent spines, deformed mouths, etc.), lack of vigor, slow rates of growth, and poor overall quality.
Culling for deformity or major weakness is self-explanatory: you never want to perpetuate such problems. But if you’re breeding a strain of one of the more popular species, particularly one with a number of strains based on color or fancy pattern, such as angelfish, swordtails or guppies, what you may consider “culls” may in fact be quite acceptable in the “pet” market, just as “pet quality” puppies are acceptable in the dog market.
Culling is an ongoing process, based on observation. One of the earliest things the aquarist can observe is the condition of the fry. Sickly, undersized, or deformed fish are often apparent long before other selection criteria can be applied. Some hatches of fry grow at such varied rates that they will ultimately “self-cull,” the larger fry preying upon the smaller. Often up to half of a large spawn can be culled early on for size or weakness, allowing more room and better growing conditions for the rest.
As the fry develop they can continue to be culled, both for overall quality and for poor color or color pattern. In killifish, such culling for color and type should always be based on the colors and patterns representative of the wild type. Occasionally a color sport may appear that is worth reproducing, such as the Gold Lineatus or the various colors of Australe. In fish that mature late, or that develop long finnage, culling for all but the most obvious problems may be delayed well into maturity, and even up to show, sale, or breeding times.
There are a number of ways to dispose of the culled fish, depending on the size and the number of culls and on your own personal situation.
Culled fish may be fed to other, larger fish (having a large, hungry cichlid or other predator in the fishroom, or having a friend who has one, does sometimes have advantages). Culls may also be humanely destroyed. Many breeders recommend freezing cull fish in a small amount of water as a painless method of euthanasia. Others suggest decapitation using a sharp razor blade. If you make your own paste or frozen fish food, healthy culls may be used in your favorite recipe as part of the protein requirement.
Releasing culls into local waters is never an acceptable alternative. “Flushing” is also to be avoided, since it subjects the culls to prolonged distress before death, and provides the (hopefully remote) possibility that the fish will survive and get into local native waters.
Culling may not be a pleasant task, but it is a necessary one that serves a number of useful purposes. Cull carefully, dispose of the culls humanely, and then sit back, relax, and enjoy watching those beautiful, healthy fish that remain.
Reference: Niedzielski, Gregory J. “A Few Words About Culling (A Necessary Evil?),” Journal of the American Killifish Association (JAKA) 31 (6) 192-195 (1998).
— G.C.K.A. Newsletter, October 1999 Return to top of page
Diapause in Annual Killifish Eggs … some observations
By Donna M. Recktenwalt
Raising the annual killifish species has always had a touch of magic about it. Where else can you spawn fish, throw the spawning medium into a plastic bag and forget about it for six months, then add water and get fry?
But–surprise, surprise–not all annual eggs must be stored in peat. Some may be quite successfully incubated in water.
All "annual" killie eggs undergo several periods of diapause (cessation of development) before hatching. However, it isn’t always necessary that development of the egg stop during these diapause periods, as usually occurs when the eggs are stored in peat. Given sufficient oxygen, some eggs will continue to develop, although they will go through all the steps of eggs that pause.
To better understand diapause, we need to consider the original habitat of the annual fishes. In most cases, annuals inhabit "temporary" pools of water–seasonal ponds, rain-filled puddles, or small wetland areas resulting from the overflow of larger watercourses. Over time, the species have adapted to their specialized environment, with extremely rapid growth and reproduction rates. The eggs are well protected against dessication, and development of the embryos can cease for long periods of time until conditions are favorable for successful hatching and growth. In some species, under certain anaerobic conditions, annual eggs have been known to remain viable for three years. (Scheel, 1962).
There are usually three distinct stages of diapause. For all killifish eggs development begins a few hours after spawning, when the fertilized egg forms a small rounded body, a blastodisk or blastule. The cells then migrate to the edge of the egg, becoming a multidisk which extends into the yolk membrane. A deep groove then usually appears around the egg’s equator (gastrulation); in time this disappears, the embryo covers the yolk, and development stops. At this point the eggs appear clear to the naked eye.Diapause I
In the wild eggs laid in mud will be in an environment low in oxygen–even anaerobic. If the egg was spawned by one of the annual species, development ceases at this point; the egg may remain unchanged for months. If the egg was spawned by non-true annuals, development may be forcibly halted by reduction of the oxygen supply, to resume when the oxygen levels rise, usually due to drying of the pool. As water evaporates, anaerobic mud becomes moist soil with a higher oxygen content. If kept anaerobic for too long, the non-annual eggs may die.
In the aquarium, development from Diapause I is affected by the spawning media, among other factors. A deep container of fine peat will become anaerobic over time, while eggs incubated in water can be kept well oxygenated.
Diapause II
When development continues, a short, thin-bodied embryo appears on the yolk membrane, usually in the yolk groove. In another day or two, the embryo half encircles the yolk. At this point, embryos from both African annual and semi-annual species may again rest, sometimes for months. These are known as “resting embryos,” or Diapause II eggs. In American annuals, this stage does not necessarily occur, with egg development apparently controlled by available oxygen. When development resumes, the blood and circulatory system appears. After this has occurred, arresting the development of the egg will kill the embryo.
Diapause III
The third phase of diapause occurs after the embryo has consumed its yolk. In annuals, these “resting fry” are characterized by the cessation of circulation, with no heart movement seen. This period of diapause may last for weeks or months. Sudden strong stimuli, such as light, may trigger a heartbeat and circulation, followed by embryo movement, but if the egg is returned to the dark, all movement stops.
Eggs from non-annuals which don’t show these characteristics of “resting fry” don’t hatch when their development is complete, however forced hatching can often be accomplished by adding additional CO2. If there are plants in the container with the eggs, the eggs will often hatch at night.
However, It Doesn’t Always Work Out That Way …
Experience in the fishroom has shown that diapause can be affected by a number of factors, and that even annual eggs don’t always have to go through the usual resting stages.
Among other factors that can affect diapause: the presence of adult fish in the tank, higher temperatures, and oxygen levels. Presumably adult fish, especially females, may produce hormones that keep eggs from leaving Diapause I. Killikeepers that specialize in the annual species have observed that warmer and wetter correlates directly with shorter incubation times, and that higher oxygen levels often result in better hatch rates.
But annual eggs can also sometimes develop continuously in water and then hatch successfully. Dr. John Wourms called these “escape eggs,” since they would be the first of any spawn that hatched. He also observed that different eggs in the same spawn often develop at different rates; this is why we can sometimes get repeat hatches from the same (redried) bag of peat.
For those who are interested, successful water incubation has been reported in the following, although this is undoubtedly only a partial list.
Austrofundulus myersi
Austrofundulus transiliis
Cynolebias belottii
Cynolebias constanciae
Cynolebias melanotaenia
Cynolebias nigripinnis
Cynolebias whitei
Fundulopanchax filamentosum
Nothobranchius guentheri
Nothobranchius korthause Red
Nothobranchius melanospilus
Nothobranchius palmqvisti
Racovia brevis
Racovia hummelincki
Pterolebias longipinnis
Terranotos dolichopterus
— G.C.K.A. Newsletter, May 1999 Return to top of page
Breeding Tanks: The European Method – Worth a Try?
Having trouble breeding some of those beautiful non-annual killies that you'd really like to keep?
Here's a method to try.
In a five or ten gallon tank place well aged water. Add a generous bottom covering of sphagnum peat moss (the long, stringy kind, not the finely ground up peat), a couple of inches deep. Fill the tank with floating plants, water sprite or hornwort, for example. You might or might not include a foam filter or an airstone. Add fish, one to several pair, depending on the species.
Feed fish as usual and do regular water changes. Eggs incubate and hatch in the sphagnum peat, and the fry have plenty of hiding places and a constant food supply from the flora that inhabit the decaying peat and the plants. Every few months conduct a major cleanup of the tank, removing most of the water and the majority of the bottom matter and replacing it with fresh.
This method will work with many of the plant and substrate spawning non-annual killifish species.— G.C.K.A. Newsletter, March 1998 Return to top of page
A few misconceptions about handling annual eggs
Those who breed annual fish encounter enough natural problems, but there are a few misconceptions that can make their task even more difficult.
“In keeping South American annuals … I have run into plenty of bad information,” says Darrell Rommie. “By experimentation I have determined where [some of] the problems lie.” He offers the following observations.
Dry the spawning peat out overnight between two newspapers. Drying out peat overnight between two layers of newspapers usually dries the peat out too much; the eggs will desiccate and die.
Seal in a plastic bag and store at room temperature. Coupled with too-dry peat, incubation of annual eggs at room temperatures (70-75F) prolongs the incubation time and allows the peat to desiccate even more, resulting in poor or nonexistent hatches.
To hatch out the eggs, dump the peat into some tank water an inch deep. Tank water contains potentially destructive bacteria and is usually also deficient in oxygen. Water one inch deep water only ensures that small water changes will be totally ineffective in reducing the effects of decaying uneaten food.
So what does Darrell now do to better his chances of success with annual fish?
He uses covered spawning containers, with boiled sphagnum peat moss as a substrate. Every week or so he removes the peat, squeezes it gently in a net, spreads it out over a layer of damp paper towels and scans carefully for eggs. After he fluffs the peat with his fingers he stores it in a plastic bag properly marked with the species name, the date, and the notation “eggs seen.” Incubation is at 80F. Darrell checks the bags in storage weekly for adequate moisture, refluffs the peat and reseals them.
After about five weeks he places in the peat in a three gallon container and pours in cold (higher in oxygen) water to the halfway mark, 5-6″ deep. The fry hatch out overnight and are fed Wardley’s Liquifry-Egglayers and newly hatched brine shrimp nauplii.
“Obviously, at the speeded up times, not all the eggs will hatch,” he says. “This is where some advice from the past comes in real handy – ‘siphon off the fry and rinse and squeeze and rebag the peat for another 4-6 weeks and then try again.’ “
Reference: Romaine, Darnel. “Too Dry, Too Long, Too Bad!” AquaTropica, volume 2 Number 1. (http://biodec.wust1.edu/~hrbek/aquatrR211.html)
— G.C.K.A. Newsletter, July 1998 Return to top of page
A Few Techniques for Hatching Eggs
By Donna M. Recktenwalt
Every aquarist has his or her own tricks or preferred way to hatch out eggs, whether from annual or non-annual species. What follows are a few of the tips gleaned from the extensive experience of a number of killikeepers.
An egg is usually ready to hatch when the iris of the eye is distinct and shiny. If a light is shone on the egg, the heart will begin to beat, and the embryo will begin to move around inside the egg. However, that doesn’t necessarily mean that the embryo will hatch.
Fish eggs are surrounded by a protein coating known as the chorion. This is exceptionally thick in killifish. When the embryo is ready to hatch it produces an enzyme, chorionase, that helps to break down the chorion, allowing the fry to more easily break through the “shell.”
One thing that helps trigger production of chorionase is CO2, hence the success of those force hatching methods which increase the CO2 level in the hatching container. Exhaled air contains 90-100 times more CO2 than does the atmosphere, thus blowing into a hatching container often speeds up hatching. Some aquarists place “reluctant” eggs into a plastic 35mm film can, blow into it, then carry it around in a pocket for a few hours. This combination of added CO2 and agitation has resulted in nearly 100% hatches.
Another method that has proven successful is to make a complete water change on eggs that are ready to hatch, using water that is 10warmer. Warmer water contains less dissolved gases than colder, so the lower CO2 levels come into play with this method, as well. Adding microworms or a small pinch of yeast to the hatching container has the added effect of providing food for the newly hatched fry.
— GCKA Newsletter, April 1997 Return to top of page
Recalcitrant Breeders?
A Few Tricks to Induce Spawning
By Donna M. Recktenwalt
At one time or another, almost every killifish breeder has encountered a pair of fish that he or she simply cannot get spawn or fry from. However, notes Mike Wilson, the aquarist can make a number of common changes that may trigger spawning in reluctant breeders. The following are a few examples.
Change things in the tank. Move mops, add a mop or two, change the location of substrate containers or filters, add or subtract plants, etc.
Change the water- part or all of it. Some species will respond well to small, daily water changes. Others will begin spawning in earnest only when the water changes are suspended for a few weeks.
Add some boiled fibrous peat (New Zealand sphagnum).
Substantially lower the water level. Be certain that there is at least an inch of water above the top of the spawning container, and leave the water level low for about 24 hours. Breeding often will increase. Be certain to add additional hiding places for the females. If the females have mature eggs, they will find the males.
Move the breeders. A new environment can start or stop breeding. Plan to breed the fish immediately upon moving them; moving older breeders can be risky, especially for annual fish, triggering rapid aging and death.
Use controlled spawning. Separate the breeders and unite them only for brief spawning periods. Be sure to provide hiding places for the females, and to monitor the spawning closely, since males are often very aggressive.
Change the air or water flow.
Add a mirror to the side of the tank. Fish often respond strongly to the presence of "other fish" and egg production will increase. Be alert to increased male aggression and add extra cover for the females.
Try group spawning. Results are not always as good with groups as with pairs, but this is still worth a try. In group settings it is usually the females that initiate spawning.
Change the feeding regimen. Increase it, decrease it, change the times or the quantities. Fish that are heavily fed with quality foods often are less aggressive. Remember that heavier feeding means more water changes. Make sure that all females get enough food. Males and dominant females may monopolize the food source.
Change the temperature..
Check the spawning mops at various times. Check mops three times a day to determine when the fish are actually breeding. Different species (and even different pairs of the same species) can surprise you. Collecting eggs at different times can also break the cycle of spawning followed by egg eating.
Try a mixture of peats. Even some of the peat spawners have been known to search for and eat their own eggs. Try a deeper substrate, a mix of fibrous and non-fibrous peat, bottom mops atop the peat, or an alternate substrate material entirely - fine glass beads, greensand, or bottom mops alone.— GCKA Newsletter, October 1997 Return to top of page
Focus on Annuals
Finding and Storing the Eggs
By Donna M. Recktenwalt
Breeding and raising species of the annual and semi-annual killifishes is a challenge for the aquarist, given their substrate spawning behavior and the subsequent storage period required by the eggs. In a previous article, we focused on the spawning media available for use by the aquarist. In this article, we assume that you have observed your annual or semi-annual fish successfully spawning.
But can you prove it by finding the eggs?
This is hardly a minor question. Actually seeing newly laid eggs can prove whether you actually have eggs to store. Later on, seeing eyed up eggs can help evaluate whether the peat is ready to wet.
Seeing even small eggs that have been freshly rinsed from clean spawning mediabe it greensand, glass beads, walnut shells, sphagnum moss or mopsis not terribly difficult, no matter the egg size. Eggs vary from the very small to the very large; from clear (transparent) to a pale amber when freshly laid.
Seeing the eggs after storage in peat, however, takes practice, since no matter their color when laid, the eggs soon become stained a dark, dull brown.
"In my opinion," says Andrew Broome of New Zealand, "the only definite way to check for eggs in peat is to do it as soon as the peat is harvested. I figure that I can spot about 10% of the at that stage clear eggs." For spotting eggs, Andrew uses overhead or side lighting, and manipulates the peat in its storage bag while looking through the open top.
Jay Exner suggests using a backlight and "getting the eggs ([on] spawning mops, plants, whatever) between your eyes and a bright light. You will see bright dots for new eggs (i.e., the optical effects of light being refracted through spherical lenses." This works for peat if you can spread it out on glass with a light behind itin a glass or clear plastic baking dish, for example.
"I'm far older and have poorer vision than most," observes Wright Huntley, "but if I can find Cynolebias fluminensis eggs [in peat] when ripe, you can find them." His tips?
Use magnification to help youa jeweler's loupe or a hand-held magnifying glass.
Use a bright desk lamp (Halogen is particularly good) positioned directly across from you.
Use a white-bottomed container, and spread the peat thin. Sometimes light reflected from the bowl beneath will shine through the eggs.
Fluff the peat, then pat it down to a fairly smooth surface. Scan over it, using your magnifier, with the lamp only a couple of feet from the surface.
No luck?
Lay a paper towel over the whole mass of peat and flip it over to look at the other side. Refluff and repeat this procedure a couple of times.
As a last resort, start at one edge of the peat and slowly separate it into "examined" and "unexamined" portions, with a gap between, using tweezers or a similar tool to move the shreds of peat fiber from one pile to the other.
With luck, you'll find eggs in that newly harvested peat and can put it into storage with confidence.Storing the Eggs
The fish have spawned. You’ve collected the spawning media and checked it for eggs. You’re ready to put the eggs away for the recommended incubation period.
But what media do you use to store those eggs, and into what do you put that material?
The nearly unanimous answer regarding the proper storage medium for incubating annual or semi-annual eggs is slightly moist, crumbled peat. It’s acidic nature helps keep bacteria growth to a minimum and helps prevent eggs from fungusing. The eggs are also effectively separated from each other so that any that do go bad don’t infect the rest.
If the eggs were spawned over peat, simply dry the spawning medium to the proper degree of moistness before storage, using paper towels or newspaper to wick away the excess moisture. If spawning was accomplished over another medium, place the eggs into the slightly damp peat and mix it before drying, to achieve an effective distribution.
By far, the consensus regarding the proper storage container for egg-laden peat is plastic bags. These may be either standard plastic fish bags, or plastic freezer bags. Both are fairly permeable to the small molecules of oxygen, but impermeable to the much larger water molecues, which means that developing eggs get sufficient oxygen without the risk of drying out. Place the egg bearing peat into a plastic bag, seal the bag, and mark it properly as to species and spawning date.
More tightly sealed containers, such as plastic petrie dishes sealed with electrician’s or similar tape, have proven acceptable for eggs that undergo only a short diapause (4 weeks or so). For eggs that require longer storage they are a poor choice, due to their impermeability to oxygen.
Once the eggs are safely in their storage container, look up the appropriate incubation period based on the temperature conditions in your fishroom, and mark down the anticipated hatch date on your calendar, on the bag itself, or on index cards filed by month.
Now put the bag of resting eggs away.
From now on it’s just a matter of time. All you have to do is … wait.
— GCK Newsletter, October 1997 Return to top of page
Fry the Easy Way — (with apologies to Julia Child)
By Donna M. Recktenwalt
A major fascination about many of the killifish we keep is the way they reproduce, but breeding them can be a lot of hard work, especially for those species that require long incubation times.
What if you want to keep and breed killifish with a minimum of effort?
With some species of plant- or mop-spawning fish, it can be done.
Start out with any size tank, densely planted or well filled with fine leaved free-growing plants such as hygrophilia, hornwort, watersprite, or java moss. Add a slow running foam filter, either external or internal. Introduce the fish, feed well and do regular partial water changes; then watch fry of various sizes begin to appear.
Sound too simple?
For some species of killifish, particularly the annuals, this approach would be next to impossible; for others it is nearly ideal: the fish fertilize the plants; the plants host a colony of microflora and fauna and provide sites for egg-laying and cover for the fry; which feed off the microscopic life.
Supposedly many European killikeepers routinely breed their mop-spawning killies this way.
“I have a 20 gallon high tank that is about 3/4 full of java moss,” says Shane Essary. “I put the pair of fish that I want to spawn into the tank …. Pretty soon the tank is full of babies that get really big without having to feed them [heavily on] baby brine shrimp.” Shane has used this method successfully for A. scheeli. For A. abacinus, he substitutes mops for the java moss but still lets the eggs hatch naturally.
“I have had good luck with A. celiae celiae, letting them breed in a well planted tank and not picking eggs or rescuing fry, also with E. dageti, using java moss,” reports Harry Kuhman.
Donna Recktenwalt has been raising A. chaytori Ngabu and A. mirabilis Takwai in separate densely planted colony setups. “I get as many fry from the tanks as I do from collecting and incubating the eggs,” she says. “I regularly find fry of various sizes in with the adults. The parents seem to ignore them entirely.”
“Almost all my Epiplatys annulatus will allow young to survive,” says Lee Harper, “and a trio of Pachypanchax omalonatus has shown tolerance for young fish. Diapterons in general will not.”
“I’ve had this experience [tolerant parent killifish] with E. fasciolatus (big time! we’re talking hundreds from four adults in a plant and algae choked 40 gallon tank), E. dageti, and (to a much lesser extent) A. australe,” says Richard Sexton.
Other species that aquarists report have shown tolerance for their fry include A. ogo ottogartneri; A. coeleste, A. aureum, and A. citripinnis.
For particularly troublesome species, a variant of this method has been used successfully by some. Instead of simply filling the tank with plants, pack it nearly full of java or sphagnum moss, leaving only a little space around the outside for the adult fish, and a small depression at center top for the fry. The parent fish will spawn in the java moss and the hatching fry will gravitate to the depression at center top, where they can easily be removed for growout in a separate container.
This doesn’t always work, but may be worth a try for fish that are highly aggressive, or that regularly prey on their young.
— G.C.K.A. Newsletter, December 1997 and March 2002 Return to top of page
Starting ’em out right – A Guide to Raising Fry
By Donna M. Recktenwalt
Getting your chosen fish to pair off and spawn may provide a real feeling of success to an aquarist, but the next step is just as critical getting the resulting fry off to a good start. There’s many a peril between a clutch of fertile eggs and a tankful of healthy, thriving adult fish.
Consensus among successful breeders is that the first tank for fry should be small. Newly hatched fish are usually unable to swim very far or very fast. A fairly small, shallow tank or container keeps fry and food in close proximity. Young fish usually have good appetites; they must eat frequently and heavily in order to grow. Fish fed only once or twice a day will rarely become large, robust breeders, so feed them as many as half a dozen times a day, if possible. As the fry grow, they can be moved to larger quarters.
The disadvantage of small quarters and frequent, heavy feedings is rapidly accumulating high levels of pollution from wastes and decaying foods. High levels of nitrogenous waste also act as growth inhibitors, so frequent partial water changes (up to 50% several times a week, or even daily) are essential to maintain healthy growth rates in the fry.
Good filtration can be as important as regular water changes. Sponge filters have proven best for use with small fry, since they can’t be sucked into them, and often feed from microorganisms growing on the filter surface. Also useful are inside, air-driven box filters. Small fry often swim into such filters and feed on the trapped food particles and microorganisms there, so it’s best to leave off the lid, to prevent fry from being trapped.
Ideally, food source(s) for the newly hatched fry should have been planned for well ahead of time. Young fish need a variety of foods to grow their best, but providing it is not as simple as it is for adult fish due to the small sizes required. Dried and powdered foods can be used, both crushed adult foods and commercially prepared food for fry, but there’s no substitute for live food. There are a number of possibilities, depending on the size of the fry and the space, skills, and capabilities of the aquarist.
For all but the smallest killifish fry, the most easily available live foods are baby brine shrimp (Artemia salina nauplii), microworms, and vinegar eels. Microworms can easily be cultured by the aquarist using cornmeal slurry or cooked oatmeal as a base; vinegar eels grow without care in a mixture of apple cider vinegar and water. Baby brine shrimp can be hatched out in the fishroom with little trouble. If available, baby daphnia may also be used.
However, some fry are so tiny that even these small foods are too big. For such babies, greenwater, infusoria, or yeast or egg yolk infusions may be necessary.
Greenwater is no more than single celled algae, and can be cultured by adding algae from the aquarium or a few drops of liquid fertilizer to aged aquarium water and leaving it in a sunny location. Feed a small measure of the liquid to the fry.
Infusoria are small, single-celled animals such as euglena, paramecia, or amoebas, or microscopic multi-celled ones such as rotifers. To aged aquarium water add crushed lettuce and the culture starter. When it gets cloudy it is ready to feed, as above.
Yeast Infusion: dissolve some baker’s yeast in aged aquarium water. Feed immediately.
Egg Yolk Infusion: Hardboil an egg and remove the yolk. Mash to a fine consistency, then swirl in water. Feed small amounts of the resulting cloudy liquid.
As fry grow, they may “graduate” to larger and more varied foods. Remember, though, that the more variety in the diet and the cleaner you can keep their quarters, the better they will do.
You’ve managed to bring the clutch of eggs and the newly hatched fry safely through the early weeks of growth, and they are coming along nicely. Now comes one of the harder tasks, sorting and culling the fry. Although some aquarists seem driven to “raise them all,” sorting and culling are necessary tasks.
Sorting. Most broods contain a few individuals that grow more quickly than the others. This disparity in size can lead to harassment and actual predation of smaller individuals by the larger ones. Sorting fry by size can thus result in more fry successfully raised. For the same reasons, it is a poor idea to mix broods of different sizes and ages together.
Culling. Few broods also fail to include some specimens that are obviously weaker, less healthy, or that are deformed. In nature, these individuals would rapidly die from predation. In the more protected environments of aquarists’ tanks, we must be responsibile for selectively destroying them. These culls can either be euthanized (one humane method is to freeze them in a small amount of water), or (if they are not diseased) to use them as feeders for other fish.
Following these general steps, you should be able to have the satisfaction of raising a group of healthy, well developed young fish. From these you can then proudly and confidently select your future breeders, your show fish, and those you will sell to other hobbyists.
References:
Marshall E. Ostrow, “Raising Healthy Fry,” Tropical Fish Hobbyist, November 1980, pp. 29-38.
Bill Volkart, “Feeding Fry: How Big is Too Big?”, Tropical Fish Hobbyist, June 1994, pp. 78-81.
— G.C.K.A. Newsletter, September/October 1998 Return to top of page
Handling Eggs – Now What?
By Donna M. Recktenwalt
Even the best and most experienced killikeepers occasionally have problems with eggs and fry. The “proper way” to incubate and hatch out killifish eggs varies, of course, with the species, their spawning habits, their incubation requirements, and the preferred techniques and proven methods of the fishkeeper.
“[Water incubated] eggs, if they are viable, will do their thing,” says Charles Harrison. But sometimes, they need a little assistance. Between spawning and hatching, all eggs are under threat. They may be eaten. They may fungus. They may hatch prematurely. Or they may simply disintegrate entirely, leaving no trace.
“If [eggs] don’t develop and hatch in a timely fashion, they will disintegrate,” Lee Harper says, discussing his experiences with Aphyosemion sjoestedti. “I collect eggs from the gravel and watch them for a few days in water. If they seem to be still alive after several days (i.e. they don’t fungus or disintegrate), I then put them on or in peat. If I want to be able to count them or watch them I put them on top of damp peat. If I don’t care I just mix them into damp peat and incubate for 6 to 10 weeks. Sometimes they don’t hatch … even though they seem viable at first.”
“I never keep my [killie] eggs in tank water for fear of a higher bacteria count, which might destroy the shells and cause premature hatching,” says Jeff Bilbrough. In premature hatching, eggs die as their shells rupture. This can happen early in the development of the embryo, where it is seen ‘oozing’ out of the shell, or at subsequent stages, leaving an incompletely developed, unprotected embryo that soon dies. In the latter stages, the tail punctures the egg membrane, but the remaining part of the embryo can’t free itself.
“I do use methylene blue,” Jeff continues, “but only for the first three days to determine unfertilized eggs; they’ll turn dark blue.” He changes water in the hatching container daily, adding a few grains of Jungle brand Binox. “I’ve [also] begun sterilizing my hatch containers with weak Clorox and drying them for a few days before using them,” he says. “Since I’ve started these procedures I seldom have premature hatching and fry dying young.”
“What has worked to a degree [for premature hatching] is changing the hatch water every few days,” reports Howard Berg. “If the fry are already prematurely out of their shell I have been able to save some by changing the hatch water every day until they have used their yolk sack and become free swimming.”
Al Anderson increases the general water hardness by adding a teaspoon of well water to each 6 oz. of hatch water. “Most of the time this works,” he says.
“Keep the water that the eggs are in as clean as you can, says Charles Harrison. “Change it every few hours if you can. Pour the eggs through a Brine Shrimp net, rinse out the hatching dish, replace the eggs/fry and keep it all clean and fresh. … The best containers are the flat bottomed ones so the eggs don’t slide together and get involved with fungus, etc.”
When eggs handled in any way are dying off, then not handling them at all is a useful strategy.
Having several breeding tanks ready offers another solution; when the adults have spawned in the first tank for two weeks they are simply moved to the next and the eggs allowed to incubate unmolested. Alternatively, you can remove the mops or peat to another container for hatching. If you have spawned your fish over gravel (especially in a tank with an undergravel filter) the regular circulation in the tank can often provide sufficient aeration for the eggs.
If one method doesn’t work for you, try another. Keep in mind, too, that not all breeding fish will always produce viable spawn. Some species are notorious for infertile eggs when young, with fertility increasing as the fish age. In other cases, a pair may be fertile for a time, then cease producing eggs altogether for a while.
— G.C.K.A. Newsletter, January 1999 Return to top of page
How Big Are They, Anyway?
As killifish breeders, we often are faced with the question of “where are the eggs?” either when picking eggs from mops, or when checking peat in which fish have recently spawned. Sometimes there is no doubt that eggs are present: with some species, they are large, clear, and obvious. With other fish, the eggs may be extremely small, discolored from their proximity to peat, and deceptively difficult to see.
So how big are the eggs of various killifish? Recently Lee Harper shared the following information with the members of the killifish e-mail list (16 May 1998). Although the following list is far from complete, it may provide some guidance to other killikeepers.
Egg SizeSpecies / mm / in. / mils / micr.
E. annulatus / 0.90 / 0.035 / 35.43 /900
N. foerschi / 0.95 / 0.037 / 37.40 / 950
A. striatum / 1.20 / 0.047 / 47.24 / 1200
A. cognatum Bandundu / 1.25 / 0.049 / 49.21 / 1250
A. rectogoense / 1.25 / 0.049 / 49.21 / 1250
A. bittaeniatum Ijebu Ode / 1.25 / 0.049 / 49.21 / 1250
A. australe #14 / 1.25 / 0.049 / 49.21 / 1250
N. voselleri Karogwe / 1.30 / 0.051 / 51.18 / 1300
A. cyanostictum (Diapteron) GBN 88/29 SAM / 1.30 / 0.051/ 51.18 / 1300
A. wachtersi Obili / 1.30 / 0.051 / 51.18 / 1300
A. fulgens (Diapteron) LEC 93/2 / 1.30 / 0.051 / 51.18 / 1300
E. chaperi Angona / 1.30 / 0.051 / 51.18 / 1300
F. gardneri nigerianum / 1.35 / 0.053 / 53.15 / 1350
P. omalonatus Nosy B‚ LM 94 / 1.45 / 0.057 / 57.09 / 1450
E. fasciolatus (embryo) / 1.50 / 0.059 / 59.06 / 1500
F. gardneri N’sukka / 1.60 / 0.063 / 62.99 / 1600
A. louessense RPC 33 / 1.60 / 0.063 / 62.99 / 1600
E. fasciolatus fasciolatus (new) / 1.60 / 0.063 / 62.99 / 1600
P. sakaramyi Joffreville LM/94 / 1.70 / 0.067 / 66.93 / 1700
A. sjoestedti / 1.80 / 0.071 / 70.87 / 1800
R. aff. holmiae / 2.10 / 0.083 / 82.68 / 2100
Ap. lineatus Gold / 2.10 / 0.083 / 82.68 / 2100
R. cryptocallus / 2.25 / 0.089 / 88.58 / 2250
Fundulus chrysotus / 2.40 / 0.094 / 94.49 / 2400
— G.C.K.A. Newsletter, Sept/Oct 1998 Return to top of page
How I Do It …
For those who breed killifish, there are always challenges ahead. In this occasional column, we’ll highlight a few breeders and their proven techniques.
"I breed killies as trios in gallon pickle jars," says Zavier Burgos of Orlando, Florida. "It works great with the spawning mop and the fish, because all they do is breed. They are not looking for each other, they know of each others’ presence in the jar and they go for it."
This jar method works well for a number of the smaller species and often yields large numbers of eggs – "I have gotten from a single pair of A. striatum 175 eggs, and all but one hatched." – but it is more time intensive since you must supervise the breeding setup.
"I condition the fish for 2-3 weeks, depending on the fullness of the female," says Zavier. He separates the sexes during this conditioning period, and feeds them up to six times a day with a variety of foods, including 40-46% protein flakes, spirulina flakes, live foods, and freeze dried tubifex and bloodworms.
About 25% of the water in the conditioning tanks is changed every other day. Xavier uses rainwater for all his breeding tanks, aged for at least two weeks before use, then filtered for a day through peat and for two more days over charcoal. "The peat has hormones that induce breeding, and it disrupts the carbonate hardness in the water so you can adjust the pH more easily," says Xavier. For each breeding setup, this pretreated, aged water is put into a gallon size pickle jar with a spawning mop and the fish added.
"Don’t use lighting while breeding," cautions Xavier. "Just place the jar in a corner at eye level." You will be able to see the fish, but not the eggs. Breeding is over when you see no more movement from the fish. This may be as little as 15 minutes to two hours or more, depending on the species. When breeding is done and the female is tired, be certain to remove her; the male can be left for another hour or so. If all has gone right, "you should have too many eggs to count and all [of them] fertile."— G.C.K.A. Newsletter, March 1999 Return to top of page
An Overview
Inbreeding in Killifish
By Donna M. Recktenwalt
Population geneticists are well aware that there is great variation in the amount of inbreeding that occurs in nature and the resulting inbreeding depression that often occurs in captivity. However, the mechanics that cause this effect are not obviously clear.
For example, in Rivulus marmoratus most individuals are completely homozygous (alike), and natural populations (with a few exceptions) are simply arrays of homozygous clones. Yet this species inhabits one of the most challenging environments in nature, mangrove swamps that endure great fluctuations of salinity, temperature, etc.
Also, peripheral populations of Xiphophorus maculatus show little inbreeding depression in the laboratory, while those from larger and more centrally located populations show strong depression effects in just a few generations of inbreeding.
There seem to be two important active variables at work here: effective population size and the extent of inbreeding in nature. Large populations tend to accumulate deleterious recessive genes; when these populations are inbred in the laboratory, these recessives (which would persist indefinitely-and probably harmlessly-in the heterozygous form) become homozygous and result in inbreeding depression. The mating systems of some populations favor inbreeding (the self-fertilization of R. marmoratus is its ultimate form). These populations gradually eliminate the deleterious recessives without much inbreeding depression in any given generation.
This leads to two questions: how big are the wild populations? And do they have mating systems that favor inbreeding? If only a few males in a large population do most of the breeding, the effective population sizes may be smaller than expected.
There is also such a thing as outbreeding depression, most often expressed as F1 sterility. This might be encountered when crossing lines derived from populations which (unknown to us) are genetically incompatible. Chromosomal changes (“mutations”) often lower the fertility of heterozygous carriers, since they can result in unbalanced chromosome sets. Other variants, such as Robertsonian translocations (another form of chromosomal transfer) also may occur. The results of such mutations usually die early. If two populations happen to be fixed for several chromosomal differences, any hybrids between them tend to be sterile. Sometimes, by normal genetic drift, a mutation can become fixed in one population but not another; hybrids between two such populations often are viable, but have reduced fertility.
Many of the West African killifish, particularly from the Fundulopanchax and Aphyosemion groups, are of interest to evolutionary geneticists because they consist of populations with different multiple Robertsonian translocations. As Joergen Scheel showed (see Rivulins of the Old World, the first edition) hybrids between these fish are often viable, but sterile. Basically, these groups of fish are arrays of chromosomally differentiated “sibling” species.
Thus the idea of “introducing new blood” by crossing wild-caught individuals into established stocks is risky. In the absence of sufficient genetic information or precise locality data, it should not be attempted.
— G.C.K.A. Newsletter, November 1998 Return to top of page
Incubating and Hatching Eggs
By Donna M. Recktenwalt
There are probably almost as many ways to incubate plant-spawning killifish eggs as are there are killifish keepers. Relevant topics for discussion always seem to include the best methods for collecting eggs, for incubating and hatching them, and for raising the resulting fry.
For fish that spawn in plants, the "natural" way is simplest. Either remove the parents to another tank after a week or two, or let the eggs hatch and fry develop in the same tank with their parents.
For killifish that have spawned in mops, egg collection is straightforward: shake the breeding mops gently to flush out any fish hiding within, remove the mops from the water and squeeze out, then search for eggs by pulling aside two or three strands at a time in bright light. The eggs may be removed from the mop for hatching in a separate container either in water or on peat, or the entire mop may be removed for hatching elsewhere, without handling the eggs at all.
One thing that may affect hatching success, and one that few people think much about, is cleanliness of their hands. Did you wash your hands before working with the fish or their eggs? Have you recently worked with materials that could have an adverse effect? Even slight chemical residues can have immediate (and sometimes fatal) consequences in aquaria.
For species that produce eggs more than usually sensitive to handling, the handling itself can be disastrous. In such cases, the entire mop can be removed to a separate hatching container or a plastic storage bag. This removal technique has proven useful for such fish as Epiplatys annulatus and some of the rainbowfishes, but can be used for more common species as well.
Whatever incubation technique you choose, fish eggs must have water to develop properly. But what water should you use? Many killikeepers use water from the parents’ tank. Some suggest using aged new water. Still others swear by distilled or reverse osmosis water. Some insist that a medicated hatch water mix is crucial to success.
Can the eggs be exposed to light, or should they be kept in darkness? Adherents from both points of view point to success in hatching eggs and raising fry.
Should You Medicate Hatch Water? "I would see if the eggs develop without using any antifungal chemical," says Ralph Taylor. Fertilized, viable eggs seem to develop no matter the conditions, surviving even fungus and neglect to produce viable fry. Aquarists have also noticed that when fish are just starting to produce eggs fertility (viable eggs) is often very low. As the fish mature, fertility improves.
For those concerned about fungus and bacterial contamination of the eggs, several hatch mix formulae have proven useful.
Methylene blue is a biological dye which a number of bacteria can use as an alternative to oxygen for respiration; when used this way, the dye turns colorless. This "favoring the bacteria" approach may put fungus at a disadvantage, thus providing some additional edge to egg development.
"I put a small amount of methylene blue in with new eggs," says Donald Nute. "This will not keep an infertile egg from fungusing, but it will slow down the poisoning of fertile eggs by the toxins the fungus produces." When eggs start to show development, he moves them into clear water.
"I see little hatch-rate difference from any dyes I add to the water," says Wright Huntley. His hatch mix? To one gallon of reverse osmosis water add a teaspoon of salt, 5 drops methylene blue, and fungus guard or Binox (made by Jungle Labs).
Donald Nute stores his incubating eggs in the drawers of a small plastic storage cabinet. The eggs are stored in methylene blue treated water and are checked daily, with fungused eggs removed. Developing eggs are moved into clear water which is changed once or twice a week.
Acriflavine. "I used to use methylene blue as an antifungal agent for all of my eggs," says Cal Him. "Then I switched to Acriflavine. I put one teaspoon salt, 4 drops Aquarisol and 1 drop Acriflavine into one gallon of water." Acriflavine Plus is an entirely different formula and contains Malachite Green, which may harmful.Incubation and Hatching
Hatching containers may be any small, lidded, easily handled glass or plastic containers that can be readily cleaned and which will keep eggs from clumping together, thus fostering direct cross-contamination. Small baby food jars, petrie dishes, margarine tubs, even the small sauce or dressing containers from restaurants have been successfully used for incubating eggs.
Some aquarists have their best hatches by using fresh aged water and changing it every day (or as often as possible). Water may be gently poured out and replaced in the original container, or the eggs may be poured through a fine mesh net and then moved into a clean container. A sprig of java moss may help keep the water cleaner.
“Clean [water] and high oxygen content seem to be the best qualities for water incubation,” says Wright Huntley, “but doing it on damp peat is even better.” Incubating the eggs on damp peat (just slightly squeezed out, so the surface is still wet but not liquid) lets you stall hatching for a few days. If you are collecting just a few eggs a day, you may be able to hatch a weeks’ spawn at once, making rearing easier. Be certain to check for bad eggs every few days, and for eye development at 3-4 weeks. Most eggs will hatch on their own when they are fully developed. Eggs that are ready to hatch, both peat- and water-incubated, will show a clear eye-ring. Eggs can be hatched out in a shoebox or other suitable container, using water from a tank with plants. After a day or two, add some snails and some duckweed or java moss.
But sometimes eggs don’t hatch. What then?
When the eggs look well developed, “I put them in a 35mm plastic film canister and carry them in my pocket” for a few hours, says Donald Nute. “Then I can pour the fry out into a container with aged water and a bit of java moss.” This procedure results in about 90% of the eggs hatching at one time. Left alone, the same eggs would probably hatch out over a period of several days.
A few alternative methods to force hatching:
Put the eggs in a small, half filled bottle, then exhale into it (there is 90-100 times the amount of CO2 in exhaled air than there is in the atmosphere). This technique has even been successfully used to encourage “reluctant” annual eggs to hatch.
Add a small measure of microworms to the hatching container.
Bring down the pH slightly by adding a peat pellet to the hatching container.
Whether it is the warmth or the decreased oxygen levels that work, try making a complete water change on the eggs using water that is 10°F warmer, or move the incubation container to a warmer location.
Sometimes, however, we just have to admit that those eggs won’t hatch.
Better luck next time!
— G.C.K.A. Newsletter – August, 1999 Return to top of page
How I Do It …Incubating and Hatching Eggs
"Most of my Fundulopanchax eggs I gather using fine sand or fine peat," says Al Anderson. He boils the peat three times to remove most of the tannins, then puts the peat into a blender and chops it until it is so fine that it passes through a homemade net made out of mosquito netting bought from a fabric store. The netting is coarse enough to allow the peat to pass through, but fine enough to hold the eggs.
Harvested eggs are rinsed off into a small bowl of clear tap water, then picked up with an eyedropper and gently rolled around on a paper towel to clean them off. For storage, Al places the eggs about a half inch apart on a sponge that has been boiled and wet with tank water that has a small amount of Acriflavine in it. The sponge is kept in a light-proof container (he uses a shoebox spray-painted black on the outside). For the first few days he removes any white eggs, then leaves the eggs alone (except for checking that there is enough water in the box to keep the humidity up) until they are eyed up.
When the eggs are ready to hatch gold eye rings are visible, and if a strong light is pointed at them, the fry will sometimes spin around inside the eggs. For hatching, the eggs are placed in a bowl of tank water about one inch deep, stirred to bring the eggs together into the center of the bowl, then a few microworms added.
The bowl is then floated in the shoebox where the fry will be raised, and the fry gently released. Al keeps the water level in the fry tanks about 2" deep for the first week or so.– G.C.K.A. Newsletter, October 1999 Return to top of page
Incubating Eggs … A Judgement Call
By Donna M. Recktenwalt
How long to incubate killifish eggs is a matter determined by the species, the incubation media and its wetness, and the temperature at which the eggs are stored. Most non-annual killi eggs incubate in water or on wet peat for 14-21 days at average household temperatures of 72-75°F, although a few take considerably longer. At higher temperatures, development is accelerated, with hatching sometimes occurring in as little as ten days.
For annual eggs stored in peat, normal incubation times are generally a consensus of experience and convenience. Some may successfully hatch after a month of water incubation; some may peat may be wet in only a few weeks; but most need to be incubated for several months, a few for as much as 8 or 10 months.
Published tables provide suggested incubation times for various annual species, but these should only be taken as guidelines. It is, however, wise to establish that the embryos are fully developed before wetting stored peat, since individual conditions may result in longer or shorter incubation times.
Temperature has a major influence on development rates, with warmer usually resulting in shorter incubation times. However, warmer and drier may slow development.
Are They Ready to Hatch?
Most non-annual killifish eggs are incubated in water or on wet peat. Development in water is easy to monitor, removal of bad eggs is simple, and when the eggs are ready, they usually hatch without complications. Eggs incubated on wet peat have a reduced risk of fungal contamination, but are more difficult to observe due to the dark coloration of the storage medium.
Especially with annual eggs, knowing whether the eggs are ready is important. Wet peat too early and you may have bellysliders or no hatch at all; wet too late and the embryos may already have died.
The best indication of an embryo’s readiness to hatch is the appearance of a dark, shiny iris in the eye. Or shine a bright light, such as a flashlight, on the embryo. If you can see the heart begin to beat, or a fiftul “swimming” activity within the egg, the embryo is ready. Both these techniques are better accomplished using an inexpensive low-power microscope, or a bright light and a strong hand-held or free-standing lens.
Knowing when annual eggs are ready is of major importance to the aquarist, but eggs in peat are notoriously difficultr to see, especially after they have been in storage for a time. Some squarists can “fluff and spread” the peat over a light colored background and spot eggs fairly easily, although most agree that the longer the eggs have been in storage the more difficult the task, since the eggs take on the peat color.
Some recommend setting a small clear-sided container of water on a sunny windowsill and sprinkling a bit of peat on the surface. The eggs sink quickly (some say “plummet”), allowing observation of the developing fry. However, some annual embryos are pale or nearly transparent, making them doubly difficult to see.
But The Eggs Won’t Hatch!
Sometimes knowing that the embryos are ready isn’t quite enough. What to do when they won’t hatch?
The outside of a fish egg consists of a tough protein coating known as chorion. This is exceptionally thick in killifish and one of the reasons annual eggs are so durable. When the embryo is ready to hatch it produces an enzyme (chorionase) that breaks down this protein, softening the outer shell so the fry can break through. The production of chorionase is triggered by rising levels of Carbon Dioxide (CO2) around the egg. In nature this occurs when soil containing fish spawn is wet by rains. Mud holds less oxygen than moist earth, signaling the developing fry that conditions are right for hatching.
This adapotation can be directly utilized by the aquarist to “force hatch” eggs, by placing them in a container and exhaling into it before carrying it around in a pocket for a time, or by adding a measure of microworms to the hatch water.
— G.C.K.A. Newsletter, Sept/Oct 1998 Return to top of page
Incubating Eggs on Peat
By Donna M. Recktenwalt
One of the most useful tools that killifish breeders have at their disposal is peat moss. We use it as a pH buffer, as a spawning medium, and as a substrate. However, many aquarists have found that it has another use: as an incubation medium for the eggs of many plant- or mop-spawning non-annual species.
"I hardly ever water incubate eggs," says Geert van Huijgevoort, "especially with species like Fundulopanchax amieti or F. puerzli, that need a longer incubation period." Even with frequent water changes, he points out that it is nearly impossible to prevent fungus from wiping out all or most of the eggs. "By far, the best way for me to incubate eggs, whether they are from Aphyosemion, Fundulopanchax, Epiplatys, or Rivulus, is to put them on top of boiled or microwaved peat."
In a petri dish or plastic container (small butter or margarine tub) with a tight lid, place a layer of peat moss that has been boiled, drained through a net and squeezed until no more water runs out. To this add water from the tank where you are harvesting the eggs, or your favorite hatching water mix. Allow the peat to soak, then pour out excess standing water.
Eggs should be placed on top of this peat without touching each other, the container covered and kept in the dark. The first week or so, check every few days and remove any unfertilized or fungused eggs. After that, check weekly to make certain there is sufficient moisutre, adding water as required.
At the same time, check for eye development. The eggs are ready to hatch when a nice gold ring is visible around the eye. Some eggs will even show movement of the embryo when stimulated by a bright light.
When the peat is wet (flooded with water) after the incubation period, all fry usually hatch within one or two days, so eggs laid over the period of a week or two can be hatched together. Any laggards can be "encouraged" by using any of the proven methods for forced hatching.
"On peat there seems to be some leeway on hatching time," says Mike Reid. His students use damp peat to incubate F. gardneri, and have found that it can double the incubation times. A. australe eggs usually take three weeks to hatch, but that time can be extended to six. The technique has also been proven with A. cameronense, A. striatum, and others.
There is a downside to the peat technique, however. "Sometimes I find little hatchlings dead on the peat," says Geert, "especially with my Chromaphyosemions," which lay eggs that sometimes hatch after nine instead of the usual twelve days.
A variation of this incubation technique is to use clean paper towels wet with your favorite hatching solution; this may not always work, but may be worth a try if you’re having problems with hatches.- G.C.K.A. Newsletter, February 1999 Return to top of page
Keep More Than One Pair!
As every "experienced" killikeeper knows, only a few basic rules must be observed when keeping killifish. Keep them clean (most of the time). Keep them covered (all the time)! Keep them well fed and happy. Given the above, killifish usually flourish and reproduce.
But there are a few other, unwritten rules that many killikeepers have learned the hard way. We’re not talking here about the finer points of animal husbandry, but of the invariable inclusion of Murphy’s Law to fishkeeping – "whatever can go wrong, will."
Rule One: Always buy more than one pair of a new fish (if they are available, and if you can afford it). Invariably, one of the pair will die soon after purchase, sometimes even before you get them home. There’s nothing more frustrating than having only half of a pair and being unable to find a replacement. Consider a second pair as insurance.Corollary One: “If you buy the only pair (in the auction, at the convention, in your area, the country, etc.) the male will kill the female within 24 hours.” (Wright Huntley)
Corollary Two: The same thing will happen down at the fishroom level. “Even if separated, one fish (usually the male) will find the one gap in the tank cover, and [become a] crispy critter.” (Cathy Carney)
By buying more than one pair you also have more breeding options to work with, which can be a plus when you are attempting to maintain genetic diversity and the health of a strain. An alternate to buying more than one pair of breeders is to buy a bag or two of eggs. Sometimes you have the option of buying both, particularly when the breeder has a number of pairs spawning. The genetic advantages of this are obvious, but another thing to keep in mind, says Scott Davis, is the fact that “fry grown out in your own water are often more durable and longer lived,” than are purchased fish, however healthy. Stock brought into your fishroom sometimes never acclimates effectively.
Corollary Three: If you buy two pair they will all get along fine and live to a ripe old age. It happens – but don’t depend on it.
Rule Two: If you successfully breed the fish, distribute some of them immediately. You may fail to breed the fish again. You may end up with fry of all one sex. Having other stock around provides you with backup insurance, and helps to keeps the species in the hobby.
– G.C.K.A. Newsletter, October 2000 Return to top of page
Killie Soup – Yum!
Thanks to Vitee Tao, via the Killifish (e-)Mail List
Here’s a new thick, chunky recipe for storing semi-annual eggs in this case, F. filamentosus (FIL).
Use a generous quantity of peat moss.
Add a small handful of java moss.
Introduce a pair or trio of FIL into a 12 inch tank (water also helps).
Add a small quantity of paramecium.
Allow the adult fish to spawn for 2-4 weeks.
Remove peat and wring out in a fine net until it’s the consistency of “not-so-sloppy mud”. Forget the smoking tobacco theory!
Place peat and eggs in a plastic container and store at a temperature of 21C (70F) for 6-8 weeks.
Dump peat into a tank or settled water at 23C (74F). Wait 3-4 hours and watch VAST quantities of paramecium (and other protozoans) emerge, along with plenty of FIL fry … complete with a ready-made food source.
— G.C.K.A. Newsletter, November 1997 Return to top of page
Lunar Effects on Egg Production?
“One factor that may influence egg production is the moon,” observes Bruce Stallsmith. “Many fish … produce eggs in a cycle tightly linked to phases of the moon. Usually they breed best on a new moon, when nights are dark. Some North American killifish are known to do this, for instance, Fundulus heteroclitus, the mummichog, … only breeds when the new moon causes a spring, or very high, tide in the marsh. The eggs remain attached to salt marsh grasses above the water line for four weeks until the next spring tide, when they hatch and the young fall into the water.”
There is no proof that other, inland killifish are so affected, but lunar influence may provide a possible correlation with cycles of egg production.
— G.C.K.A. Newsletter, November 1998 Return to top of page
More On Greensand
By Donna M. Recktenwalt
Variously known as green marl or Jersey Green Sand, greensand is a fine, altered dark-green sedimentary material (probably of oceanic origin, since shark’s teeth are sometimes found in it) that is mined and sold as a holistic fertilizer and soil conditioner for plants. It differs from silica sand by being extremely fine grained and extremely dense; its high iron content gives it a blue-green/black color.
Eggs are collected from greensand either by stirring the spawning material and then netting out the floating eggs, or by slowly pouring the sand through a fine sieve into a second container of water.
A number of breeders have used greensand for spawning their annual killifish. The benefits of the material are that you know exactly how many eggs are contained; and you know when the eggs should be ready to hatch.
But greensand has its downside, as well. It is a fine, powdery sand, difficult to clean and messy to work with. Even after numerous vigorous washings, it tends to cloud the water, and it continually leaches salts and minerals. Its use can also result in egg loss or damage from abrasion, and in lowered viability, possibly due to interference with fertilization or to leachates in the water.
For some time, Wright Huntley used greensand extensively for spawning Fundulopanchax occidentalis, Nothobranchius guentheri and N. korthausae. He had what he considered mixed success with the spawning medium, and encountered apparent species specific infertility. With N. guentheri the fertility rate was about 95%; with F. occidentalis 80%; and for N. korthausae it dropped to 5%. Over peat, spawning fertility for the same pair of KOR was about 80%.
Barry Cooper has also used greensand extensively to spawn Nothos, and has also observed that fertility varies. He puts eggs collected from greensand on peat for a period of observation before storage, and has seen fertility rates as high as 95% and as low as 20% when using the material. “What is difficult to compare is what the fertility is in peat,” he says. “When eggs are collected from greensand you see every egg. That’s not the case when [eggs are] spawned in peat, although you can get an indication by looking for abundance at collection time, and looking again some weeks later.”
There apparently is a cost in terms of fertility to spawning over greensand, but the causative agent is as yet unclear: it might be something in the sand itself; it might be abrasion, either at spawning time when the egg is soft and particularly vulnerable, or later during harvest. It also may be a problem of fertilization in the closely packed sand.
“My advice,” says Barry Cooper, “is not to depend exclusively on greensand for [spawning] critical species.”
— G.C.K.A. Newsletter, January 1998 Return to top of page
Focus on the Mud Spawners
Selecting the Spawning Media
By Donna M. Recktenwalt
Although the annual and semi-annual killifishesthe Nothobranchius of Africa and the Cynolebias of South America, as well as some of the Fundulopanchax species, and otherscontain some of the most beautiful and interesting members of the killifish kingdom, they are also among the most challenging to breed, given their substrate spawning behaviors and the need for their eggs to go through a sometimes extended incubation period. But the beauty and the fascinating reproductive behaviors of the annual and semi-annual fishes are reasons enough to accept the challenges of keeping them.
Of course you supply the preferred water and temperature conditions for the species, and you properly condition the breeding fish. But what do you use for a spawning medium? How do you know if you have eggs? And then what is the best way to store those eggs for the required diapause period?
Although peat is the usual substrate of choice for annual and semi-annual species, some breeders have been highly successful using other media in its placefine glass beads, greensand, or bottom mops.
Peat moss comes in several forms and varieties.
Canadian Sphagnum peat (nursery grade) may be purchased by the bale at your local garden store, usually wrapped in plastic. It may contain fine rootlets, twigs, and other solid matter, but some species seem to prefer this roughage.
Compressed peat pellets, the “gourmet” approach for smaller amounts, are more uniform and wet well. The resulting peat is quite fine from the #703 pellets, and somewhat coarser from “Gro Brix.” Be certain to purchase the pellets without additives.
Finally, there is “New Zealand” peat. This product appears more like dried java moss, and is very fibrous, with very little particulate matter.
Al Anderson, who has bred a number of “divers”, suggests running regular peat through a blender to create”super-fine peat”. Although this extremely fine material will cause clouding in the spawning tank, it’s easy to harvest the eggs by rinsing the peat through a fine net or sieve.
Whatever type of peat you use, it must first be boiled and cooled. For small amounts this may be effectively accomplished in a microwave; for larger amounts, a non-metal pan on the stove will do the job. Try to schedule the task on a day when you can have the doors open, or when the rest of the family will be elsewhere. Boiling peat has a distinctiveand to some, unpleasantodor.
There are several recommendations for storing prepared peat. One method recequires squeezing out the excess water, then storing at room temperature in a sealed glass jar or in the refrigerator in a plastic bag .For larger amounts, store peat in water in a 5-gallon pail with a small hex undergravel filter running. Wet peat in still water will rapidly turn sour.
Greensand (or “green marl”) is a fine sand/clay blend of dusts released by glacial ice melt. It may (or may not) be available from garden stores or hydroponic suppliers. Since greensand is in part fine clay silt, it easily clouds tank water, and the rough edged sand grains can cause abrasion damage. To use, place about 3/4″ in the bottom of a container. To harvest the eggs, fill another dish with clear water, set a fine sieve on top of it, and gently wash the greensand through the sieve.
Fine glass beads are used for polishing, cleaning and deburring, and sandblasting. Coated with reflective material, they are used for road signs and the reflective stripes on roads. They make a good spawning medium, since they have no rough edges and will damage neither fish nor spawn. Place the beads in a small square container and locate in the darkest corner of the tank. To harvest, stir the beads gently in a circular motion. The eggs will rise and gather near the center.
Spawning mops are a traditional killifish standby, used to both to provide cover for the fish and as a site for spawning. Although traditionally used for plant spawners, mops can be used for diving and plowering spawners as well.
Wright Huntly reports that he has successfully spawned diving species in mop-filled pots slightly taller than the length of the male fish. He notes that you may need to train the fish for a couple of weeks, using peat beneath the mops, but the fish will learn to use them. As Derek Fairbrother points out, “collecting [the eggs] is very simple. All you do is pull the container out, leaving a bit of water … then run the yarn through your hands slowly. Because most of the eggs for powers and divers aren’t very “sticky”, the eggs simply fall into the water.” For fish that like a deep bed to dive into when they spawn, try using mops or a thick layer of sphagnum moss on top of the peat. The fish will seek and spawn in the peat; the mops or sphagnum layer will help prevent spillage, and keep the fish from finding and eating their own eggs.
Whatever medium you choose, most breeders agree that confining the spawning material to a container of some sort is usually preferable to having it loose on the bottom of a tank. Plastic containers work well, but often must be weighted with marbles or small stones to keep them on the bottom. A better answer, suggests Ted Klotz, is glass containers with screw lids. These are particularly effective for media that tends to cloud the water. Select an appropriately sized jar, add the media of choice, then fill to the top with water and screw on the lid. Place the whole container in the spawning tank. When the spawning media has settled, carefully unscrew and remove the lid. To collect the spawning material, carefully screw the lid back on and then remove the jar. This procedure eliminates spillage, and the weight of the jar itself keeps it firmly on the bottom.
To reduce peat spillage, Steve Halbasch suggests putting peat into a rectangular container and partially filling it with water, then covering the entire container with a “sleeve” cut from old pantyhose . With the “sleeve” in place, gently introduce the container into the tank. An hour or so later, when the peat has settled, slide the “sleeve” partway back. The fish often will spawn beneath the covered side of the bowl. When it’s time to remove the peat, simply slide the “sleeve” back into place and carefully lift the container from the tank.
— G.C.K.A. Newsletter, June 1997 Return to top of page
Sex Ratios in Aphyosemion zygaima
There has been considerable discussion in aquatic circles about the factors that may or may not influence gender determination in developing fry.
Now some evidence from the direct experience of at least one killikeeper has tested the link between water conditions and gender in killifish.
In September 1996, Gary Elson, writing on the electronic Killifish Mailing List, wrote that he was spawning a young trio of Aphyosemion zygaima. In the spring, water conditions were pH 7.0, hardness approximately 90 ppm and 18-19C. With fry mortality of near zero, he got an even sex ratio.
During the summer, with the same fish, spawnings were larger, although egg production “was awful” above 23C. With water parameters of pH 7.4, hardness 140 ppm, and temperature 22-24C, he got almost 100% males. Gary also noted that he had a high mortality rate, so that it was possible that the larger males were killing the smaller females.
In February, Gary posted again on this subject, with the following: “Back in September, I posted a question about skewed Aphyosemion zygaima sex ratios (15-1 males). I was hatching and rearing my fish at pH 7.4, 120-140 ppm. Following the excellent debate, which led to a lot of fascinating information about post hatch sex determination, I started raising some batches in pH 6.5, 30 ppm filtered rainwater mixed with tapwater.
“The results are in! 9 to 1 females!”
— GCKA Newsletter, April 1997 Return to top of page
Skewed Sex Ratios – Not Just an Interesting Question
By Donna M. Recktenwalt
After extensive effort, you’ve finally succeeded in spawning and hatching out fry from that “special fish.” You’ve even successfully raised a number of them past the wriggler stage and into juveniles. You’re expecting the more or less normal sex ratio of 50/50.
But when they sex out you have almost all males – or all females.
What happened?
Such occurrences in a hobbyist’s tanks are disappointing and discouraging. But in the commercial aquaculture industry they pose a real problem with major economic effects. This has lead to research, and results that seem to indicate that in most freshwater fish, the combination of lower temperature and higher pH normally results in the production of more females than males.
Other evidence, both anecdotal and from research, seem to indicate that developing fry may be influenced by a number of outside factors – the age of the spawning adults, the temperature, pH, and DH of the water; light levels; and the subtle mix of chemicals in the water, resulting from the presence of other fish, of rotting vegetation, and the like.
Working with the Atlantic silverside (Menida menida), B.E. Kynard noted “The sex ratio in fishes that normally have separate sexes can be influenced by the environment.” He noted that sex determination of the fry was under the control of both genotype and temperature during a specific period of larval development. In addition, the spawn from different females varied in their responsiveness to the variable (temperature).
Uwe Romer, working with Apistogramma species, found that keeping fry at higher temperatures usually resulted in the production of more males, and that higher pH values resulted in more females. His research indicates that the best sex ratio distributions for the study species occurred when water temperatures were maintained at about 78°F (26°C), with pH levels about 6.8, during the first month of life.
Friedrich Bitter, speaking at Killie Revue ’96, observed that among the Rivulus species, the gender of fry is determined not at conception, but some time during the first few days or weeks of life. The most balanced ratios of male to female seemed to result when the fry were hatched and raised in water from the parents’ tank.
The above findings may or may not be valid for all killifish.
However, the next time you find yourself with a badly skewed sex ratio among a batch of developing fry, it might be worthwhile to try adjusting the pH or temperature level in subsequent hatches to see if that has any positive effect on the outcome.
— G.C.K.A. Newsletter, February 1997 Return to top of page
Skewed Sex Ratios
By Donna M. Recktenwalt
It’s a problem that many killikeepers encounter from time to time: badly skewed sex ratios in a batch of fry.
What causes these apparent shifts from the norm? And more importantly, what can we do about them?
Even the experts admit that there is much about fish embryology and development that they do not know, including the time that gender is determined. “Are killie eggs ‘differentiated’ as to gender when they are laid?” asks one hobbyist. Is the spawning water a factor? Are fertilization rates higher under certain conditions?
“Apparently … [the determination of sex] can happen at various times in the [development] process, depending on environmental conditions,” says Brian Watters. Environmental conditions, including population density and water temperature, pH, and hardness may all have their effects.
In Apistogramma species, studies have linked sex ratios to temperature, especially during the first month. Cynolebias may react similarly. It appears that pH matters more to fish from narrow temperature ranges, such as rainforest Aphyosemions.
“I don’t think that the answer to skewed sex ratios lies in a single simple factor such as pH (or temperature),” says Brian Watters. “While I have … not conducted controlled experiments, … one observation that I have been able to make consistently … is that if I raise a large batch of Nothos (or South American annuals, for that matter) under fairly crowded conditions, I invariably get a large dominance of males. In contrast, smaller batches raised under less cramped conditions usually (but not always) produce more even sex ratios.”
Lee Harper got contrasting sex ratios in different Diapteron species under similar conditions. Lee uses unaltered tap water with a pH that drifts naturally toward the acid in his tanks. Makeup water is at pH 7.4, but the tanks tend to be 6.0 to 6.5. “For Rivulus xiphidius … I lowered the hardness by about 1/2 by the addition of rainwater. This … produced more females than males.”
Foreign breeders, too, have observed the phenomena of skewed sex ratios. In “Soderjanie i razvedenie aquariumnih rib” (Russian, 1991) A. S. Polonskii wrote: “The temperature and chemical composition of the water can affect sex ratio in the fry of Cyprinodontidae. For example, at 22-25°C most of the fry will be females; if the temperature is not constant, most of the fry will be males. At the same pH =6.0, Epiplatys dageti will have more female offspring (>90%) in the soft water (dH about 5 degrees); in the hard water (dH 24) about 90% of the fry will be males. Aphyosemion gabunense in the acidic water (pH 5.0) will give more females, at higher pH=6.5 more males.”
“There has [also] been some recent work looking at various chemicals that mimic female hormones,” says Andrew Broome. These have been shown to cause sex ratio skewing in salmonids and may even be responsible for a decreased sperm count in humans. “The males that do appear are increasingly ‘female-like,’ [with] some even producing proteins that are normally only associated with egg production.” The suspect chemicals apparently come from a variety of sources and seem effective at very low concentrations.
Others, too, have linked skewed sex ratios to pH. Supposedly Dr. Joanne Norton did some experiments that showed that platies’ sex was a function of pH.
But sometimes there is no apparent cause for what we see in our tanks.
“I have … two maturing colonies of Aphyosemion (Diapteron) abacinum and georgiae,” Lee Harper wrote a year ago. “Each is about 30 or so members. The abacinum are almost all males; the georgiae all look like females at the same age and size. They were hatched and raised under close to identical conditions-water, temperature, etc. In this case it seems to be genetically controlled. The parents in both cases were young pairs.”
“Certain species of Rivulus … [regularly] throw predominantly one sex,” reports John Boylan, who got skewed sex ratios ranging from 4:1 (male to female), to as much as 20:1 among the various species he keeps. R. agilae showed a 10:1 ratio at room temperature; which reversed to 1:10 when the eggs and fry (until sexable) were kept at 75°F (24°C) or higher. R. sp. Isle of Pines showed a consistent 4:1 ratio unaffected by changing water parameters, from soft/acidic to hard/alkaline at a constant temperature. “Interestingly, some of my other Rivulus species breed with even sex ratios.”
What to do if faced with a skewed sex ratio in a batch of fry? Generally there’s nothing you can do. But on the next batch, try changing the pH of the water; raising or lowering the water temperature; changing the water more often; or giving the fry more room.
There’s no guarantee that any of these will work for you, but it might be worth a try.
Additional Killie-related Reading:
Bitter, F. JAKA 22(3): 96-102 (May 1989)
Boylan, John, “Sex Ratios with Rivulus,” AquaTropica, Vol. 1, No. 4.
Chroke, Sharno. JAKA 17(6): 232-233 (Nov 1984)
De Boer, R. JAKA 23(5): 155-157 (Sep 1990)
Morenski, R. JAKA 17(1): 10-12 (Jan 1984)
Wilson, Mike. JAKA 18(1): 31-39 (Jan 1985)
Tropical Fish Hobbyist, January 1997.
Additional Non-killie Reading:
Conover, D.O. & B.E. Kynard, 1981. “Environmental Sex Determination: Interaction between Temperature &
Genotype in a Fish.” Science 213: 577-579.
Conover, D.O. and D.A. Van Voorhees, 1990. “Evolution of a Balanced Sex Ratio by Frequency-dependent
Selection in Fish.” Science 250: 1556-1558 (Dec 14, 1990).
Rubin, D.A., 1985. “Effect of pH on Sex Ratio in Cichlids and a Poeciliid (Teleostei).” Copeia 233-235, 1985.
Marx, Jean, 1995. “Tracing how the Sexes Develop.” Science 269 (29 Sep 1995) 1822-1824. (Fruit flies and
nematodes)
Sullivan, J.A. & R.J. Schultz, 1986. “Genetic and Environmental Basis of Variable Sex Ratios in Laboratory Strains
of Poeciliopsis lucida. ” Evolution 40, 152-8.
— G.C.K.A. Newsletter, December 1998 Return to top of page
Some Views On … Skewed Sex Ratios in Killifish
Imbalanced sex ratios in killifish fry are nothing new; hobbyists have long been encountering this problem in various species.
"In humans sex is determined by specific sex chromosomes, an x for female and a y for male," Allen Johnson reminds us. For reptiles gender is largely determined by environmental conditions (pH, temperature, various hormonal exposure during egg development). In fish, sex may be at least partially environmentally determined, with developing eggs and fry affected by pH, chemical, or temperature cues. Specific application of temperature and pressure on developing Tilapia eggs nearly always guarantees males for the aquaculture industry.
But for killifish, exactly when is sex determined? At the time of fertilization? During embryonic development? Post hatching, but before maturation?
"I personally believe temperature or pH are the predominant factors in sex ratio," Allen says. However, "in my experience ... females are most active in laying eggs immediately after being fed. ... It may be that eggs laid in the morning will be exposed to a longer light period and may develop into one sex, while those laid late in the day may develop into the other."
The age of parents may also be a factor on the sex ratio of the fry, observes Joe Weber. "In a number of species (in the Aphyosemion genus) I seem to get more even ratios as time passes" and the breeders age.
"Water temperature, pH, and hardness will all change the sex ratios of several species I keep," observes John Wubbolt. "For example, I raise Epiplatys annulatus. If I raise the fry in soft acidic water I get a majority of males. If I harden the water so that it's slightly alkaline and moderately hard, I will get more females. Same goes for Fundulopanchax gardneri N'sukka and Udi, and for Epiplatys dageti."
"In my trials with Aphyosemion zygaima," Gary Elson says, the results "boiled down to ... alkaline hardish (140ppm) water produced males ... while soft (but not necessarily acidic) water gave a closer to even sex ratio.... My A. ogoense 80/24 also give more females in tap/rainwater mixes than in straight tap water."
Those who have raised some of the Rivulus or Pterolebias species know of the "two per container" technique. Raised together, fry from some of these species will be heavily gender biased. Raised for the first few days/weeks of their lives two or three to a small container, then put back together for growout, many hatches are more evenly balanced. This may be due to chemical signals which cause the fixing of gender at some point, or there may be unobserved predation by the slightly stronger fry.
So there may be several things to try if you're getting hatches that are heavily of one gender. Change the pH or the temperature of the water in the breeding tank and/or the hatching and growout containers. Try different pairings of breeders. Or take the extra effort to raise the newly hatched fry for the first few weeks in pairs. You may find an answer that works for you. – G.C.K.A. Newsletter, March 2001 Return to top of page
Storing Eggs
By Ken Harsh
This article first appeared in the GCKA Newsletter in March 1986.
Consider the number of ways that eggs can be stored before they are hatched. If you are like me, then they are stored in petri dishes, with water from the breeding tank, along with some acriflavine (enough to make a light yellow color). I add a drop of acriflavine to another cup or dish, and then add a drop or two from that to the petri dish. I store the eggs in a dark place, away from the light. This seems to work best for two-week top spawners.
For the 30 day bottom spawners I either place the eggs in water, as in the above manner, and incubate for 30-35 days, until they are eyed up very well. I then add a pinch of microworms, and let them hatch out. Alternatively, I place peat in a petri dish and place the eggs on top of the very moist peat. When the time is up, I either pick the eggs off the peat and hatch them out, or hatch the whole mess.
I have also been known to incubate whole mops in a tank with some acriflavine for sensitive species, and to remove the parents. With some touchy kinds I incubate the eggs in partial distilled water, or with a pinch of peat in the water. Some eggs will not hatch no matter what I seem to do for them, so I pass them along to someone else.
Besides petri dishes, I have also seen butter dishes, Tupperware, and other items used for egg storage. Some people have used the small plastic tool cabinets, the kinds with the little drawers. Well, I thought those were kind of neat a year or two ago. I placed the only seven (7) eggs I ever got from a pair of Aphyosemion aureum into one of the drawers. It was drier than usual in the fishroom, and you guessed it, when I looked at the eggs a week later they had dried up. Sorry, I missed it again.
Many people store bottom spawner eggs in moist peat in plastic bags until they are ready to hatch. Others use green sand. I will not comment on either, since I do very poorly with annual type fish and stick pretty much to top spawners and 30 day bottom spawners.
I buy plastic petri dishes from wholesale supply houses. The price is … very reasonable [but] you usually have to buy four dozen or so dishes at a time…. I used to use glass petri dishes, but they were a pain to wash, rinse, and sterilize, and I was always dropping them, so in my case the plastic dishes are the solution.
I do not put more than 12-20 eggs in a petri dish for water incubation (top spawners). It seems if I get too many in a dish then the percentage of eggs that fungus is higher.
Sometimes some batches of eggs seem to be bad, and you get a terrible percentage hatch, while at other times the rate is very good.
Remember to take the eggs when you get them; some fish produce eggs on an irregular schedule.
— G.C.K.A. Newsletter, Sept/Oct 1998 Return to top of page
Vacation Care – A Trick for Fry
We all know the concerns of going away and leaving our fish: what if some emergency happens while I'm gone? Will they be all right without being fed?
Most experts agree that adult fish can manage quite well for up to several weeks without being fed, as long as their water and temperature conditions remain within acceptable ranges. But what about fry?
"Unless your tanks are completely sterile, they're full of living creatures that the fish can graze on," says Oleg Kiselev. "Java moss is a veritable treasure trove of rotifers, protozoans, infusoria, and so on. Fish can live like this for quite a while."
Lee Harper has found an alternative that works well for him. "For smaller containers (shoe box type) which contain fry of various sizes, I do the following," he says.
1. Before leaving, feed live foods that will continue to live in the water (daphnia, microworms, etc.)
2. Add a piece of lettuce leaf and some pond snails in each container, plus the usual Java moss.
3. Don't put all your fry (of a species) in the same container; divide them among several. That way, if one container goes bad, you won't lose them all.
"The lettuce leaf procedure works so well that I keep reminding myself that I should do it when I'm not away, as an alternative to daily feedings and semi-daily water changes," says Lee.
Wright Huntley offers some additional tricks for leaving a few babies for a couple of weeks. Float a few mosquito egg rafts on the water. Or put the fry in an oversized tank with lots of Java moss and a strong culture of daphnia, infusoria, etc. A few small snails will help keep the culture going. But "moving the babies to plastic shoe- or sweater boxes and leaving them with an experienced aquarist who is routinely feeding baby brine shrimp is the best way."- G.C.K.A. Newsletter, February 1999 Return to top of page
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