Desiccation effects in A. bellottii

VanDoorenVarelaFig1Another manuscript out in the open:

“Desiccation plasticity and diapause in the Argentinian pearlfish Austrolebias bellottii”

We (Irma Varela Lasheras and myself) did detailed demographic work on A. bellottii embryos and found that these respond relatively little to being incubated in either water or air with high humidity. The eggs were incubated in multiwell plates, and this allows a very easy follow-up of the embryos, and a good yield of developed embryos. I am happy with the results and they take me all the way back to the start of my PhD thesis, when I read Prof. Michio Hori‘s thesis on Cicindela japonica, in which he did a demographic study of three stages of tiger beetle larvae with graphs similar to the one I show in this post.



Evolutionary rates AustrolebiasThis week I posted a manuscript on bioRxiv. Here is the link to it:

“A case for sympatric speciation by cannibalism in South-American annual killifish (Austrolebias)”

We demonstrate that large Austrolebias evolved at least three times from small ones. For one case, we argue that piscivory evolved starting from the evolution of cannibalism on conspecifics.





Floral foam

A. wolterstorffi floral foamA new incubation trick: I cut slices from a block of floral foam and push small depressions in them, the size of a coin and about 1 cm deep. These pits are filled with cocopeat, from the container where my killies have been laying eggs. I drip some rainwater on the foam to keep it wet, and I also cut a thin lid. Eat pit gets four fresh eggs on top of the cocopeat, to be incubated. Then the wafer goes into a 250ml plastic container for storage and to avoid desiccation. Here some A. wolterstorffi eggs after one month of storage. The white patch is fungus: I lost one egg out of the four in this pit. As you might notice, the other three contain well-developing embryos. The scratches around the pit are from stabilizing my usb microscope on the foam.

A. wolterstorffi embryoThis afternoon I checked a small sample of A. wolterstorffi (Velasquez) eggs, which I photograph regularly. In a few embryos, the developing heads and eyes can be seen. In this egg, the two darkers dots above the yolk droplet are the eyes. USB-camera, near IR light. I also made a short video, where you can see that there is a heartbeat.

Photographing eggs

Yesterday I colllected Austrolebias wolterstorffi (Velasquez) eggs. I just decided to make some pictures of them, using a DinoLite USB microscope with UV (395nm) and near IR (940nm) leds. Here are the results on a fresh egg (well maybe not from the last few days) that I just put straight from the peat onto a piece of paper on my desk. The result is simple to achieve and really encouraging. So I will try to get pictures of all main developmental stages with this lighting. Left: UV; Right: IR.

wolterstorffi egg (UV)wolterstorffi egg (IR)




1000 islands

Filter plant islandTo keep tanks clean(er), I decided to use plants that grow well and root well in water as filters. Here on the photograph three different plants on small floating islands in my fish basins, situated in an unheated greenhouse. They all grow through winter. Leftmost: Vietnamese coriander (Persicaria odorata), middle: Japanese parsley (Oenanthe javanica), right: watercress (Nasturtium officinale). What’s really great is that the plants need to be kept growing to make them take up nutrients. What helps well for that is harvesting them, and all three are edible. Aquaponics in its simplest form.

Egg drop out

A. cheradophilus (La Paloma 2016)In 2005 I received eggs of A. elongatus “Ezeiza” incubated inĀ  Sphagnum magellanicum and eggs of A. vandenbergi “Talon Cansado” in the same material last year. I decided to give this spawning material some extra attention and offered it to fish as spawning material. For one of the samples collected like this and currently seven months old, I tried to determine the state the embryos were in. To my surprise, that was very easy! After some drying, eggs started rolling out of the moss. For example, the one on the photograph, which is A. cheradophilus “La Paloma”. Will definitely experiment with this spawning substrate further!

0.5 L Culture

Jar culture (2016)I’m experimenting with cultures that I can basically pour into the fish tank entirely, once “mature”. The culture I mean. For now I’m trying things in 0.5 glass jars at home on a shelve with moderate daylight. I’ve added tap water, a few Daphnia and small detritus worms that I scooped from a rainwater drum in my garden. On the bottom there are some specks of cocopeat and I fed the system a few times with dried yeast. So far, the Daphnia multiply, the worms too, the yeast seems to persist. Things go well. One jar had a solution with some drops of coffee creamer standing in it for a week before the water was replaced and the other animals were added. The cream had created a film on the glass in which the worms indulge. Next challenge: Massively increase the surface where the film can settle which hopefully increases the worm fraction. The nice thing about that is that I won’t pour the surface covered with film into the fish tanks with the feed. On the photo: few of the worms. If you magnify them a bit, you can see their chaetae.

Fry outside

The last few weeks I hatched fry from several species. In some cases, there are lots. Due to space limitations, I decided to put a few groups outdoors. Before that, they were fed twice with Artemia. Then they were put in 70 liter tanks with a stock of invertebrates that I fished and sieved from a ditch. Things seems to go great! There are some casualties of course, as the photo shows, but overall everything goes well and the fry even seem to grow faster than ever. The fry are from Austrolebias prognathus “Salamanca”, and charrua “Ruta1316”. If all goes well, I might do this more systematically.

Hatching on a sinus (2)

More on the experiment done in the ECOLAB. The results indicate that hatching does not depend in a simple and obvious manner on the temperature pattern. What the data do suggest is that the proportion of alevins which swim well and which are still alive after two weeks is largest, when water is added at the lowest temperature in the cycle, so that it increases steadily for 12 hours after wetting.