gonadal differentiation in reptiles exhibiting environmental sex determination
TRANSCRIPT
E-Mail [email protected]
Sex Dev DOI: 10.1159/000358892
Gonadal Differentiation inReptiles Exhibiting EnvironmentalSex Determination
S. Kohno a–c B.B. Parrott a–c R. Yatsu e, f S. Miyagawa e, f B.C. Moore d
T. Iguchi e, f L.J. Guillette Jr. a–c
a Department of Obstetrics and Gynecology, and b Marine Biomedicine and Environmental Sciences Center,Medical University of South Carolina, and c Hollings Marine Laboratory, Charleston, S.C. , d School ofBiological Sciences, Louisiana Tech University, Ruston, La. , USA; e Okazaki Institute for Integrative Bioscience,National Institute for Basic Biology, National Institutes of Natural Sciences, and f Department of Basic Biology,Faculty of Life Science, Graduate University for Advanced Studies, Okazaki , Japan
Sertoli cells and granulosa cells of the developing testis and ovary, respectively, and the mechanisms by which gene ex-pression is regulated during TSD events. Further, reptilian sentinels and their mechanisms of gonadogenesis will likely remain important indicator species for environmental health. Thus, ongoing and new investigations need to tie molecular information to gonadal morphogenesis and func-tion in reptiles. Such data will not only provide important information for an understanding of the evolution of these phenomena in vertebrates, but could also provide an im-portant understanding of the health of the environment around us. © 2014 S. Karger AG, Basel
Reptiles, including crocodilians, turtles, lizards and snakes, exhibit a variety of life histories, including various sex determination systems [Organ et al., 2009]. In those reptilian species in which sex is determined by the inher-itance of sex chromosomes, both XY/XX and ZZ/ZW sys-tems have been reported. In contrast, environmental fac-
Key Words
CYP19 aromatase · DNA methylation · Endocrine-disrupting contaminants · Estrogen signal · Germ cell migration · Germ cell specification · Oogenesis · Spermatogenesis · Temperature-dependent sex determination
Abstract
As temperature-dependent sex determination (TSD) and ho-mozygote or heterozygote genetic sex determination (GSD) exist in multiple reptilian taxa, they represent sex determina-tion systems that have emerged de novo. Current investiga-tions have revealed that the genetic mechanisms used by various reptilian species are similar to those used by other vertebrates. However, the recent completion or near com-pletion of various reptilian genome projects suggests that new studies examining related species with and without TSD could begin to provide additional insight into the evolution of TSD and GSD in vertebrate ancestors. Major questions still remain concerning germ cell migration and specification, the differentiation of gonadal accessory cells, such as the
Published online: March 13, 2014
Satomi Kohno, Department of Obstetrics and Gynecology Marine Biomedicine and Environmental Sciences Center Hollings Marine Laboratory, Medical University of South Carolina 331 Fort Johnson Road, Charleston, SC 29412 (USA) E-Mail kohno @ musc.edu
© 2014 S. Karger AG, Basel1661–5425/14/0000–0000$39.50/0
www.karger.com/sxd
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
2
tors, such as temperature and/or humidity, determine sex in other species [Packard et al., 1987; Janzen and Paukstis, 1991; Kohno and Guillette, 2013]. Both sex determina-tion systems can be ‘overridden’ by exposure of the devel-oping embryo to sex steroid hormone exposures or chem-icals with these properties. Thus, reptiles are sensitive to environmental factors, such as climate change and expo-sure to endocrine disrupting compounds [Hamlin and Guillette, 2011]. Indeed, in ovo exposure studies have demonstrated that sex ratios of American alligators (Al-ligator mississippiensis) [Milnes et al., 2005] and red-eared sliders (Trachemys scripta) [Willingham and Crews, 1999] are sensitive to environmental contaminants with endocrine disrupting activity (EDCs). Here, we review reptilian gonadogenesis with an emphasis on environ-mental aspects.
Evolution of Sex Determination Systems
One of the central pillars of evolutionary biology is the investigation of the diverse sex-determining systems found among taxa. Differences in the strategies of sex de-termination can lead to speciation or extinction by influ-ences at the genetic, physiological, behavioral, and popu-lation levels. Temperature-dependent sex determination (TSD) is a mechanism in which the sex of an individual is directly determined by specific thermal cues during em-bryonic development [Ezaz et al., 2006]. Currently, the underlying molecular mechanisms controlling TSD are not fully understood. In the wild, TSD reptiles are report-ed to trend toward a female biased sex ratio at hatch [Fer-guson, 1985], but long-term studies of multiple popula-tions examining this issue do not exist. Organisms with TSD exhibit 3 major patterns, which are female to male (F-M), female to male to female (F-M-F) and male to fe-male (M-F) patterns ( fig. 1 ). In TSD, the thermal range is well defined, sometimes by a thin margin of only 2–3 ° C. In the case of the American alligator, a shift from 30 to 33 ° C is sufficient to switch the sex of the resulting hatch-lings from 100% females to 100% males [Deeming and Ferguson, 1989].
The occurrence of TSD is not rare in vertebrates ( fig. 2 ). The phenomenon occurs regularly among reptiles and fish, and its variants can be observed in amphibians as well. In reptiles, the sphenodontian (tuatara) and croc-odilian (alligator and crocodile) groups have the distinc-tion of being entirely composed of species exhibiting TSD. Curiously, the temperature ranges for crocodilian egg incubation are much narrower compared with the rest of the reptiles [Deeming, 2004]. Crocodilians, unlike other reptiles, belong to the archosaurian lineage; thus, they are more closely related to birds, organisms exhibit-ing genetic sex determination (GSD), than to other rep-tiles. As for other reptiles, all sea turtles studied to date exhibit TSD, whereas other chelonians display GSD or TSD [Organ et al., 2009]. Among squamates (lizards, snakes and geckos), TSD is restricted to only a few clades [Organ et al., 2009; Pokorna and Kratochvil, 2009]. Inter-estingly, the lizard Gekko japonicus possesses heteromor-phic sex chromosomes, yet at the same time exhibits TSD [Tokunaga, 1985].
In contrast to reptiles, TSD is less common in the fish species studied to date. The Atlantic silverside (Menidia menidia) , a teleost fish, has been demonstrated to exhib-it TSD in the wild [Baumann et al., 2012]. However, a major general difference exists, as unlike reptiles, fish sex ratios (based on sex determination) can be altered by a
Fig. 1. Three different patterns of temperature-dependent sex de-termination: F-M (top), F-M-F (middle) and M-F (bottom) pat-terns. For example, species with the F-M pattern produce females at lower egg-incubating temperatures, whereas males are pro-duced at higher egg-incubating temperatures.
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
3
multitude of environmental factors such as pH, stress and malnutrition, in addition to temperature. Further-more, while reptilian sexual development is irreversible, fish are reported to retroactively alter their sex [Kobaya-shi et al., 2005]. The labile ambiguity of this sex-deter-mining mechanism could imply that fish TSD involves a very different series of mechanisms than that found in reptiles.
GSD is the more common mechanism among verte-brates. It is displayed in mammals, along with birds, am-phibians, and selected reptiles and fish. Basic mecha-nisms of GSD revolve around the presence of heritable sex-biased gene dosage, and an individual’s sex is defined at fertilization. The majority of vertebrate GSD can be broadly categorized into either XX/XY, male heteroga-metic system (XX = female; XY = male) or ZW/ZZ, fe-male heterogametic system (ZW = female; ZZ = male). Both systems are usually thought of in terms of a simple Mendelian model and are expected to generate a balanced sex ratio in theory.
Whereas TSD and GSD mechanisms were tradition-ally approached as discrete phenomena, the contempo-rary view suggests the 2 systems exist as polar ends of a continuum. For example, many species having hetero-morphic sex chromosomes and also showing thermal sensitivity to sex have now been identified, indicating that GSD and TSD systems of sex determination can co-exist. In reptiles, GSD is more or less inferred from extensive karyotyping as well as assessment of temperature effects on sex determination (or lack thereof). XX/XY, ZW/ZZ chromosome systems have been identified in many groups, although a few remain undefined; that is, homo-morphic sex chromosomes or unconventional hetero-morphic sex chromosomes could be present [Badenhorst et al., 2013]. Moreover, reptilian sex determination mech-anisms are not well understood, and the exact genotype and loci that would prompt sex determination is yet to be elucidated. Even the sexual development of some GSD-based organisms can be influenced by temperature in a profound manner. Though not considered to be ‘true’
Fig. 2. Phylogeny of amniotes with PGC specification, PGC migration, sex determi-nation and birth system noted. Crocodil-ians and tuatara exhibit only TSD, whereas lizards and turtles exhibit both mecha-nisms (GSD and TSD) as well as display all 3 patterns of TSD (see fig. 1). Crocodilians and birds form Archosaura and are sister groups; however, they exhibit different sex determination patterns – TSD and GSD. Figure data is derived from Ezaz et al., [2006], Bachvarova et al. [2009b], Organ et al. [2009], Pokorna and Kratochvil [2009], and St John et al. [2012].
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
4
TSD, temperature-induced sex reversal has been reported in teleosts (e.g. zebrafish, ricefish) and amphibians (e.g. Iberian ribbed newt, wood frog). In these cases, an organ-ism displays a balanced sex ratio at normal ambient tem-peratures. However, exposure to an extreme temperature during the embryonic period can override the individu-al’s genotypic sexual development. In typical cases, high heat exposure of around 30 ° C can lead to masculinization [Dournon et al., 1990; Wallace et al., 1999; Uchida et al., 2002; Ospina-Alvarez and Piferrer, 2008].
GSD has been classified by identifying a consistent ge-netic difference between males and females. Tradition-ally, sexually dimorphic chromosomes (heteromorphic sex chromosomes) have been identified by karyotyping [Cole, 1971]. Fragments of chromosomes associated with one sex, identified by chromosome painting, have also proved successful at revealing loci associated with GSD [Valleley et al., 1994]. More recently, restriction site-as-sociated DNA sequencing has been applied to discover allelic differences showing a strong association to a spe-cific sex [Chapus and Edwards, 2009]. In addition, poten-tial effects of incubation temperature on sex determina-tion and the pattern of TSD can be identified by the max-imum likelihood model selection [Godfrey et al., 2003].
From the early days of TSD research, many have con-templated its evolutionary significance. After all, a sex ra-tio prone to environmental influence hardly seems effi-cient for population growth, yet TSD is still prevalent in many long-lived and evolutionarily robust species to this day. The Charnov-Bull model proposes that certain incu-bation temperatures result in differential fitness (survival × fertility) to one sex [Charnov and Bull, 1977]. In this model, TSD is advantageous relative to GSD when an en-vironment is patchy and results in biased fitness toward a particular sex; that is, the incubation temperature would serve as a cue to the environmental conditions for the hatchling. If the environment elevates or suppresses fit-ness toward one sex, the TSD mechanism allows adjust-ment for an offspring sex ratio with higher overall fitness. In addition, it has been proposed that in order for TSD to retain an evolutionary advantage over GSD, a species must be sufficiently long-lived so that yearly fluctuations in the sex of offspring do not negatively affect the sex ra-tio of the breeding population [Bull and Bulmer, 1989]. While the Charnov-Bull model remains an attractive model to explain the evolutionary pressures leading to TSD, empirical evidence supporting the model has been challenging to produce. That said, experiments in fish have produced data that agree with this model [Conover, 1984]. More recently, measuring the lifetime reproduc-
tive success of sex-reversed Jacky dragons (Amphibolurus muricatus) produced the first empirical evidence sup-porting the Charnov-Bull model in amniotes [Warner and Shine, 2008]. Males resulting from incubations at an intermediate temperature resulted in higher reproductive success when compared to pharmacologically derived males incubated at either of the higher or lower female-producing temperatures. In contrast, females produced from incubation at the extreme temperatures displayed higher reproductive success than those incubated at the intermediate temperature.
The early view on TSD (and environmental sex deter-mination – ESD) suggested that with the advent of gono-chorism, hermaphroditic ESD organisms gradually evolved GSD [Bull, 1980]. Subsequent modern phyloge-netic analysis, however, indicates ancestral GSD with a transition to TSD in some vertebrates [Janzen and Krenz, 2004]. Furthermore, the transition between TSD and GSD has occurred repeatedly in vertebrates and in a rela-tively short evolutionary time. Sister taxa, such as croco-dilians (TSD) and birds (GSD), are such an example [Shaffer et al., 2013].
In turtles, phylogenetic assessments suggest that TSD is the ancestral mode of sex determination. Subsequent GSD in turtles seems to have been independently derived multiple times, with both XX/XY and ZW/ZZ GSD sys-tems. Similarly, the squamate ancestral sex determination mechanism is suggested to have been TSD, but modern extant lizards predominantly exhibit GSD, unlike turtles. Parsimony-based comparative analysis shows the loss of TSD occurring in at least 6 major clades in turtles and in-dependently developing in 3 major squamate clades [Po-korna and Kratochvil, 2009]. Detailed analysis indicates highly frequent TSD/GSD turnover within some of those clades [Pokorna and Kratochvil, 2009]. For example, the Gekkonidae (geckos) family possesses XX/XY and ZW/ZZ GSD as well as TSD during a short evolutionary pe-riod. The diversification of sex determination seems to be easily induced, and even the same species (e.g. yellow mud turtle, Kinosternon flavescens ) can display an alter-native TSD pattern (at least alterative response to tem-perature) in different geographical population [Ewert et al., 1994; Godfrey et al., 2003]. These observations indi-cate significant variation in reptilian sex determination, and the frequent transition between TSD and GSD points towards malleable properties in the mechanisms associ-ated with sex determination in this group.
Many of the orthologues of key sex differentiation-re-lated genes have also been discovered in TSD and GSD reptiles, and its comprehensive genetic analyses seem to
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
5
suggest that both mechanisms follow a fundamentally similar differentiation cascade [Urushitani et al., 2011; Matsumoto and Crews, 2012]. Despite its biological sig-nificance, sex determination mechanisms display fre-quent evolutionary transitions, and a multitude of char-acteristic and genetic variations in TSD and GSD. In real-ity, sex determination seems likely to be more complex than a ‘simple’ bidirectional spectrum. A better under-standing of the intricate relation and origin of GSD and TSD requires a deeper comparative understanding of go-nadogenesis in amniotes.
Specification and Migration of Germ Cells in Reptiles
The specification and migration of primordial germ cells are critical events in gonadogenesis. In vertebrates, the primordial germ cells (PGCs) arise outside of the go-nadal tissue and exhibit 2 distinct specification modes [Extavour and Akam, 2003]. In one mode of specification (predetermined mode, sometimes called preformation), germ cells are predetermined by an accumulation of de-terminants in a region of the embryonic cytoplasm, called the germ plasm before or just after fertilization, and even-tually gives rise to germ cells after cell division and/or cel-lularization [Extavour and Akam, 2003]. In the second mode (inductive mode, sometimes called epigenesis), germ cells are specified by inductive signals from neigh-boring cells, and this inductive mode is a basal mode in reptiles [Extavour and Akam, 2003; Bachvarova et al., 2009b]. These modes of specification can be identified by the localization of the PGCs during the early somite stag-es (anterior vs. posterior of extra-embryonic endoderm) in reptiles [Extavour and Akam, 2003; Bachvarova et al., 2009b], and there are 2 different pathways by which PGCs migrate into the genital ridge (through the bloodstream vs. dorsal mesentery) [Hubert, 1969; Fujimoto et al., 1979; Gilbert, 2010].
At the early somite stages, PGCs are in the junctional endoderm, lying just outside the embryo in the lateral posterior region [Bachvarova et al., 2009a]. The PGCs travel to the genital ridge and arrive in the bipotential go-nad at the onset of gonadogenesis. The pathway by which the PGCs arrive in the gonad differs among species and has the potential to be associated with the specification mode of the germline [Bachvarova et al., 2009b].
The predetermined mode of PGC specificity has been reported in snakes and birds, and these data have been extended to suggest crocodilians specify their germline using this mode, given that birds and crocodilians are the
only remaining archosaurs. PGCs are observed at the on-set of gastrulation in the lower hypoblast layer of birds, anterior to the blastopore, and are carried forward to form an anterior crescent. This extra-embryonic region containing the PGCs is called the germinal crescent [Tsunekawa et al., 2000; Bachvarova et al., 2009b]. The PGCs subsequently migrate into the genital ridge through the bloodstream in birds, but no record is yet available for reptiles. The PGCs are derived from epiblast cells that mi-grate from the central region of the area pellucida to a crescent-shaped zone in the hypoblast at the anterior bor-der of the area pellucida [Eyal-Giladi et al., 1981; Gins-burg and Eyal-Giladi, 1987]. The avian PGCs arrive in the genital ridge at the embryonic stage exhibiting 25 so-mites, which is equivalent to the second stage of crocodil-ian embryonic development described by Ferguson [1985] and occurs within 24 h of oviposition [Bachvarova et al., 2009b]. The localization of PGC at this early stage has not been evaluated in depth due to the difficulty of obtaining specimens and/or the lack of genetic tools, such as antibodies or gene sequence information. Future re-search examining this process in various reptilian species is needed to further elucidate the evolutionary dynamics of PGC specification and migration.
The inductive mode has been observed in some turtles (Testudines) and lizards (Lacertoids). In these species, the germline is specified by interaction with neighboring cells at the posterior horseshoe-shaped region of extra-embry-onic endoderm during the early somite stage, whereas in the predetermined mode the germ cells are specified by the germ plasm at the hypoblast and accumulated at the anterior germinal crescent on extra-embryonic endo-derm at early somite stage [Bachvarova et al., 2009b]. The germ cells are located during early embryonic develop-ment in a posterior crescent or horseshoe-shaped region of extra-embryonic endoderm and then migrate by inter-stitial movement through the gut and dorsal mesentery to the gonad [Extavour and Akam, 2003]. Although the ini-tial locations of PGCs in undifferentiated embryonic bird and turtle gonads are similar, no record of PGCs migrat-ing through the bloodstream has been reported for tur-tles. Although germ cell origin and migration in turtles has not been thoroughly characterized, one study report-ed that PGCs migrate through the dorsal mesentery to colonize the gonad similarly to the mouse gonad [Pieau and Dorizzi, 2004]. As in birds, it appears that germ cells do not play an essential role in embryonic gonadal mor-phogenesis in turtles, as a depression in number of PGCs does not induce alterations in gonadal morphology [Di-Napoli and Capel, 2007].
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
6
The PGC specification modes in other reptilian species have not been elucidated to date, although Bachvarova et al. [2009b] classified them as belonging to the circumfer-ential group. The circumferential group has been report-ed in reptiles such as lygosomines (lizards), chamaeleo-nids (chameleons), sphenodontids (tuatara), and anguids (slow worms) [Bachvarova et al., 2009b]. Embryos exhib-iting this pattern have PGCs located in the junctional zone of extra-embryonic endoderm circumferentially around the early somite embryo, and there are more PGCs in the anterior region of the extra-embryonic me-soderm than the posterior region, with a gradient of den-sity of PGCs [Bachvarova et al., 2009b]. Due to the similar localization of PGCs to the predetermined specification at early somite stages, and the same pathway of PGCs mi-gration as the predetermined mode [Bachvarova et al., 2009b] (see fig. 2 for summary), PGCs in this group of animals may be specified by the predetermined mode. However, further investigations are required to evaluate this hypothesis.
Mechanisms of Temperature-Dependent Sex
Determination in Reptiles
Many of the genes required for gonadogenesis are broadly conserved among vertebrates. However, stand-ing in stark contrast to the conservation of this core ge-netic machinery is a remarkable divergence by which sex is initially determined. In many reptiles, including some squamates, many turtles, tuatara, and all crocodilians studied thus far, the temperature at which the egg is incu-bated determines or dramatically influences sex. In spe-cies undergoing TSD, temperature during incubation must be translated into a biological switch that differen-tially directs gonadal development towards either a testis or an ovary. Despite many studies aimed at elucidating the molecular nature of this trigger, our understanding remains incomplete.
One of the initial revelations into the mechanism un-derlying TSD was that sex determination in reptiles is ex-quisitely sensitive to endogenous, as well as exogenous, hormone signaling. For example, across 3 different orders of reptiles and regardless of the TSD pattern (F-M, F-M-F and M-F; see fig. 1 ), eggs incubated at the male-producing temperature (MPT) and treated exogenously with the en-dogenous sex steroid hormone estradiol-17β produce fe-males instead of males [Bull et al., 1988; Crews et al., 1989; Lang and Andrews, 1994]. Aromatase is the enzyme re-sponsible for the final conversion of androgens to estro-
gens in vertebrates [Evans et al., 1986; McPhaul et al., 1988; Tanaka et al., 1992; Gabriel et al., 2001; Murdock and Wibbels, 2003] and consistent with a role for endog-enous estrogens in TSD; multiple studies have demon-strated a dramatic increase in the gonadal expression of aromatase in embryos incubated at the female-producing temperature (FPT) [Crain et al., 1997; Gabriel et al., 2001; Milnes et al., 2002; Murdock and Wibbels, 2003]. Fur-thermore, experiments in which alligator gonads were cultured ex vivo have demonstrated increased estrogen production, and thus, increased aromatase activity, when gonads are cultured at FPT relative to MPT [Smith and Joss, 1994; Smith et al., 1995]. While this sufficiency of estrogen to induce male-to-female sex reversal has been well established across reptiles and vertebrates in general, a de facto requirement for estrogen in specifying the fe-male sex remains less clear. Estrogen signals via estrogen receptor alpha (ERα or ESR1), not ERβ (ESR2), drive male-to-female sex reversal in American alligators [Koh-no et al., unpubl. data].
Studies utilizing pharmacological approaches have shown that in a few turtle species, treating eggs incubated at FPT with compounds that inhibit aromatase activity results in female-to-male sex reversal [Dorizzi et al., 1994; Rhen and Lang, 1994; Wibbels and Crews, 1994; Richard-Mercier et al., 1995]. However, similar experi-ments incorporating aromatase inhibitors in alligators and at least one other turtle species have yielded findings that are more difficult to interpret [Lance and Bogart, 1992; Jeyasuria et al., 1994]. In these studies, partial fe-male-to-male sex conversions were observed in some cases, whereas in others, female development was either only partly disrupted or an effect on sex determination was completely absent. Further complicating the inter-pretation of these results is that aromatase inhibition is thought to result in reduced estrogen levels, but by block-ing the aromatization of androgens, treatment can also result in increased androgen levels. Thus, it is not clear if the observed masculinizing effects of aromatase inhibi-tion are mediated by a reduction in estrogen signaling or increased androgen levels. Treatment with an androgen that is unable to be aromatized, 5α-dihydrotestosterone, has been reported to skew sex ratios towards male devel-opment in turtle eggs incubated at pivotal temperatures [Wibbels et al., 1992; Wibbels and Crews, 1995]. How-ever, 5α-dihydrotestosterone treatment is not sufficient to induce female-to-male sex reversal in turtles or alliga-tors incubated at FPT [Lance and Bogart, 1992; Rhen and Lang, 1994]. Why is 5α-dihydrotestosterone able to in-fluence sex determination at pivotal temperatures, but
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
7
not at FPT? This could reflect the nature of the underlying molecular switch responding to incubation temperature. Intersex animals are typically not produced from incuba-tions at intermediate temperatures resulting in mixed sex ratios, suggesting that a bimodal threshold is reached during normal sex determination. However, an underly-ing continuous spectrum of sex appears to be present, as those embryos incubated at pivotal temperatures appear to be more sensitive to endocrine signals than those raised at temperatures producing all males or females. Although rare, it should be noted that some reports have indicated that intersex animals were produced at intermediate tem-peratures in the red-eared slider turtle and the European pond terrapin [Pieau et al., 1998; Barske and Capel, 2010]. Studies examining how the balance between androgen and estrogen signaling interact with temperature to affect sex determination are needed to resolve their respective contributions. In addition, further studies examining the role of endogenous hormone signaling are needed to ful-ly understand how they function in sex determination.
Soon after it was realized that steroid hormone signal-ing critically influences sex in reptiles, investigators turned their attention towards identifying genes involved in sex determination. Simple, yet elegant, studies began to reveal a temporal cascade of gene expression that could be differentially directed by incubation temperature [see review by Shoemaker and Crews, 2009]. These genes were primarily identified using a candidate approach in which the expression of genes with critical roles in sex determi-nation/differentiation of humans and traditional labora-tory model animals were tested at key points during the thermo-sensitive period (TSP). Perhaps not surprisingly, many of the genes involved in mammalian sex determi-nation are conserved in reptiles (not surprising that mam-mals use genes that are conserved from a reptilian ances-tor), with a few key exceptions. Sex-determining region Y (SRY) , for example, is the male-determining gene on the Y chromosome of mammals, but it appears to be absent in nonmammalian vertebrates, including all reptiles stud-ied to date. Despite this absence, the gene directly down-stream of SRY in mammalian sex determination, SRY-box 9 (SOX9) , is present and is expressed during the TSP in many reptiles with TSD [Moreno-Mendoza et al., 1999; Western et al., 1999; Valleley et al., 2001; Shoemaker et al., 2007; Parrott et al., 2014]. SOX9 is a high-mobility group-box containing transcription factor that is respon-sible for initiating Sertoli cell differentiation and subse-quent testis formation in mammals [Foster et al., 1994; Morais da Silva et al., 1996; Vidal et al., 2001]. This up-stream role in sex determination has been demonstrated
by studies in which the ectopic expression of SOX9 in chromosomally female (XX) mice leads to sex reversal and male development [Vidal et al., 2001]. However, while many of the mammalian sex-determining genes are present in reptiles, their functional conservation and po-sition within the genetic pathway leading to sex determi-nation is not well established.
Anti-Müllerian hormone (AMH) is also a key regula-tor of Sertoli cell differentiation in mammals and is ex-pressed during the TSP in many reptiles [Tran et al., 1977; Western et al., 1999; Shoemaker et al., 2007]. The expres-sion of SOX9 precedes AMH in mammals, and it is thought to directly induce the expression of AMH [De Santa Barbara et al., 1998]. However, conservation of the temporal and spatial dynamics of SOX9 and AMH ex-pression is not clear across reptiles. For example, in the alligator, it has been reported that AMH mRNA expres-sion is detected before that of SOX9 mRNA, indicating that SOX9 is not the primary determinant of AMH ex-pression [Western et al., 1999; Urushitani et al., 2011] . This temporal deviation from mammalian sex determi-nation, with AMH expression preceding that of SOX9 , has also been reported in the chicken and could represent an evolutionary divergence in the mechanism regulating AMH expression [Oreal et al., 1998]. In the olive ridley sea turtle (Lepidochelys olivacea), SOX9 is expressed at both MPT and FPT until the end of the TSP at which point its expression decreases dramatically in the pre-sumptive ovary [Moreno-Mendoza et al., 1999; Torres Maldonado et al., 2002]. Consistent with this report, it was also reported that in the red-eared slider turtle, sexu-ally dimorphic expression of SOX9 was not established until late in the TSP [Spotila et al., 1998; Barske and Capel, 2010]. However, a more recent report investigating the detailed spatial expression of SOX9 has shown clear di-morphism in the bipotential gonad near the beginning of the TSP [Shoemaker et al., 2007]. A study examining SOX9 expression in the leopard gecko, Eublepharis macu-larius , suggests a pattern similar to that of the olive ridley, but a more detailed analysis investigating the presence of spatial dimorphism during early stages of the TSP is still needed [Valleley et al., 2001]. Taken together, these stud-ies implicate a conserved role for SOX9 and AMH in the early stages of testis development in reptiles with TSD. Although the localization of SOX9 and the effects of es-trogen signaling on it were well investigated in the devel-oping gonad of the red-eared slider turtle [Barske and Ca-pel, 2010], the timing and spatial dynamics of SOX9 and AMH expression are still not completely resolved in rep-tiles. Studies examining the colocalization of SOX9,
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
8
AMH, and other key genes involved in sex determination during key stages of the TSP, could aid in elucidating their regulatory relationships. Small interfering RNA of SOX9 successfully reduced SOX9 expression at MPT on gonad-al organ culture of L. olivacea [Sifuentes-Romero et al., 2013], and this technique will allow a detailed investiga-tion of the mechanisms of TSD.
Studies examining gene expression during the TSP have revealed a temperature-dependent temporal hierar-chy of gene expression. In addition to SOX9 and AMH , investigators have identified a suite of genes involved in the sex determination/differentiation of other taxa and that are also expressed during the TSP of reptiles with TSD. These include genes such as: DMRT1 (doublesex and mab-3 related transcription factor 1), SF1 (steroido-genic factor 1), RSPO1 (R-spondin 1), and FOXL2 (fork-head box L2). A comprehensive list of genes and the tim-ing of their expression have been reviewed in detail else-where [Yao and Capel, 2005; Shoemaker and Crews, 2009] ( fig. 3 ). Yet, our knowledge of the mechanisms that influence the transcriptional activity of these genes is based almost exclusively on studies in traditional labora-tory model species. Furthermore, our understanding of how temperature influences the post-transcriptional reg-ulation and function of these genes remains incomplete. These gaps can primarily be attributed to the technical challenges of working on nontraditional species. How-ever, despite the challenges, a few studies are starting to provide insights into how these genes are regulated.
The alternative splicing of exons from pre-mRNAs originating from a single locus can give rise to multiple gene products with different functions. In the fruit fly, Drosophila melanogaster , the genetic regulatory network responsible for sex determination is dependent on the di-morphic alternative splicing of multiple genes. In short, a sex chromosome-counting mechanism results in the al-ternative splicing of the master gene, sex-lethal (sxl) . Whereas sxl transcripts in males harbor a third exon that contains a stop codon and results in a truncated, non-functional product, sxl undergoes an alternative splicing event in XX females that produces a function protein [Bell et al., 1988]. Interestingly, Sxl is itself a splicing factor and ultimately influences the dimorphic splicing of the dou-blesex (dsx) transcript [Burtis and Baker, 1989]. Dsx is thought to be the evolutionary precursor to the DMRT1 gene found in mammals and other vertebrates, including reptiles [Raymond et al., 1998; Smith et al., 1999]. The expression of DMRT1 is dimorphic during the TSP in many reptiles with TSD. For example, in both the red-eared slider and the olive ridley turtles, the gonadal ex-
Fig. 3. Gene expression profiles during sex determination in American alligator (Alligator mississippiensis) and red-eared slider turtle (Trachemys scripta) at MPT and FPT. Relative expression levels are indicated by the darkness of the bar. The red box indi-cates the thermo-sensitive period for sex determination in each species: SOX9 , AMH, WT1, DMRT1, SF1, DAX1, CYP19, RSPO1 , and FOXL2 . Data for this figure was derived from Morrish and Sinclair [2002], Yao and Capel [2005], Shoemaker and Crews [2009], and Urushitani et al. [2011].
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
9
pression of DMRT1 is dimorphic during the early stages of the TSP in embryos incubated at MPT or FPT [Torres Maldonado et al., 2002; Shoemaker et al., 2007]. In the Indian mugger, Crocodylus palustris , the expression of DMRT1 is dimorphic during the TSP and has also been shown to undergo alternative splicing [Anand et al., 2008]. However, each isoform appears to be expressed more highly in the gonad of embryos incubated at the MPT relative to those incubated at FPT and, thus, would suggest a more critical role for transcriptional regulation in establishing dimorphic activity.
In a similar study, it was demonstrated that SOX9 un-dergoes alternative splicing in the gonad of Indian mug-gers during the TSP [Agrawal et al., 2009]. In contrast to DMRT1 , alternative splicing of SOX9 transcripts appears to be dimorphic at MPT or FPT during the early stages of the TSP. Whereas transcripts containing the DNA-bind-ing domain are detected in the gonad at both tempera-tures, transcripts containing the transactivation domain are only detected in gonads from embryos incubated at the MPT [Agrawal et al., 2009]. SOX9 protein is not de-tected in the gonad of embryos incubated at FPT and sug-gests that the SOX9 transcript produced in the female go-nad is not translated into a full-length protein. Thus, while the transcriptional activity of SOX9 appears to be similar, the functional product is only observed in the go-nads incubated at MPT. These studies have raised the question of how widespread this phenomenon is across taxa. At least in the mouse, alternative splicing of SOX9 appears to be conserved as sexually dimorphic splice vari-ants in the transactivation domain are detected [Agrawal et al., 2009]. Does the expression of SOX9 at both FPT and MPT observed during the early stages of the TSP in the olive ridley and the leopard gecko [Moreno-Mendoza et al., 1999; Valleley et al., 2001] give rise to functional prod-ucts? This apparent conservation of SOX9 splicing in the mouse allows for studies in a genetically tractable model by which the specific mechanism leading to this dimor-phism can be teased apart. In addition, the recent se-quencing of the alligator and turtle genomes should fa-cilitate comparative studies in which the presence of di-morphic splice variants in key genes involved in sex determination can be investigated in additional species with TSD [St John et al., 2012; Shaffer et al., 2013].
Epigenetic mechanisms have long been invoked as key mediators between the environment and an organism’s genome. Modifications influencing transcriptional activ-ity, such as DNA methylation and small RNA, could be involved in translating temperatures into the distinct ge-netic pathways that determine sex. In the chicken, DMRT1
is required for testicular development, and its gonadal ex-pression is regulated by an intricate DNA methylation-dependent mechanism [Teranishi et al., 2001; Smith et al., 2009]. The male hypermethylated (MHM) region is not methylated in female chicks and produces noncoding RNAs that silence DMRT1 expression [Yang et al., 2010]. In males, DMRT1 expression persists, as the MHM region is hypermethylated and subsequently does not produce the noncoding RNAs associated with the locus. Given the relatively close evolutionary relationships between croco-dilians and birds, it would be interesting if a MHM region or perhaps a similar mechanism was present in crocodil-ians. If so, it remains plausible that it could be tempera-ture sensitive (see fig. 4 ).
DNA methylation in the aromatase (CYP19A1) pro-moter has been linked to sex determination in fish [Con-tractor et al., 2004; Navarro-Martín et al., 2011]. Sex de-termination in the European sea bass is influenced by both genetic factors and temperature. Raising fish at higher temperatures skews sex ratios towards males [Ospina-Alvarez and Piferrer, 2008]. These higher tem-peratures were shown to result in increased methylation of the CYP19A1 promoter and decreased CYP19A1 ex-pression in the sea bass [Navarro-Martín et al., 2011]. In-terestingly, recent work has also demonstrated that the CYP19A1 promoter is hypermethylated in the gonads of American alligator embryos incubated at MPT relative to FPT [Parrott et al., 2014]. Furthermore, the methylation status of other genes involved in sex determination that are expressed before CYP19A1 were examined, and it was observed that the SOX9 promoter is also differentially methylated at MPT and FPT in this species. In contrast to the CYP19A1 promoter, the SOX9 promoter is hyper-methylated in the gonads of embryos incubated at FPT relative to MPT [Parrott et al., 2014]. Recent studies inthe red-eared slider have also shown that the putative CYP19A1 promoter is methylated at elevated levels in those embryos incubated at MPT relative to those at FPT [Matsumoto et al., 2013]. This reported dimorphic meth-ylation originates during the TSP, suggesting that it plays an integral role in sex determination. Thus, it appears that the effect of temperature on CYP19A1 promoter meth-ylation could be conserved across fish and reptiles. It will be interesting to see if thermo-sensitive methylation is a conserved aspect of TSD across other reptiles as well. Un-derstanding how other key genes involved in sex determi-nation are regulated, at both the pre- and post-transcrip-tional levels, should yield insights into the influence of temperature on sex determination.
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
10
Where and what molecule is the initial trigger for TSD? Studies utilizing the ex vivo culture of gonadal tissues have demonstrated that they are able to induce the appropriate sex-specific response to temperature cues in isolation [Moreno-Mendoza et al., 2001; Pieau and Dorizzi, 2004; Shoemaker-Daly et al., 2010]. Despite this observed suffi-ciency of gonads to independently respond properly to temperature cues, it cannot be entirely ruled out that distal temperature-dependent processes act in vivo to influence sex determination. If gonadal tissue is capable of the tem-perature response, then what could be the thermo-sensi-tive trigger? In American alligators, one of the most well-studied thermo-sensitive factors, heat-shock proteins (HSPs), was examined: sexual dimorphic patterns were ob-served for gonadal HSP27 and HSP70A mRNA levels [Kohno et al., 2010]. Although further study is required to evaluate the function of HSPs in sex determination, it is intriguing to think that they or perhaps other thermo-sen-sitive proteins could be acting as the trigger for TSD ( fig. 4 ).
Next-generation sequencing technologies and the newly available genomes of several reptiles are now pro-
viding feasible approaches that will broaden the scope and depth of our current knowledge of TSD. Comparing the gonadal transcriptomes of embryos incubated at ei-ther MPT or FPT will reveal the extent of dimorphic gene expression on both a qualitative and quantitative level. Are there genes with dimorphic expression that have not been previously identified through traditional candidate approaches? Furthermore, aligning sequences resulting from RNA-seq experiments to reference genomes will al-low for the identification of temperature-dependent splice variants. Studies investigating DNA methylation patterning are now open to the entire genome. Cis-regu-latory elements can be identified using comparative ap-proaches, and epigenetic modifications can subsequently be probed. For investigators seeking insights into the mechanistic underpinnings of TSD, the future is certain-ly bright. Indeed the current technology was successful in revealing the phylogenic position of turtles among rep-tiles [Wang et al., 2013]. However, optimism in the field should be tempered by the crucial and ongoing challenge of testing the functionality of these discoveries in nontra-ditional model organisms.
Peculiarities of Reptilian Folliculogenesis
Despite species-specific differences in gene expression timing, function or dosages within developmental net-works regulating gonadal differentiation in reptiles, the overall resulting ovarian follicle structures and functions display strong similarity with general vertebrate patterns [Hernández-Franyutti et al., 2005; DeFalco and Capel, 2009]. Reptilian ovarian morphology shows conservation of discrete cortex-medulla compartments with oocytes developing from oogonial nests in the epithelial cortex and interacting with stromal cells of the underlying me-dulla during follicle formation and maturation. However, variations in this morphological development progres-sion result from species-specific differences between oviparous (egg-laying) and viviparous (live birth) repro-ductive strategies [Hernández-Franyutti et al., 2005] and also in regards to differing patterns of seasonal cyclicity.
The assembly of ovarian follicles is crucial to adult fer-tility, and there are considerable differences in the timing of initial follicular somatic-germ cell interactions among reptiles. Some reptiles form follicles as embryos [Forbes, 1956]; for example, follicle assembly occurs prior to hatching in many species of turtles [Pieau and Dorizzi, 2004]. Similar to birds assembling follicles during the post-hatching period [Callebaut, 1968a, b], the American
Fig. 4. A model of potential mechanisms underlying TSD and dif-ferentiation in reptiles. Depending on egg-incubating temperature (MPT or FPT), an undifferentiated bipotential will become a testis or ovary. The mechanistic mystery of thermo-sensitivity (TS) still exists between temperature and genes, which induces sex determi-nation and differentiation, and they could exhibit cross-talk. We speculate (within the black box) that DNA methylation patterning, alterative splicing, small RNA and heat-shock proteins could pro-vide some of the missing parts driving TSD.
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
11
alligator assembles follicles over a relatively extended post-hatching period of months [Moore et al., 2008, 2010b].
Further, reptiles retain the ability to produce new fol-licles throughout adult life. While mammals and birds transform all oogonia into primary oocytes during em-bryonic development or the beginning of post-natal life, reptiles retain a reserve of oogonia into reproductive adulthood in cortical germinal beds. Oogonia of these germinal beds continue to undergo mitosis, producing a continuous source of new germ cells. The number of ger-minal beds and morphology varies among reptilian spe-cies [Jones et al., 1982], but do not change over the life-time of an individual [Radder and Shine, 2007]. Discrete single or multiple germinal beds have been characterized in squamate ovaries [Guraya and Varma, 1976; Hernán-dez-Franyutti et al., 2005; Moodley and van Wyk, 2007], whereas crocodilians and turtles possess clusters of oogo-nia (nests) distributed throughout the entire ovarian cor-tex, adjacent to previtellogenic follicles [Uribe et al., 1996; Uribe and Guillette, 2000; Calderón et al., 2004; Moore et al., 2010b; Pérez-Bermúdez et al., 2012]. As observed across squamate phylogenies, the number of germinal beds per ovary correlates with clutch size and resulting fecundity [Jones et al., 1982; Radder et al., 2008].
Across reptilian species, follicular cell layers display differences in granulosa cells structure and putative function(s). Turtle and crocodilian ovaries assemble fol-licles with a simple, single-layered, homogeneous granu-losa layer that transforms from cuboidal cells in small fol-licles to low columnar cells with progression of folliculo-genesis, though it remains a monolayer throughout [Calderón et al., 2004]. In contrast, lizards and snakes produce follicles with a stratified, heterogeneous mix of granulosa morphologies and substantially modify the structure of follicular cells during folliculogenesis. Initial-ly, follicles possess a single granulosa layer, but during previtellogenic development, the follicular layer differen-tiates into 3 cell types: small, intermediate and pyriform cells [Uribe et al., 1996; Gómez and Ramírez-Pinilla, 2004]. Most notable in the polymorphic stage of squa-mate follicular cells is the consistent development of large clear pyriform cells; however, the morphology of these pyriform cells varies between species [Vieira et al., 2010]. These cells possess distinctive cytoplasmic projections that cross the notched zona pellucida and directly con-nect the pyriform cells with the oocyte cytoplasm. Cur-rently, a complete understanding of the functions of these bridges is unknown, though pyriform and intermediate cells synthesize and transfer RNA into the oocyte via
these junctions [Motta et al., 1995] and a sustenance func-tion of directly supplying macromolecules and organelles to the oocyte had been observed [van Wyk, 1984; Hernán-dez-Franyutti et al., 2005; Moodley and van Wyk, 2007]. With the initiation of yolk deposition, the stratified fol-licular layer transforms by way of pyriform and interme-diate cells regressing, leaving only a simple granulosalayer of small cells during vitellogenesis. Apoptosis of pyriform and intermediate cells with absorption of cyto-plasmic materials by the oocyte and of the nuclear rem-nants being disposed by the remaining small granulosa cells has been observed [Motta et al., 1995].
Squamate follicular thecal cell layers, derived from the ovarian stroma, can be morphologically homogeneous [Moodley and van Wyk, 2007] or differentiated into mul-tiple cell types [Guraya and Varma, 1976]. In crocodil-ians, turtles, and birds, stromal tissues differentiate into a collagenous lacunar system with chordae and vascular el-ements defining large lymphatic spaces [Calderón et al., 2004; Moore et al., 2010b; Pérez-Bermúdez et al., 2012]. Lacunae morphology varies within and between turtle and crocodilian species with differences in the size of la-cuna and histochemical staining characteristics [Pérez-Bermúdez et al., 2012]. In alligators, fibroblast tissues from this compartment initiate the formation of vascular-ized thecal interna and collagen-rich theca externa, which includes bundles of smooth muscle putatively involved in supporting the developing follicles [Uribe and Guillette, 2000]. Functionally, lacuna could serve as a mechanism to accommodate substantial follicle expansion during de-velopment [Callebaut et al., 1997].
Since early oogenesis and folliculogenesis are highly regulated by endocrine signaling, they must be sensitive to the environment. Indeed, developmental exposure(s) to estrogenic compounds of pharmaceutical (e.g. diethyl-stilbestrol) or environmental (e.g. estrogenic pesticides) origin is associated with higher incidence of the multioo-cytic follicle (polyovular follicle) which correlates with fertilization and/or embryonic survival rates in rodents [Iguchi et al., 1986; Guillette et al., 1994]. The mechanism underlying this pathology has been hypothesized as a dis-ruption of the gonadotropin-estrogen-inhibin/activin signaling cascade [Guillette and Moore, 2006], but it is still poorly understood. Although further investigations on this signaling pathway have begun to elucidate the mechanisms of normal and abnormal folliculogenesis [Moore et al., 2010a, c, d, 2012], reptilian folliculogenesis requires further study. This process could be one of the sensitive endpoints for investigations on the role of con-taminant exposure early in life (see below).
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
12
Reptilian Spermatogenesis
Spermatogenesis begins with progenitor cells, sper-matogonia, that differentiate into spermatozoa via mito-sis and meiosis in seminiferous tubules. In reptiles, the testicular architecture and germ cell arrangement within the seminiferous tubules is similar to that described in other amniotes. Gribbins [2011] recently published an at-las of the morphology and ultrastructure of reptilian sper-matogenesis; however, the molecular mechanisms of rep-tilian spermatogenesis are poorly understood when com-pared with studies in mammals. Reptilian testes have a permanent population of Sertoli cells and a stable blood- testis barrier, the tunica albuginea of the testis [Gribbins, 2011]. Although the basic components of the testis (germ, Sertoli and Leydig cells) are the same among amniotes, reptiles lack a consistent spatial development of germ cells like that observed in birds and mammals. Reptilian germ cell development exhibits seasonal cyclicity like am-phibians, birds and many mammals [Gribbins, 2011]. In reptiles, the majority of germ cell generations progress through the stages of spermatogenesis as a single popula-tion with accompanying cytoplasmic bridges. Mitotic, meiotic and resting spermatogonia reside in the basal compartment depending on the breeding season [Grib-bins et al., 2006].
In seasonal breeders, such as those reptiles found in temperate environments, a single major spermiation event occurs during the breeding season, and only sper-matogonia are present in the testis as it enters the quies-cent period during the nonbreeding season [Gribbins, 2011]. In contrast, new generations of germ cells enter spermiogenesis and spermiation with each wave of sper-matogenesis in continuous breeding reptiles [Gribbins, 2011]. FSH induces recrudescence of spermatogenesis in temperate reptiles either before or after mating [Masson and Guillette, 1985; Licht et al., 1989], whereas FSH stim-ulates mitosis of type A spermatogonia during early sper-matogenesis in mammals [Waits and Setchell, 1990]. The resting spermatogonia in reptiles could be the same as the stem cell spermatogonia in mammals, based on morpho-logical characteristics [Russell, 1990]. Reptilian resting spermatogonia acting as stem cells would provide sper-matogonia populations during each breeding season after their survival during hibernation [Gribbins, 2011]. In wall lizards (Hemidactylus flaviviridis) , a microarray analysis has profiled the transcriptome of 3 different sea-sonal conditions of the testis. The genes expressed in the testis during regression, recrudescent and the active phase of reproductive cycle were associated with specific tes-
ticular morphologies during these phases [Gautam et al., 2013]. This transcriptome profile indicated that the genes associated with maintenance of spermatogonial stem cell were expressed in the regressed phase testis of this lizard such as the C-X-C chemokine receptor type 4. Thus, the results revealed that genes involved in fundamental tes-ticular functions were conserved from reptiles to mam-mals. This transcriptome profile has begun to provide deeper insight into the regulation of reptilian spermato-genesis during the reproductive cycle. However, further investigation is required as this initial study is just begin-ning to explain the morphological and physiological changes observed in reptilian testes. Although American alligators exhibit a distinct testes at 1 month after hatch-ing, only spermatogonia can be identified on histological analysis [Moore et al., 2010b] ( fig. 5 ). The molecular mechanism of the initiation of spermatogenesis is still un-known in the American alligator.
The first and second stages of reptilian spermatogen-esis, like that in mammals, are the mitotic stage and the meiotic stage, respectively. The second meiotic division produces haploid gametes. Prophase 1 of meiosis begins with the spermatogonia B and progressively generates leptotene, zygotene, pachytene, and diplotene spermato-cytes. This phase takes roughly 4 weeks in the reptiles in general, which is longer when compared with other ver-tebrates [Russell, 1990]. The ‘deleted in azoospermia-like’ (DAZL) gene is a germ cell marker and is strongly ex-pressed in the spermatogonia near the basement mem-brane in 6-month-old red-eared slider turtles. In contrast, DAZL mRNA levels were low in the prospermatogonia and proliferating spermatogonia of the hatchling. DAZL mRNA was not detected in the spermatocytes, spermatids or somatic cells including Sertoli and Leydig cells in 6-month-old red-eared sliders [Bachvarova et al., 2009b]. Since DAZL expression was observed in turtle spermato-gonia and DAZL promotes meiosis in mammals [Shen et al., 2012], further analyses examining reptilian spermato-genesis and the function of DAZL and DAZL-related fac-tor are expected to provide a better understanding of these processes.
The third stage, the spermiogenic stage, involves sper-matid morphogenesis and is extensively supported by Sertoli cells. Reptilian spermiogenesis takes 5–8 weeks which is longer than the 4 weeks typically observed in mammals. After differentiation, a highly hydrodynamic motile cell, the spermatozoon, is obtained. The 3 major structures of the reptilian spermatozoon are similar to that of mammals and include the acrosome complex, the nucleus and the flagellum [Gribbins, 2011]. Although the
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
13
number of steps to produce a spermatid is reported to vary among mammalian species (12–19 steps), all reptiles studied to date appear to have a more conserved number of steps, 7–8, to complete spermiogenesis [Gribbins, 2011]. Few studies have examined the details of spermio-genic development at the ultrastructural or genetic level in reptiles. The process of elongation during reptilian spermiogenesis exhibits 2 patterns, with squamates ex-hibiting a filamentous condensation in their chromatin, whereas non-squamate reptiles show a granular conden-sation in the elongating spermatid nucleus [Gribbins, 2011]. However, the contributions of these differences in the elongation process of spermiogenesis on any physio-logical function have not been elucidated as well as the association between the elongation process in spermio-genesis and the different specification/migration of PGCs among the species.
Sexual Determination and Differentiation as
Markers of Environmental Health
In the early 1990s, reports began to appear describing the endocrine disrupting effects of various environmental contaminants [Colborn et al., 1993; Guillette et al., 1994; Tyler et al., 1998]. It became rapidly clear that studies of wildlife – especially studies examining the gonadal devel-opment and differentiation in wildlife – would be critical for an understanding of the role of EDCs in wild popula-tions. Studies of reptilian sex determination and gonadal differentiation have been central to discussions of envi-ronmental health and risk [Iguchi et al., 2001; Hamlin and Guillette, 2011]. Our group has hypothesized that embry-onic exposure to contaminants capable of acting as endo-crine disruptors could induce permanent organization-al – epigenetic – changes in the developing organism [Guillette and Moore, 2006; Guillette and Iguchi, 2012]. In order for this hypothesis to have a factual basis, em-bryos and subsequent neonatal and juveniles must be ex-posed to contaminants that are endocrinologically active and exposed at levels that are physiologically and envi-ronmentally relevant. A growing literature indicates that both of these prerequisites appear to be fulfilled in various populations from fish to humans [Bergman et al., 2012].
We have used the American alligator to address many of these questions and have documented a series of devel-opmental alterations from gene expression to gonadal morphology and function [Guillette et al., 2000; Milnes and Guillette, 2008]. A number of contaminants identi-fied in alligator eggs [Heinz et al., 1991; Giroux, 1998] and
serum [Guillette et al., 1999] exhibit an affinity for the al-ligator estrogen and/or progesterone receptors [Vonier et al., 1996; Guillette et al., 2002] and will activate these re-ceptors in vitro to induce gene expression [Katsu et al., 2004, 2008, 2010]. These data suggest that many of the contaminants found in the embryonic or juvenile envi-ronment have the potential to be EDCs. Further, eggs and juveniles from Lake Apopka, Fla., a lake with many alliga-tors exhibiting gonadal abnormalities, have higher con-
Fig. 5. Three-dimensional reconstruction and histological sections of the male gonad-adrenal-mesonephros (GAM) complex in a 1-month-old American alligator. The 3D-reconstruction (left pan-el) of GAM revealed that the gonad (III, blue), adrenal gland (I, yellow) and mesonephros (II, wired) are tightly connected to each other by connective tissue (light blue). Histological sections (bot-tom panel) of the GAM revealed that the medulla of the gonad was differentiated totally and seminiferous tubules were distinct at one month after hatching. The mesonephros also has differentiated into the epididymis. Histological sections (right panel) were pre-pared by using common methods and were stained with a modi-fied Masson’s trichrome staining. Stained sections were scanned with a digital camera on the light microscope. The pictures were aligned and reconstructed using Reconstruct software [Fiala, 2005]. The figure was modified from Kohno and Guillette [2013].
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
14
centrations of a number of these EDCs when compared to similar samples obtained from alligators living in refer-ence site, Lake Woodruff National Wildlife Refuge, Fla. [Guillette et al., 1999]. Importantly, these chemicals, when combined, exhibit additivity or greater than addi-tivity in the estrogen receptor competitive bindings as-says [Vonier et al., 1996]. Affinity for a receptor does not guarantee that a contaminant has a steroid mimicking ef-fect, as it could equally act as a hormone antagonist [Kelce et al., 1995; Gray et al., 1996]. Experimental testing of these compounds for endocrine disrupting ability is re-quired to further examine these hypotheses.
Various environmentally common pesticides or pesti-cide metabolites can override the temperature-sensitive sex determination mechanisms in alligator or turtle em-bryos, demonstrating that these compounds could act in a manner similar to natural estrogens [Bergeron et al., 1994; Matter et al., 1998; Willingham and Crews, 1999; Milnes et al., 2005]. Contaminants measured in the egg yolk of alligators and capable of altering sex (male to fe-male) in alligator embryos include o,p ′-DDE, p,p ′-DDE, 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin, indole-3-carbinol, trans-nonachlor, and p,p ′-DDD. Further, p,p ′-DDE, a commonly bioaccumulated metabolite of the pesticide DDT, exhibits a variety of actions, estrogenic, anti-andro-genic or neither, depending on the species and endpoint examined [Guillette and Iguchi, 2003; Guillette et al., 2006]. Both isoforms of DDD, also metabolites of DDT, are biologically active, having antagonistic activity on ad-renal steroidogenesis in some species [Brown et al., 1973; Guillette, 2006]. The compound trans-nonachlor is a component of technical grade chlordane, a pesticide used extensively in the past in the US for the treatment of ter-mites, but currently not in use. This compound has a high affinity for the alligator estrogen receptor [Vonier et al., 1996; Guillette et al., 2002] and is capable of altering sex determination in the American alligator [Rooney, 1998]. Each of these compounds are important contaminants in biological systems as they readily bioaccumulate and bio-magnify in the food chain.
The concentrations reported to cause sex reversal, and alterations in gonadal gene expression and steroidogen-esis, are well within the range of concentrations measured in alligator eggs from Lake Apopka, Fla. For example, trans-nonachlor and p,p ′-DDD induce sex reversal at a dose as low as 100 ppb (μg/kg) [Rooney, 1998]. In a previ-ous study, Heinz et al. [1991] observed that alligator eggs collected from Lake Apopka in 1984 and 1985 had great-ly elevated concentrations of p,p ′-DDE: 5.8 ppm wet weight (1984: n = 3 eggs; range = 3.4–7.6 ppm) and 3.5
ppm wet weight (1985: n = 23 eggs; range = 0.89–29 ppm). These concentrations are similar to those observed by Gi-roux [1998] 10 years later (n = 29 eggs; mean ± SE = 4.10 ppm ± 1.27) and are similar to alligator liver concentra-tions measured recently [Garrison et al., 2010]. In addi-tion to p,p ′-DDE, alligator eggs (n = 23) collected in 1985 from Lake Apopka had detectable levels of p,p ′-DDD (ND–1.8 ppm), dieldrin (0.02–1.0 ppm), and cis-chlor-dane (ND–0.25 ppm) [Heinz et al., 1991]. These concen-trations are elevated compared to eggs collected on sev-eral other lakes. Not surprisingly, we have observed that these chemicals, as well as trans-nonachlor, mirex and endrin, are present at ppb concentration in the serum of juvenile alligators from Lake Apopka [Guillette et al., 1999]. If the eggs from Lake Apopka have elevated levels of compounds that can act estrogenically, one could hy-pothesize that the population would exhibit a skewed sex ratio, with a preference for females. We have observed such a phenomenon in a laboratory study [Milnes et al., 2005, 2008], but we do not have data for wild populations.
Additional studies of male reptiles indicate that treat-ment with the fungicide methyl thiophanate damages the seminiferous epithelium and decreases steroid receptor expression, making the exposed lizards infertile [Car-done, 2012]. Similarly, we also have observed a variety of sexual and reproductive alteration associated with envi-ronmental contaminants in male American alligators liv-ing in Florida lakes [Guillette et al., 2000; Guillette, 2006]. Although few molecular studies have been conducted un-til recently, due to the lack of genetic tools for reptiles, a number of reptilian genome projects have been complet-ed or are near completion: the green anole, soft-shell tur-tle, green sea turtle, painted turtle, and American alligator [Alföldi et al., 2011; St John et al., 2012; Shaffer et al., 2013; Wang et al., 2013]. Recent technological advances, such as next-generation sequencing, could help identify many of the molecular mechanisms associated with reptilian sex determination and differentiation and the role of en-vironmental factors on these phenomena.
Conclusions
Reptilian gonadogenesis is an attractive area for re-search as these organisms exhibit a variety of sex determi-nation systems. Further, as TSD and GSD (homozygote or heterozygote) exist in multiple reptilian taxa, they rep-resent sex determination systems that have emerged de novo. Current investigations have revealed that the ge-netic mechanisms used by various reptilian species are,
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
15
not surprisingly, similar to those used by other verte-brates. However, the recent completion or near comple-tion of various reptilian genome projects suggests that new studies examining related species with and without TSD could begin to provide additional insight into the evolution of TSD and GSD in vertebrate ancestors. Novel technologies, such as next-generation sequencing, will be available to examine the naturally occurring evolutionary experiments that have occurred in the various lineages of reptiles. Major questions still remain concerning germ cell migration and specification, the differentiation of go-nadal accessory cells, such as the Sertoli cells and granu-losa cells of the developing testis and ovary, respectively, and the mechanisms by which gene expression is regu-lated during TSD events. Further, reptilian sentinels and their mechanisms of gonadogenesis will likely remain im-portant indicator species for environmental health. Thus, ongoing and new investigations need to tie molecular in-
formation to gonadal morphogenesis and function in reptiles. Such data will not only provide important infor-mation for an understanding of the evolution of these phenomena in vertebrates, but could also provide an im-portant understanding of the health of the environment around us.
Acknowledgements
We thank the colleagues in our laboratories and are grateful for the support from a variety of agencies. This work was funded by the following grants: The Gulf Research Initiative (S.K. and L.J.G.); the National Institute for Basic Biology (T.I.); a Grants-in-Aid for Scientific Research 24370029 (T.I.) from the Ministry of Educa-tion, Culture, Sports, Science and Technology of Japan; a UK-Ja-pan collaboration grant (T.I.) from the Ministry of Environment, Japan; and grants from the Howard Hughes Professors Program and the CoEE Center for Marine Genomics (L.J.G.).
References
Agrawal R, Wessely O, Anand A, Singh L, Aggar-wal RK: Male-specific expression of Sox9 dur-ing gonad development of crocodile and mouse is mediated by alternative splicing of its proline-glutamine-alanine rich domain. FEBS J 276: 4184–4196 (2009).
Alföldi J, Di Palma F, Grabherr M, Williams C, Kong L, et al: The genome of the green anole lizard and a comparative analysis with birds and mammals. Nature 477: 587–591 (2011).
Anand A, Patel M, Lalremruata A, Singh AP, Agrawal R, et al: Multiple alternative splicing of Dmrt1 during gonadogenesis in Indian mugger, a species exhibiting temperature-de-pendent sex determination. Gene 425: 56–63 (2008).
Bachvarova RF, Crother BI, Johnson AD: Evolu-tion of germ cell development in tetrapods: comparison of urodeles and amniotes. Evol Dev 11: 603–609 (2009a).
Bachvarova RF, Crother BI, Manova K, Chatfield J, Shoemaker CM, et al: Expression of Dazl and Vasa in turtle embryos and ovaries: evi-dence for inductive specification of germ cells. Evol Dev 11: 525–534 (2009b).
Badenhorst D, Stanyon R, Engstrom T, Valen-zuela N: A ZZ/ZW microchromosome system in the spiny softshell turtle, Apalone spinifera , reveals an intriguing sex chromosome con-servation in Trionychidae. Chromosome Res 21: 137–147 (2013).
Barske LA, Capel B: Estrogen represses SOX9 during sex determination in the red-eared slider turtle Trachemys scripta . Dev Biol 341: 305–314 (2010).
Baumann H, Casian JAR, Conover DO: Contrast-ing latitudinal variations in vertebral number and sex determination in Pacific versus Atlan-tic silverside fishes. Copeia 2012: 341–350 (2012).
Bell LR, Maine EM, Schedl P, Cline TW: Sex-le-thal, a Drosophila sex determination switch gene, exhibits sex-specific RNA splicing and sequence similarity to RNA binding proteins. Cell 55: 1037–1046 (1988).
Bergeron JM, Crews D, McLachlan JA: PCBs as environmental estrogens: turtle sex determi-nation as a biomarker of environmental con-tamination. Environ Health Perspect 102: 780–781 (1994).
Bergman A, Heindel J, Jobling S, Kidd K, Zoeller RT (eds): State of the science of endocrine dis-rupting chemicals 2012, in WHO/UNEP (Ge-neva 2012).
Brown RD, Nicholson WE, Chick WT, Strott CA: Effect of o,p’-DDD on human adrenal steroid 11β-hydroxylation activity. J Clin Endocrinol Metab 36: 730–733 (1973).
Bull JJ: Sex determination in reptiles. Quart Rev Biol 55: 3–21 (1980).
Bull JJ, Bulmer MG: Longevity enhances selection of environmental sex determination. Heredi-ty (Edinb) 63: 315–320 (1989).
Bull JJ, Gutzke WH, Crews D: Sex reversal by es-tradiol in three reptilian orders. Gen Comp Endocrinol 70: 425–428 (1988).
Burtis KC, Baker BS: Drosophila doublesex gene controls somatic sexual differentiation by producing alternatively spliced mRNAs en-coding related sex-specific polypeptides. Cell 56: 997–1010 (1989).
Calderón ML, De Pérez GR, Ramíres Pinilla MPR: Morphology of the ovary of Caiman crocodi-lus (Crocodylia: Alligatoridae). Ann Anat 186: 13–24 (2004).
Callebaut M: [ 3 H]Uridine incorporation during previtellogenesis and early vitellogenesis in oocytes of chick (Gallus gallus) . J Embryol Exp Morphol 20: 169–174 (1968a).
Callebaut M: Extracorporal development of quail oocytes. Experientia 24: 1242–1243 (1968b).
Callebaut M, Van Nassauw L, Harrisson F: Com-parison between oogenesis and related ovari-an structures in a reptile, Pseudemys scripta elegans (turtle) and in a bird Coturnix cotur-nix japonica (quail). Reprod Nutr Dev 37: 233–252 (1997).
Cardone A: Testicular toxicity of methyl thio-phanate in the Italian wall lizard (Podarcis sicula) : morphological and molecular evalua-tion. Ecotoxicology 21: 512–523 (2012).
Chapus C, Edwards SV: Genome evolution in Rep-tilia: in silico chicken mapping of 12,000 BAC-end sequences from two reptiles and a basal bird. BMC Genomics 10(suppl 2):S8 (2009).
Charnov EL, Bull J: When is sex environmentally determined? Nature 266: 828–830 (1977).
Colborn T, vom Saal FS, Soto AM: Developmen-tal effects of endocrine-disrupting chemicals in wildlife and humans. Environ Health Per-spect 101: 378–384 (1993).
Cole CJ: Karyotypes of the five monotypic species groups of lizards in the genus Sceloporus . Am Mus Novit 2450: 1–17 (1971).
Conover DO: Adaptive significance of tempera-ture-dependent sex determination in a fish. Am Nat 123: 297–313 (1984).
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
16
Contractor RG, Foran CM, Li S, Willett KL: Evi-dence of gender- and tissue-specific promoter methylation and the potential for ethinyl-estradiol-induced changes in Japanese meda-ka (Oryzias latipes) estrogen receptor and aro-matase genes. J Toxicol Environ Health A 67: 1–22 (2004).
Crain DA, Guillette LJ Jr, Rooney AA, Pickford DB: Alterations in steroidogenesis in alliga-tors (Alligator mississippiensis) exposed natu-rally and experimentally to environmental contaminants. Environ Health Perspect 105: 528–533 (1997).
Crews D, Wibbels T, Gutzke WHN: Action of sex steroid hormones on temperature-induced sex determination in the snapping turtle (Chelydra serpentina) . Gen Comp Endocrinol 76: 159–166 (1989).
Deeming DC: Prevalence of TSD in crocodilians, in Valenzuela N. Lance VA (eds): Tempera-ture-Dependent Sex Determination in Verte-brates (Smithsonian Books, Washington DC 2004).
Deeming DC, Ferguson MWJ: Effects of incuba-tion temperature on the growth and develop-ment of embryos of Alligator mississippiensis . J Comp Physiol B 159: 183–193 (1989).
DeFalco T, Capel B: Gonad morphogenesis in vertebrates: divergent means to a convergent end. Annu Rev Cell Dev Biol 25: 457–482 (2009).
De Santa Barbara P, Bonneaud N, Boizet B, Des-clozeaux M, Moniot B, et al: Direct interac-tion of SRY-related protein SOX9 and ste-roidogenic factor 1 regulates transcription of the human anti-Müllerian hormone gene. Mol Cell Biol 18: 6653–6665 (1998).
DiNapoli L, Capel B: Germ cell depletion does not alter the morphogenesis of the fetal testis or ovary in the red-eared slider turtle (Trache-mys scripta) . J Exp Zool B Mol Dev Evol 308: 236–241 (2007).
Dorizzi M, Richard-Mercier N, Desvages G, Gi-rondot M, Pieau C: Masculization of gonads by aromatase inhibitors in a turtle with tem-perature-dependent sex determination. Dif-ferentiation 58: 1–8 (1994).
Dournon C, Houillon C, Pieau C: Temperature sex-reversal in amphibians and reptiles. Int J Dev Biol 34: 81–92 (1990).
Evans CT, Ledesma DB, Schulz TZ, Simpson ER, Mendelson CR: Isolation and characteriza-tion of a complementary DNA specific for hu-man aromatase-system cytochrome P-450 mRNA. Proc Natl Acad Sci USA 83: 6387–6391 (1986).
Ewert MA, Jackson DR, Nelson CE: Patterns of temperature-dependent sex determination in turtles. J Exp Zool 270: 3–15 (1994).
Extavour CG, Akam M: Mechanisms of germ cell specification across the metazoans: epigenesis and preformation. Development 130: 5869–5884 (2003).
Eyal-Giladi H, Ginsburg M, Farbarov A: Avian primordial germ cells are of epiblastic origin. J Embryol Exp Morphol 65: 139–147 (1981).
Ezaz T, Stiglec R, Veyrunes F, Marshall Graves JA: Relationships between vertebrate ZW and XY sex chromosome systems. Curr Biol 16:R736–R743 (2006).
Ferguson MWJ: Reproductive biology and em-bryology of the crocodilians, in Gans C, Billett FS, Maderson PFA (eds): Biology of the Rep-tilia: Development, vol 14, pp 329–491 (J. Wi-ley and Sons, New York 1985).
Fiala JC: Reconstruct: a free editor for serial sec-tion microscopy. J Microsc 218: 52–61 (2005).
Forbes TR: The development of the reproductive system of a lizard, Anolis carolinensis . Am J Anat 98: 139–157 (1956).
Foster JW, Dominguez-Steglich MA, Guioli S, Kwok C, Weller PA, et al: Campomelic dys-plasia and autosomal sex reversal caused by mutations in an SRY -related gene. Nature 372: 525–530 (1994).
Fujimoto T, Ukeshima A, Miyayama Y, Horio F, Ninomiya E: Observations of primordial germ cells in the turtle embryo (Caretta caret-ta) : light and electron microscopic studies. Dev Growth Differ 21: 3–10 (1979).
Gabriel WN, Blumberg B, Sutton S, Place AR, Lance VA: Alligator aromatase cDNA se-quence and its expression in embryos at male and female incubation temperatures. J Exp Zool 290: 439–448 (2001).
Garrison AW, Guillette LJ Jr, Wiese TE, Avants JK: Persistent organochlorine pesticides and their metabolites in alligator livers from Lakes Apopka and Woodruff, Florida, USA. Int J Environ Anal Chem 90: 159–170 (2010).
Gautam M, Mathur A, Khan MA, Majumdar SS, Rai U: Transcriptome analysis of spermato-genically regressed, recrudescent and active phase testis of seasonally breeding wall lizards Hemidactylus flaviviridis . PLoS One 8:e58276 (2013).
Gilbert SF: Developmental Biology, ed 9 (Sinauer Associates, Sunderland 2010).
Ginsburg M, Eyal-Giladi H: Primordial germ cells of the young chick blastoderm originate from the central zone of the area pellucida irrespec-tive of the embryo-forming process. Develop-ment 101: 209–219 (1987).
Giroux DJ: Lake Apopka Revisited: A Correla-tional Analysis of Nesting Anomalies and DDT Contaminants, thesis (University of Florida, Gainesville 1998).
Godfrey MH, Delmas V, Girondot M: Assess-ment of patterns of temperature-dependent sex determination using maximum likelihood model selection. Ecoscience 10: 265–272 (2003).
Gómez D, Ramírez-Pinilla MP: Ovarian histology of the placentotrophic Mabuya mabouya (Squamata, Scincidae). J Morphol 259: 90–105 (2004).
Gray LE Jr, Monosson E, Kelce WR: Emerging is-sues: the effects of endocrine disrupters on re-productive development, in DiGiulio RT, Monosson E (eds): Interconnections between Human and Ecosystem Health, pp 45–82 (Chapman and Hall, London 1996).
Gribbins KM: Reptilian spermatogenesis: a histo-logical and ultrastructural perspective. Sper-matogenesis 1: 250–269 (2011).
Gribbins KM, Elsey RM, Gist DH: Cytological evaluation of the germ cell development strat-egy within the testis of the American alligator, Alligator mississippiensis . Acta Zool 87: 59–69 (2006).
Guillette LJ Jr: Endocrine disrupting contami-nants – beyond the dogma. Environ Health Perspect 114(suppl 1): 9–12 (2006).
Guillette LJ Jr, Iguchi T: Contaminant-induced endocrine and reproductive alterations in reptiles. Pure Appl Chem 75: 2275–2286 (2003).
Guillette LJ Jr, Iguchi T: Ecology. Life in a con-taminated world. Science 337: 1614–1615 (2012).
Guillette LJ Jr, Moore BC: Environmental con-taminants, fertility, and multioocytic follicles: a lesson from wildlife? Semin Reprod Med 24: 134–141 (2006).
Guillette LJ Jr, Gross TS, Masson GR, Matter JM, Percival HF, Woodward AR: Developmental abnormalities of the gonad and abnormal sex hormone concentrations in juvenile alligators from contaminated and control lakes in Flor-ida. Environ Health Perspect 102: 680–688 (1994).
Guillette LJ Jr, Brock JW, Rooney AA, Woodward AR: Serum concentrations of various envi-ronmental contaminants and their relation-ship to sex steroid concentrations and phallus size in juvenile American alligators. Arch En-viron Contam Toxicol 36: 447–455 (1999).
Guillette LJ Jr, Crain DA, Gunderson MP, Kools SAE, Milnes MR, et al: Alligators and endo-crine disrupting contaminants: a current per-spective. Am Zool 40: 438–452 (2000).
Guillette LJ Jr, Vonier PM, McLachlan JA: Affin-ity of the alligator estrogen receptor for serum pesticide contaminants. Toxicology 181–182: 151–154 (2002)
Guillette LJ Jr, Kools S, Gunderson MP, Bermu-dez DS: DDT and its analogues: new insights into their endocrine disruptive effects on wildlife, in Norris DO, Carr JA (eds): Endo-crine Disruption: Biological Basis for Health Effects in Wildlife and Humans, pp 332–355 (Oxford University Press, New York 2006).
Guraya SS, Varma SK: Morphology of ovarian changes during reproductive-cycle of house lizard, Hemidactylus flaviviridis . Acta Mor-phol Neerl Scand 14: 165–191 (1976).
Hamlin HJ, Guillette LJ Jr: Embryos as targets of endocrine disrupting contaminants in wild-life. Birth Defects Res C Embryo Today 93: 19–33 (2011).
Heinz GH, Percival HF, Jennings ML: Contami-nants in American alligator eggs from Lake Apopka, Lake Griffin and Lake Okeechobee, Florida. Environ Monit Assess 16: 277–285 (1991).
Hernández-Franyutti A, Uribe Aranzábal MC, Guillette LJ Jr: Oogenesis in the viviparous matrotrophic lizard Mabuya brachypoda . J Morphol 265: 152–164 (2005).
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
17
Hubert J: Origin and development of oocytes, in Gans C (ed): Biology of the Reptilia, vol 14, pp 43–50 (Wiley, New York 1969).
Iguchi T, Takasugi N, Bern HA, Mills KT: Fre-quent occurrence of polyovular follicles in ovaries of mice exposed neonatally to diethyl-stilbestrol. Teratology 34: 29–35 (1986).
Iguchi T, Watanabe H, Katsu Y: Developmental effects of estrogenic agents on mice, fish, and frogs: a mini-review. Horm Behav 40: 248–251 (2001).
Janzen FJ, Paukstis GL: A preliminary test of the adaptive significance of environmental sex determination in reptiles. Evolution 45: 435–440 (1991).
Janzen FJ, Krenz JG: Phylogenetics: which was first, TSD or GSD, in Valenzuela N, Lance VA (eds): Temperature-Dependent Sex Determi-nation in Vertebrates, (Smithsonian Books, Washington DC 2004).
Jeyasuria P, Roosenburg WM, Place AR: Role of P-450 aromatase in sex determination of the diamondback terrapin, Malaclemys terrapin . J Exp Zool 270: 95–111 (1994).
Jones RE, Swain T, Guillette LJ Jr, Fitzgerald KT: The comparative anatomy of lizard ovaries, with emphasis on the number of germinal beds. J Herpetol 16: 240–252 (1982).
Katsu Y, Bermudez DS, Braun EL, Helbing C, Mi-yagawa S, Gunderson MP, et al: Molecular cloning of the estrogen and progesterone re-ceptors of the American alligator. Gen Comp Endocrinol 136: 122–133 (2004).
Katsu Y, Ichikawa R, Ikeuchi T, Kohno S, Guil-lette LJ Jr, Iguchi T: Molecular cloning and characterization of estrogen, androgen, and progesterone nuclear receptors from a fresh-water turtle (Pseudemys nelsoni) . Endocrinol-ogy 149: 161–173 (2008).
Katsu Y, Matsubara K, Kohno S, Matsuda Y, To-riba M, et al: Molecular cloning, characteriza-tion, and chromosome mapping of reptilian estrogen receptors. Endocrinology 151: 5710–5720 (2010).
Kelce WR, Stone CR, Laws SC, Gray LE, Kemp-painen JA, Wilson EM: Persistent DDT me-tabolite p,p ′-DDE is a potent androgen recep-tor antagonist. Nature 375: 581–585 (1995).
Kobayashi Y, Sunobe T, Kobayashi T, Nagahama Y, Nakamura M: Promoter analysis of two aromatase genes in the serial-sex changing gobiid fish, Trimma okinawae . Fish Physiol Biochem 31: 123–127 (2005).
Kohno S, Guillette LJ Jr: Endocrine disruption and reptiles: Using the unique attributes of temperature-dependent sex determination to assess impacts, in Matthiessen P (ed): Endo-crine Disrupters: Hazard Testing and Assess-ment Methods (John Wiley & Sons, Hoboken 2013).
Kohno S, Katsu Y, Urushitani H, Ohta Y, Iguchi T, Guillette LJ Jr: Potential contributions of heat shock proteins to temperature-depen-dent sex determination in the American alli-gator. Sex Dev 4: 73–87 (2010).
Lance VA, Bogart MH: Disruption of ovarian de-velopment in alligator embryos treated with
an aromatase inhibitor. Gen Comp Endocri-nol 86: 59–71 (1992).
Lang JW, Andrews HV: Temperature-dependent sex determination in crocodilians. J Exp Zool 270: 28–44 (1994).
Licht P, Denver RJ, Pavgi S: Temperature depen-dence of in vitro pituitary, testis, and thyroid secretion in a turtle, Pseudemys scripta . Gen Comp Endocrinol 76: 274–285 (1989).
Masson GR, Guillette LJ Jr: FSH-induced gonadal development in juvenile lizards, Eumeces ob-soletus . J Exp Zool A Comp Exp Biol 236: 343–351 (1985).
Matsumoto Y, Crews D: Molecular mechanisms of temperature-dependent sex determination in the context of ecological developmental bi-ology. Mol Cell Endocrinol 354: 103–110 (2012).
Matsumoto Y, Buemio A, Chu R, Vafaee M, Crews D: Epigenetic control of gonadal aro-matase (cyp19a1) in temperature-dependent sex determination of red-eared slider turtles. PLoS One 8:e63599 (2013).
Matter JM, McMurry CS, Anthony AB, Dicker-son RL: Development and implementation of endocrine biomarkers of exposure and effects in American alligators (Alligator mississippi-ensis) . Chemosphere 37: 1905–1914 (1998).
McPhaul MJ, Noble JF, Simpson ER, Mendelson CR, Wilson JD: The expression of a function-al cDNA encoding the chicken cytochrome P-450arom (aromatase) that catalyzes the for-mation of estrogen from androgen. J Biol Chem 263: 16358–16363 (1988).
Milnes MR, Guillette LJ Jr: Alligator tales: new les-sons about environmental contaminants from a sentinel species. Bioscience 58: 1027–1036 (2008).
Milnes MR, Roberts R, Guillette LJ Jr: Effects of incubation temperature and estrogen expo-sure on aromatase activity in the brain and gonads of embryonic alligators. Environ Health Perspect 110(suppl 3): 393–396 (2002).
Milnes MR, Bryan TA, Medina JG, Gunderson MP, Guillette LJ Jr: Developmental altera-tions as a result of in ovo exposure to the pes-ticide metabolite p,p ′-DDE in Alligator mis-sissippiensis . Gen Comp Endocrinol 144: 257–263 (2005).
Milnes MR, Bryan TA, Katsu Y, Kohno S, Moore BC, et al: Increased posthatching mortality and loss of sexually dimorphic gene expres-sion in alligators (Alligator mississippiensis) from a contaminated environment. Biol Re-prod 78: 932–938 (2008).
Moodley GK, van Wyk JH: Folliculogenesis and ovarian histology of the oviparous gecko, Hemidactylus mabouia (Sauria: Gekkonidae). Afr J Herpetol 56: 115–135 (2007).
Moore BC, Uribe-Aranzabal MC, Boggs AS, Guil-lette LJ Jr: Developmental morphology of the neonatal alligator (Alligator mississippiensis) ovary. J Morphol 269: 302–312 (2008).
Moore BC, Hamlin HJ, Botteri NL, Guillette LJ Jr: Gonadal mRNA expression levels of TGFβ superfamily signaling factors correspond with post-hatching morphological develop-
ment in American alligators. Sex Dev 4: 62–72 (2010a).
Moore BC, Hamlin HJ, Botteri NL, Lawler AN, Mathavan KK, Guillette LJ Jr: Posthatching development of Alligator mississippiensis ova-ry and testis. J Morphol 271: 580–595 (2010b).
Moore BC, Kohno S, Cook RW, Alvers AL, Ham-lin HJ, et al: Altered sex hormone concentra-tions and gonadal mRNA expression levels of activin signaling factors in hatchling alliga-tors from a contaminated Florida lake. J Exp Zool A Ecol Genet Physiol 313: 218–230 (2010c).
Moore BC, Milnes MR, Kohno S, Katsu Y, Iguchi T, Guillette LJ Jr: Influences of sex, incubation temperature, and environmental quality on gonadal estrogen and androgen receptor mes-senger RNA expression in juvenile American alligators (Alligator mississippiensis) . Biol Re-prod 82: 194–201 (2010d).
Moore BC, Roark AM, Kohno S, Hamlin HJ, Guillette LJ Jr: Gene-environment interac-tions: the potential role of contaminants in somatic growth and the development of the reproductive system of the American alliga-tor. Mol Cell Endocrinol 354: 111–120 (2012).
Morais da Silva S, Hacker A, Harley V, Goodfel-low P, Swain A, Lovell-Badge R: Sox9 expres-sion during gonadal development implies a conserved role for the gene in testis differen-tiation in mammals and birds. Nat Genet 14: 62–68 (1996).
Moreno-Mendoza N, Harley VR, Merchant-La-rios H: Differential expression of SOX9 in go-nads of the sea turtle Lepidochelys olivacea at male- or female-promoting temperatures. J Exp Zool 284: 705–710 (1999).
Moreno-Mendoza N, Harley VR, Merchant-La-rios H: Temperature regulates SOX9 expres-sion in cultured gonads of Lepidochelys oliva-cea , a species with temperature sex determi-nation. Dev Biol 229: 319–326 (2001).
Morrish BC, Sinclair AH: Vertebrate sex determi-nation: many means to an end. Reproduction 124: 447–457 (2002).
Motta CM, Scanderbeg MC, Filosa S, Andreuc-cetti P: Role of pyriform cells during the growth of oocytes in the lizard Podarcis sicula . J Exp Zool 273: 247–256 (1995).
Murdock C, Wibbels T: Cloning and expression of aromatase in a turtle with temperature-de-pendent sex determination. Gen Comp Endo-crinol 130: 109–119 (2003).
Navarro-Martín L, Viñas J, Ribas L, Díaz N, Gutiérrez A, et al: DNA methylation of the gonadal aromatase (cyp19a) promoter is in-volved in temperature-dependent sex ratio shifts in the European sea bass. PLoS Genet 7:e1002447 (2011).
Oreal E, Pieau C, Mattei MG, Josso N, Picard JY, et al: Early expression of AMH in chicken em-bryonic gonads precedes testicular SOX9 ex-pression. Dev Dyn 212: 522–532 (1998).
Organ CL, Janes DE, Meade A, Pagel M: Geno-typic sex determination enabled adaptive ra-diations of extinct marine reptiles. Nature 461: 389–392 (2009).
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Kohno /Parrott /Yatsu /Miyagawa /Moore /Iguchi /Guillette Jr.
Sex DevDOI: 10.1159/000358892
18
Ospina-Alvarez N, Piferrer F: Temperature-de-pendent sex determination in fish revisited: prevalence, a single sex ratio response pattern, and possible effects of climate change. PLoS One 3:e2837 (2008).
Packard GC, Packard MJ, Miller K, Boardman TJ: Influence of moisture, temperature and sub-strate on snapping turtle eggs and embryos. Ecology 68: 983–993 (1987).
Parrott BB, Kohno S, Cloy-McCoy JA, Guillette LJ Jr: Differential incubation temperatures re-sult in dimorphic DNA methylation pattern-ing of the SOX9 and aromatase promoters in gonads of alligator (Alligator mississippiensis) embryos. Biol Reprod 90:2 (2014).
Pérez-Bermúdez E, Ruiz-Urquiola A, Lee-González I, Petric B, Almaguer-Cuenca N, et al: Ovarian follicular development in the hawksbill turtle (Cheloniidae: Eretmochelys imbricata L.). J Morphol 273: 1338–1352 (2012).
Pieau C, Dorizzi M: Oestrogens and temperature-dependent sex determination in reptiles: all is in the gonads. J Endocrinol 181: 367–377 (2004).
Pieau C, Dorizzi M, Richard-Mercier N, Desvages G: Sexual differentiation of gonads as a func-tion of temperature in the turtle Emys orbicu-laris : endocrine function, intersexuality and growth. J Exp Zool 281: 400–408 (1998).
Pokorna M, Kratochvil L: Phylogeny of sex-deter-mining mechanisms in squamate reptiles: are sex chromosomes an evolutionary trap? Zool J Linn Soc-Lond 156: 168–183 (2009).
Radder R, Shine R: Germinal bed condition in a polyautochronic single-clutched lizard, Bas-siana duperreyi (Scincidae). Amphib-reptil 28: 159–162 (2007).
Radder RS, Pizzatto L, Shine R: Morphological correlates of life-history variation: is lizard clutch size related to the number of germinal beds in the ovary? Biol J Linn Soc Lond 94: 81–88 (2008).
Raymond CS, Shamu CE, Shen MM, Seifert KJ, Hirsch B, et al: Evidence for evolutionary con-servation of sex-determinng genes. Nature 391: 691–695 (1998).
Rhen T, Lang JW: Temperature-dependent sex determination in the snapping turtle: manip-ulation of the embryonic sex steroid environ-ment. Gen Comp Endocrinol 96: 243–254 (1994).
Richard-Mercier N, Dorizzi M, Desvages G, Gi-rondot M, Pieau C: Endocrine sex reversal of gonads by the aromatase inhibitor Letrozole (CGS 20267) in Emys orbicularis , a turtle with temperature-dependent sex determination. Gen Comp Endocrinol 100: 314–326 (1995).
Rooney AA: Variation in the endocrine and im-mune system of juvenile alligators: environ-mental influence on physiology. Thesis, Gainesville, University of Florida (1998).
Russell LD: Histological and Histopathological Evaluation of the Testis, ed 1 (Cache River Press, Clearwater 1990).
Shaffer HB, Minx P, Warren DE, Shedlock AM, Thomson RC, et al: The western painted turtle genome, a model for the evolution of extreme physiological adaptations in a slowly evolving lineage. Genome Biol 14:R28 (2013).
Shen W, Park BW, Toms D, Li J: Midkine pro-motes proliferation of primordial germ cells by inhibiting the expression of the deleted in azoospermia-like gene. Endocrinology 153: 3482–3492 (2012).
Shoemaker C, Ramsey M, Queen J, Crews D: Ex-pression of Sox9 , Mis , and Dmrt1 in the gonad of a species with temperature-dependent sex determination. Dev Dyn 236: 1055–1063 (2007).
Shoemaker CM, Crews D: Analyzing the coordi-nated gene network underlying temperature-dependent sex determination in reptiles. Se-min Cell Dev Biol 20: 293–303 (2009).
Shoemaker-Daly CM, Jackson K, Yatsu R, Matsu-moto Y, Crews D: Genetic network underly-ing temperature-dependent sex determina-tion is endogenously regulated by tempera-ture in isolated cultured Trachemys scripta gonads. Dev Dyn 239: 1061–1075 (2010).
Sifuentes-Romero I, Merchant-Larios H, Milton SL, Moreno-Mendoza N, Díaz-Hernández V, García-Gasca A: RNAi-mediated gene silenc-ing in a gonad organ culture to study sex de-termination mechanisms in sea turtle. Genes 4: 293–305 (2013).
Smith CA, Joss JMP: Steroidogenic enzyme activ-ity and ovarian differentiation in the saltwater crocodile, Crocodylus porosus . Gen Comp En-docrinol 93: 232–245 (1994).
Smith CA, Elf PK, Lang JW, Joss JMP: Aromatase enzyme-activity during gonadal sex-differen-tiation in alligator embryos. Differentiation 58: 281–290 (1995).
Smith CA, McClive PJ, Western PS, Reed KJ, Sin-clair AH: Conservation of a sex-determining gene. Nature 402: 601–602 (1999).
Smith CA, Roeszler KN, Ohnesorg T, Cummins DM, Farlie PG, et al: The avian Z-linked gene DMRT1 is required for male sex determina-tion in the chicken. Nature 461: 267–271 (2009).
Spotila LD, Spotila JR, Hall SE: Sequence and ex-pression analysis of WT1 and Sox9 in the red-eared slider turtle, Trachemys scripta . J Exp Zool 281: 417–427 (1998).
St John JA, Braun EL, Isberg SR, Miles LG, Chong AY, et al: Sequencing three crocodilian ge-nomes to illuminate the evolution of archo-saurs and amniotes. Genome Biol 13: 415 (2012).
Tanaka M, Telecky TM, Fukada S, Adachi S, Chen S, Nagahama Y: Cloning and sequence analy-sis of the cDNA encoding P-450 aromatase (P450arom) from a rainbow trout (Oncorhyn-chus mykiss) ovary; relationship between the amount of P450arom mRNA and the produc-tion of oestradiol-17β in the ovary. J Mol En-docrinol 8: 53–61 (1992).
Teranishi M, Shimada Y, Hori T, Nakabayashi O, Kikuchi T, et al: Transcripts of the MHM re-gion on the chicken Z chromosome accumu-late as non-coding RNA in the nucleus of fe-male cells adjacent to the DMRT1 locus. Chromosome Res 9: 147–165 (2001).
Tokunaga S: Temperature-dependent sex deter-mination in Gekko japonicus (Gokkonidae, Reptilia). Dev Growth Differ 27: 117–120 (1985).
Torres Maldonado LC, Landa Piedra A, Moreno Mendoza N, Marmolejo Valencia A, Meza Martínez A, Merchant Larios H: Expression profiles of Dax1 , Dmrt1 , and Sox9 during temperature sex determination in gonads of the sea turtle Lepidochelys olivacea . Gen Comp Endocrinol 129: 20–26 (2002).
Tran D, Muesy-Dessole N, Josso N: Anti-Mülle-rian hormone is a functional marker of foetal Sertoli cells. Nature 269: 411–412 (1977).
Tsunekawa N, Naito M, Sakai Y, Nishida T, Noce T: Isolation of chicken vasa homolog gene and tracing the origin of primordial germ cells. Development 127: 2741–2750 (2000).
Tyler CR, Jobling S, Sumpter JP: Endocrine dis-ruption in wildlife: a critical review of the evi-dence. Crit Rev Toxicol 28: 319–361 (1998).
Uchida D, Yamashita M, Kitano T, Iguchi T: Oo-cyte apoptosis during the transition from ova-ry-like tissue to testes during sex differentia-tion of juvenile zebrafish. J Exp Biol 205: 711–718 (2002).
Uribe MCA, Guillette LJ Jr: Oogenesis and ovar-ian histology of the American alligator Alliga-tor mississippiensis . J Morphol 245: 225–240 (2000).
Uribe MDCA, Portales GL, Guillette LJ Jr: Ovar-ian folliculogenesis in the oviparous Mexican lizard Ctenosaura pectinata . J Morphol 230: 99–112 (1996).
Urushitani H, Katsu Y, Miyagawa S, Kohno S, Ohta Y, et al: Molecular cloning of anti-Mül-lerian hormone from the American alligator, Alligator mississippiensis . Mol Cell Endocri-nol 333: 190–199 (2011).
Valleley EM, Harrison CJ, Cook Y, Ferguson MW, Sharpe PT: The karyotype of Alligator mississippiensis , and chromosomal mapping of the ZFY/X homologue, Zfc . Chromosoma 103: 502–507 (1994).
Valleley EMA, Cartwright EJ, Croft NJ, Markham AF, Coletta PL: Characterisation and expres-sion of Sox9 in the leopard gecko, Eublepharis macularius . J Exp Zool 291: 85–91 (2001).
van Wyk JH: Ovarian morphological changes during the annual breeding cycle of the rock lizard Agama atra (Sauria: Agamidae). Navor Nasione Museum 4: 237–275 (1984).
Vidal VPI, Charboissler MC, de Roolj DG, Schedl A: Sox9 induces testis development in XX transgenic mice. Nat Genet 28: 216–217 (2001).
Vieira S, de Pérez GR, Ramírez-Pinilla MP: Ultra-structure of the ovarian follicles in the placen-totrophic andean lizard of the genus Mabuya (Squamata: Scincidae). J Morphol 271: 738–749 (2010).
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M
Gonadal Differentiation in Reptiles Determining Sex by Environment
Sex DevDOI: 10.1159/000358892
19
Vonier PM, Crain DA, McLachlan JA, Guillette LJ Jr, Arnold SF: Interaction of environmental chemicals with the estrogen and progesterone receptors from the oviduct of the American alligator. Environ Health Perspect 104: 1318–1322 (1996).
Waits GMH, Setchell BP: Physiology of the mam-malian testis, in Lemming GE (ed): Marshall’s Physiology of Reproduction Reproduction in the Male, vol 2, pp 1–105 (Churchill Living-stone, London 1990).
Wallace H, Badawy GMI, Wallace BMN: Am-phibian sex determination and sex reversal. Cell Mol Life Sci 55: 901–909 (1999).
Wang Z, Pascual-Anaya J, Zadissa A, Li W, Nii-mura Y, et al: The draft genomes of soft-shell turtle and green sea turtle yield insights into the development and evolution of the turtle-specific body plan. Nat Genet 45: 701–706 (2013).
Warner DA, Shine R: The adaptive significance of temperature-dependent sex determination in a reptile. Nature 451: 566–568 (2008).
Western PS, Harry JL, Graves JA, Sinclair AH: Temperature-dependent sex determination: upregulation of SOX9 expression after com-mitment to male development. Dev Dyn 214: 171–177 (1999).
Wibbels T, Crews D: Putative aromatase inhibitor induces male sex determination in a female unisexual lizard and in a turtle with tempera-ture-dependent sex determination. J Endocri-nol 141: 295–299 (1994).
Wibbels T, Crews D: Steroid-induced sex deter-mination at incubation temperatures produc-ing mixed sex ratios in a turtle with TSD. Gen Comp Endocrinol 100: 53–60 (1995).
Wibbels T, Bull JJ, Crews D: Steroid hormone-induced male sex determination in an amni-otic vertebrate. J Exp Zool 262: 454–457 (1992).
Willingham E, Crews D: Sex reversal effects of en-vironmentally relevant xenobiotic concentra-tions on the red-eared slider turtle, a species with temperature-dependent sex determina-tion. Gen Comp Endocrinol 113: 429–435 (1999).
Yang X, Zheng J, Xu G, Qu L, Chen S, et al: Exog-enous cMHM regulates the expression of DMRT1 and ER α in avian testes. Mol Biol Rep 37: 1841–1847 (2010).
Yao HHC, Capel B: Temperature, genes, and sex: a comparative view of sex determination in Trachemys scripta and Mus musculus . J Bio-chem 138: 5–12 (2005).
Dow
nloa
ded
by:
Nat
iona
l Lib
rary
of M
edic
ine
13
0.14
.254
.26
- 4/
7/20
14 3
:25:
30 P
M