PhD week 65: DMHF

Male (left) and female (right) weevil parts mounted in DMHF.

The internal morphology of insects is a veritable gold mine of interesting characters. Obviously, in order to find investigate these characters it is necessary to dissect specimens. Unfortunately, this results in disarticulated beetle bits that one needs to store somehow in order to look at them again in the future.

The classic method of storing dissected materials is in very small vials that can be kept on the pin that holds the remainder of the specimen. This can be useful, but is a bit fiddly to remove the pieces from the vial when one needs to look at them again. In addition, it's become difficult to buy smaller glass vials, and I'm not a fan of the polyurethane vials that are readily available.

An alternative method that I've been exploring this past week is using the mounting medium DMHF. This mountant is soluble in water and dries crystal clear. The method that I've been using is to put a drop of DMHF on a card, immediately place the parts into the medium, add sufficient DMHF to cover the parts entirely, and leave for a couple of days to set. The card is then pinned below the specimen, and the parts are both protected and readily viewable (see picture above). I haven't tried it yet, but I understand that removing the parts is as simple as placing the card in a small dish of water and waiting a few minutes for the DMHF to dissolve.

A frustrating part of working with DMHF is that it forms a skin soon after exposure to air (c. 30 seconds). This skin can make it a little tricky to manoeuver pieces after placing them into the medium. After a little bit of experience though, one can usually get pieces in without needing to do too much messing around with them after the fact. I've had a positive first experince with the stuff, and am intending to carry on using it for the time being at least.

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Websites:
Logic Matters
NetKnots—Tautline Hitch
Arctic Terns breeding in Netherlands migrate via Australia
LaTeX Stack Exchange: What fonts are installed on my box?

Listened:
Norma Jean—Disconnecktie: The Faithful Vampire

Watched:
Star Trek (Original Series) Season One

PhD week 10: First gel

This week's milestone is pictured above. It is a picture of an agarose electrophoresis gel. These gels are made from a seaweed derivative, and are used to separate DNA of different lengths. Samples of DNA are placed into holes in the gel, and an electric current is passed through it. The DNA moves through the gel towards the positive electrode, but longer pieces move more slowly than shorter pieces. The gel is then stained with a chemical that binds to the DNA and fluoresces under UV light, allowing one to see the result. The white dashes in the picture above show where DNA has ended up. The obvious ones at either side are the DNA ladder, a mixture of DNA of known lengths, which allows one to figure out the lengths of DNA in the samples.

It so happens that this particular gel is the first one of many that I will run over the course of my PhD. It is the result of a PCR amplification of a specific fragment of DNA for a number of weevil specimens. The reason you can tell that it is genuine is that it is hideous as far as agarose gel pictures go, and the only reasons for showing anyone a picture of this quality is to point out the problems with it, or for sentimental reasons. It is shown here for both purposes. For starters, I left it running for too long, and the ladder has run off the end. Secondly, only two of eighteen samples actually worked (the ones that have been ringed). One of these is a positive control, a DNA sample that is known to have worked under the same conditions previously.

So, all in all, it's a somewhat disappointing result. However, optimising PCR protocols is a routine (though annoying) part of getting DNA sequences from a number of specimens. What this gel does show clearly is that I shall have to go through that process before I can routinely sequence DNA from my weevils.

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Read:
   Posadas P. 2012. Species composition and geographic distribution of Fuegian Curculionidae (Coleoptera: Curculionoidea). Zootaxa 3303: 1–36
   Wilson D. 2010. The People's Bible. The Remarkable History of the King James Version. Oxford: Lion
   McCulloch D. 2010. A History of Christianity: The First Three Thousand Years London: Penguin
   Psalms 52–54

Websites:
Marcus Ardern's blog and website
gitHub smart HTTP support details
gitHub: Forking repositories
Geographx free downloads
Using near IR in microscopy
Setting up screensavers in Precise Pangolin
AviAtlas

Listened:
Kevin Johansen—Sur O No Sur
Color Tango—Con Estilo Para Bailar

Watched:
Wallace's standardwing (Semioptera wallacei) mating dance
National Geographic TV Profile of Jim Frazier
Star Trek: Deep Space Nine Season 3

PhD week 6: Laser Scanning Confocal Microscopy

Bone cells viewed by laser confocal microscopy. Courtesy of the Wellcome Images photostream. License: CC: BY-NC-ND

This week, I had the opportunity to learn more about laser scanning confocal microscopy (LSCM). This type of microscopy uses lasers to excite fluorescent compounds in a very thin layer of the sample, allowing very clear images to be produced. Images can be made at a ranges of depths in the sample, and can then be merged together to give an idea of the 3D structure of the subject being imaged, such as the proturan leg shown in this video.

While it is frequently held that chitin can fluoresce to a degree, it appears that it is rather the presence of proteins and other sclerotizing agents in the cuticle that is primarily responsible for any autofluorescence seen. In the celebrated case of scorpions, it is believed that an alkaloid and a coumarin compound causes the fluorescence, while in froghoppers, it is resilin.

If the chitin is not fluorescing adequately, there are a number of stains that can be used. Eosin Y was judged to be most useful of a number of stains in one study, however in the test I did this week, I judged that it primarily stained the membranes, and not so much the sclerotised structures. A few other stains that may or may not prove useful include calcofluor, primulin and Congo red.

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Read:
Koerner L, Gorb SN, Betz O. 2012. Functional morphology and adhesive performance of the stick-capture apparatus of the rove beetles Stenus spp. (Coleoptera, Staphylinidae). Zoology 115: 117-127

Grant PR, Grant BR. 2008. How and Why Species Multiply. The Radiation of Darwin's Finches. Princeton: Princeton University Press

McCulloch D. 2010. A History of Christianity: The First Three Thousand Years London: Penguin

Psalms 37–38

Websites:
New Zealand Tango Festival, 19–26 June 2012

OsiriX an open-source volume rendering program.

Suraj Gupta—How R Searches and Finds Stuff

Steveko's Blog—10 things I hate about Git

BeetleBase and FlyBase genomic databases.

Installing Arial fonts in Wine

WineHQ information about Leica LAS AF Lite

Volume 3 of the NZ Inventory of Biodiversity published

Listened:
Demon Hunter—45 Days

Zao—Awake?

Foo Fighters—Wasting Light

Watched:
Star Trek: Deep Space Nine Season 3

Further Seems Forever—Light Up Ahead music video

Microscopy mounting media II

Today I found a brilliantly written, hugely informative, and well-illustrated guide to Safe Microscope Techniques for Amateurs, written by Walter Dioni. It is a series of four articles, originally published in Micscape, the magazine of the Microscopy UK organisation.

"Safe Microscope Techniques for Amateurs" discusses liquid media, solid media, mixed media and glycerin jellies. All techniques are explained clearly and several suggestions for equipment or readily available appropriate subsitutes are given. In summary, the series is an excellent introduction to microscope mounting techniques and gives some very useful tips for making good microscope mounts.