Category Archives: Research Methods

Noose Pole Poll

We anolologists (and herpetologists generally) are a devoted bunch, particularly when it comes to our field equipment. It is therefore very troubling to learn that an essential component of our field kit is being discontinued. Perhaps most chilling is the thought losing access to our beloved [1] [2Cabela’s Panfish Poles. A recent series of tweets between AA stalwart James Stroud and Cabela’s customer service revealed noose poles are currently out of stock and may not return:

We have experienced the disappearance and return [1] [2] [3] of these poles before and, despite our best efforts, have not found a good alternative. With this essential tool at risk, I am taking up the effort to convince Cabela’s it is worthwhile to continue producing panfish poles. I would like to present them with the economic argument that many herpetologists use, and will continue to buy, this product.  I created a Twitter poll below and will present the results to Cabela’s customer service in making our case. Please take a moment to share your thoughts using the poll and in the comments. Thanks!

SICB 2017: A Field Based Approach to Study Behavioral Flexibility


Levi Storks explains his project in New Orleans.

Most animal learning studies have been conducted in the lab with the assumption that those findings are representative of behavior in the field. However, assessing behavior in the field increases ecological relevance. In addition, birds and mammals have received much of the attention in cognitive studies. Yet we on Anole Annals know that these lizards can be quite clever.

Levi Storks, a Ph.D. student in Manuel Leal’s lab at Mizzou, set out to address these issues by designing a method for testing behavioral flexibility in brown anoles (Anolis sagrei). Wild lizards in the Bahamas were allowed to feed unrestricted on a maggot placed in the middle of a testing apparatus in order to acclimate lizards to the structure. Storks then used a clear plastic tube to block the direct route to food, requiring lizards to move to either end to gain access. Lizards that successfully completed this task were then tested to see if they could associate unique patterns on the ends of the tube with single openings.

Storks found that a subset of lizards could successfully complete the first detour task, and lizards made fewer errors over the course of solving the detour task. These findings suggest brown anoles can learn and exhibit behavioral flexibility. Stay tuned for more of Levi’s work as he’ll be applying these methods to assess differences in behavioral flexibility between populations that vary in ecology!   


Is There a Crisis in Anolis Taxonomy? Part 2


In a (somewhat) recent blog post entitled “Is there a crisis in Anolis taxonomy?”, Julian Velasco invited discussion on a perceived decline in the number of new anole taxonomists.  While it was a fun look at the dynamics of anole taxonomy over time, I couldn’t help but feel like there is a more pressing taxonomic crisis going on right now, and it affects many of the researchers that frequent this blog.

I fear too many species of Anolis are being described based on questionable evidence.  While this problem is not unique to anoles (a common term for it is “taxonomic inflation”; Isaac et al. 2004), a number of recently described anole species may be the result of overzealous taxonomic splitting.  I will give some examples below and then briefly discuss two lines of evidence that I believe are often used to divide species inappropriately.  Before I do so, it’s worth stating up front that I’ll focus on the work of Dr. Gunther Köhler and colleagues. This shouldn’t be surprising, as Dr. Köhler is the most prolific living describer of anole species.  The following criticisms should not be seen as personal, as Köhler is not unique on any of the points I discuss below.  But with many cryptic species described or resurrected over the past 10-15 years, his work has the largest impact on anole taxonomy and the science that depends on it.

I’ll start with the revision of the Anolis tropidonotus complex published in Mesoamerican Herpetology (Köhler et al. 2016).  Below I provide a quick breakdown of the paper.  I hope that others will contribute their own views on this work in the comments.  The A. tropidonotus group is one that I am well-acquainted with, having spent months of field time collecting individuals across the distribution of the group.  Köhler et al. (2016) raise a subspecies (A. tropidonotus spilorhipis) to species status while describing two new species, A. wilsoni and A. mccraniei.  Unfortunately, the data presented–morphology and DNA–do not appear to strongly support the recognition of any new species level taxa.  I argue that the inference of four species within A. tropidonotus sensu lato should require stronger evidence than that presented.


The authors sequenced 16S mitochondrial DNA for molecular analyses and present a consensus tree from Bayesian analyses of these data. This tree recovers four well-supported and geographically circumscribed mtDNA haplotype clades that correspond with the four new species. A table following the tree reveals the genetic distances between putatively new species topped out at 4.5%. This level of mitochondrial divergence is significantly less than intraspecific variation observed in other anoles (Malhotra & Thorpe 2000; Thorpe & Stenson 2003; Ng & Glor 2011). Moreover, Köhler et al.’s (2016) sampling map reflects sparse sampling of molecular data.

Based on Figure 3, morphology (other than perhaps hemipenes, which I discuss below) does not provide any support for delimitation of those populations characterized by distinct mtDNA haplotypes. The dewlap differences reported are slight and appear to fall within the type of variation observed within and among other populations of species in this group (see photos at the top of this post for an example of two spilorhipis males that came from the same locality; photos courtesy Luke Mahler). Bottom line–we see several populations with mitochondrial haplotypes that cluster together geographically with little to no morphological evidence for divergence.

The phylogenetic and morphological patterns displayed in Köhler et al. (2016) are consistent with patchy sampling of a widespread and continuously distributed species with potentially locally-adapted populations. The authors cite “the high degree of genetic distinctiveness… as evidence for a lack of gene flow, and conclude that these four lineages represent species-level units” (Köhler et al. 2016). This assumption is questionable, as researchers have long known of the pitfalls of using mtDNA to determine gene flow (Avise et al. 1983; Avise et al. 1984; Funk & Omland 2003) and supporting evidence from morphology is lacking. The different hemipenial types represent the strongest evidence for recognizing the lineages mtDNA haplotype groups. Below I will discuss the utility of those traits for species delimitation.

Finally, the authors did not compare their purported new tropidonotus-like species to Anolis wampuensis, a morphologically indistinguishable (McCranie & Kohler 2015) form that is potentially codistributed with the new species A. mccraniei. This should have been done to avoid the possibility that A. wampuensis is conspecific with one of the newly named forms.

Another example of taxonomic inflation in Anolis is from a 2014 monograph in Zootaxa (Köhler et al. 2014). Continue reading Is There a Crisis in Anolis Taxonomy? Part 2

Seeking Input for a Child-Friendly Research Project


In my science lab with my little green friend. This photo will actually be on the back cover of my upcoming book!

As a regular reader of Anole Annals and a subscriber to the Twitter feed, I am honored to have the opportunity to write this post. For those who might remember, I am the elementary school science teacher in Princeton, NJ who made international news (and a mention on Anole Annals) when one of my kindergarten students brought me a juvenile Anolis carolinensis that her mother found in a bundle of salad greens. I am happy to report that “Green Fruit Loop” is still doing well in a spacious terrarium, and I have considered the logistics of returning her to the wild once she’s fully grown. Of course, from what I’ve been reading about her place of origin (south Florida), I’ll have to make sure I find a spot with tall trees, to make sure she has refuge from Anolis sagrei.

Green Fruit Loop

I’ve gotten into the habit of referring to Green Fruit Loop as a “she,” but perhaps an anole specialist could make an accurate determination?

My students continue to be enthralled with our surprise classroom companion, and I have been considering ways to include these children in a scientific investigation on color change We have a second terrarium of adopted Anolis carolinensis (my momentary fame made me a magnet for unwanted pets), and even though I have told my students that anoles don’t assume specific colors to blend in with their backgrounds, this group was almost exclusively green when housed with plants, but since a fungal disease eliminated all vegetation over the winter, these anoles now remain perpetually brown among the rocks and woodwork.


Green Fruit Loop definitely doesn’t look green here!

These observations, which my students have used as evidence that Carolina anoles do, in fact, change color to camouflage (contrary to what their teacher tells them), have prompted me to consider a long-term study, in which several basking platforms will be painted different colors and anoles that use them will be photographed at multiple intervals per day. For example, one platform might be green, one brown, one white, and one black, and a camera on a timer will take photographs of each platform hourly. We could then compare these photographs over time, determine which individuals are exhibiting certain colors on certain platforms, and possibly draw conclusions from what we observe. I recently obtained a grant from the American Society of Plant Biologists to build two large habitats for tropical plants, so this would be an ideal location to house additional groups of anoles for this experiment to proceed.

If anybody has suggestions for the colors and materials that we might use for basking platforms (I am planning on four per habitat, each under its own light), as well as any possible modifications to this experiment for greater scientific merit, please feel free to comment on this post or write to me at Of course, animal welfare is always the highest priority in any of my educational projects, and my group of adopted anoles will never be housed with any field-collected specimens (like Green Fruit Loop) to minimize possible spread of parasites and disease.

Once this experiment gets going, please check in and see what my students are learning on Twitter @markeastburn or at my website Thank you for reading!


Shipping Live Lizards via Cargo from the Dominican Republic

Assuming you can’t get your lizards to fly themselves to your lab, you might want to read this information on how to transport them home. Photo from

After years of transporting live anoles from the Caribbean to my lab in the United States in my checked luggage, this summer in the Dominican Republic, a Delta Airlines agent refused to accept our cooler full of lizards as luggage for our plane. After pursuing every avenue we could think of, it became clear that our only remaining option was to ship the lizards as cargo. We spent several days working out this process, and after making a number of mistakes, we finally arrived at a relatively smooth procedure. To prevent others from having to learn these steps on their own, if such a situation arises for other researchers, we’ve written out the steps that worked for us below. The details provided are for the airport in Santo Domingo, but this general approach may be helpful in other locations as well. (And, if you find yourself in the Dominican Republic in the near future, I’d be happy to give you the contact information for all of the folks listed below.)

Continue reading Shipping Live Lizards via Cargo from the Dominican Republic

BSA of Norops lineatopus

Geometric Morphometric Analysis of the Shoulder of Jamaican Anoles

garmani mating trivers IIxBirds are lovely animals. Our avian friends swoop through the air, defecate on field equipment, and consume lizards. What’s not to like?! Well, their shoulder region, for example. Lost interclavicle, reverted muscle pathways, and so many other anatomical adaptations that appear crucial for the modern avian life style, but that are hard to explain in a gradual-evolutionary context. Reconstructing the structural evolution of the avian shoulder remains a challenging task to students of biomechanics and kinematics. When I left my European homestead to enter the Canadian realm of biological sciences, I was hoping to solve the evolutionary mystery of the avian shoulder, at least in part. Alas, the discovery of anoles sent me on a much more convoluted journey.

Here is the first tale that resulted from that endeavour (Tinius & Russell 2014).

Continue reading Geometric Morphometric Analysis of the Shoulder of Jamaican Anoles

How Many Lamellae Are on this Toepad?

One of the age old questions in anole morphology is at what point do you stop counting lamellae on the toepad?

Without giving any more information on various techniques or methods, I thought it would be interesting to ask the AA community their personal opinions. Below I have attached a flatbed scan of a toepad. Could people please fill out the corresponding poll below, and I will present the results in a follow up post!

alt text

Lamellae numbered 1-51 on the 4th digit of an Anolis lizard hindfoot

Fluorescent Lizard Skeletons Used to Precisely Measure Growth

Several weeks ago, Anole Annals highlighted a recent paper that uncovered the molecular bases of craniofacial dimorphism in the carolinensis clade of Anolis lizards (for full disclosure, I am the lead author of that paper). Hidden deep within that research is a relatively new technique for precisely measuring rates of skeletal growth that may be of interest to the community. I briefly introduced this technique several years ago in a post about methods of skeletal preparation, but with the details of this method now available it is worth highlighting once more.

Double labeled facial skeleton of A. carolinensis. Green label (calcein) and red label (alizarin complexone) separated by 30 days.

Because some images shouldn’t be lost in the supplementary materials. Double labeled facial skeleton of A. carolinensis. Green label (calcein) and red label (alizarin complexone) separated by 30 days.

Growth in body size can often be measured using calipers or a ruler. But in some situations a finer-scale analysis may be necessary, such as when differences in growth rate may be subtle, within the range of error associated with those manual methods. Fluorescent calcium chelators provide the precision needed to measure differences on the order of microns per day. In the recent paper, this technique was used to measure facial elongation in sexually mature green anoles, which was only ~8um per day in males and ~4um per day in females. These compounds are stable, are not highly toxic to animals, are relatively inexpensive, and can be easily used in the field or the lab. They can also be applied to adults or hatchlings with little modification to the protocol as injection volumes are typically 10-20ul depending on size. Ultimately, there is a lot of versatility to the way in which this method can be applied.

Dimorphism in facial growth rates between male and female A. carolinensis. Modified from Sanger et al. 2014.

Dimorphism in facial growth rates between male and female A. carolinensis. Modified from Sanger et al. 2014.

While new to herpetology, this technique was adopted from the biomedical literature on fracture repair where precise spatiotemporal measure of bone deposition is required. The general experimental framework is that pulses of chelators with different fluorescent properties are delivered at distinct intervals, the skeleton prepared, and the distance between the labels recorded from digital photographs. Calcium chelators are available that fluoresce under many of the standard filters used in modern microscopy – including green (calcein), red (alizarin complexone), orange (xylenol orange), and blue (calcein blue and oxytetracycline) – offering great experimental flexibility. Once incorporated into the bone, their signature remains strong for at least 30-45 days, until it is remodeled away as the living skeleton continues to grow and reshape itself. In the recent paper on craniofacial dimorphism, fluorescence in the facial skeleton could be observed following simple removal of the skin because the face has little to no overlying connective tissue. Measuring growth of the vertebrae or limbs is also possible, but may require careful sectioning of the bone using either plastic or paraffin protocols. Ultimately I think that there is a lot of potential with this method that has yet to be explored in the context of organismal biology. I hope that by highlighting this method here more people become aware of its utility and give it a try.

Available Now: A New, Large Phylogeny of Anoles

BEAST estimated phylogeny of anoles. Circles on nodes represent posterior probability, black > 0.95, grey > 0.90, white > 0.70.

BEAST estimated phylogeny of anoles. Circles on nodes represent posterior probability, black > 0.95, grey > 0.90, white > 0.70.

In the course of our recent study on sex chromosome evolution in anoles (Gamble et al. in press) [AA post] we assembled a 216-species mitochondrial DNA phylogeny of anoles, the largest published to date (at least that we know of), yet containing only a little more than half of all recognized species. Although we collected new sequences for some species, our dataset is largely built on the hard work of others who collected and published on sequences from across the genus, such as Jackman et al. 1999, Poe 2004, Nicholson et al. 2005,  Mahler et al. 2010 [AA post], and Castañeda & de Quieroz 2011 [AA post].  Without access to data from these and other studies, we would have had a far less complete and robust tree for our comparative analyses.

There is a big debate going on now regarding what, where and how much data should be shared in association with publishing academically. I personally feel that providing easy access to those data used and generated during a study serves to accelerate the rate and increase the quality of scientific discovery. I am heartened that more and more journals are making data deposition a requirement for publication, although often this means little more than dumping sequence data to GenBank. Sites like Dryad, Figshare, and GitHub now provide open, permanent, and citable access to raw data, figures and, most importantly in my view, research products like alignments, code and analysis logs. In an effort to make our data as accessible and useful as possible we have archived our alignment, MrBayes and BEAST consensus trees as well as as the BEAST posterior distribution on the digital data repository Dryad [doi link]. It is our hope that other anolologists can use and improve upon these data to ask new, interesting questions and to build a larger, more complete view of the evolution of anoles.

The History Of Lizard Noosing

Time honored anole field technique. But since when?

Here at AA, we’ve frequently discussed the art and practice of lizard noosing, such as posts on the best material to use to construct a noose, as well as the variety of suitable poles commercially available. Recently, I was asked a question for which I did not have an answer. To wit, what is the history of lizard noosing? Did our herpetological forebears use nooses? I’m aware that at least some herpetologists in the 70’s were doing so. What about earlier than that? Did Stan Rand noose lizards? Ernest Williams in his younger days? Barbour?

Everyone’s aware that when looking for information, if you can’t find it on Google, it’s not worth knowing. This, however, would seem to be an exception. Wikipedia has no entry on lizard noosing, nor does a Google search on the relevant terms turn up any answers (such a search does, however, turn up a plethora of websites and Youtube videos offering lizard noosing tutorials).  So, I put it to you, AA readers: who can enlighten us on the history of anole noosing?

Anole Skeletal Preparation: Useful And Beautiful

Recently, we had a post on the cool bark anole embryo photographs produced by Catherine May at Arizona State. Catherine has now done this one better by producing a series of photographs, along with explanatory text, detailing the process by which skeletal preparations are made via the old method of clearing-and-staining. As the photo reveals, the resulting products are not only scientifically informative, but quite beautiful. And while on the topic of anole skeletal preparation, check out Thom Sanger’s Halloween-themed post on the same from 2011.

Anolis conspersus, UV Dewlap Photos And Anoles As House Geckos


On a recent trip to Grand Cayman I was interested in the UV reflecting dewlap of Anolis conspersus. The dewlaps of these lizards appear blue to our visual system but are maximally reflective in the ultraviolet. While anoles have 4 cone types (ultraviolet, blue, green and red sensitive), humans have only 3 and cannot see UV light so to understand what these lizards look like in the UV, we have to use specialized camera equipment.  The photo to the right shows what a displaying A. conspersus looks like to our camera system when imaged in the human visual spectrum as commercially available digital cameras also have only three channels corresponding to the three human cone types.  Presumably if we were also able to see in the ultraviolet as many other animals can, our cameras would be designed with a separate channel for ultraviolet.



These images of the lizard in the UV show clearly the regions of the dewlap and that are highly UV reflective and the pattern of UV reflectance in other areas.  One somewhat interesting finding is that while the dewlap scales are highly reflective across the human visual spectrum (which is why they appear white to our eyes) they reflect very little UV light.  The lower photo is a monochromatic image (both the red and blue channels in this camera are sensitive to UV so the raw image appears purple) that makes it a bit easier to see brighter areas as white.  Note how bright the dewlap appears relative to the reflectance standard, when imaged in the human visual spectrum a similar monochromatic image of the dewlap would appear very dark.  I believe this shows the potential value of UV photography when studying Anolis dewlap patterns.  While the UV nature of the A. conspersus dewlap is uniform, it’s likely that other species have patterns visible in the UV we’ve previously missed.  We have also used this UV photography setup in SE Asia to image Draco flying lizards and other species, some of which have patterns that are visible only in the UV band.  The goal of this project is to make a camera system with pixel channels similar to the four cone types found in Anolis lizards and birds to image whole organisms and really “see” the patterns organisms experience with their visual system as they would see them.  As Anolis visual pigments and their associated oil droplets appear to be fairly conserved, this seems to be achievable.

photo (2)

Another surprise (to me) was the large number of A. conspersus on Grand Cayman using lights at night to feed.  I’ve spent many months doing fieldwork in SE Asia and Central America and can’t recall seeing this sort of thing with other diurnal lizard species, but on Grand Cayman it was quite common in A. conspersus.  I observed one A. conspersus male chase away a Hemidactylus that got too close to the light, showing that the anoles at least occasionally displaced the group I typically associate with feeding around lights.  A check of the literature shows this has occasionally been documented on other Caribbean islands, but as far as I can tell no one has published on this in mainland species.  What diurnal lizard species have others observed using lights to feed at night?

ASU Green Anole Genome Reannotation Now Available on Ensembl

Green anole (Anolis carolinensis). Photo courtesy of Karla Moeller.

Green anole (Anolis carolinensis). Photo courtesy of Karla Moeller.

Ensembl Release 71 includes many updates for Anolis carolinensis, including the addition of the Arizona State University (ASU) Anole Genome Project annotation recently published in BMC Genomics (Eckalbar et al., 2013). This release includes an updated Ensembl gene set and aligned RNA-Seq data from a number of tissues, including embryo, lung, liver, heart, dewlap, skeletal muscle, adrenal gland, ovary, and brain, which have been added to the track viewer. These RNA-Seq data from individual tissues and from the ASU reannotation or the “Anole Genome Project” can be viewed just below the Ensembl gene tracks, as in this example.

Advice Needed: GPS Tags For Giant Bronze Geckos?

Here’s a question for AA readers from Nancy Bunbury, from the Seychelles Island Foundation, who is conducting some exciting work on large gecko interactions, ecological roles, and niche separation in the palm forests of the Seychelles:

Giant-bronze-gecko-on-tree“The main species in question is Ailuronyx trachygaster (first field study on this amazing species) and one thing we would love to do is look at movements and territory size (also because we suspect it’s the main pollinator for the coco de mer which has huge conservation and inevitably commercial value). We are looking into GPS tags for the geckos (which are about 150g in weight) but it seems the technology for such a small tag requiring GPS and remote downloading is not yet available. Do you happen to know if such tags have yet been developed and who I might be able to contact for them (I’ve tried the standard larger companies for animal tracking devices)?”

Any suggestions?

Possible Cage For Lizard Field Experiments

IMG_1720On a recent trip to Toronto, eminent bee-man and pollination biologist James Thomson showed me his lab, including a cage used for bee pollination studies.  The cardboard box is a “box of bees” that can be bought commercially and the experiment involves training bees to go to containers with different colors. Despite being fascinated by the research, my mind couldn’t help but wandering to thinking about how useful such a contraption could be to set up in the field for ecological or behavioral anole studies. As you can see, the cage is big enough that it could house a number of anoles at natural densities, and the mesh lets sunlight and rain through. James kindly informed me that the cages can be purchased at Bioquip; the largest they stock is 6′ (h) x 6′ (w) x 12′ (l), but James told me that larger models can be custom-ordered, and that they are very hardy in the field. Someone should try this!

Anolis: The Most Written About Lizard Genus?

In the era of Big Data, we can ask questions that would have been inconceivable just a few years ago.  Consider the types of questions we can ask using Google’s Ngram Viewer, which uses full-text searches of >4% of all books ever printed to characterize relative word or phrase usage over time (this approach was initially described in a 2011 Science paper about “Quantitative analysis of culture using millions of digitized books“).

Among the most important questions one might ask with the Ngram Viewer is “What is the most written-about lizard genus?”  I did some preliminary scouting to assess the relative usage of some of the lizard genera that I guessed would be the most popular. I quickly narrowed my queries to the five taxa – Anolis, Sceloporus, Varanus, Lacerta, and Gekko – that I think give the most interesting graphs for discussion. I excluded other potentially popular genera from my queries for for a few reasons. Iguana is very popular, but I eliminated it because it is often used colloquially to refer to lizards that don’t necessarily belong to the genus Iguana. Eumeces never appears as frequently as the other genera in my searches. Pogona is immensely popular as a pet, but usage of this genus name is still far below the others in my list.

Ngrams_1800_1900Lacerta jumps out to a big early lead and maintains a strong lead throughout the 19th century, thanks to its widespread use in Latin-language literature from the 19th century and countless books about the European fauna (Ngrams Viewer even provides links to the books or articles containing the phrase of interest!).

Ngrams_1900_2000In the early 20th century, Anolis joins the competition as one of the most popular lizard genera, and opens up a sizeable lead by the 1980s that it maintains until the turn of the 20th century.  Although Anolis is briefly surpassed by Varanus in the 2000s, it nudges back into the lead by the end of 2008!



There you have it folks, quantitative proof of the popularity of Anolis!  Have I failed to consider some genera that might be competing with Anolis in the lizard genus popularity contest?

Tissue for genetic material: options other than tail tips?

I was hoping to get suggestions from the readers of AA about methods of tissue collection for genetic work other than tail tips. I’ve been working with the agamid lizard Sitana ponticeriana, and my work is now taking decidedly genetic directions. It remains unclear whether or not these lizards regenerate their lost tails–while they seem to lose tails easily, I didn’t see any lizards with noticeably regenerated tails in the field. Given this, I am a little uncomfortable with the idea of taking tail tips as tissue for genetic work. Are there other common and easy options for sampling tissue from lizards? Many thanks in advance for your responses!

(Feel free also to weigh in with whether or not you think it acceptable to collect tail tips in a species that certainly autotomizes its tail but does not grow it back–it seems like a grey area to me).

A male Sitana ponticeriana near Pune, India.

Considered breeding anoles in situ?

If you are lucky enough to live in the tropics then you can do away with incubators, endless tinkering with temperature, humidity and light regimes and let nature do it for you. I have been breeding A. apletophallus in Panama for almost two years and thought I should share with you my take on breeding anoles “in situ.”

Left: Mesh cages hanging in the shade house. Right: Newly emerged hatchling.

I use a very basic shade house that is situated on the edge of the forest. The temperature and humidity are similar to the forest where the lizards live. In the shade house, the lizards are housed individually in mesh cages that I constructed from pop-up laundry hampers and mesh bags. Each cage is outfitted with three branches and a plastic leaf. Females have a shallow soil plate to lay eggs in, which they happily do. I feed adult lizards every three days and at the same time check for eggs. All eggs are removed and placed in a plastic cup with water and cotton wool, which is then placed inside a ziplock bag. Eggs are “incubated” at ambient temperature (~45 days). When the eggs hatch the hatchlings are transferred to plastic boxes with a mesh lid. These “baby boxes” also contain three small branches and a plastic leaf. Hatchlings and subadults are feed every other day. All lizards are sprayed daily with water. Although there is probably room for improvement, this has been a successful and economical strategy to breeding anoles in the tropics. For anyone who wants more details I have posted this on my webpage under “animal husbandry.”

Color Catalogue For Field Biologists

Anole biologist Gunther Köhler has produced a handy manual, available from Herpeton publishers, to help describe colors of specimens, especially in field situations. The book’s introduction can explain better than I what it is used for and why it was written:

The accurate description of the coloration in life of organisms represents an important component of the work of any field biologist. Subtle differences in the coloration in life, such as in the color of the iris, the lining of the mouth cavity, or the tongue are diagnostic for certain species and have been used by taxonomists to differentiate among species.

Whereas many aspects of the external morphology of scientific specimens can be preserved with proper fixation methods, there is still no way to assure the long-term conservation of the coloration in life in such specimens. This is especially true for animals traditionally fixed with the help of formalin and ethanol, such as fishes, amphibians, and reptiles, and then stored as a wet collection. Colors such as red, yellow, and orange disappear rapidly once the specimen is placed in the preservative. Green-colored amphibians and reptiles can turn blue, lavender, purple, or black within a short time after preservation.

The Catalogue also provides definitions and examples of different phenotypic characteristics.

Of course, taking photographs of animals helps to document the coloration in life. Possible drawbacks to this technique are incorrectly adjusted white balances, which cause colors not to be reproduced accurately. Also, photographs often do not show coloration of hidden body parts. Therefore, biologists have a long tradition of recording colors by making written descriptions. Since individuals see colors differently and because it not easy to define, for example, different shades of brown or green in words, having a color standard helps to produce more objective and detailed descriptions that also have a greater chance of being reproducible. Such a reference can be used to compare descriptions made by different persons at different times and places. For decades, field biologists have utilized the “Naturalist’s Color Guide” by Frank B. Smithe (1975-1981) as the standard reference for color descriptions. However, for many years now, this important reference has been out of print and is no longer available.

I have used Smithe’s “Naturalist’s Color Guide” (called “Smithe Guide” from here on) extensively during the past 20 years, and my copy now clearly shows signs of this intensive usage under field conditions over the years. With no hope of being able to obtain a copy in good shape to replace my old one, I decided to produce a new reference to fill the gap left by the now unavailable Smithe Guide.

The resulting “Color Catalogue for Field Biologists” you are holding in your hand is not a duplicate of the Smithe Guide. Continue reading Color Catalogue For Field Biologists

More On Nicholson et al. 2012: Let’s Look At Their Methodology

ResearchBlogging.orgMost people who have commented on the blog about Nicholson et al. 2012 have focused on whether is it really necessary to name all these inferred clades as genera. I agree with those who state it is completely unnecessary and disruptive, and that there are alternative ways (e.g., assigning names to relevant clades independent of the genus rank) to describe the diversity of Anolis. That said, I would like to direct the discussion towards the methodology used. Yes, there are a lot of missing ND2 data in their dataset (e.g., all of the new data presented in Castañeda and de Queiroz 2011 is missing), but I think it is more relevant to consider how they treated the data they did include. First, the molecular partition of their DNA: the protein coding gene ND2 was not partitioned into codon positions, which has been shown to be the best strategy (e.g., Schulte and de Queiroz, 2008; Torres-Carvajal and de Queiroz, 2009; Castañeda and de Queiroz, 2011), and instead, they chose to set a different partition for each of the tRNAs included (five) and one more for the origin for the light strand replication piece (which is ~30 bases long). As the Bayesian analysis requires a large-enough number of characters to estimate the parameter values for the model selected, I thought it was recommended to have partitions of more than ~300 bases (and I can’t think from the top of my head for a specific citation here). Neither the OL nor any of the tRNAs is close to this size (and the AICc, the corrected Akaike Information Criterion, intended for small sample sizes should have been used to select the best fitting model here instead of the regular AIC).(For more on partition selection and consequences of under– or overparameterization, check Brown and Lemmon, 2007 and Li et al. 2007). This should raise an eyebrow about the thoroughness of the analyses. However, in reality, I think this would have little effect on the actual phylogeny. Those clades that are strongly supported would be robust enough to withstand model and partition misspecifications.

On the other hand, the treatment of the morphological characters might have more serious effects on the resulting topology. Nicholson et al. explain that they used Poe’s 2004 morphological data as is, but without the complex coding system he used for continuous and polymorphic characters, and instead considering all possible characters to be equally weighted. (To be fair, Poe did use equal weighting for characters in his analyses; the cost of changes between states within a single character is what is different). Poe coded continuous characters using a gap-weighting method, which divides the range of a continuous character into discrete segments, maintaining information on the order of the character states and the magnitude of the difference between them, and he coded polymorphic characters using a frequency method, which keeps track of the fraction of individuals within the sample that shows a given state. From what I understood, Nicholson et al. considered all changes to be of equal cost, so transitioning from the smallest head to the largest head, or from having all individuals showing condition x to all individuals showing condition y (where some taxa exhibit both conditions), will cost 26 steps, which is the cost of changing from state a to state z (as recognized by Poe). This means, in the combined parsimony analysis, a transition between the two extreme states in a continuous or polymorphic morphological character is equivalent to [single] DNA substitutions at 26 different positions [characters]. Moreover, changes in those morphological characters that were not continuous or polymorphic would cost only a few steps. This weighting scheme (in the parsimony context) will actually give a higher weight to some morphological characters, which is exactly the opposite of what the authors were aiming for (i.e., equal weights). The effects of this unbalanced weighting on the resulting topology? Not sure, but I’m going to guess not insignificant!

One last thing. Several of their proposed genera (Dactyloa, Deiroptyx, Chamaelinorops and Xiphosurus) are not monophyletic on their combined data tree, the one that supposedly serves as the basis for their taxonomy…

KIRSTEN E. NICHOLSON, BRIAN I. CROTHER, CRAIG GUYER & JAY M. SAVAGE (2012). It is time for a new classification of anoles (Squamata: Dactyloidae) Zootaxa, 3477, 1-108