Vol.5 - Issue 4 Winter, 2015 - Southwestern Center for

SWCHR BULLETIN
Volume 5, Issue 4
Winter 2015
ISSN 2330-6025
Conservation – Preservation – Education – Public Information
Research – Field Studies – Captive Propagation
The SWCHR BULLETIN is published quarterly by the
SOUTHWESTERN CENTER FOR HERPETOLOGICAL RESEARCH
PO Box 624, Seguin TX 78156
www.southwesternherp.com
email: admin@swchr.org
ISSN 2330-6025
OFFICERS 2015-2016
COMMITTEE CHAIRS
PRESIDENT
Tim Cole
AWARDS AND GRANTS COMMITTEE
Gerald Keown
VICE PRESIDENT
Gerry Salmon
COMMUNICATIONS COMMITEE
Gerald Keown
EXECUTIVE DIRECTOR
Gerald Keown
ACTIVITIES AND EVENTS COMMITTEE
[Vacant]
BOARD MEMBERS AT LARGE
Toby Brock
D. Craig McIntyre
Benjamin Stupavsky
Robert Twombley
Bill White
NOMINATIONS COMMITTEE
Gerald Keown
BULLETIN EDITOR
Chris McMartin
MEMBERSHIP COMMITTEE
[Vacant]
CONSERVATION COMMITTEE
Robert Twombley
ASSOCIATE EDITOR
Ben Stupavsky
ABOUT SWCHR
Originally founded by Gerald Keown in 2007, SWCHR is a
501(c)(3) non-profit association, governed by a board of directors
and dedicated to promoting education of the Association’s
members and the general public relating to the natural history,
biology, taxonomy, conservation and preservation needs, field
studies, and captive propagation of the herpetofauna indigenous to
the American Southwest.
THE SWCHR LOGO
JOINING SWCHR
There are several versions of the SWCHR logo, all featuring the
Gray-Banded Kingsnake (Lampropeltis alterna), a widely-recognized
reptile native to the Trans-Pecos region of Texas as well as
adjacent Mexico and New Mexico.
For information on becoming a member please visit the
membership page of the SWCHR web site at
http://www.southwesternherp.com/join.html.
ON THE COVER: Southwestern Speckled Rattlesnake, Crotalus mitchellii pyrrhus,
Yuma County, AZ (Bill White). With this photograph, Bill won the SWCHR 2014
H. F. Koenig Award for Excellence in Herpetological Photography.
BACKGROUND IMAGE: Elephant Tusk, Big Bend NP, TX (Chris McMartin)
©2015 Southwestern Center for Herpetological Research. The SWCHR
Bulletin may not be reproduced in whole or in part on any web site or in any other
publication without the prior explicit written consent of the Southwestern Center
for Herpetological Research and of the respective author(s) and photographer(s).
SWCHR Bulletin
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Winter 2015
TABLE OF CONTENTS
A Message from the President, Tim Cole
46
Enhanced Health in Captive Snakes Begins with Proper Husbandry, Mark L. Heinrich MS, DVM
47
Husbandry Techniques for a Large Colony of Whiptail Lizards, Genus Aspidoscelis
(Lacertilia: Teiidae), David Jewell et al.
53
A Two-headed Sidewinder (Crotalus cerastes) and Review of Axial Bifurcation in Snakes
(Serpentes: Viperidae), Robert Twombley
57
Photographing Reptiles and Amphibians on a White Background, Nathan Hall
61
Notes on a Clutch of Eggs from a Buttermilk Racer, Coluber constrictor anthicus
(Serpentes: Colubridae), John Williams
63
A CALL FOR PAPERS
Are you a field herpetologist or a herpetoculturist working with species native to the American Southwest? Do you have a paper or an
article you have written for which you would like to find a permanent repository? Want to be assured you will always be able to share it
with the world? Submit it to the SWCHR Bulletin for possible publication. Submitted manuscripts from SWCHR members, as well as nonmembers, will be considered. There are NO page charges to have your articles appear in the SWCHR Bulletin, as some other publications
are now requiring.
To be accepted for publication, submissions must deal with herpetological species native to the American Southwest. Such topics as field
notes, county checklists, range extensions, taxonomy, reproduction and breeding, diseases, snake bite and venom research, captive breeding
and maintenance, conservation issues, legal issues, etc. are all acceptable. For assistance with formatting manuscripts, search ‘scientific
journal article format’ on the internet and tailor the resultant guidance to suit.
Previously published articles or papers are acceptable, provided you still hold the copyright to the work and have the right to re-publish it.
If we accept your paper or article for publication, you will still continue to be the copyright holder. If your submission has been previously
published, please provide the name of the publication in which it appeared along with the date of publication. All submissions should be
manually proofed in addition to being spell checked and should be submitted by email as either Microsoft Word or text documents.
Send submissions to swchrbulletin@swchr.org.
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Winter 2015
A Message from the President
We at SWCHR hope everyone had a great 2015! Since winter is generally a slow time for field herping, we close this
year’s volume of the SWCHR Bulletin with a focus on captive husbandry. When kept under appropriate conditions,
captive herps can provide insight useful in broadening our knowledge of various species. We have some very
interesting and informative articles in this issue:
- Mark Heinrich provides a primer of baseline considerations for maintaining healthy animals, with an emphasis on
snakes. But first, to put you in the proper frame of mind, you need to be listening to Mark’s Alterna Rush music CD
while reading the first article! Not only is Mark an accomplished herp veterinarian and herp keeper, but a talented
musician also. In addition to Alterna Rush and his follow-up Glass of Milk albums, Mark has also provided the music
for the annual Sanderson Snake Days dinner these past several years.
- Next, David Jewell and colleagues give us a glimpse of a colony of whiptails used for groundbreaking research into
the complex world of the various sexual and asexual (parthenogenic) species found in the SWCHR region of
interest. It's impressive to read about such a large colony of lizards being kept for research, and the majority of
them were produced in the facility.
- Who doesn’t want to see a two-headed Sidewinder! Robert Twombley shares an interesting article about a recent
specimen as well as a survey of axial bifurcation observations and discussion.
- For anyone who’s tried the “Meet Your Neighbors” style of photography, tips and tricks shared by professional
photographer Nathan Hall will be greatly appreciated.
- Most of us have taken in that wild herp for either photography, documentation, or that special breeding
project. It’s an added bonus when that animal lays a clutch of eggs or gives live birth while in our care. In our final
article this month, John Williams relates a brief natural history note not normally available on the Buttermilk Racer.
Enjoy this issue and thanks to our editor Chris McMartin for putting it together. We’re all looking forward to a
fruitful 2016 with many great herping adventures—both in the field and domestically!
SWCHR Bulletin
Enhanced Health in Captive Snakes Begins
with Proper Husbandry
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breeders or hobbyists with many animals may choose a simpler
setup employing wood shavings or paper. Whatever one
chooses one must consider the time constraint and labor
involved in sufficiently cleaning the enclosure in question.
by Mark L. Heinrich MS, DVM
Maintenance and care of snakes in captivity has become a
lifelong endeavor for many. Knowledge of life histories has
flourished over the last few decades making once difficult-tomaintain species readily available to the general public today.
Therefore, the number of people maintaining and interacting
with these animals is on the rise. Fortunately, in conjunction
with this increased interest, these animals are better understood
for their necessary role in the natural world. Continued
advances in captive husbandry and care are needed to further
solidify the rights of individuals who have a genuine passion for
these animals and wish to maintain them in captivity. Even
though snakes are more respected than in the past, more
appreciation is warranted for these deserving majestic creatures.
When maintaining a particular species, one must first understand
the optimal environmental and nutritional variables required by
the particular animal in question.
Habitat (cage setup),
temperature, humidity, and feeding parameters must be closely
matched to the natural environment in which a particular species
has evolved. Many snakes presented to veterinarians suffer from
ailments related to deficits in captive husbandry techniques.
This is a simple setup with paper substrate and a glass water bowl. Such a setup
is inexpensive, easy to clean, and increases efficiency while maintaining a number
of animals. Photo by the author.
Like most ‘herp people,’ I don’t remember not having an affinity
for reptiles, particularly ophidians. As a result I have maintained
various herps in captivity for much of my life. The following
tips and opinions were birthed from techniques that have
worked for me throughout the years. My philosophies are in no
way all-inclusive or represent the only way to do things, but they
are certainly laden with my personal experiences and biases.
Husbandry Basics
Proper husbandry begins with thorough and consistent cage
cleaning. Maintaining a consistently clean captive environment
can thwart many adverse health issues. I employ an initial wash
and rinse using dishwashing detergent followed by chlorhexidine
disinfection and another rinse. A weak sodium hypochlorite
solution, one part Clorox to twenty to thirty parts water, also
works well for disinfection. Make sure all particulate material is
removed from water bowls and cage surfaces—simply spraying a
disinfectant on fecal material does not eliminate all potential
disease-causing organisms. An automatic dishwasher also works
well for water bowls and small cages.
Various substrates are available today for use in enclosures.
Preferences depend upon the mission of the particular hobbyist
or professional. Zoological institutions may choose a very
elaborate setup with the purpose of an esthetically pleasing
display. A hobbyist with one or two animals may elect to
maintain a complex, elaborate setup as well. Professional
A plastic box, with an appropriately-sized hole in the lid, works well as a nest
box. Utilization of a plastic shoebox or sweater box is sufficient as a nesting area
for ophidian reproduction. Damp sphagnum moss works well as an egg-laying
substrate. The author has also used shredded computer paper and newspaper
successfully. Photo by the author.
In general, the more simple the setup, the less laborious the
effort when thoroughly and consistently cleaning captive
enclosures. I personally prefer simple setups. Paper products
provide a simple alternative when choosing a functional
substrate. One is not as tempted to only spot-clean when using
paper. Spot-cleaning shavings, dirt, or other three-dimensional
substrates risks leaving behind potential pathogenic bacteria,
viruses, or parasites. Mites are also more likely to flourish in a
substrate that provides three-dimensional structure for refuge
SWCHR Bulletin
and reproduction. Newspaper, napkins, or paper towels work
well. Empty paper towel or toilet paper rolls can be used for
enrichment and then discarded when soiled. Small cardboard
boxes may also be used as hides. Glass water bowls are easily
cleaned as are plastic ones, which can double as a hide.
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Nutrition
Proper nutrition is a hallmark of sustainable ideal health.
Feeding frequency and food types depend upon the time of year,
the species, and the individual’s body condition, health, and
reproductive status. Both underfeeding and overfeeding can
lead to adverse health effects. A thin individual, or ovulating or
pregnant female, may require increased frequency feeding
and/or larger meals. Food items must also be maintained on a
nutritionally sound diet. Improperly fed food animals (e.g., mice)
may lead to deficiencies and subsequent health issues in the
animals consuming such prey items. Feeding obese food items
may lead to major organ problems, including fatty liver.
Biological magnification may be an issue if contaminated food
items are routinely fed. Heavy metals and insecticides may
accumulate to toxic levels. After eating many meals containing
small traces of toxins, over a period of time, the toxin builds up
in the consumer’s tissues to a point where severe adverse health
conditions or death ensues.
A Black Pine Snake (Pituophis melanoleucus lodingi) laying eggs. Providing a small
amount of damp sphagnum moss is sufficient for this snake’s oviposition. Photo
by the author.
Environmental conditions need to be optimized when
maintaining a particular species in captivity. When choosing an
animal one can maximize the variables that have been
evolutionarily embraced by choosing a snake native to your
location, or one that lives in an area similar to your own. In
doing so, one also minimizes the time, money, and effort needed
for proper maintenance. All snakes are poikilothermic and
require varying degrees of heat depending on their species and
activity cycles. Temperate species as a general rule do well in
many environments. Some desert species, however, will not
thrive in tropical/subtropical conditions and vice versa. Know
the animal and its requirements well so the experiences of both
keeper and kept will be positive.
The Gray-banded Kingsnake (Lampropeltis alterna) has evolved to embrace a
desert environment. Photo by the author.
Food types are also a consideration. Know your species and
what they primarily eat in their free-living state. Some can be
conditioned to eating an unnatural or infrequently consumed
item. Take care that such an item will not adversely affect your
animal. Food size should also be considered. Small snakes will
potentially try to consume impossible-to-swallow large food
items, causing undue stress and possibly death. Deceased food
items pose less traumatic risk to your snake. If you choose to
feed live adult/subadult rodents, supervise your feeding process,
as severe mutilation and even death may result.
Assist-feeding may be warranted in certain situations. Some
snakes may lose their appetite at various times throughout their
yearly cycle or when diseased. Hatchlings may be healthy but
unwilling to take prey. Scenting with lizards, toads, fish, soaps,
etc. may entice a picky eater to take a meal. Sectioning a food
item or rubbing brain tissue on the head of a pinkie may
stimulate an animal to consume its prey.
This Western Hog-nosed Snake (Heterodon nasicus) hatchling would not consume
a toad- or fish-scented pinkie. It finally ate when offered a brain-scented pinkie.
Photo by the author.
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Winter 2015
Employing a small mosquito hemostat to assist-feed a hatchling Western Hognosed Snake (Heterodon nasicus). Start a well-lubricated (brain tissue or water)
newborn mouse headfirst into the snake’s mouth. Then grasp the mouse’s body
with a blunt-ended hemostat. Carefully and slowly advance the prey item into
the esophagus beyond the snake’s head. Gently loosen the hemostat’s grasp.
Pull it back and re-grasp the food item, advancing it ever so slowly until the item
is fully in the esophagus. This technique, as with any method of force-feeding,
requires great care to avoid damaging internal structures, especially the
esophagus. A torn esophagus will always lead to death. Photos by the author.
This hatchling California Kingsnake (Lampropeltis getula californae) would not
consume unscented or scented pinkies. It ate well after being offered a
longitudinally-sectioned fuzzy. Photos by the author.
Assist-feeding by force is sometimes needed especially in
diseased animals with no appetite. Various syringe/extruder
type instruments (‘pinkie pumps’) exist. Stainless steel feeding
tubes also provide an effective alternative to force-feeding.
Blunt small hemostats can also be employed when force-feeding
whole food items. Various finely ground nutritional aids are
available and work well when presented with the need to forcefeed. When using tube instruments, food items must be well
macerated or blended/liquefied to prevent internal trauma when
administering the meal. Whatever is used, be sure the
instrument is blunt and smooth so as not to cause harm to the
snake’s esophagus. A torn esophagus translates to a dead snake.
Force-feeding instruments. Top and bottom—pinkie pumps. Middle—stainless
steel feeding tube. Photo by the author.
Reproduction
One of the most fulfilling experiences in keeping snakes is the
occurrence of successful reproduction by animals under one’s
care. If the animals are maintained correctly and provided with
optimal nutrition and care then reproduction is inevitable.
Depending on the species, pairs are placed together at the
appropriate time, usually following a period of
brumation/hibernation. The author has recently allowed gravid
females outside access to bask in unadulterated ultraviolet (UV)
radiation provided by natural sunlight.
Hypothetically,
reproductive health may benefit due to increased vitamin D3
production necessary for proper calcium metabolism.
A gravid Great Plains Ratsnake (Pantherophis guttatus emoryi) basking naturally
outside. Photo by the author.
SWCHR Bulletin
As mentioned earlier, a nest box containing moist substrate (e.g.,
sphagnum moss) is necessary for oviposition. Eggs can then be
collected and moved to an appropriately-sized plastic container
containing moistened vermiculite. Place one or two small holes
in the lid of the container to prevent suffocation of neonates
after hatching. The container is then placed in an incubator and
eggs are incubated at the appropriate temperature—generally in
the mid 80s Fahrenheit. Livebearing herps do not require a nest
box per se but a hide box should be provided, at least for a sense
of security.
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Winter 2015
are considered normal flora in some snakes. Some of these
organisms are zoonotic and can be transferred from snakes to
humans. This fact dictates the need for proper hygiene in
conjunction with handling snakes. One must insist on hand
disinfection after handling snakes or any reptile, especially when
children and/or immunocompromised individuals are involved.
One should also adhere to this principle when interfacing with a
personal collection. Wash hands in between handling individual
animals or groups of animals to prevent the spread of disease.
Diseases can be transmitted directly from animal to animal, or
indirectly via your hand, instrument, poorly-cleaned water bowl
or cage, or even the transfer of an uneaten mouse from one
snake to another. Diseases may also be transmitted by bloodsucking invertebrates, including various insects, ticks, and mites.
Bacteria can cause disease locally (in the case of a localized
abscess) or systemically, in which the organisms flourish
throughout the diseased animal’s body. Various external visible
symptoms, including mouth rot, may only be part of a more
extensive systemic disorder. Once a particular bacterial infection
is diagnosed the attending veterinarian can prescribe the
appropriate antibiotic.
Milksnake (Lampropeltis triangulum) and Trans-Pecos Ratsnake (Bogertophis
subocularis) hatchlings. Photos by the author.
Disease
Pathogens include organisms that can be spread from animal to
animal, causing disease in susceptible individuals. When adding
a newly acquired snake to one’s collection it is recommended to
provide at least a two-month quarantine in a separate room.
While not always practical, one should adhere to a strict
quarantine to minimize the sometimes-disastrous results of
infectious disease spread. Potential pathogens include bacteria,
viruses, fungi, and various parasites. The presence of these
organisms in healthy animals does not always result in disease.
Some potential pathogens, including Salmonella and Pseudomonas,
New Mexico Milksnake (Lampropeltis triangulum celaenops). A) Cloacal abcess in a
NM milksnake. B) Caseated material surgically removed. This is the typical
appearance of a bacterial abscess in snakes. Photos by the author.
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Winter 2015
As with other potential pathogens, virus isolation from a snake
does not always discern etiology. Viruses, however, including
the inclusion body disease (IBD) virus and paramyxovirus, have
been documented to cause severe devastation in captive snake
populations. It is strongly recommended to seek out veterinary
advice and diagnostics if one suffers serious losses or senses a
serious illness in their collection. Unlike bacteria, viruses invade
an animal’s living cells and reproduce within them. Because of
this, they cannot be treated with antibiotics used to treat
bacterial, fungal, or parasitic diseases. Antiviral drugs are
expensive and not curative; therefore, treating viral infections
poses a tremendous—and many times ineffective—effort. If the
animal’s immune system cannot neutralize a given virus then a
terminal outcome may be inevitable.
This radiograph of a Mohave Rattlesnake (Crotalus scutulatus) shows severe bony
degenerative changes in the vertebral column. Bacteria, including Salmonella,
have been implicated in causing osteomyelitis lesions throughout skeletal tissues
in snakes. Photograph by the author.
Adverse conditions associated with old age and unknown
etiological agents also abound in captive-raised snakes. The
author maintains several colubrids in their twenties, and also
recently lost a Corn Snake (Pantherophis guttatus guttatus) which
lived to the age of thirty-two. Like other vertebrates, snakes can
exhibit senility changes including cataract formation, skin color
changes, and behavior changes consistent with neurological
degeneration. Neoplasia (cancer) is commonly encountered,
especially in older individuals. Some cancers are surgically
remedied and some prove to be fatal. Some may have a viral
etiology and some carry a cause that is unknown.
A young Ball Python (Python regius) with mouth rot. Mouth rot be a primary
infection, more commonly it is secondary to a more extensive pathological
condition. Photo by the author.
Fungal organisms are commensally associated with snakes and
are not generally a problem in captive collections. Parasites,
both external (e.g., mites) and internal (e.g., worms and
protozoans), are generally simple to diagnose by a veterinarian.
Specific drug and treatment protocols can then be utilized
successfully.
A 32-year-old Corn Snake (Pantherophis guttatus guttatus). Note the opacity of the
lens (mature cataract) and the loss of scale pigment from this once brilliantlycolored orange snake. Photo by the author.
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Winter 2015
Radiograph of the swelling. This snake also exhibited exostosis in its vertebral
column, possibly due to a nutritional issue while a young growing animal. Photo
by the author.
(Upper) This 15-year-old Gray-banded Kingsnake (Lampropeltis alterna) was
suffering from pericardial effusion. (Lower) The fluid was removed from around
the heart and the snake did well thereafter. No etiological agent was found in the
fluid. This condition was thought to be a result of trauma from a dislodged rock
that pinned the snake in its enclosure. Photos by the author.
Diagnosis: biliary adenocarcinoma, an aggressive tumor associated with the liver.
Photo by the author.
Mid-body swelling in a Sonoran Mountain Kingsnake (Lampropeltis pyromelana).
Photo by the author.
This 20-year-old axanthic Bullsnake (Pituophis catenifer sayi) was presented thin,
with a mid-body swelling. Photo by the author.
SWCHR Bulletin
Surgery on the bullsnake revealed a large mass associated with the intestine.
Histopathological diagnosis was an intestinal adenocarcinoma. Photos by the
author.
Predators, including the family cat or dog, can be a potential
hazard to your snake if allowed to interface with one another.
The author has seen several cases involving pet mammals
maiming or killing a pet snake. It is not recommended to allow
your cat or dog to interact with your snakes. Make sure your
cages are escape-proof to eliminate the possibility of a disastrous
meeting.
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Winter 2015
An unusual Trans-Pecos Ratsnake (Bogertophis subocularis) morph, consistent with
chimerism.
This specimen shows both normal and recessive blonde
morphological traits. Photo by the author.
Conclusion
Snakes provide an interesting alternative when considering a pet.
Many species may outlive our conventional mammalian pets. If
one properly cares for such an animal and seeks out veterinary
care when necessary, good health will abound, and the animal
will provide many years of enjoyment. Who knows? Maybe a
new morph will be conceived under your watch.
Husbandry Techniques for a Large Colony of
Whiptail Lizards, Genus Aspidoscelis
(Lacertilia: Teiidae)
by David Jewell, Alex Muensch, Christina Piraquive, Kristy
Winter, Richard Kupronis, Diana P. Baumann
Stowers Institute for Medical Research, Kansas City, Missouri
This escaped hatchling Pale Milksnake (Lampropeltis triangulum multistriata) had a
most unusual encounter with a Black Widow Spider (Latrodectus sp.) in the reptile
room. The neonate was removed from the web alive but died shortly thereafter.
Photo by the author.
Introduction
Whiptail lizards are fast, diurnal, insectivorous lizards that
inhabit desert grassland and shrubs. They are typically active for
only two to five hours per day during three to four months of
the year, depending on species and geographical location.
Within this genus there are approximately 50 species and many
subspecies, roughly a third of which reproduce via
parthenogenesis. All known parthenogens in this family have
arisen by hybridization of different sexually reproducing species.
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Winter 2015
The Desert Grassland Whiptail (Aspidoscelis uniparens), one of several species
maintained. Photo courtesy Stowers Institute for Medical Research.
Captive Colony
Currently our colony consists of four sexually reproducing
species (Aspidoscelis burti, A. gularis, A. inornata, and A. tigris), six
parthenogenetic species (A. exsanguis, A. neavesi, A. neomexicana,
A. sonorae, A. tesselata, and A. uniparens), and various hybrids.
Years of hard work, detailed observations, and husbandry
refinements have led to some species in the colony reaching the
12th generation in less than twelve years. This has significantly
reduced the need to capture animals from the wild. As of this
writing, the colony only contains approximately 3% wild caught
individuals. The colony size averages 750 animals.
Two of the three types of enclosures used: pens (children’s wading pools) and
runs (melamine wood). Photo courtesy Stowers Institute for Medical Research.
The third type of enclosure employed is shown in this view of 54-gallon
Rubbermaid® tubs. Photo courtesy Stowers Institute for Medical Research.
Hatchlings and juveniles are housed in 54-gallon Rubbermaid®
tubs. A mercury vapor bulb provides heat and UV light at one
end of the enclosure, creating a thermal gradient. Water is
provided in a three- or five-inch plastic saucer placed at the cool
end of the enclosure. The basking area is maintained between
110 and 120 degrees Fahrenheit during the day. Three or more
hiding places, in the form of pieces of cardboard or egg crate,
are provided for all enclosures and placed throughout the
thermal gradient.
Tub setup showing hide areas, water dish, basking spot, and egg-deposition PVC
tube. Photo courtesy Stowers Institute for Medical Research.
Housing
Three different types of housing are utilized: tubs, pens and
runs. Each type of enclosure is multi-functional, allowing
housing of lizards of different sizes. All enclosures are opentopped, eliminating the problem of ultraviolet (UV) rays being
filtered through enclosure lids. The enclosure types were
developed to house species that would not fare well in traditional
glass cages due to their strong flight response.
Lizards are housed at a density of up to 10 animals per tub.
Tubs are placed in rows on a sealed concrete floor. Tubs for
hatchlings are placed on rack shelving stacked two high. As
soon as animals become large enough to reproduce, a black
polyvinyl chloride (PVC) tube approximately 8 inches long and 3
inches in diameter is filled with moist sand and placed
horizontally in the enclosure for egg deposition.
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Winter 2015
Larger lizards are maintained in either a pen or a run depending
on the requirements of these particular animals. Pens are
constructed of two children’s wading pools, with one on the
ground and the other inverted, resting on the rim of the other,
with the bottom cut out. These two pools are held together with
cable ties or screws. This housing method works well with
lizards that have a tendency to run into walls when frightened.
The flexible plastic of the wading pool pens reduces the risk of
rostral abrasions, while allowing normal flight behaviors.
Melamine run setup. Photo courtesy Stowers Institute for Medical Research.
Lighting
Looking into a pen made from two children’s wading pools. Photo courtesy
Stowers Institute for Medical Research.
The runs are built of coated melamine wood with movable
dividers allowing flexibility in housing size. Both pens and runs
allow for the same types of water receptacle, hides, and general
configuration as the tubs, but they all have at least two basking
spots and a PVC tube filled with moistened sand for egg
deposition and additional humidity. These tubes are made of
black PVC (approximately 10 inches long with a 4-inch
diameter) with sanded edges. Tubes are positioned in the
enclosure with one end towards a basking spot and the other
towards the cooler end of the enclosure. This allows a gravid
female the ability to choose an oviposition site within a
temperature gradient within the tube, as well as giving all lizards
in an enclosure an area of higher humidity to aid in the shedding
process. The basking areas for runs and pens are kept at 120 to
130 degrees Fahrenheit. The warmer temperature allows gravid
females better thermoregulation options. All enclosures are
misted once daily to aid in shedding and to prevent dehydration.
Overhead fluorescent lighting provides a 12:12 on/off
photoperiod, with mercury vapor bulbs above the enclosures
coming on for nine hours during the ‘daylight’ hours. Room
temperatures are set to 83 degrees Fahrenheit during the day,
dropping to 70 degrees Fahrenheit at night. The colony is
housed in a room having windows on three sides, allowing
natural photoperiod changes throughout the year. The bulbs are
attached to metal struts with threaded rods suspended over the
enclosures. This design allows the basking lights to be adjusted
up or down to create the desired thermal gradient. Every
mercury vapor bulb’s UV output is tested prior to use with a
SpectraSuite® Spectrometer.
Seven whiptails (Aspidoscelis sp.) basking under a merucy vapor lamp. The lamp
height is adjustable to maintain the desired temperature gradient. Photo courtesy
Stowers Institute for Medical Research.
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Winter 2015
Feeding and Supplementation
Gut-loaded crickets are the primary food source with
mealworms being offered once a week. Feeding frequency
varies by enclosure; different colored laboratory tape is used to
indicate the size and quantities of food items to be fed in each
enclosure. This allows for efficient feeding, performed in the
morning while the lizards are most active.
A rotation of Miner-All, Herptivite™, and Rep-Cal® is used,
with each of these supplements being fed at least weekly.
Crickets (Acheta domestica) or mealworms (Tenebrio molitor) are
placed in a plastic bag with the supplement and gently shaken to
coat the insects prior to being fed to the lizards. Mealworms are
fed out approximately once a week to the all of the lizards that
would normally eat half-inch or larger size crickets. Waxworms
(Pyralidae) may be offered to thin or injured individuals to assist
with appetite restoration and weight gain.
A juvenile Aspidoscelis sp. Photo courtesy Stowers Institute for Medical Research.
Under these parameters, the fertile egg hatch rate is about 90%,
with eggs taking approximately 55 days to hatch. Hatchlings
weigh between 0.5 and 2 grams, with a snout-vent length of 1.1 2.3 inches and a total length of 3.1 - 4.7 inches, varying by
species. Hatchlings are weighed, measured, and photographed
on the day they hatch.
Another successful hatching event. Photo courtesy Stowers Institute for Medical
Research.
Eggs, Incubation, and Hatchlings
Every enclosure housing a female of reproductive size receives
an egg-laying tube which is emptied and refilled every four days.
This schedule was selected to balance minimizing the stress of
disturbing animals during the laying process with needing to add
new, moist sand, as the contents can become desiccated in five
days. When eggs are found they are sealed in an 8-ounce deli
cup half filled with medium grade vermiculite, mixed with water
1:1 by weight. Each deli cup is labeled with parental
information. The deli cups are placed in a large laboratory-grade
incubator set to 82 degrees Fahrenheit and 95% relative
humidity.
Side of tub showing cage cards for individual identification and management.
Photo courtesy Stowers Institute for Medical Research.
Animal Identification and Data Management
Cage cards are created for each animal recording the following
information: identification number, species, generation, hatch
date, and any unique identifying characteristics. Cage cards are
then paired with a photograph of each lizard and placed on the
outside of each enclosure to facilitate identification. As lizards
grow, the photographs are updated regularly. For species where
markings are too indistinct for visual identification, lizards may
be tagged with either a passive integrated transponder (PIT) or
visible implant elastomer (VIE) tag. A comprehensive Microsoft
Access database has been designed in-house to store all
SWCHR Bulletin
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Winter 2015
information relating to each animal, including health and
location logs, growth rates, genotyping information,
photographs, and other procedural data, in addition to tracking
parentage, lineage, and progeny.
A Two-headed Sidewinder (Crotalus cerastes)
and Review of Axial Bifurcation in Snakes
(Serpentes: Viperidae)
by Robert Twombley
Herein I describe the first reported case of axial bifurcation in
the Mohave Sidewinder (Crotalus cerastes cerastes). The mother of
the affected individual originated from the Mohave Desert in the
vicinity of Apple Valley, San Bernardino County, California at an
elevation of 2,946 feet (898 m) on April 17, 2015. The female C.
c. cerastes showed no sign of being gravid upon her capture.
When first placed within the enclosure, she went immediately to
the water dish where she spent an estimated two to three
minutes drinking. Such dehydration in C. c. cerastes is a direct
result of the drought conditions Apple Valley was experiencing
at the time of capture.
Two-headed Mohave Sidewinder (Crotalus cerastes cerastes). The neonate did not
survive. Photo by the author.
While the female gestated, she was given an ambient daytime
temperature of 80 degrees Fahrenheit, with a nighttime drop to
65 degrees Fahrenheit. A basking bulb created a hotspot of 100
degrees Fahrenheit. While she was offered prey items once
every seven days, she only consumed two meals during her
gestation period; one on May 14, 2015 and another on July 29.
On August 25, 2015 at 11:24 a.m. (birth took place sometime
between 8:00 am and 11:23 a.m.) I observed a total of seven
neonates, six of which had no noticeable malformation and are
considered to be heathy to this date. Upon observation of the
neonates, I opened up the enclosure to transfer the neonates to a
separate cage. All neonates successfully broke free of their
embryonic sac, save for one. When I carefully removed the sac
from this individual the teratological condition of axial
bifurcation became apparent. The neonate was then placed in its
own enclosure and monitored for approximately 57 minutes.
During this time no movement was observed and it became
apparent the animal was stillborn.
Materials and Methods
Ventral scales were counted according to Dowling (1951).
Anomalous half-ventrals were not counted, and partially divided
ventrals were counted as one scale. Overall length was obtained
using a piece of string which was then measured three times.
Head length was measured from the quadrate-articular jaw joint
to tip of snout, whereas snout length was measured from tip of
snout to anterior margin of orbit, using a flexible ruler.
Dorsal view of the specimen. Photo by the author.
Specimen Description
The axially-bifurcated C. c. cerastes individual’s overall condition
includes axial bifurcation, fusion of the ventral scales at the
fourth ventral scale, fusion at the neck with the ventral scale, and
fusion at the twenty-first ventral scale which caused the fusion of
the abdomen. A total of 74 ventral scales were observed and
counted.
The overall length is 1 7/8 inches (no snout-vent length was
taking due to inability to locate the vent). The left head was
under developed; its length was 1/4 inch and snout length was
3/8 inch. The left head was positioned directly under the right
head, which was fully developed. The length of the right head
was 5/16 inch and the snout length was 7/16 inch. Dorsal
coloration was gray with blotches of light brown along the dorsal
midline.
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Winter 2015
aquatic diapsid) fossil found within the Yixian Formation of
northeastern China (Buffetaut et al., 2007).
Another dorsal view of the neonate. The ‘button’ at the end of the tail is visible
coiled in the center. Photo by the author.
Types of Axial Bifurcation
The aberrancy of two heads within ophidian species is
scientifically referred to as axial bifurcation, dicephalism, and
somatodichotomy (Wallach, 2007). Smith and Pérez-Higareda
(1987) proposed the following seven terms to categorize axial
bifurcation in ophidians:
craniodichotomy, prodichotomy,
proarchodichotomy,
urodichotomy,
opisthodichotomy,
amphidichotomy, and holodichotomy. An explanation of each
follows:
- A craniodichotomous specimen has two incompletely divided
heads, a single atlas and axis, and a single body and tail.
- A prodichotomous snake has two complete heads, each with
an atlas and axis; either a single or two short necks; and a single
body and tail.
- A proarchodichotomous specimen has two heads, two long
necks, and a single body and tail.
- A urodichotomous snake has one head and body but two tails.
- An opisthodichotomous specimen has one head, two bodies,
and two tails.
- An amphidichotomous snake has two heads, a single body, and
two tails.
Holodichotomy refers to a pair of twins from a single egg,
usually healthy and normal but reduced in size in comparison
with their siblings (Wallach and Salmon, 2013).
Causes of Axial Bifurcation
Axial bifurcation is not an unfamiliar condition within
herpetofauna. The first known example of axial bifurcation
within Reptilia is of a 120 million year old choristoderan (semi-
Lateral/ventral view of the deceased snake. Photo by the author.
Eleven possible causes have been proposed for the teratological
abberancy known as axial bifurcation:
1) Incomplete division of a single embryo. Wilder (1908)
considered axial bifurcation to occur in this manner.
2) Partial fusion of two embryos.
Attributed cases
include Vipera berus (Dorner, 1873), Crotalus durissus (Vanzolini,
1947), and Thamnophis sirtalis (Wallach, 2007).
3) Abnormally low or high temperatures during incubation or
gestation. Cases resulting from known suboptimal incubation
temperatures include Elaphe schrencki (Bakken and Bakken, 1987),
Natrix natrix (Allen, 1990), Bothrops moojeni (de Andrade and Abe,
1992), Crotalus atrox (Muir, 1990), and C. viridis (Pendlebury,
1976). Conversely, cases implicating elevated temperatures
include Natrix natrix (Riches, 1967), Python molurus (Vinegar,
1973), Nerodia fasciata (Osgood, 1978), Lampropeltis getula (Zweifel,
1980), and Pantherophis alleghaniensis (Ball, 1995). Gutzke and
Packard (1987) found abnormalities in Pituophis sayi to be eight
times as common at 32 degrees Celsius than at 27 degrees
Celsius.
4) Tornier (1901) and Thomson (1935) suggest axial bifurcation
may occur due to regeneration after an embryonic lesion.
5) Anoxia during embryonic development. Stockard (1921)
demonstrated that anoxia (low oxygen supply) during the
gestation period of fishes will result in the production of twoheaded and double-bodied offspring, as well as twin individuals
from a single egg.
6) Toxic effect of metabolic secretions during a prolonged
sojourn in the oviduct.
Cases include Platyceps
SWCHR Bulletin
florulentus (Heasman, 1933) and Pantherophis guttatus (Wallach,
2007).
7) Inbreeding depression from small population gene pools,
back-crossing, designer morphs, and albinos, such as in captive
collections. A Pantherophis guttatus produced from a cross
between albino and hypomelanistic parents was found among 22
siblings that also included albino, hypomelanistic, and
anerythristic hatchlings. Another dicephalic Pantherophis guttatus
originated from the breeding of two ‘snow’ morphs (Alex Hue,
pers. comm.). Two albino dicephalic specimens resulted from
matings with a pair of albino parents. “Medusa III” was
produced from two amelanistic albino Pantherophis obsoletus
(Wallach, 2007). Another, “Hammerhead,” originated from two
albino Crotalus atrox (Muir, 1990). The mating of a pair of albino
sibling Lampropeltis getula californiae produced a clutch of eight
eggs; the four which hatched were two albino and two normallypigmented
individuals,
one
of
the
latter
being
opisthodichotomous (Wallach, unpubl.). A pair of identical
female conjoined twin Pantherophis guttatus born with their hearts
outside the body were the progeny of a cross between a ‘reverse
Okeetee’ morph and a striped heterozygous albino (Wallach,
unpubl.). The 16 progeny from the cross of a ‘creamsicle’ and
‘snow’ Pantherophis guttatus were three ‘snows’ and 13
‘creamsicles,’ one of which was dicephalic (Tim Curran, pers.
comm.). A dicephalic Lichanura trivirgata was born from the
mating of two 100% heterozygous Limburg-strain albino
Lichanura trivirgata (Wallach, 2007).
8) Hybridization. Newman (1917) demonstrated this in fish,
through hybridization within the infraclass Teleostei, Fundulus
heteroclitus x Fundulus diaphanus, which resulted in axial bifurcation
(known colloquially as “double monsters”). Other examples
from hybridization in Serpentes are Lampropeltis alterna x L.
mexicana which resulted in an opisthodichotomous snake, with a
single head but double bodies and tails (Ball, 1995). Another
example is Lystrophis pulcher x L. mattogrosssensis (Wallach, 2007).
A Lamprophis lineatus x L. fuliginosus cross also resulted in two
dicephalic offspring and one set of twins in 2000 (Donny
Herring, pers. comm.). An additional specimen resulted from a
Lampropeltis mexicana thayeri x L. ruthveni cross (Wallach, 2007).
9) Environmental pollution. Gray et al. (2001) observed
various developmental anomalies in wild Thamnophis sirtalis from
an 80-acre hazardous waste site used as a landfill from 1941 to
1981, in Millcreek Township near Erie, Pennsylvania. Additional
examples exist in captive-bred Thamnophis sirtalis from the same
area (Gray et al., 2003).
10) Chemical toxins in captivity. Unpublished reports of
dicephalism occurring after use of chemicals in captivity include
Lamprophis fuliginosus with ‘Mr. Clean,’ Pantherophis guttatus with
‘Shell No-Pest Strips,’ and Thamnophis sirtalis with ‘Vapona
Insecticide Strips’ (Wallach, 2007).
11) Exposure to radiation. The spontaneous mutation rate in
vertebrates is doubled with 30-60 roentgens of radiation,
whereas the lethal dose is 800-1000 roentgens (Sachsse, 1983;
Wallach, 2007).
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Winter 2015
Several axially-bifurcated lower vertebrates have been produced
within a laboratory setting using these methods. While the
majority of produced specimens have been fish (Kellicott, 1916;
Newman, 1917; and Werber, 1915 and 1916), one lizard has also
been produced (Wallach, 2007). There has been a single,
secondhand report of an axially-bifurcated ophidian being
artificially produced at the Cincinnati Zoological Gardens facility
(Abe, 1952). However, Wallach (2007) was unable to confirm
this statement and would like to learn if it is true.
Healthy litter-mates of the two-headed Mohave Desert Sidewinder (Crotalus
cerastes cerastes). Photo by the author.
Within the genus Crotalus there have been 34 reported instances
across 11 species documented as having axial bifurcation to
date: Crotalus adamanteus, C. atrox, C. basiliscus, C. durissus, C.
horridus, C. lutosus, C. mitchelli, C. oreganus, C. scutulatus, C. tigris, and
C. viridis (McAllister and Wallach, 2006). With this publication
C. cerastes may now be included in that list, which brings the
count up to 35 reported instances across 12 species within the
genus. It is estimated that only 1 in 50,000 wild rattlesnakes are
affected by this mutation (Allen, 1956).
References
Allen, A. “Two-headed Snakes.” Aquarist & Pondkeeper, August
1990, p. 20.
de Andrade, D. V., and A. S. Abe. “Malformações em Ninhadas
de Caiçaca, Bothrops moojeni (Serpentes: Viperidae). Memórias do
Instituto de Butantan 54(2), 1992, pp. 61-67.
Ball, J. C. 1995. “Axial Bifurcation. Case Study: A Two-headed
Yellow Rat Snake.” Reptiles and Amphibians Magazine 32, 1995, pp.
36-43.
Bakken, D. J. and L. A. Bakken. “Dicephalism in the Russian
Rat Snake, Elaphe schrencki schrencki.” Bulletin of the Chicago
Herpetological Society 22(1), 1987, p. 2.
SWCHR Bulletin
Buffetaut E., Jianjun L., Haiyan T., and He Z. “A Two-headed
Reptile from the Cretaceous of China.” Biology Letters 2007(3),
pp. 80-81.
Dorner, H. “Eine Kreuzotter mit Zwei Köpfen.” Der Zoologische
Garten 14(11), 1873, pp. 407-410.
Dowling, H. G. “A Proposed Standard System of Counting
Ventrals in Snakes.” British Journal of Herpetology 1, 1951, pp. 9799.
Gray, B., H. M. Smith, J. Woodling, and D. Fischer. “Some
Bizarre Effects on Snakes, Supposedly from Pollution at a Site in
Pennsylvania.” Bulletin of the Chicago Herpetological Society 26(7),
2001, pp. 144-148.
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Winter 2015
Vago and G. Matz (eds.). Centre National de la Recherche
Scientifique, Université d’Angers, 1983, pp. 197-205.
Smith, H. M. and G. Perez-Higareda. “The Literature on
Somatodichotomy in Snakes.”
Bulletin of the Maryland
Herpetological Society 23(4), 1987, pp. 139-153.
Stockard, C. R.
“Developmental Rate and Structural
Expression:
An Experimental Study of Twins, ‘Double
Monsters’ and Single Deformities, and the Interaction among
Embryonic Organs during Their Origin and Development.”
American Journal of Anatomy 28(2), 1921, pp. 115-266.
Thomson, J. A. Biology for Everyman, Vol. 1. New York: E. P.
Dutton & Co., 1935.
Gray, B., H. M. Smith, and D. Chiszar. “Further Anomalies in
the Litters of a Garter Snake from a Hazardous Waste Site.”
Bulletin of the Chicago Herpetological Society 38(1), 2003, pp. 4-6.
Tornier, G. “Überzählige Bildungen und die Bedeutung der
Pathologie für die Biontotechnik.”
Verhandlungen des
internationalen Zoologen—Kongresses, 1901, pp. 491-492.
Gutzke, W. H. N. and G. C. Packard. 1987. “Influence of the
Hydric and Thermal Environments on Eggs and Hatchlings of
Bull Snakes Pituophis melanoleucus.” Physiological Zoology 60(3), 1987,
pp. 9-17.
Vanzolini, P. E. “Notas Sôbre um Deródimo de Crotalus durissus
terrificus (Laur.).” Papéis Avulsos de Zoología 8(24), 1947, pp. 273283.
Heasman, W. J. “The Anatomy of a Double-headed Snake.”
American Journal of Anatomy 67(2), 1933, pp. 331-345.
Vinegar, A. “The Effects of Temperature on the Growth and
Development of Embryos of the Indian Python, Python molurus
(Reptilia: Serpentes: Boidae).” Copeia 1973(1), pp. 171-173.
Kellicott, W. E. “The Effects of Low Temperature upon the
Development of Fundulus.” American Journal of Anatomy 20(3),
1916.
Wallach, Van. “Axial Bifurcation and Duplication in Snakes.
Part I. A Synopsis of Authentic and Anecdotal Cases.” Bulletin of
the Maryland Herpetological Society 43(2), 2007, pp. 57-95.
McAllister, Chris T. and Van Wallach. “Discovery of a
Dicephalic Western Diamondback Rattlesnake, Crotalus atrox
(Serpentes: Viperidae), from Texas, with a Summary of
Dicephalism Among Members of the Genus Crotalus.” Arkansas
Academy of Science Journal 60, 2006, pp. 67-73.
Wallach, Van, and Gerald T. Salmon. “Axial Bifurcation and
Duplication in Snakes. Part V. A Review of Nerodia sipedon
Cases with a New Record from New York State.” Bulletin of the
Chicago Herpetological Society 48(8), 2013, pp. 102-106.
Muir, J. H. “Three Anatomically Aberrant Albino Crotalus atrox
Neonates.” Bulletin of the Chicago Herpetological Society 25(3), 1990,
pp. 41-42.
Newman, H. H.
“On the Production of Monsters by
Hybridization.” The Biological Bulletin 32(5), 1917, pp. 306 -321.
Osgood, D. W. “Effects of Temperature on the Development
of Meristic Characters in Natrix fasciata.” Copeia 1978(1), pp. 3347.
Pendlebury, G. B. 1976. “Congenital Defects in the Brood of a
Prairie Rattlesnake.” Canadian Journal of Zoology 54, 1976, pp.
2023-2025.
Riches, R. J. “Notes on a Clutch of Eggs of the Viperine Snake
(Natrix maura).” British Journal of Herpetology 4(1), 1967, pp. 14-16.
Sachsse, W. “Inheritance and Environment as Causes for
Teratogenesis in Amphibians and Reptiles.” Proceedings of the First
International Colloquim on Pathology of Reptiles and Amphibians, C.
Werber, Ernest I. “Is Defective and Monstrous Development
Due to Parental Metabolic Toxaemia?” The Anatomical Record
9(1), 1915.
_____. “Experimental Studies Aiming at the Control of
Defective and Monstrous Development. A Survey of Recorded
Monstrosities with Special Reference to the Ophthalmic
Defects.” The Anatomical Record 9(7), 1915, pp. 561-562.
_____. “Experimental Studies on the Origin of Monsters. I. An
Etiology and an Analysis of the Morphogenesis of Monsters.”
Journal of Experimental Zoology 21, 1916, pp. 485-584.
Wilder, H. H. “The Morphology of Cosmobia; Speculation
Concerning the Significance of Certain Types of Monsters.”
American Journal of Anatomy 8, 1908, pp. 355-440.
Zweifel, R. G. “Aspects of the Biology of a Laboratory
Population of Kingsnakes.” Reproductive Biology and Diseases of
Captive Reptiles, J. B. Murphy and J. T. Collins (eds.). Ithaca:
Society for the Study of Amphibians and Reptiles, 1980, pp. 141152.
SWCHR Bulletin
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Winter 2015
Photographing Reptiles and Amphibians on a
White Background
by Nathan Hall
Photographing reptiles and amphibians on a white background is
easy. Unfortunately, photographing those subjects on a white
background and keeping the background pure white is
exceedingly difficult, unless one is proficient in background
removal techniques like deep etching, clipping path, and
compositing the subject/image from the background. Whether
showcasing available specimens for sale or simply for the
purpose of displaying them on a pure white background, several
steps should be followed to ensure the specimens are exposed
properly and the background is pure white.
I created this tutorial to address the issues some photographers
have when photographing herps on a white background, without
having to use tedious techniques to make the background pure
white. The background will probably be a shade of light to dark
gray if the subject is properly exposed, or the subject will be
completely blown out if the background is clipped to pure white.
I’ve spent many photo sessions banging my head against the wall
in frustration because I just couldn't get the background pure
white without overexposing the subject. After years of trial and
error, I finally figured out a way to keep the subject properly
exposed and the background perfectly white.
Now, one can always outsource the image to a plethora of
clipping-path companies that will gladly deep etch your image
and make the background whatever color you want for a fee.
Pure white is the standard, but they can always match the
background to your web site or any other final destination.
When shooting products for catalogs, eBay, Amazon, or any
other company requiring a pure white background, I use a
clipping-path company to deep etch my images. I do not want
shadows on the products, and my technique doesn't work as well
with completely removing shadows. Conversely, I believe
shadows are important when showcasing specimens, simply to
give them dimension so they do not look completely flat. I see a
lot of reptile and amphibian images where the shadows are
completely removed in post-production, and in my opinion the
images often look heavily processed.
Equipment and Software
There’s absolutely no need to have all of the more expensive
equipment I use in order to achieve great results. I’m a
professional photographer, and it’s my livelihood. I’m simply
sharing what materials and computer programs I use to shoot
reptiles and amphibians on a white background.
An example of the results achievable using techniques presented in this article.
The flighty nature of many herp subjects, such as these Texas Banded Geckos
(Coleonyx brevis), necessitates handheld photography to maintain focus. Photo by
the author.
I’m a Canon guy, and I’ve always been a Canon guy because I
started my photography career with Canon equipment and am
most comfortable using it. When shooting macro photos of
herps, one will need a good camera body and macro lens. I
shoot with a Canon 5D Mark III and Canon EF 100mm f/2.8L
Macro IS USM lens.
An off-camera flash with a large diffuser is fundamentally
important when doing this technique. I use an Elinchrom
BXRi-500 strobe with an Elinchrom 53-inch Midi Octa softbox.
The strobe has a built-in wireless trigger, so I can fire the strobe
wirelessly. The strobe is 500 watts, and the softbox creates
beautiful wrapping soft light, which is important when shooting
subjects on a white background—it minimizes hard shadows. I
started my photography career with Paul C. Buff’s strobes and
diffusers/modifiers, including AlienBees and White Lightning
strobes. Paul’s products are relatively inexpensive and perfect
for those venturing into studio photography.
I use a C-stand and boom/extension arm for the strobe and
modifier (I’ll go into more detail later). The boom/extension
arm is important so the modifier can be placed over the
subject(s). The only other materials needed are a shooting table
(I use a 4 foot by 2 foot Rubbermaid table, but a sheet of
plywood and two plastic sawhorses work just as well), a 4 foot
by 8 foot sheet of white Formica, two A-clamps, and two 8-to12-foot light stands. I use Adobe Camera RAW (Lightroom has
the same Develop Module) and Photoshop CS6 to post-process
all of the RAW images.
SWCHR Bulletin
Preparation
First, I set up the shooting table, and clamp the white Formica
sheet to the two light stands with the A-clamps to form a
‘sweep.’ The stands are positioned about 3 feet apart since the
shooting table is 2 feet wide. The light stands are about 6.5 feet
high. Once clamped, the Formica sweep covers the table to the
end. If the light stands are much lower, there will be excess
Formica not supported by the table, and it may crack.
Setup the Formica sheet using light stands and A-clamps to form a sweep.
Photo by the author.
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Winter 2015
Now that the table and white sweep are set up, it’s time to
position the light and modifier/diffuser. I position the modifier
over the sweep, slightly angled to illuminate the background.
The modifier is about 2 feet from the sweep on the low end and
about 3 feet on the high end. The slight angle creates just
enough fill to minimize shadows and heat up the background.
I’ve tried positioning the light at the classic 45/45 commonly
used with studio portraiture photography, but I’ve found that a
lesser angle works better. I’ve also tried utilizing additional
lights, with limited success. There are unlimited ways to shoot
macro in the studio—this is how I do it. My technique is simple
and effective and can be replicated by any home photographer
on a tight budget.
The modifier is carefully positioned to provide optimum lighting. The angle is
different than that used for standard portrait photography (see text). Photo by
the author.
Photography
From this angle, it is readily apparent the curvature of the Formica eliminates
unsightly corners (and shadows) in your photographs. Photo by the author.
With setup complete, it’s time to shoot! The subject of this
shoot and tutorial is the Texas Banded Gecko (Coleonyx brevis).
It’s a challenging little gecko to photograph, so I’m going to
make it even more difficult by photographing an adult and
juvenile at the same time—I’m a glutton for punishment. I
crank the strobe all the way up to full power since I’m typically
stopped down to f/16-f/22 (f/18 for this shoot). That’s 500
watts of light! I want detail throughout the subject, which is why
I’m shooting at such a small aperture. My shutter speed is 1/125
second. I shoot reptiles and amphibians handheld, because
they're constantly on the move, and a tripod is impractical when
shooting moving subjects. I want as clean an image as possible,
so ISO is always 100. I always shoot RAW opposed to JPEG to
have a lossless file. I feather the light. I place the subject on the
sweep so the light hits the subject from the edge of the modifier
and not the hotspot in the middle where the bulb is.
SWCHR Bulletin
Processing
63
Winter 2015
are many ways to achieve similar results. I hope this tutorial is
helpful. Now get out there and shoot!
After I upload the images to my computer, I open them in
Adobe Camera RAW CS6 (Lightroom has the same Develop
Module, but it’s called the Basic Panel in ACR) and begin by
adjusting the White Balance Temperature slider.
That’s
fundamentally important to the overall success of the image. I
know my Elinchrom strobes are about 5050-5150K, but that
won’t cut it. I use a gray card when shooting herps in the studio
to nail white balance. I use the White Balance Eyedropper Tool
in ACR to sample the Camera RAW White Balance (18% gray)
swatch on my gray card. It works like a charm, and I nail white
balance every time. I rarely adjust the Tint slider.
The finished product—a portrait of adult female and juvenile Texas Banded
Geckos (Coleonyx brevis). Photo by the author.
Gray card used for white balance. “Camera RAW White Balance” (18% gray) is
in the lower left. Photo by the author.
Next, I work down the sliders in the Basic Panel (Develop
Module in Lightroom). After the White Balance sliders is the
Exposure slider. I always try to do as much in-camera as
possible, but I sometimes need to adjust exposure in post. The
next slider I adjust is the Highlights slider. I usually crank the
Highlights slider almost all of the way to the right (100) to begin
blowing out the background. Again, the goal is to have the
subject properly exposed and completely blow out the
background to pure white. The next slider down is the Shadows
slider. I adjust that slider if I need to open up any shadows and
add some fill light.
Notes on a Clutch of Eggs from a Buttermilk
Racer, Coluber constrictor anthicus
(Serpentes: Colubridae)
by John Williams
On the evening of May 28th, 2008, I collected an adult female
Buttermilk Racer (Coluber constrictor anthicus) beneath discarded
trash in northeastern Harris County, Texas. With fading light, I
decided to keep the snake overnight for release at the site in the
morning after taking photographs. I placed the snake in a small
Tupperware-type box with loose bark and a coconut fiber
substrate.
The next slider is the Whites slider. This is the most important
slider when trying to get the background pure white. I crank the
slider to the right until the entire background clips completely
without losing any detail in the subject. The Blacks slider is next,
and I usually move it to the left to bring back the blacks washed
out when I cranked the other sliders. Moving down, the next
slider is Clarity, and I use it sparingly. A setting of +10 is about
as high as I go with that slider. The last slider I adjust is the
Vibrance slider, and I’m pretty conservative with that slider as
well. I crop the image, then output sharpen for web or print.
I learned these methods by experimenting with different
workflows over the years, and I’ve found this workflow to be the
quickest, easiest, and most reliable when photographing reptiles
and amphibians on a white background. As mentioned, there
Female Buttermilk Racer (Coluber constrictor anthicus) with her recently-laid clutch
of 12 eggs. Photo by the author.
SWCHR Bulletin
64
Winter 2015
The next morning I discovered the snake had laid 12 eggs
overnight beneath one of the pieces of loose bark. After quick
photographs I subsequently released her at the site of capture. I
kept the eggs in an outdoor storage closet. Though not
measured, temperatures were likely in the low 80s Fahrenheit. I
periodically misted the coconut fiber when it dried out.
A hatchling racer slowly makes its way out of the egg. Photo by the author.
The entire clutch with a U.S. quarter for size comparison. Photo by the author.
On July 18th and 19th (51-52 days after being laid), 10 of the 12
eggs successfully hatched, most within two hours of each other.
The remaining two eggs were examined and appeared to contain
late stage stillborns. Babies measured between 9 and 11 inches
total length. The following week the hatchlings were released
beneath the trash where the female was discovered.
A recently-hatched Buttermilk Racer. Photo by the author.
A hatchling racer pips through its eggshell. Photo by the author.
Another view of the hatchling. Photo by the author.
SWCHR CODE OF ETHICS
As a member of the Southwestern Center for Herpetological Research, I subscribe to
the Association’s Code of Ethics.
Field activities should limit the impact on natural habitats, replacing all cover objects,
not tearing apart rocks or logs and refraining from the use of gasoline or other toxic
materials.
Catch and release coupled with photography and the limited take of non-protected
species for personal study or breeding use is permitted. The commercial take and sale
of wild-caught animals is not acceptable.
Collecting practices should respect landowner rights, including but not limited to
securing permission for land entry and the packing out of all personal trash.
Captive-breeding efforts are recognized as a valid means of potentially reducing
collection pressures on wild populations and are encouraged.
The release of captive animals including captive-bred animals into the wild is
discouraged except under the supervision of trained professionals and in accordance
with an accepted species preservation or restocking plan.
The disclosure of exact locality information on public internet forums is discouraged in
most circumstances. Locality information posted on public internet forums usually
should be restricted to providing the name of the county where the animal was found.
When specific locality data is provided to one in confidence, it should be kept in
confidence and should not be abused or shared with others without explicit
permission.
Other members of the Association are always to be treated cordially and in a respectful
manner.
SWCHR
PO BOX 624
SEGUIN TX 78156