The Making of a Blackwater Tank

December 29, 2010


After having unsuccessfully attempted to extensively breed Sailfin tetras for over a year, I got annoyed, and felt that it was time to get serious. Inspired by setups used to breed amazonian catfish, I decided to set up a blackwater tank, which I’d then subject to seasonal variations in conductivity, pH, temperature, food accessibility and flow.

The idea was that imitating the seasonal floods of the forests from which the tetras came, would trigger spawning.


* 30 liter (about 8 gallon) tank

* A block of unprocessed, low-humic (not degraded), peat.

* Access to a water distillery (a reverse osmosis, a.k.a. RO, unit will do).

* Plants (mainly java moss, Pistia and Salvinia), oak leaves, and a hollow piece of driftwood.

* Air pump, heater, and a small circulation filter (360 liters per hour).

* Lighting: 11 watt compact fluorescent.


Figure 1. First I cut a 4 cm (1.5″) thick slice out of the peat block.


Figure 2. The peat was “low humic”, meaning the peat moss it is made of is not severely degraded, but very fibrous, and that pieces of the plant are still clearly identifiable. I chose this because I hoped it would hold together better in water than fine-particle high-humic peat – and it did!


Figure 3. The tank. An old battered 30 liter breeder. The slice of peat has been pressed into place, and water added. The peat holds together so well that even fresh after filling water there’s not much debris in the water. Most of what there was floated and could easily be scooped out.


Figure 6. Finished result. The layer of leaves turned out to be highly appreciated by the sailfins.



As it turned out, the tank functioned extremely well. The pH varied between 4.5 – 5.8, and I did get the sailfins to breed. The low pH meant the leaves degraded extremely slowly. The plants grew like crazy, especially the Javamoss, which almost took over the tank. The peat still held together perfectly, even after a year, and still colored the water, but no longer appreciably affected pH (which towards the end had drifted over 6.5).

But all wasn’t great. Downsides of using peat was that the water was very strongly colored, at its peak we’re talking more like coffee than tea! Also “dust” from the peat would clog the filter requiring frequent cleaning, and the lower parts of the peat turned anoxic, so when I removed it… well, let’s just say it didn’t smell like roses. For more details on how this tank operated, see my Breeding Sailfin Tetras article.

For a breeding tank intended to operate for a year or so, I think this set up worked very well indeed, but I’m not sure I’d use this approach in a more permanent aquarium.


ADDENDUM: Ron Last has used a very similar set-up and had it running for four years, and mailed this extremely interesting (Swedish language) link to me, where he details his experiences:




Hatching Artemia the easy way.

November 14, 2009

1) Take an empty plastic ice-cream box or similar. Cut the lid in half. Press one half down into the box, approximately at the middle of the box, so the box is divided into two halves. It should be a tight fit to the walls of the box, but a few millimeters free between the bottom and the edge of the lid. You’ll probably need a few pieces of tape to hold the lid in place.

2) Dissolve one heaping tablespoon (about 20 grams) of table salt, with or without iodide, in one liter of tap water, then pour the salted water into the ice-cream box until it’s about 1/2 – 2/3 full.

3) Put about one milliliter (about 1/4 teaspoon – don’t use too much, 1/4 teaspoon is plenty!) of Artemia-eggs on one side of the “barrier”, and place the container where it’s light (improves hatching rate) and the temperature is between 20 – 30° Celsius (25 – 27°C is ideal).

4) At 25°C the eggs hatch after about 24 hours. You’ll be able to see the Artemia like little brownish, jumping, “dots”. If you see no hatched Artemia after 24 hours, don’t worry! Some strains take a bit longer to hatch.

5) Artemia are attracted to light, so light a lamp by one corner of the ice cream box, on the other side of the barrier from where the eggs are.

6) The Artemia will now swim towards the light, under the barrier, and congregate in a dense swarm closest to the light.

7) You can now easily siphon out the artemia into an artemia sieve or reusable nylon coffee filter, while the empty eggshells remain on the other side of the barrier. What you see in the sieve below is about 1/10th of the total amount of Artemia I got this batch.

8) Wash the Artemia (not strictly necessary, it’s to get rid of salt & bacteria) by gently rinsing them, still in the sieve/filter, with tap water. If you use a flat-bottomed sieve (like in the picture) it helps to tilt the sieve a little, to concentrate the Artemia to one corner of the sieve.

8) Done! All you have to do now is flush the Artemia from the sieve/filter down to your fish!

Repeat as necessary!

The Artemia should be used as soon as possible after hatching, as it quickly loses nutritional value. As all eggs hatch within a day or two, you’ll need to start anew, with fresh water and eggs, every two days. By having two hatching boxes running with one days interval, you can have constant access to freshly hatched Artemia.

Breeding Yamato (Amano) shrimp

November 11, 2009

Breeding amano shrimp (Caridina multidentata)

This is the shortened version of my overly long three-hour epic “Breeding Amano Shrimp”, originally published in 2005.


Obviously, you must have both males and females to breed Amano-shrimp. While there is a definite difference in size (males are much smaller) and body shape (females have broader tails), there is a much easier way of telling the sexes apart: you look at the second row of spots along the side of the shrimp. Females have elongate spots, more a broken line than individual spots really, while males have round spots.

Once you have determined you have males and females, you just have to make sure the animals are well-fed, and wait. Soon the females gonads will start swelling, looking like a yellowish-greenish ‘filling’ dorsally in the females – up to one third of the females volume will eventually be occupied by developing eggs!

As the female is getting ready to breed, she releases pheromones into the water, and the males become frantic; aquarists have named this phase the “shrimp race”. Prior to mating the males “swarm” around, much like moths around a lamp, trying to copulate with everything which moves, including each other and fish!

Copulation normally takes place late in the evening, with the male first swimming around, then landing on top of, the female. If she does not fight him off, he then climb in under her, belly to belly, to deposit the sperm. The whole procedure is over in a matter of seconds. If the female is not ready, or if there are many males trying to mate with her, fights may break out. With many males present, one may even get “communal” spawnings, with one female surrounded by numerous of males.

A couple of days after copulation, the female lays the eggs, gluing them onto her abdominal swimmerets. Literature suggests the female may lay as many as 2000 eggs. When freshly laid, the eggs are a dark moss-green color, but become progressively lighter and more khaki in color as they mature. The female carries the eggs for about 5 weeks before they hatch. The color of the eggs can be used as a rough guide to when they’re going to hatch, but a better estimate can be obtained by looking closely at the eggs (e.g. with a hand lens, or, as I do, via a macro photo): if you can see the eyes of the developing embryos through the eggshell, then the eggs will hatch within a few days.

When the eggs are close to hatching, the eyes of the embryos are visible through the eggshell.

I’d recommend you remove the female when hatching draws close and place her in a jar or small breeding tank, where the eggs may hatch in safety. The larvae are positively phototactic (swim towards light) and can be collected by shining a flashlight into the aquarium at night, and you can siphon out the larvae as they are drawn to the light.

The eggs do not all hatch at once. In my experience, most hatch during the night, and the remainder throughout the next day.

You should return the female, who does not eat her young, to the main aquarium as soon as the eggs have hatched. Within days of releasing the larvae, the female will have mated, moulted, and be carrying a new batch of eggs. Mature females are always producing eggs, and nearly always carrying some as well.

Note: do NOT place the female in brackish water! While I found out that adults survive quite high salinities, the eggs fail to hatch if the water is brackish – I lost two batches this way.


The newly hatched young are whitish and exceedingly small, approximately 1.8 mm long. The larvae are planktonic, positively phototactic, and hatch as early Mysid stage larvae. For the first weeks they swim in a curious vertical, head-down, position, after which they adopt a more horizontal swimming position.

In nature, adult Amano-shrimp live in mountain streams and the larvae are washed out into the sea where they feed on marine plankton and grow. After metamorphosis they migrate back up into the streams. We must mimic this cycle in order to breed the shrimp. Therefore the larvae have to be transferred to salt water as soon as possible, at the latest at the 8th day after hatching, because after that they become unable to live in fresh water. There is no need to gradually increase salinity, the larvae have no problem being unceremoniously dumped straight into salt water. To make the salt water, I would suggest using either filtered natural seawater, or a quality commercial salt mix intended for coral reef aquaria, e.g. Instant Ocean, which should be aerated vigorously prior to use.


There is some controversy surrounding what the optimal salinity is for the developing larvae. Hayashi & Hamano report total failure to survive in salinities up to and including 8.5 ppt; optimal survival at 17 ppt (80% survival rate); and suboptimal survival (11%) at salinities up to 35 ppt (full marine salinity). This is quite different from what I found, with zero survival at salinities lower than 25 ppt, suboptimal survival at 25 ppt (3 larvae, out of at least 200, survived for four weeks without metamorphosing at which point the attempt was aborted), and high survival at 30-35 ppt (an estimated 80% reaching postlarval stage). Other people who’ve experimented with salinities have come to the same conclusion: any salinity below 30 ppt or over 35 ppt will result in heavy mysid-stage larval losses!

However, after metamorphosis the postlarval shrimp no longer tolerate full-strength seawater and should be moved to fresh water. Alternatively losses might be avoided by lowering salinity to 17 ppt when the first larvae start to metamorphose.

I’ve used an airstone with reduced flow for circulation. There may be problems with flotation of larvae if aeration is too vigorous.


The larvae require fine-particulate food for the first weeks, after which they are capable of accepting larger food particles. Judging by published reports, the larvae can subsist on pretty much any food small enough – successful rearing have been reported using e.g. fine-crushed Spirulina-flake, and dry yeast.

I fed mine about 5 times per day, initially with small amounts of brewers yeast, Baby Star II (50-100 microns microencapsulated feed for egg-laying freshwater fish), and after a week also Golden Pearls (microencapsulated food for marine invertebrate larvae, 50-100microns). There was also quite a lot of diatom algae growing in the tank, you can see it as brownish “dust” in the photos, and I’ve since learned that diatoms is an important food for shrimp larvae.

Seriously, I think access to diatoms is absolutely necessary. Hayashi & Hamano report that larvae did not begin to feed until the fourth day (first ecdysis), after which they successfully raised their larvae on a diet of “adhesive diatoms, Cymbella and Navicula, or a grated artificial diet, mixed with a small amount of rice bran”, and note that the larvae were capable of foraging on the tank walls and bottom. Hayashi & Hamano failed to raise larvae on a diet of just rice bran, or a diet of Chlorella phytoplankton + rotifers.

Diatoms. You need them. Remember I told you so.

Once they could eat it the golden pearls was a good feed – the larvae eagerly accept it, grow fast, and the strong color of the feed makes it easy to see that the larvae are eating (the guts turn brownish red, and eventually the whole larva turns orange).

The larvae will both catch food drifting in the water, and browse on surfaces.

I tried feeding the shrimp freshly hatched Artemia, but although I could see them catch & eat some artemia, most of the artemias quickly grew too big for the shrimp to eat, and actually started reproducing in the rearing tank. As the Artemia are clearly competing with the larvae for food, I’d advice against feeding Artemia to Amano-larvae.


My shrimp larvae started metamorphosing to postlarvae at day 30 to 60. The settled postlarvae are about 8 mm long. My experiences suggest they are capable of being transferred to fresh or brackish water immediately after metamorphosis, and will die within a few days if left in full marine salinity.


The larvae do not metamorphose into adult form all at the same time. It will take several weeks until the last has metamorphosed.  An interesting observation: Whenever I change water in the aquarium, the postlarvae start swimming around like crazy, and continue doing so for a few hours. Presumably the water change triggers their migratory instincts.

At day 127 after hatching, the largest of my shrimp were about as big as the smallest Amanos one see in shops, i.e. about 25 mm, and ready to sell. With heavier feeding, and a larger grow-out tank, the shrimp would have reached marketable size faster.


An amazingly good page with information on the keeping and breeding of this shrimp:

Another very good report on a successful breeding:


Hayashi, K-I. and T. Hamano. 1984. The complete larval development of Caridina japonica De Man (Decapoda, Caridea, Atyidae) reared in the laboratory. Zoological Science 1:571–589. CSA