Research round-up

Unfortunately there has been little activity on the blog of late, mainly due to the small matter of getting my PhD thesis handed in, submitting manuscripts to journals, and finding a job etc! Having said that, I have been somewhat busy in other parts of the Web. Boopboops now has a sister Twitter feed for science related things (@boopsboops), and I have now coded up a Website promoting my CV, publications, and research skills etc, etc.

So, in absence of anything better, and as I've been meaning to do for a while, I thought I'd write about my favourite fish papers of 2009, 2010, and 2011.

(1) Larmuseau et al. (2009) To see in different seas: spatial variation in the rhodopsin gene of the sand goby (Pomatoschistus minutus). Molecular Ecology 10.1111/j.1365-294X.2009.04331.x

I like the idea of looking at how organisms adapt to their surroundings. This study compared variation in the rhodopsin visual pigment locus with phylogeographic patterns in "neutral" mitochondrial and microsatellite markers (i.e. likely to detect any population-genetic structure), and found that in the sand goby, the two were discordant. Variation in the rhodopsin gene (RHO/RH1/RHOD) was partitioned differently and corresponded to photic environment (light penetration, water turbidity etc). There were also signs of positive selection at sites coding for amino acid changes relevant to spectral adaptation.

It's also interesting to note that rhodopsin is a commonly used marker for phylogenetic studies, which is probably due to early studies on vertebrate visual systems providing easy to use primer sets. However, I would be cautious about its use now, as these apparent convergences due to environmental conditions may not give a good indication of common ancestry for a species tree!

(2) Lavoué et al. (2011) Remarkable morphological stasis in an extant vertebrate despite tens of millions of years of divergence. Proceedings of the Royal Society B 10.1098/rspb.2010.1639 

If you've ever kept a tropical aquarium, you may have seen the African butterfly fish (Pantodon buchholzi) lurking in the oddball tanks. They're indeed a strange fish and are great fun to keep, clinging to the surface and greedily snapping up any insects that you feed them. Pantodon buchholzi is the species in a monotypic genus and family, known from the Niger and Congo basins.

When their mitochondrial genomes were sequenced, the researchers estimated that the two isolated populations had diverged over 50 million years ago, despite looking almost identical in terms of shape and meristics!

Evolution is taking place on the DNA clearly, but not on the external anatomy it seems. The reasons as to why and how this has happened are fascinating. The authors state "Proposed mechanisms of morphological stasis include stabilizing selection, ecological niche conservatism and genetic and developmental constraints". I look forward to further studies on this.

(3) Mims et al. (2010) Geography disentangles introgression from ancestral polymorphism in Lake Malawi cichlids. Molecular Ecology 10.1111/j.1365-294X.2010.04529.x

The cichlid flocks of the African Rift Lakes are an almost extreme opposite example to the one presented above. There is huge phenotypic diversity, but often very little in the way of molecular differences. The mbuna cichlids Labeotropheus fuelleborni and Metriaclima zebra, are quite different in appearance, but share mitochondrial DNA haplotypes typical of very recently diverged, or hybridising species. The authors also report "greater mtDNA differentiation among localities than between species".

Information from the nuclear genome can help in these situations of understanding levels of gene flow, but can have limited resolving power when not used in sufficient number. Enter NGS. Modern sequencing methods can now provide orders of magnitude more data, and with a large SNP (single nucleotide polymorphism) set, here the authors report that the two species are indeed genetically distinct, and that recent hybridisation among the two species is unlikely. Certainly a useful tool for exploring these questions further.

A method of photographing and preserving fishes for molecular studies

Voucher specimens are important in molecular studies, almost maybe as important as for morphological studies. A good voucher will be useful to both molecular and morphological research for many years to come; a good voucher will allow any misidentified specimens to be easily corrected, and will permit any interesting molecular results to be effectively corroborated with morphology.

But generating good vouchers in molecular studies is hard. Formalin, the fixative chemical of choice for ichthyologists, degrades DNA and makes extraction/PCR difficult (but see Zhang, 2010). Instead, ethanol can be used as a fixative, but ethanol fixed specimens are often brittle, faded, and of poorer long-term quality.

It's often best to take a tissue sample from your specimen, store this in ethanol, and formalin fix the rest of the fish as a voucher. This is fine, but you'll want to know which tissue sample comes from which specimen, and for small fishes it's not possible to permanently attach the label to the specimen without causing damage. Of course, you could put them all in individual jars, but you could soon run out of jars or space. Transporting them is a big problem too, and this is where you really need to save space.

So, after trying out some quite unsatisfactory methods, and ruining many good specimens, I have developed a nice method of generating quality molecular vouchers:

Step 1. Fill vials for tissue samples with high-grade 100% ethanol. Label the tubes internally with pencil on archive quality "goatskin" paper, and externally with marker pen. The vouchers can be kept separate using small polythene zip-seal bags. They need to be perforated first, however, with a paper hole punch (do several at a time). They should also have their bottom corners cut off to allow the bags to drain. Place another label in the bag.

Step 2. Get everything ready in advance. Here I have:

• Latex gloves
• 10% formalin (clearly labelled)
• MS-222 (fish anaesthetic)
• Spirit burner to decontaminate tools
• Variety of forceps and scalpel
• Pencil
• Squares of cardboard to use as a clean surface for tissue preparation. 
• Vials for tissue samples
• Bags for voucher

Step 3. Assemble your light source and photo rig. Here I use an adjustable microscopy light (halogen desk lamps can be substituted) and a shallow white tray. I used a piece of folded graph paper as a scale for these photos. Now, mix up your MS-222 (overdosed) and water into a shallow clear tray (the lid of a tube rack), and the fish can now be added (wait for 10 mins to ensure death). Make sure the fish is only just covered.

Step 4. Adjust the light angle and photograph the left-hand side of the fish, always adding the label. Remember to set your camera's white balance correctly (usually using the custom mode). The picture can then be cropped and the file name changed.

Step 5. Take the fish out of the solution and place on the card sheet. Use the scalpel to carefully excise a tissue sample from the right-hand side of the fish. Pectoral fin clips can also be taken to cause less damage, but on small fishes this won't yield much tissue, and using mitochondrion rich muscle may reduce the likelihood of numts (Song et al., 2008).

Note: don't cut from the caudal peduncle area if characters such as caudal peduncle scale counts may be important for identifying your fish.

Step 6. Next, place the fish into the plastic bag with the forceps, and place into the formalin. The position of the fish and fins can be manipulated through the holes in the bag with the forceps. This ensures the fish is not bent and the fins are not folded down.

Step 7. Throw away the card sheet and replace with new. Clean the implements with a wet tissue and then sterilise with the spirit burner. Repeat process for rest of specimens.

Step 8. Leave vouchers in formalin for approximately three days (longer for larger fishes). After three days, remove from formalin and wash thoroughly with water. Leave in water for 24 hours to dilute remaining formalin. Place into weak 35% alcohol (ethanol or clear methylated spirit) solution for three days before final storage in 70% alcohol. The voucher will have lost a lot of its colour by now, but can be photographed again to document the preserved colour pattern.

Of course, these bags have not been tested for long-term (i.e. indefinite) storage, and are only recommended as a temporary (<5yr) storage or transport solution.

In addition, although I haven't yet tested it, this method could hopefully be adapted for use in the field.