Category: Research Methods Page 6 of 9

An A. distichus futiley searching for possible escape routes.

A cage within a cage

Most butterfly cages are easily collapsable (though not always, as this picture indicates), making them convenient for storage.

We have at least six butterfly cages (seen collapsed and stacked here).

When handling lizards in the lab, it’s important to minimize the chances of a crafty critter escaping.  Collapsable “butterfly cages” are a convenient and thrifty way to ensure the security of your lizards during cage maintenance, egg checks, lizard-handling, and feeding.   Before removing the egg laying cups to see if any eggs were laid, for example, we generally place the Kritter Keeper housing breeding animals into a butterfly cage to ensure that any animals springing out don’t get very far (this practice, of course, will prove unnecessary once our room is entirely converted to new custom cages).  In addition to their many uses in the lizard colony, we also use butterfly cages extensively in the field as temporary housing or to sort animals sampled throughout the course of a day.  The cages we use are only around $20 each and they’re well worth the investment for any lizard lab.

A copy of the toe clipping scheme resides inside the egg log as a quick reference when clipping or ID-ing babies.

In a recent post on marking methods for field studies, Yoel made mention of the technique we use here in the lab: toe clipping.  It is true, as Yoel stated, that this is not an ideal method for lizards in field studies due to the difficulty of identifying the numbers from afar and the chance loss of toes in a rough lizard life.  However, for the purposes of the lab, toe clipping has proven to be an easy and effective method of identification.  After looking into a few schemes used by other researchers, I settled on a pattern that allows for numbers 0-1999 and involves clipping at most two toes on each foot.  With such high egg production over the past year in the lab, it is looking like the next round of breeding will require an adjustment to allow for numbers 0-9999, but the original scheme is serving its purpose for the moment.

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Anolis species in HerpNet wordcloud, name size is plotted to be proportional to the number of records on HerpNet for that species

Because I’m a big fan of obtaining data from public databases I’m writing another post on availability of anole data from huge bioinformatic databases.  This time, I’ll discuss the NSF-funded database of amphibian and reptile museum records known as HerpNet.  I found an astonishing 142,225 unique Anolis specimen records on HerpNet, including 602 unique binomials.  The over-abundance of names relative to recognized species is due largely to lots of misspellings (I found five different spellings for vermiculatus and four for valencienni).  An interesting side note on how errors in electronic databases can propagate themselves: One individual of Anolis sagifer appears in the MCZ (catalog #45484).  You can see the original catalog entry here.  This entry was mis-transcribed, likely when the database was digitized.  That in term led to it’s presence on the MVZ website and HerpNet, and also spawned search pages on GBIF and ITIS.

Many of the five most common species on HerpNet are also among the most common on GenBank; A. sagrei (13040), distichus (8944), carolinensis (8270), cristatellus (7126), cybotes (7106).  A. krugi (number 2 in terms of sequences on Genbank) falls to #22 on the HerpNet list.  Lots of interesting questions could be addressed using these HerpNet records.  For example, we could use these records to thoroughly investigate how new anole names have accumulated and been used over time.  Has species discovery/description been leveling off?  HerpNet records could also be used to consider how the anole research community’s interests have changed over time and how specific policies have impacted anole collecting?  How, for example, has the US embargo of Cuba impacted collection of Cuban specimens?

The more interesting applications of the HerpNet database will come from a careful consideration of the data associated with individual specimen records.  A number of efforts, for example, are already underway to use the thousands of georeferenced locality records for anoles included in the HerpNet database to address questions about geographic range and community evolution.

Incubating eggs

As Dan Warner mentioned in a recent post, moisture availability is extremely important to the development and survival of anole embryos. Throughout our time breeding anoles in the lab, we have experimented with different methods of incubating eggs, including different substrates (potting soil, a mixture of soil and vermiculite, and just vermiculite), differing proportions of water and vermiculite, and supplementing substrate with water throughout incubation. We have now settled on a recipe that seems to minimize death, mould, and desiccation in our Anolis distichus eggs.

We currently start by mixing a recipe that combines 220g of vermiculite with 380g of water. According to a water potential curve for vermiculite, this mixture has a water potential of -150kPa. We then put 130g of the mix into clear small deli cups which have pre-punched holes on the side for ventilation. The clear cups make it easy to quickly monitor eggs and to spot new hatchlings. We store these cups in our lizard room at a temperature of about 29C.  As eggs get older they are gradually rotated closer to a small fan that is set on a timer to run 10 mins twice daily to increase airflow. We’ve found that the recipe we’re using does not require addition of more water during incubation.

I’ve heard and read about many different ways that people are treating anole eggs and would love if AA readers shared how they take care of their anole eggs!

The core of my dissertation involves assessing genome-wide patterns of gene flow during anole speciation. For a variety of reasons, I ultimately want to acquire DNA sequences from throughout the genome.  As a first pass, however, I’ve been using Amplified Fragment Length Polumorphisms (AFLPs).  As a molecular technique, AFLPs are experiencing a comeback of sorts [1] [2].  Popular in the early 2000’s, AFLPs went out of favor as Sanger sequencing became cheaper and easier.  The resurgence AFLPs has largely been due to the realization that evolutionary patterns often vary throughout genomes and therefore methods that survey as much of the genome as possible are preferable to those that look at one or a few regions.

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A highly technologically advanced egg-laying location: a used bulk-size yogurt container (generously donated by various staff in the department) filled with moist vermiculite. In this case the lid is removed so that you can see the vermiculite inside.

While Anolis distichus may have plenty of options for where to lay eggs in the field, we needed to do a bit of experimentation before landing on a good place for them to lay their eggs in the laboratory environment.  Other anole facilities tend to allow their lizards to lay eggs in the soil of potted plants, and we originally had our lizards laying in the soil substrate of their cages.  However, eggs that spend time in soil tend to desiccate quickly, sometimes even before they’re discovered.  Given that we knew our production would be limited, we wanted to avoid this risk during our breeding experiments.  We arrived at a solution that is a combination of breakfast and gardening: yogurt cups filled with moist vermiculite.

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