Protocols for Husbandry and Embryo Collection of a Parthenogenetic Gecko, Lepidodactylus lugubris (Squamata: Gekkonidae)

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Herpetological Review 49(2), 2018
230 TECHNIQUES
Herpetological Review, 2018, 49(2), 230–235.
© 2018 by Society for the Study of Amphibians and Reptiles
Protocols for Husbandry and Embryo Collection
of a Parthenogenetic Gecko, Lepidodactylus lugubris
(Squamata: Gekkonidae)
Lizards and snakes (squamate reptiles) have become
increasingly used in developmental biology research, resulting
in the establishment of several “model” lizard clades or species
(e.g., Sanger et al. 2008; McLean and Vickaryous 2011; Diaz et
al. 2017; Infante et al. 2018; Londono et al. 2017; Sanger and
Kircher 2017). When choosing a species or clade to study for
developmental questions, several criteria must be met. First,
a species or group of species must be identified which exhibit
the genotype or phenotype of interest. Second, practical criteria
must also be considered. The species or group of species must be
available for experimentation or observation in the laboratory.
Preferably, the species would be available to purchase or easy
to obtain from wild populations, easily housed in a laboratory
setting with standardized husbandry protocols, have a high
fecundity, and have additional resources for investigating
developmental questions, such as a sequenced genome or
transcriptomes.
The gecko bauplan (overall collection of morphological
features) is largely considered plesiomorphic (similar to the
ancestral form) among squamates (Conrad 2008). Yet geckos
also exhibit extremely derived morphologies, such as numerous
independent evolutions of adhesive toe pads (e.g., Gamble et al.
2012; Russell et al. 2015). This combination of morphological
conservation and novelty, as well as geckos’ utility as a system
to study convergent evolution (Gamble et al. 2012, 2015a,
2015b), makes them an excellent model clade for developmental
evolutionary biology. Their phylogenetic position as the sister
clade to all other lizards and snakes, with the possible exception
of the Dibamidae (Zheng and Wiens 2016), means that evo-devo
studies that include a gecko and almost any other lizard or snake
species will have encompassed the phylogenetic breadth of all
squamates.
Of more than 1700 described gecko species (Uetz et
al. 2017), Leopard Geckos (Eublepharis macularius) and
Madagascar Ground Geckos (Paroedura picta) have been used
to study developmental questions (Noro et al. 2009; McLean
and Vickaryous 2011). However, another gecko species stands
out as an ideal model to study developmental questions, the
Mourning Gecko (Lepidodactylus lugubris). Lepidodactylus
lugubris is small-bodied (approximately 40–44 mm snout–vent
TECHNIQUES
TECHNIQUES
AARON H. GRIFFING*
Department of Biological Sciences, Marquette University,
P.O. Box 1881, Milwaukee, Wisconsin 53201, USA
THOMAS J. SANGER
Department of Biology, Loyola University in Chicago,
1032 W. Sheridan Road, Chicago, Illinois 60660, USA
ITZEL C. MATAMOROS
Department of Biological Sciences, Marquette University,
P.O. Box 1881, Milwaukee, Wisconsin 53201, USA
STUART V. NIELSEN
Department of Biological Sciences, Marquette University,
P.O. Box 1881, Milwaukee, Wisconsin 53201, USA
TONY GAMBLE
Department of Biological Sciences, Marquette University,
P.O. Box 1881, Milwaukee, Wisconsin 53201, USA
Bell Museum of Natural History, University of Minnesota,
Saint Paul, Minnesota 55108, USA
Milwaukee Public Museum, 800 W. Wells Street, Milwaukee,
Wisconsin 53233, USA
*Corresponding author: e-mail: aaron.gring@marquette.edu
Fig. 1. Phylogenetic relationships of gekkotan families. Red mark in-
dicates the evolution of hard-shelled eggs. Chronogram is scaled to
millions of years and modified from Gamble et al. (2015b).

Herpetological Review 49(2), 2018
TECHNIQUES 231
length [SVL; Röll 2002]), with a widespread native distribution
(India, Sri Lanka, southeast Asia, Indonesia, the Philippines,
and nearly all Pacific islands, including the Hawaiian Islands;
Bauer and Henle 1994) and multiple introduced populations in
the Neotropics (Florida, USA; Central America; northern South
America; and the Galapagos; Schauenberg 1968; Henderson et
al. 1976; Hoogmoed 1989; Krysko et al. 2011; Hoogmoed and
Avila-Pires 2015). Lepidodactylus lugubris is parthenogenetic,
that is, an all-female species that reproduces in the absence of
males, with mothers producing genetically identical daughters
(Cuellar and Kluge 1972). Several L. lugubris clonal lineages have
been described and each is thought to derive from a unique
hybridization event between Lepidodactylus moestus and an as
of yet undescribed Lepidodactylus species (Radtkey et al. 1995).
Occasional backcrosses between the diploid L. lugubris (2n=44)
and one of the parental species results in triploid L. lugubris
clones (3n=66; Moritz et al. 1993; Radtkey et al. 1995). Between
five and 16 clonal lineages have been described (Ineich 1988;
Moritz et al. 1993; Yamashiro et al. 2000; Ineich 2015) based on
dorsal pattern variation, karyotypes, and allozyme variation
(Ineich 1988; Moritz et al. 1993). While the exact mechanisms
of reproduction in L. lugubris remain unknown, in other
parthenogenetic lizards the number of chromosomes doubles
prior to meiosis leading to mature diploid oocytes (Darevsky et al.
1985; Lutes et al. 2010). Although L. lugubris is parthenogenetic,
male phenotypes are occasionally encountered in the wild and
in captivity (Schauenberg 1968; Cuellar and Kluge 1972; Ineich
and Ota 1992; Brown and Murphy-Walker 1996; Röll and von
Düring 2008; Trifonov et al. 2015). However, these occasional
males appear to be infertile, either lacking mature spermatozoa
or possessing deformed spermatozoa (Yamashira and Ota 1998;
Röll and von Düring 2008).
Parthenogenetic organisms are ideal laboratory animals
for developmental studies because there is no need for
mate-pairing, every individual is reproductively active, and
individuals within clonal lineages are genetically identical.
However, few parthenogenetic reptiles are routinely bred and
maintained in laboratory settings (Cole and Townsend 1977;
Darevsky et al. 1985; Maslin 1971; Kearney and Shine 2004;
Lutes et al. 2011). Lepidodactylus lugubris has other desirable
characteristics including high fecundity, ease of captive care,
and fast maturation. Furthermore, this species is easily available
via targeted field collection or through the pet trade, space-
efficient to keep, and lays hard-shelled eggs (Fig. 1), making it
tractable to perform embryological work. Herein we describe
detailed methods for the laboratory maintenance of captive
L. lugubris and embryo collection to serve as resources for
researchers investigating developmental morphology, sexual
development, parthenogenesis, and cytogenetics. These
protocols will serve as a foundation for laboratory research on L.
lugubris to accompany forthcoming genetic and embryological
resources.
huSbandry
Source of animals.—To establish a laboratory colony, we
collected 20 wild adults from populations in the Hawaiian
Islands (USA) in 2009 and 2012 under permit from State of Hawaii
Division of Forestry and Wildlife (Permits: EX09-06, EX12-08).
Clones A, B, and C are present in Hawaii (Fig. 2; Stejneger 1899;
Cuellar 1984; Zug 2013); however, C clones are markedly rarer
(Moritz et al. 1993). Lepidodactylus lugubris of either A clone or B
clone varieties are also readily available in the U.S. and European
pet trade (pers. obs.).
Housing, humidity, temperature, and cleaning.—We keep
between 2–5 adults housed together in a single enclosure.
Because this species is parthenogenetic, issues concerning sex
ratio are nonexistent, and thus, any combination of individuals
of roughly the same size can be housed together. However, we
suggest placing individuals of the same clone types together,
which will allow for easier allocation of clone types for clone-
specific research. Furthermore, we have observed that A clones
may behave aggressively toward spotted clone B types resulting
in weight loss in the B clones and the need to keep those clones
separately. Lepidodactylus lugubris can exhibit intraspecific
aggression comparable to behaviors observed in other small
gecko species (e.g., Hemidactylus frenatus; Brown and Sakai
1988; Brown et al. 1991; Petren et al. 1993) although they do not
appear to engage in the near-lethal battles of some sexual gecko
species (e.g., Eublepharis macularius; Mason and Gutzke 1990;
Brillet 1993). Despite this, aggression between captive L. lugubris
can occur, and we suggest immediately isolating any geckos that
exhibit bite marks or excessive weight loss due to intraspecific
aggression or reproductive stresses. Brown and Sakai (1988)
demonstrated that isolated gravid individuals exhibit lower
fecundity than those in social groups. We therefore recommend
against keeping individuals in isolated enclosures if the aim is
to maximize egg production. Brown and O’Brien (1993) provide
evidence that keeping two individuals per enclosure allows for
the highest reproductive output because subordinates in L.
lugubris dominance hierarchies typically take longer to reach
sexual maturity.
Fig. 2. Color patterns of typical laboratory lineages of Lepidodactylus
lugubris. A) Clone A, B) Clone B (speckled lineage), C) Clone B
(spotted lineage).

Herpetological Review 49(2), 2018
232 TECHNIQUES
Geckos can be kept in a variety of cage designs including glass
aquaria with screen lids or plastic cages with ventilated tops. We
chose to use oversized ventilated deli cups as they are space
efficient and maintain relatively high humidity. Maintaining
high humidity (~50–80%) is important when ambient humidity
is low, such as arid regions or in winter in temperate regions.
Each enclosure consists of an oversized plastic deli container (9-
7/8 inches × 5-1/2 inches; 25.08 cm × 13.97 cm; purchased from
www.superiorshippingsupplies.com) with 12 4-mm diameter
holes in the side and a 10-inch matching plastic lid (Fig. 3A, B).
These enclosures easily can be kept on a wire or stainless-steel
shelf (Fig. 3C). We use approximately 15 mm of loose coconut
fiber (Exo Terra®) as a substrate. After rehydrating coconut fiber
from its typical commercial “brick” form, we allow coconut fiber
to dry out until no water drips out when squeezed firmly by hand.
This typically provides the ideal amount of substrate moisture
and cage humidity. Fragments of pulp fiber egg cartons can be
stacked on top of each other to provide hiding spots and shelter
(Fig. 3A, B). Cages are cleaned every two weeks. We transfer
geckos to a clean cage and then empty out the dirty enclosure
and wash/disinfect the old cages in bulk. The washing process
proceeds as follows: scrub with dish soap and water, rinse with
clean water, soak in a 5% bleach solution for five minutes, and
rinse with water again prior to air-drying.
Enclosure temperature and humidity for both postnatal
individuals and eggs should range between 24.0–28.0°C and 30–
40%, respectively. If needed, an additional heat source, such as
a heat pad (FlexWatt Heat Tape or Ultratherm Heat pad), can be
Fig. 3. A) A typical enclosure for captive Lepidodactylus lugubris showcasing coconut fiber substrate, cardboard egg carton shelter, a water
dish, and a dish of calcium supplement. B) The same enclosure from an aerial view. C) A space efficient collection of 23 L. lugubris enclosures
collectively housing 68 individuals. D) A clutch of L. lugubris eggs adhered to the wall on the enclosure.

Herpetological Review 49(2), 2018
TECHNIQUES 233
placed underneath approximately one third of the container to
ensure a temperature gradient is established. The additional heat
source should not be placed underneath the water dish, which
would result in superfluous humidity within the enclosure. If the
enclosure becomes too humid, wipe condensation off of the sides
of the enclosure as needed. Furthermore, the heat source should
never cover more than ~30% of the cage bottom to allow natural
thermoregulation within the cage and avoid over-heating.
Different clone types vary in their temperature preferences
(Bolger and Case 1994) with spotted B clones preferring lower
temperatures than A and other B clones. Lepidodactylus
lugubris in both captive and natural settings are nocturnal but
occasionally forage during the day (Oliver and Shaw 1953; Perry
and Ritter 1999). We expose our colony of L. lugubris to 14 h of
ambient light, from 32-watt fluorescent ceiling light fixtures,
each day. This species, like other nocturnal gecko species, does
not require special UV-A or UV-B lighting (de Vosjoli et al. 1998).
Food and water.—Water is available ad libitum in 2-oz plastic
cups (aka soufflé cups – SOLO®) and changed weekly or more
frequently if necessary. Geckos are fed two to three times/week.
Because L. lugubris are both insectivorous and nectivorous
(Perry and Ritter 1999), we alternate diets every feeding. A typical
feeding of insects consists of approximately 7–10 small insects
per gecko, either two-week-old crickets (Gryllodes sigillatus
or Acheta domestica) or small mealworms (Tenebrio molitor).
A typical feeding of a fruit-based prepared diet consists of
approximately 0.5 ml per individual gecko of liquid diet such as
Repashy Crested Gecko Meal Replacement (Repashy Ventures
Inc.) or Pangea Gecko Diet Breeding Formula (Pangea Reptile
LLC). Prepared gecko diets are offered in small SOLO soufflé cups
and any uneaten food should be removed from the enclosure
the following day. Lepidodactylus lugubris use large amounts of
calcium when egg-laying so a small cup of powdered calcium
supplement with vitamin D3, e.g., SuperCal HyD (Repashy
Ventures Inc.), should be made available at all times to prevent
metabolic bone disease (de Vosjoli et al. 1998). Lepidodactylus
lugubris, indeed most gecko species, will eat the calcium directly
out of the cup (deVosjoli et al. 1998; pers. obs.).
Oviposition, egg collection, and juvenile care.—Lepidodactylus
lugubris, like other gekkonids, lay hard-shelled eggs with a
fixed clutch size of two eggs via synchronous single ovulation
from each ovary (Kluge 1967; Bustard 1968; Jones et al. 1978;
Fig. 3D). Lepidodactylus lugubris occasionally oviposit a single
egg (Sabath 1981). Although no empirical data exist for ovarian
cycle length for L. lugubris, egg-laying occurs year-round
(Oliver and Shaw 1953; Jones et al. 1978). Individuals adhere
or “glue” eggs to a variety of surfaces (Oliver and Shaw 1953).
We most often find eggs adhered to the walls of the enclosures
(Fig. 3D) and underneath pulp fiber egg cartons. If cork bark
fragments are available, geckos preferentially lay eggs in the
cracks in the bark. Lepidodactylus lugubris occasionally eat
their own eggs (Miller 1979); therefore, we separate eggs from
the adults frequently and in tandem with cage cleaning. Upon
cleaning, we set any enclosures with eggs aside, with coconut
fiber remaining to provide humidity. Egg enclosures remain at
the same temperature as adult and juvenile enclosures. Besides
being checked every day for hatched juveniles, egg enclosures
can be left alone until hatching or embryo collection. Incubation
time post-oviposition exhibits an inverse linear relationship
with incubation temperature (Brown and Duffy 1992). Brown
and Duffy (1992) demonstrated that eggs from a single L.
lugubris can vary in incubation times, extremes being between
approximately 65 days post-oviposition (dpo) at an average
temperature of 25.5°C and 103 dpo at 22.0°C. Upon hatching,
juveniles are approximately 16–21 mm SVL and can be housed
with clutch-mates. Housing, humidity, temperature, feeding
schedule, and cleaning schedule of juveniles are consistent with
those of adults although live food items are smaller, e.g., pinhead
crickets, newly hatched mealworms, and flightless fruit flies
(Drosophila melanogaster or D. hydei).
Euthanasia.—Although we are keeping and breeding L.
lugubris for embryo production it is necessary to have a protocol
available for euthanasia in the event of an emergency, i.e., a lizard
has been injured or is in severe distress. Lizards, both postnatal
and near full-term embryos, can be euthanized by injection with
tricaine methanesulfonate (MS-222), an approved method for
reptile euthanasia in the 2013 AVMA Guidelines on Euthanasia
(Leary et al. 2013), using the two-stage procedure from Conroy
et al. (2009).
table 1. Preservation and associated storage methods for Lepidodactylus lugubris embryos for assorted laboratory techniques. As different or
additional methods for DNA or RNA storage may be required depending on the reason for the tissue being collected, refer to Gamble (2014).
EtOH, ethanol; MeOH, methanol; PFA, Paraformaldehyde; RT, room temperature (23°C).
Technique Preservation Method Storage Method
Specimen preparation Fix overnight in 10% formalin Dehydrate to 70% EtOH,
store at RT
Immunohistochemistry Fix 2 hours in 4% PFA at 4°C Dehydrate to 100% MeOH,
store at -20°C
Electron microscopy 1% glutaraldehyde at 4°C 1% glutaraldehyde, store at 4°C
Histology Fix overnight in 10% formalin Dehydrate to 70% EtOH,
store at RT
In situ hybridization Fix overnight in 4% PFA at 4°C Dehydrate to 100% MeOH,
store at -20°C
DNA extraction 95% EtOH 95% EtOH, store at RT
RNA extraction Trizol or snap freeze in liquid nitrogen Store at -80°C

Herpetological Review 49(2), 2018
234 TECHNIQUES
embryo ColleCtion, preServation, and Storage
The eggs of L. lugubris are often firmly adhered to the side of
the cage or on the underside of items within the cage. Eggs that
are not fully adhered to a surface by the female often desiccate
several days after laying. Regardless of how the egg is positioned
by the female, the embryo rotates to the uppermost surface of the
eggshell and can easily be visualized by shining a light through
the shell from the backside (i.e., “candling”). The eggs cannot be
easily removed from their surface by hand without cracking the
shell and damaging the embryo inside. To remove an intact egg,
we use a #11 scalpel to cut the egg off of the surface it is adhered
to. The egg can then be handled by hand or by using a perforated
spoon.
To dissect the embryo from its shell, we first submerge the
egg in 1X phosphate buffered saline (PBS) in a glass culture
dish. Once submerged the eggshell can be gently cracked and
the pieces removed using #5 watchmaker’s forceps. This leaves
an intact bolus of yolk and embryo surrounded by translucent
membranes. Using a pair of #5 forceps we free the embryo from
the yolk and membranes. The embryos can now be easily moved,
once again using a perforated spoon, to a clean dish of DEPC-
treated (i.e., RNAase free) PBS. To separate the primary nutrient
stalk from the embryo without damaging the viscera, we create a
clean cut by pinching the stalk with one set of forceps and then
sliding the second set along its length, crossing the stalk.
The method of embryo preservation and storage ultimately
depends on the long-term use of the specimen. In our labs, we
use L. lugubris embryos for morphological analysis, nucleic
acid extraction, histology, scanning electron microscopy, im-
munohistochemistry, and in situ hybridization of mRNA lo-
calization. We have summarized the preservation and storage
techniques used for L. lugubris embryos in Table 1.
Compared to working with adult specimens, there are
several critical differences to processing embryonic material.
First, moving between aqueous and alcohol-based solutions
needs to occur gradually. Rapid changes in tonicity, such
as when moving between PBS and alcohol, will lead to
dramatic wrinkling of the outer epithelium and may deform
the embryonic morphology. We suggest a three step gradual
transition between aqueous and alcohol-based solutions (e.g.,
25%, 50%, 70% ethanol). In addition, fixing tissue at 4oC helps
to reduce the chances of tissue degradation during fixation.
Finally, because of the small size of the embryos and variable
levels of gene expression, all solutions should be maintained in
sterile or RNAase-free conditions.
ConCluSionS
Using the protocols described in this paper, we have
successfully maintained a colony of Lepidodactylus lugubris
in a space-efficient and inexpensive manner. Furthermore,
our collection of nearly 60 individuals is capable of producing
approximately 50 eggs per month, making L. lugubris an ideal
amniote for embryological research. These protocols were
created with a basic knowledge of gecko captive care and natural
history, and, with additional optimization, can be applied to
other similar gekkonid species. Lepidodactylus lugubris is an
ideal model to study vertebrate evolutionary developmental
biology and parthenogenesis in a laboratory setting. With these
protocols, we hope to set a foundation to make this emerging
model species more accessible.
Acknowledgments.—We gratefully acknowledge the Marquette
University and Loyola University at Chicago Institutional Animal Care
and Use Committees (IACUC) for their hard work and approval of the
methods described in this paper. All work conducted at Marquette
University as it relates to this paper was done under an approved
Marquette University IACUC protocol. Marquette University has an
AAALAC accredited Animal Care and Use Program and has the PHS
Assurance on file with the Office of Laboratory Animal Welfare. We
thank the employees of both Gamble and Sanger Lab captive animal
rooms for their tireless effort and help optimizing care protocols.
We also thank the State of Hawaii Division of Forestry and Wildlife
for permission to collect Lepidodactylus lugubris and D. Zarkower
and C. Matson for help in the field. This manuscript benefitted
from the helpful comments and suggestions from three anonymous
reviewers. The collection and maintenance of these animals was
funded in part by Marquette University and the National Science
Foundation (DEB1657662).
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