Marsupials are born with undifferentiated gonads, and their reproductive organs differentiate consecutively, not simultaneously as in eutherian mammals. Thus, in the main marsupial model, the tammar wallaby, Macropus eugenii, the testis forms cords 2 days after birth, the ovary develops cortex and medulla about 8 days after birth, the Wolffian duct enlarges from day 10, the prostate begins to form prostatic buds about 25 days after birth, and the phallus does not become sexually dimorphic until after 50 days postpartum (pp). The brain responses also become sexually dimorphic relatively late in development, after day 25 pp. This relatively elongated period of differentiation has allowed experimental manipulation at each stage of development to induce often dramatic sex reversal of both internal and external genitalia.

Marsupials are remarkable models for the study of sexual differentiation, because all differentiation happens after birth when the young are in a pouch or firmly attached to teats in the pouchless marsupials. The presence of a pouch is not a universal characteristic of marsupials, and many species in both Australian and South American species are pouchless. In addition, in marsupial males the scrotum is cranial to the penis - the opposite anatomical arrangement to that seen in scrotal eutherian mammals. Even more useful for studies of sexual development is the fact that each of the structures differentiate at separate times postnatally [reviewed in Renfree et al., 1995]. Marsupials are born with undifferentiated gonads, and in the tammar, testes cords are first detected at around day 2 postpartum (pp), Wolffian ducts start to regress in the female by day 10, and Müllerian ducts in the male similarly begin to regress at around day 10 [Renfree et al., 1996]. The prostate begins to form prostatic cords at around day 25 [Lucas et al., 1997; Ryhorchuk et al., 1997]. The phallus does not become sexually dimorphic until after day 50-60 [Butler et al., 1999; Leihy et al., 2004, 2011] (fig. 1). This separation of morphological changes has allowed us to investigate the hormonal and molecular control of each of these structures during their differentiation by administration of androgens, oestrogens and inhibitors, and also by castration and gonadal transplantation.

Fig. 1

The timing of virilisation in the tammar wallaby. In the tammar the development of most male (blue boxes) and female (pink boxes) characteristics occurs after birth at a time which is easily accessible (bottom panel timeline). At this time, plasma testosterone remains at similar levels in both sexes [Renfree et al., 1992]. However, gonadal testosterone is higher in males compared to females until day 50 when a sharp decline occurs. Interestingly, the scrotum and mammary primordia are differentiated well before the gonads are capable of any hormone secretion [adapted from Renfree et al., 1987, 2006].

Fig. 1

The timing of virilisation in the tammar wallaby. In the tammar the development of most male (blue boxes) and female (pink boxes) characteristics occurs after birth at a time which is easily accessible (bottom panel timeline). At this time, plasma testosterone remains at similar levels in both sexes [Renfree et al., 1992]. However, gonadal testosterone is higher in males compared to females until day 50 when a sharp decline occurs. Interestingly, the scrotum and mammary primordia are differentiated well before the gonads are capable of any hormone secretion [adapted from Renfree et al., 1987, 2006].

Close modal

Pioneers of mammalian sexual differentiation in the 1940s, RK Burns, CR Moore and CF Morgan in the USA, working on the Virginia opossum, Didelphis virginiana, and A Bolliger in Australia working on the brush-tailed possum, Trichosurus vulpecula, recognised the importance of marsupials for understanding how male and female mammals develop. Marsupials provide access to what is effectively an exteriorised fetus that could be treated with hormones without the confounding effects of the placenta or the mother's physiology [reviewed in Burns, 1961]. This led to attempts to modify the development of the sex organs with the application of (sometimes massive doses and lengthy treatments with) sex hormones [reviewed in Burns, 1961]. They achieved some dramatic effects on the internal and external genitalia, but the significance of the lack of effects on the scrotum and mammary glands were missed. In this review, we will first discuss how sex reversal has been achieved in the gonads, prostate, phallus, and on testicular descent and then revisit the reason for the inability to sex reverse the pouch and scrotum in the last section of our review.

Postnatal Treatments in vivo

Treatment of male tammars from the day of birth to day 25 pp with 17β-oestradiol causes testicular dysgenesis with necrotic germ cells and gross changes in the urogenital sinus. However, the tunica albuginea becomes thickened, very similar to the surface epithelium of the ovary [Shaw et al., 1988]. Testes do not descend into the scrotum, which itself remains completely unaffected, and no mammary glands develop. Treating female tammars with testosterone propionate does not change ovarian morphology, but their Wolffian and Müllerian ducts are hypertrophic [Shaw et al., 1988]. In the North American opossum Didelphis, after ‘painting' the abdomen with oestradiol dipropionate for 30 days after birth, the external and internal genitalia become fully sex-reversed such that an ovotestis ‘indistinguishable from ovaries' form and they have dilated Wolffian ducts [Burns, 1961]. In the grey short-tailed opossum Monodelphis there are less dramatic effects but the dysgenetic testes remain intra-abdominal, development of the prostate gland is inhibited, and the phallus is feminised [Fadem and Tesoriero, 1986]. Interestingly, female Monodelphis young are apparently unaffected by treatment with testosterone.

The extent of the effects of androgens or oestrogens administered for 25 days from the day of birth and examined at day 50 pp is determined by the length of gestation and by when treatments start. Oestrogen-treated neonatal males have abnormal testicular morphology as described above by Shaw et al. [1988], but if the young are born prematurely and oestrogen is administered at day 25 instead of day 26, the testes are fully sex-reversed, with a morphology similar to that of normal ovaries with cortex and medulla, and perhaps most surprising of all, meiotic XY germ cells [Coveney et al., 2001]. The germ cells of treated males become surrounded by follicle-like cells and enter meiosis like the female germ cells of the same chronological age (day 50 pp). Male germ cells do not enter meiosis normally at this stage of development and are in mitotic arrest in the testis. Thus, not only are the somatic cells sex-reversed, but somewhat unusually, the germ cells are as well [Coveney et al., 2001; Renfree et al., 2001]. This result is identical to that reported by Burns [1939a, b, 1942] in the Virginia opossum, in which complete gonadal sex reversal also occurred in ‘early born' (assumed to be premature) litters.

Germ cells have an important function in the development of gonadal morphology. Loss of female germ cells after birth is associated with the formation of Sertoli-like cells that secrete anti-Müllerian hormone (AMH) in seminiferous-like cords in a developing ovary, but loss of male germ cells in both tammars and mice has no effect on testis formation [Burgoyne, 1988; McLaren, 1991a, b; Whitworth et al., 1996; Whitworth, 1998]. Similarly, in mice, male germ cells will enter meiosis if they are cultured with disaggregated lung cells [McLaren and Southee, 1997]. In mice, it is loss of meiotic germ cells that induces cord-like structures [McLaren and Southee, 1997; Adams and McLaren, 2002]. In tammars, the development of seminiferous cords after transplantation of ovaries under the abdominal skin of males or after culture with AMH appears to be a consequence of loss of mitotic germ cells [Whitworth et al., 1996], recently confirmed in the mouse [Rios-Rojas et al., 2016]. Thus it is the germ cells' presence or absence and not the mitotic/meiotic stage that is important. Our hypothesis is that the default condition for somatic tissue is the formation of cords, and the default condition for germ cells is the progression into meiosis [Renfree and Shaw, 2001]. Recent data from mutant We/We mice show that the testis can still form in the absence of germ cells but the cords are irregular and Sertoli cell numbers are reduced [Rios-Rojas et al., 2016].

Prenatal Treatments in vitro

In order to avoid the difficulty of obtaining the rare premature births, we developed a gonadal culture system. Gonads from fetuses at day 25 of the 26.5 day gestation, when cultured for 5 days in media containing oestrogen, are sex-reversed both morphologically and molecularly [Pask et al., 2010]. Oestrogen has a powerful role in controlling sexual differentiation in non-mammalian vertebrates [Nakamura, 2010], but it has always been somewhat disregarded as a factor in early mammalian sexual differentiation although it is recognised that oestrogen still remains a critical factor in maintaining ovarian somatic cell fate [Britt and Findlay, 2003]. Nevertheless, it is clear that oestrogen can sex reverse marsupial gonads in vivo and in vitro. Tammar germ cells and somatic cells express oestrogen receptors α and β (ESR1 and ESR2) from the indifferent gonad stages through testis and ovary differentiation and in adults [Calatayud et al., 2010]. In cultured gonads, oestrogen receptor mRNA is not altered by the addition of oestrogen to the culture medium, but the receptors translocate to the nucleus.

In developing tammar testes, the male-specific genes AMH and sex-determining region Y (SRY)-box 9 (SOX9) are upregulated before the peak of SRY expression, well before testicular cord formation [Pask et al., 2010]. In the ovaries, there is early expression of forkhead-box L2 (FOXL2) and wingless-type MMTV integration site family, member 4 (WNT4), as in the mouse. Most interestingly, tammar gonads cultured in the presence of oestradiol are molecularly and morphologically sex-reversed. Exogenous oestrogen downregulates SRY and AMH (fig. 2), but even though SOX9 expression is unchanged, oestrogen treatment blocks its entry into the nucleus of the testis and SOX9 remains cytoplasmic [Pask et al., 2010]. The XY gonad takes on an ovarian-like appearance after this oestrogen treatment during the 5 days of culture, just as in the in vivo experiments above. This mechanism by which oestrogen can regulate nucleocytoplasmic shuttling of SOX9 may be an ancestral pathway that promotes ovarian development [Pask et al., 2010].

Fig. 2

The impact of hormonal signalling on the gonad and phallus. A Oestrogen treatment in vitro decreases SRY and AMH expression in the gonad, but there are no significant changes in SOX9. BSOX9 expression levels are consistently higher in the testis compared to the ovary. C Androgen decreased SHH and BMP4 expression, but in this normalised (% of control) data, GLI2 was not significantly lower. D Castration at day 25 does not significantly change SHH, GLI2, or BMP4 expression. A and B adapted from Pask et al. [2010], C and D adapted from Chew et al. [2014].

Fig. 2

The impact of hormonal signalling on the gonad and phallus. A Oestrogen treatment in vitro decreases SRY and AMH expression in the gonad, but there are no significant changes in SOX9. BSOX9 expression levels are consistently higher in the testis compared to the ovary. C Androgen decreased SHH and BMP4 expression, but in this normalised (% of control) data, GLI2 was not significantly lower. D Castration at day 25 does not significantly change SHH, GLI2, or BMP4 expression. A and B adapted from Pask et al. [2010], C and D adapted from Chew et al. [2014].

Close modal

In the tammar, the mesonephric ducts, which become the Wolffian ducts, are open to the urogenital sinus by late gestation. Wolffian ducts start to regress in females about day 10 pp, are rudimentary by day 25, and are fully regressed in untreated females before day 34 [Renfree et al., 1996]. Female tammar neonates treated with the potent androgen 5α-androstane-3α,17α-diol (5α-adiol) by mouth from day 10 to day 35 retain their Wolffian ducts, whilst the control females do not [Shaw et al., 2006]. There is evidence suggesting that Wolffian duct development depends on the delivery of androgens directly down the Wolffian duct from the testis [Tong et al., 1996]. However, when testes from neonatal males are grafted beneath the flank skin of female neonates, Wolffian ducts are retained in most testis graft recipient females [Renfree et al., 2009]. The effect of the transplanted gonads was not unilateral, since Wolffian ducts ipsilateral or contralateral to the grafts do not differ.

Male neonates treated from day 10 to day 35 with 4MA (17-β-n, n-diethylcarbamoyl-4-methyl-4-aza-5-alpha-androstan-3-one), an inhibitor of 5α-reductase, have significantly smaller Wolffian ducts than control males (fig. 3), suggesting that adiol is locally converted to the active androgen dihydrotestosterone (DHT). Müllerian ducts in males normally regress by day 13 pp, but unexpectedly, 4MA-treated males retained their Müllerian ducts, whilst control males had highly regressed Müllerian ducts. The ducts are massively hypertrophied, similar to those of males treated with oestradiol [Shaw et al., 1988; Coveney et al., 2002a], suggesting that blockage of 5α-reductase may allow some testosterone to be aromatized to oestrogen. These hypertrophied Müllerian ducts were also observed by Burns [1939a, b] when he treated Didelphis young with oestrogen. Clearly, marsupial Müllerian ducts are highly sensitive to steroid alterations and are either retained or become hypertrophic. Regardless of androgen inhibition or oestrogen stimulation, in contrast to Wolffian ducts, true sex reversal of Müllerian ducts has not been achieved in marsupials.

Fig. 3

Treatment with androstanediol preserves Wolffian ducts (measured by diameter). Adiol-treated females have significant (p < 0.002) increases in Wolffian duct diameter whilst 4MA (5α-reductase inhibitor)-treated males have significantly (p < 0.0005) smaller Wolffian duct diameters. Asterisks indicate level of statistical significance. Redrawn from Shaw et al. [2006].

Fig. 3

Treatment with androstanediol preserves Wolffian ducts (measured by diameter). Adiol-treated females have significant (p < 0.002) increases in Wolffian duct diameter whilst 4MA (5α-reductase inhibitor)-treated males have significantly (p < 0.0005) smaller Wolffian duct diameters. Asterisks indicate level of statistical significance. Redrawn from Shaw et al. [2006].

Close modal

The male urogenital tract is differentiated by androgens in all mammals, and this steroid hormone is crucial for the correct prostate and phallus development. However, in the tammar, we discovered that neither testosterone nor DHT were sexually dimorphic in the plasma [Renfree et al., 1992; Wilson et al., 1999]. A similar situation was evident in another marsupial, the grey short-tailed opossum [Fadem and Harder, 1992a, b; Xie et al., 1998]. Thus, there was clearly another androgen involved which was responsible for virilisation of the prostate and phallus.

We investigated the androgen metabolism pathway in the tammar testis and discovered an alternative pathway of testosterone metabolism via the production of 5α-adiol, which is back-convertible to dihydrotestosterone, and this was responsible for virilisation [Wilson et al., 2002a, b; 2003a, b]. This alternate pathway has now been demonstrated in several other mammals [Mahendroo et al., 2004; Wilson et al., 2005] and can explain the virilisation seen in girls with P450 oxidoreductase deficiency or those with 21-hydroxylase deficiency [Arlt et al., 2004; Homma et al., 2006; Speiser et al., 2010; Reisch et al., 2013].

There is strong evidence that 5α-adiol is critical for the development of the urogenital sinus in the tammar. Exogenously administered 5α-adiol is equally as potent as dihydrotestosterone in virilisation, and both hormones induce prostate bud formation in female pouch young [Leihy et al., 2001]. Whilst testosterone is not sexually dimorphic at any stage of development, 5α-adiol is higher in males compared to females at least between day 20-40 pp, a critical stage in tammar virilisation (fig. 4). Tissue incubations show it is produced in the target tissues and is the key circulating androgen needed for the virilisation of the prostate and phallus [Wilson et al., 2003a].

Fig. 4

Schematic representation of testosterone conversion during virilisation of the urogenital sinus (top panel). Disruption of the testosterone signalling by the inhibitors flutamide (AR inhibitor) and finasteride (5α-reductase inhibitor) results in the absence of a prostate or phallus and a normal female urogenital opening (bottom left panel). The presence of androgen is required for the development of the prostate and the phallus. Adiol or testosterone treatment of females masculinises both prostate and phallus. A supra-physiological prostate is formed with male-like morphology (bottom right panel). Redrawn from Renfree et al. [2002], Wilson et al. [2002b, 2003b].

Fig. 4

Schematic representation of testosterone conversion during virilisation of the urogenital sinus (top panel). Disruption of the testosterone signalling by the inhibitors flutamide (AR inhibitor) and finasteride (5α-reductase inhibitor) results in the absence of a prostate or phallus and a normal female urogenital opening (bottom left panel). The presence of androgen is required for the development of the prostate and the phallus. Adiol or testosterone treatment of females masculinises both prostate and phallus. A supra-physiological prostate is formed with male-like morphology (bottom right panel). Redrawn from Renfree et al. [2002], Wilson et al. [2002b, 2003b].

Close modal

Prostate Differentiation and Sex Reversal

The mammalian prostate is a segmented and compact structure in humans, while the mouse has a multi-lobed structure. The tammar prostate has segmented regions which more closely resemble that of the human prostate, making the marsupial prostate a relevant model for understanding patterning of the human prostate. There is a high degree of conservation in all phases of prostate development and in the expression of key patterning genes, FOXL2, SOX9, and NK3 homeobox1 (NKX3.1) at all phases of tammar prostate development from early patterning to budding and final differentiation [Gamat et al., 2015]. However, the formation of the prostate is under the control of hormones which are easily manipulated in the tammar.

The initiation of prostate development and subsequent growth of the male phenotype in the tammar requires 5α-adiol. Orally administered and injections of the 5α-reductase inhibitor finasteride or the androgen receptor antagonist flutamide inhibit the differentiation of the urogenital sinus into a prostate and phallus [Lucas et al., 1997; Ryhorchuk et al., 1997]. Conversely, after low doses of 5α-adiol or 5α-adiol enanthate administered to female pouch young, a ‘normal' urogenital sinus develops. This process is time dependent, and it is clear that initiation and development of prostate patterning requires a brief exposure to androgens [Lucas et al., 1997]. When high doses of 5α-adiol are given to female pouch young between day 20 and day 150 pp, their urogenital sinuses are sex-reversed and a supra-physiological prostate develops which is larger than that of males of the same age (fig. 4). Thus, prostatic development can be induced in females over a wide period of time. Collectively, these data support the idea that adiol is the main circulating hormone in the tammar for inducing virilisation during development [Leihy et al., 2001; Wilson et al., 2002a, 2003b].

Sex Reversal of the Phallus and Hypospadias

Differentiation of the penis and the male urethra from the genital tubercle is a gradual process of elongation of the penile shaft and progressive closure of the urethral groove so that the urethral closure progresses from proximal to distal along the shaft. Closure of the urethra may fail at any point, and when it does, hypospadias results [Baskin et al., 2001; Cunha et al., 2015]. Virilisation of the phallus is under the influence of hormones in all mammalian species, and disruptions of this process can cause hypospadias which occurs in 1 in 125 live births worldwide. Disruptions in the androgen signalling pathways have similar outcomes in both man and marsupial, since androgen signalling is essential for penile urethral closure.

The phallus is the last of the urogenital system structure to become sexually dimorphic in the tammar wallaby. Male and female phalluses are virtually identical until around day 50 pp when the penis begins to elongate and differentiate (fig. 5). A unique feature of the tammar model is that the phallus development can easily be manipulated with hormones to induce sex reversal and hypospadias during this 50-day period while the young is accessible in the mother's pouch.

Fig. 5

The effect of androgens on phallus growth. A A normal day 150 pp female and male phallus. B Castration of a male pouch young results in growth of a smaller more female-like genitalia. C Adiol-treated females develop a male-like phallus growth but with partial closure of the urethral seam and a shift in the urethral opening. D The normal growth curve of a male (closed squares) and female (open circles). E Castration (thick line closed triangles) retards the length of the phallus and results in a growth curve which resembles that of the normal female phallus. F Adiol treatment (thick lines, closed inverted triangles) of the female pouch young induces a rapid increase in growth and a more male-like growth pattern for the duration of the treatment (in this case from 40 to 80 days pp). ug = Urogenital groove, uo = urogenital opening. Redrawn from Leihy et al. [2001, 2002, 2004].

Fig. 5

The effect of androgens on phallus growth. A A normal day 150 pp female and male phallus. B Castration of a male pouch young results in growth of a smaller more female-like genitalia. C Adiol-treated females develop a male-like phallus growth but with partial closure of the urethral seam and a shift in the urethral opening. D The normal growth curve of a male (closed squares) and female (open circles). E Castration (thick line closed triangles) retards the length of the phallus and results in a growth curve which resembles that of the normal female phallus. F Adiol treatment (thick lines, closed inverted triangles) of the female pouch young induces a rapid increase in growth and a more male-like growth pattern for the duration of the treatment (in this case from 40 to 80 days pp). ug = Urogenital groove, uo = urogenital opening. Redrawn from Leihy et al. [2001, 2002, 2004].

Close modal

In the tammar, castration of male pouch young at day 25 prevents closure of the urethral groove to induce a hypospadias-like morphology, but partial closure of the urethral groove occurs in males castrated at day 40, a far less severe phenotype, suggesting androgen imprinting occurs well before day 40. The timing of androgen signalling is critical, as castration after day 40 results in normal closure of the urethral groove [Leihy et al., 2004]. Transplantation of testes or the addition of androgens to females induces virilisation and the development of a male phenotype phallus. Thus, this period between day 20-40 represents the critical window of androgen sensitivity.

The early prenatal development of the phallus is androgen independent but is followed by an androgen-dependent phase. Interestingly, after day 50 the major growth and development of the phallus occurs at a time when the circulating androgen is at its lowest [Renfree et al., 1992; Leihy et al., 2004]. This drop in circulating androgens correlates with the time that the phallus first becomes sexually dimorphic in length [Shaw et al., 2000], suggesting that androgen withdrawal is necessary for penile growth. This suggestion is supported by the expression of the key patterning genes including sonic hedgehog (SHH), bone morphogenetic protein 4 (BMP4), fibroblast growth factor 8 (FGF8), and GLI family zinc-finger 2 (GLI2) which are involved during the androgen independent phase of development and so are regulated by the levels of circulating androgens [Chew et al., 2014]. SHH and its effector GLI2 influence mesenchymal differentiation adjacent to the urethral epithelium [Haraguchi et al., 2007]. BMPs increase apoptosis and downregulate Fgf8 and Wnt5a in the genital tubercle [Suzuki et al., 2003; Yamada, 2005; Yamada et al., 2003, 2006]. In tammar males, there is an upregulation of these key patterning genes at day 50 when testicular androgens are low and treatment with androgen downregulated their expression [Chew et al., 2014]. Interestingly, when pouch young are castrated at day 25, there are no significant changes in the levels of the key patterning genes at day 50, but there is an increase in gene expression if castration occurred at day 28 pp (fig. 2). Whilst many studies have investigated the influence of key genes involved in the androgen independent phase or androgen-dependent signalling, there is very little information on the impact of androgens on these pathways in phallus growth patterning.

Females can be masculinised, and the degree of masculinisation is dependent on the presence and timing of hormonal signalling. Similarly, hypospadias can be induced in males, and the severity of hypospadias depends on when and how long the treatment is administered. The window of sensitivity in the tammar initially appears to be somewhere between 20-40 days [Leihy et al., 2004] but may in fact be more restricted to within 20-30 days pp [Chew et al., 2014]. The difference in response after castration at day 25 compared to the increased gene expression after castration at day 28 suggests that the window of androgen insensitivity may be very brief indeed, as little as 3-5 days. Thus urethral closure depends on androgen acting in a time window well before closure starts [Leihy et al., 2002, 2004], and there is a dual action of sonic hedgehog protein (SHH) in this process [Chew et al., 2014].

Androgens are essential for testicular descent in eutherian mammals [Hutson et al., 1997]. Testicular descent occurs in 2 phases: transabdominal migration and inguinoscrotal migration. Transabdominal migration is accompanied by the swelling and outgrowth of the gubernaculum and the regression of the cranial suspensory ligament. During inguinoscrotal migration, the gubernaculum thins and elongates, reaching the base of the scrotum [Hutson et al., 1997]. In humans, unlike in any other mammal so far described, the process of testicular descent is completed by closure by fusion of the epithelium of the processus vaginalis to the inguinal canal: failure of this process leads to inguinal hernia and scrotal hydrocoel [Hutson et al., 2000]. Interestingly, this process also occurs in the tammar wallaby [Renfree et al., 1996; Coveney et al., 2001]. There is, however, a small lumen that persists, but the testes cannot pass back into the abdomen, prevented by a membranous flap or valve that covers the internal inguinal ring [Coveney et al., 2001]. As in humans, if inguinal closure fails in the tammar, it predisposes the affected animal to inguinal hernia, and as in humans, macropodid marsupials have an upright posture. During their hopping mode of locomotion, this would cause considerable pressure on the inguinal canal and susceptibility to inguinal hernias, suggesting the reason for closure in this marsupial group as in man [Coveney et al., 2001] (fig. 6).

Fig. 6

Effects of oestrogen on testicular descent. A Schematic diagram of testicular descent in the tammar wallaby. The testis migrates from the abdominal cavity via the inguinal canal to the scrotum as the animal develops. K = Kidney, T = testis. B The inguinal canal is fused in the tammar and humans but open in the brush tail possum and in all other species. C Oestrogen treatment shifts the testicular position more dorsal relative to the lumbar position. Redrawn from Shaw et al. [1988] and from Renfree et al. [2001].

Fig. 6

Effects of oestrogen on testicular descent. A Schematic diagram of testicular descent in the tammar wallaby. The testis migrates from the abdominal cavity via the inguinal canal to the scrotum as the animal develops. K = Kidney, T = testis. B The inguinal canal is fused in the tammar and humans but open in the brush tail possum and in all other species. C Oestrogen treatment shifts the testicular position more dorsal relative to the lumbar position. Redrawn from Shaw et al. [1988] and from Renfree et al. [2001].

Close modal

We have shown that inguinal closure is dependent on androgens [Coveney et al., 2002b]. All male pouch young treated with the anti-androgen receptor flutamide develop inguinal hernias [Lucas et al., 1997; Coveney et al., 2002b]. Calcitonin-gene related peptide (CGRP) secreted into neurones of the dorsal root ganglia and the genitofemoral nerve (that innervates the gubernaculum) is responsible for the process of inguinal closure in humans [Hutson et al., 1997, 2000]. Female tammar young have fewer CGRP-positive neurones than males. Similarly, there are significantly fewer cell bodies expressing CGRP in flutamide-treated tammar males and their inguinal canals do not close although their testes descend into the base of the scrotum, showing that androgen is not essential for the initial phases of descent but is needed for inguinal closure [Coveney et al., 2002b].

Perhaps most interesting is the effect of oestrogen treatment on testicular descent. Treatment of male pouch young with oestradiol benzoate from day 10 to day 25 pp and from day 0 to day 25 results in dysgenetic testes, long thin gubernaculae, open inguinal canals, and significant inguinal hernias. Testicular descent fails in these animals, and the testes remain in an abdominal position similar to the location of the ovaries in female pouch young (fig. 6). Since the differentiation of the testes is not normal, these effects appear to be caused by both abnormal or deficient AMH and androgen secretion and possibly also by direct effects of oestrogen on other target tissues including the gubernaculum and inguinal canal [Coveney et al., 2002a].

The extended period of development ex utero in marsupials has provided an opportunity to understand the origin of sexual behaviour. The tammar wallaby has a multi-male mating system, and although reproduction in females is highly synchronised, controlled by lactational stimuli and photoperiodic ones that depend on the time of the year, the males are capable of fertilising females at any time of the year [Inns, 1982; Hynes et al., 2005]. Males do have a seasonal peak in testosterone [Inns, 1982] when the females enter their postpartum oestrus in late January [Rudd, 1994], but there is also another minor peak of plasma testosterone in October when the female young leave their mother's pouches and enter puberty [Williams et al., 1998]. Male young do not become sexually mature until 18-24 months after birth [Williamson et al., 1990].

Mating behaviour is under the influence of androgens, as in all male mammals. Castration at 25 days after birth (before most of the reproductive system is sexually dimorphic) abolishes male-type copulatory behaviour as adults, while adult females treated with a testosterone implant show male type sexual behaviour, indistinguishable from that of intact males [Rudd et al., 1996].

The oestradiol\LH positive feedback response is also sexually dimorphic in marsupials as in eutherian mammals. Females have a surge in plasma LH after injection of oestradiol that persists in ovariectomised females. However, it does not occur in adult males or in females with a testosterone implant [Rudd et al., 1999]. Interestingly, the male pouch young castrated at day 25 have a plasma LH surge after an oestradiol challenge. Thus, castrating males at day 25 makes the male brain respond to gonadotrophins like the brain of females. These experiments confirm that the sensitivity to steroid hormones is not restricted to the gonads and reproductive tract but includes brain sex, which is at least partially sex reversible.

This review began with the experiments of Burns and Bolliger attempting to sex reverse neonatal marsupials with exogenous hormones. Whilst they achieved dramatic sex reversal of the internal and external genitalia and showed that oestrogen blocked testicular differentiation, their treatments failed to affect the development of the scrotum, mammary glands, or pouch. Unfortunately, they did not recognise the significance of this discovery. However, we now know that the scrotal and mammary primordia, the processus vaginalis, and the gubernaculum all develop completely independently of hormones [O et al., 1988; Renfree and Short, 1988; Shaw et al., 1988]. We showed that on the day of birth genetic male tammars all have scrotal bulges but no mammary primordia, whilst genetic females all have mammary primordia but no scrotal bulges. We concluded that there is a primary genetic control of the differentiation of these structures [O et al., 1988]. Treatment of neonatal young with androgens or oestrogen has no effect on the scrotum and mammary glands, so these structures develop independently of hormonal stimulation [Shaw et al., 1988]. Indeed, hormone-independent sexual dimorphisms are now known to occur in eutherian mammals as well as in birds [Renfree et al., 2014]. Examining the relatively few known intersexes, we now know that these are under the control of the X chromosome, so that 1 X codes for a scrotum, and 2 XXs for mammary glands and a pouch [Renfree and Short, 1988]. Thus, a marsupial Klinefelters-type syndrome male with an XXY karyotype has a pouch and penis (with normal abdominal testes), whilst a Turners-type marsupial female with an XO karyotype has an empty scrotum but female internal genitalia. However, despite the availability of several marsupial genomes [Mikkelsen et al., 2007; Renfree et al., 2011; Murchison et al., 2012] and with several more currently underway, we do not yet know the identity of this X-linked gene. Its discovery is awaited with interest.

Marsupials are unique models for understanding the differentiation of the internal and external genitalia because of the temporal and sequential development of each of the structures and for the relative ease with which they can become sex-reversed by hormonal manipulation. We have successfully sex-reversed each of the key structures: the testis (but less so the ovary), the prostate and urogenital sinus, the penis and clitoris, and even the brain by treatment with hormones over specific or sequential time windows. The tammar wallaby has proven to be a tractable model for studies of sex reversal. With the availability of new marsupial-specific genomic resources, the characterisation of the underlying molecular controls will provide novel perspectives for further understanding the evolution of sexual differentiation of the internal and external genitalia.

We thank our many colleagues, collaborators, and graduate students, especially Geoff Shaw, Jean D Wilson, Roger Short and Andrew Pask, who have contributed to the studies summarised here, and to the Australian Research Council and the National Health and Medical Research Council for financial support.

The authors have no conflicts of interest to declare.

1.
Adams IR, McLaren A: Sexually dimorphic development of mouse primordial germ cells: switching from oogenesis to spermatogenesis. Development 129:1155-1164 (2002).
2.
Arlt W, Walker EA, Draper N, Ivison HE, Ride JP, et al: Congenital adrenal hyperplasia caused by mutant P450 oxidoreductase and human androgen synthesis: analytical study. Lancet 363:2128-2135 (2004).
3.
Baskin LS, Erol A, Jegatheesan P, Li Y, Liu W, Cunha GR: Urethral seam formation and hypospadias. Cell Tissue Res 305:379-387 (2001).
4.
Britt KL, Findlay JK: Regulation of the phenotype of ovarian somatic cells by estrogen. Mol Cell Endocrinol 202:11-17 (2003).
5.
Burgoyne PS: Role of mammalian Y chromosome in sex determination. Philos Trans R Soc Lond B Biol Sci 322:63-72 (1988).
6.
Burns RK: The differentiation of sex in the opossum (Didelphis virginiana) and its modification by the male hormone testosterone propionate. J Morphol 65:79-119 (1939a).
7.
Burns RK: Sex differentiation during the early pouch stages of the opossum (Didelphys virginiana) and a comparison of the anatomical changes induced by male and female sex hormones. J Morphol 65:497-547 (1939b).
8.
Burns RK: Hormones and the experimental modification of sex in the opossum. Biological Symposia 10:125-146 (1942).
9.
Burns RK: Role of hormones in the differentiation of sex, in Young WC (ed): Sex and Internal Secretions, Vol 1, pp 76-158 (Williams and Wilkins, Baltimore 1961).
10.
Butler CM, Shaw G, Renfree MB: Development of the penis and clitoris in the tammar wallaby, Macropus eugenii. Anat Embryol 199:451-457 (1999).
11.
Calatayud NE, Pask AJ, Shaw G, Richings NM, Osborn S, Renfree MB: Ontogeny of the oestrogen receptors ESR1 and ESR2 during gonadal development in the tammar wallaby, Macropus eugenii. Reproduction 139:599-611 (2010).
12.
Chew KY, Pask AJ, Hickford D, Shaw G, Renfree MB: A dual role for SHH during phallus development in a marsupial. Sex Dev 8:166-177 (2014).
13.
Coveney D, Shaw G, Renfree MB: Estrogen-induced gonadal sex reversal in the tammar wallaby. Biol Reprod 65:613-621 (2001).
14.
Coveney D, Shaw G, Renfree MB: Effects of oestrogen treatment on testicular descent, inguinal closure and prostatic development in a male marsupial, Macropus eugenii. Reproduction 124:73-83 (2002a).
15.
Coveney D, Shaw G, Hutson JM, Renfree MB: The development of the gubernaculum and inguinal closure in the marsupial Macropus eugenii. J Anat 201:239-256 (2002b).
16.
Cunha GR, Sinclair A, Risbridger G, Hutson J, Baskin LS: Current understanding of hypospadias: relevance of animal models. Nat Rev Urol 12:271-280 (2015).
17.
Fadem BH, Harder JD: Estrogen in peripheral plasma during postnatal development in gray short-tailed opossums. Physiol Behav 52:613-616 (1992a).
18.
Fadem BH, Harder JD: Evidence for high levels of androgen in peripheral plasma during postnatal development in a marsupial: the gray short-tailed opossum (Monodelphis domestica). Biol Reprod 46:105-108 (1992b).
19.
Fadem BH, Tesoriero JV: Inhibition of testicular development and feminization of the male genitalia by neonatal estrogen treatment in a marsupial. Biol Reprod 34:771-776 (1986).
20.
Gamat M, Chew KY, Shaw G, Renfree MB: FOXA1 and SOX9 expression in the developing urogenital sinus of the tammar wallaby (Macropus eugenii). Sex Dev 9:216-228 (2015).
21.
Haraguchi R, Motoyama J, Sasaki H, Satoh Y, Miyagawa S, et al: Molecular analysis of coordinated bladder and urogenital organ formation by Hedgehog signaling. Development 134:525-533 (2007).
22.
Homma K, Hasegawa T, Nagai T, Adachi M, Horikawa R, et al: Urine steroid hormone profile analysis in cytochrome P450 oxidoreductase deficiency: implication for the backdoor pathway to dihydrotestosterone. J Clin Endocrinol Metab 91:2643-2649 (2006).
23.
Hutson JM, Hasthorpe S, Heyns CF: Anatomical and functional aspects of testicular descent and cryptorchidism. Endocrine Rev 18:259-280 (1997).
24.
Hutson JM, Albano FR, Paxton G, Sugita Y, Connor R, et al: In vitro fusion of human inguinal hernia with associated epithelial transformation. Cells Tissues Organs 166:249-258 (2000).
25.
Hynes EF, Rudd CD, Temple-Smith PD, Sofronidis G, Paris D, Shaw G, Renfree MB: Mating sequence, dominance and paternity success in captive male tammar wallabies. Reproduction 130:123-130 (2005).
26.
Inns RW: Seasonal changes in the accessory reproductive system and plasma testosterone levels of the male tammar wallaby, Macropus eugenii, in the wild. J Reprod Fertil 66:675-680 (1982).
27.
Leihy MW, Shaw G, Wilson JD, Renfree MB: Virilization of the urogenital sinus of the tammar wallaby is not unique to 5α-androstane-3α,17β-diol. Mol Cell Endocrinol 181:111-115 (2001).
28.
Leihy MW, Shaw G, Renfree MB, Wilson JD: Administration of 5alpha-androstane-3alpha, 17beta-diol to female tammar wallaby pouch young causes development of a mature prostate and male urethra. Endocrinology 143:2643-2651 (2002).
29.
Leihy MW, Shaw G, Wilson JD, Renfree MB: Penile development is initiated in the tammar wallaby pouch young during the period when 5α-androstane-3α,17β-diol is secreted by the testes. Endocrinology 145:3346-3352 (2004).
30.
Leihy MW, Shaw G, Wilson JD, Renfree MB: Development of the penile urethra in the tammar wallaby. Sex Dev 5:241-249 (2011).
31.
Lucas JC, Renfree MB, Shaw G, Butler CM: The influence of the anti-androgen flutamide on early sexual differentiation of the marsupial male. J Reprod Fertil 109:205-212 (1997).
32.
Mahendroo M, Wilson JD, Richardson JA, Auchus RJ: Steroid 5α-reductase 1 promotes 5a-androstane-3α,17β-diol synthesis in immature mouse testes by two pathways. Mol Cell Endocrinol 222:113-120 (2004).
33.
McLaren A: Development of the mammalian gonad: the fate of the supporting cell lineage. BioEssays 13:151-156 (1991a).
34.
McLaren A: Sex determination. The making of male mice. Nature 351:96 (1991b).
35.
McLaren A, Southee D: Entry of mouse embryonic germ cells into meiosis. Dev Biol 187:107-113 (1997).
36.
Mikkelsen TS, Wakefield MJ, Aken B, Amemiya CT, Chang JL, et al: Genome of the marsupial Monodelphis domestica reveals innovation in non-coding sequences. Nature 447:167-177 (2007).
37.
Murchison EP, Schulz-Trieglaff OB, Ning Z, Alexandrov LB, Bauer MJ, et al: Genome sequencing and analysis of the Tasmanian devil and its transmissible cancer. Cell 148:780-791 (2012).
38.
Nakamura M: The mechanism of sex determination in vertebrates-are sex steroids the key-factor? J Exp Zool A Ecol Genet Physiol 313:381-398 (2010).
39.
O WS, Short RV, Renfree MB, Shaw G: Primary genetic control of somatic sexual differentiation in a mammal. Nature 331:716-717 (1988).
40.
Pask AJ, Calatayud NE, Shaw G, Wood WM, Renfree MB: Oestrogen blocks the nuclear entry of SOX9 in the developing gonad of a marsupial mammal. BMC Biol 8:113 (2010).
41.
Reisch N, Hogler W, Parajes S, Rose IT, Dhir V, et al: A diagnosis not to be missed: nonclassic steroid 11β-hydroxylase deficiency presenting with premature adrenarche and hirsutism. J Clin Endocrinol Metab 98:1620-1625 (2013).
42.
Renfree MB, Shaw G: Germ cells, gonads and sex reversal in marsupials. Int J Dev Biol 45:557-567 (2001).
43.
Renfree MB, Short RV: Sex determination in marsupials: evidence for a marsupial-eutherian dichotomy. Philos Trans R Soc Lond B Biol Sci 322:41-53 (1988).
44.
Renfree MB, Shaw G, Short RV: Sexual differentiation in marsupials, in Haseltine FP, McClure ME, Goldberg EH (eds): Genetic Markers of Sex Differentiation, pp 27-41 (Plenum Press, New York 1987).
45.
Renfree MB, Wilson JD, Short RV, Shaw G, George FW: Steroid hormone content of the gonads of the tammar wallaby during sexual differentiation. Biol Reprod 47:644-647 (1992).
46.
Renfree MB, Harry JL, Shaw G: The marsupial male: a role model for sexual development. Philos Trans R Soc Lond B Biol Sci 350:243-251 (1995).
47.
Renfree MB, O WS, Short RV, Shaw G: Sexual differentiation of the urogenital system of the fetal and neonatal tammar wallaby, Macropus eugenii. Anat Embryol 194:111-134 (1996).
48.
Renfree MB, Coveney D, Shaw G: The influence of estrogen on the developing male marsupial. Reprod Fertil Dev 13:231-240 (2001).
49.
Renfree MB, Wilson JD, Shaw G: The hormonal control of sexual development. Novartis Found Symposium 244:136-152; discussion 152-136, 203-136, 253-137 (2002).
50.
Renfree MB, Pask AJ, Shaw G: Sexual development of a model marsupial male. Aust J Zool 54:151-158 (2006).
51.
Renfree MB, Fenelon J, Wijiyanti G, Wilson JD, Shaw G: Wolffian duct differentiation by physiological concentrations of androgen delivered systemically. Dev Biol 334:429-436 (2009).
52.
Renfree MB, Papenfuss AT, Deakin JE, Lindsay J, Heider T, et al: Genome sequence of an Australian kangaroo, Macropus eugenii, provides insight into the evolution of mammalian reproduction and development. Genome Biology 12:R81 (2011).
53.
Renfree MB, Chew KY, Shaw G: Hormone-independent pathways of sexual differentiation. Sex Dev 8:327-336 (2014).
54.
Rios-Rojas C, Spiller C, Bowles J, Koopman P: Germ cells influence cord formation and leydig cell gene expression during mouse testis development. Dev Dyn 245:433-444 (2016).
55.
Rudd CD: Sexual behaviour of male and female tammar wallabies (Macropus eugenii) at post-partum oestrus. J Zool 232:151-162 (1994).
56.
Rudd CD, Short RV, Shaw G, Renfree MB: Testosterone control of male-type sexual behavior in the tammar wallaby (Macropus eugenii). Horm Behav 30:446-454 (1996).
57.
Rudd CD, Short RV, McFarlane JR, Renfree MB: Sexual differentiation of oestradiol-LH positive feedback in a marsupial. J Reprod Fertil 115:269-274 (1999).
58.
Ryhorchuk AR, Shaw G, Butler CM, Renfree MB: Effects of a 5 alpha-reductase inhibitor, finasteride, on the developing prostate and testis of a marsupial. J Androl 18:123-130 (1997).
59.
Shaw G, Renfree MB, Short RV, O WS: Experimental manipulation of sexual differentiation in wallaby pouch young treated with exogenous steroids. Development 104:689-701 (1988).
60.
Shaw G, Renfree MB, Leihy MW, Shackleton CH, Roitman E, Wilson JD: Prostate formation in a marsupial is mediated by the testicular androgen 5α-androstane-3α,17β-diol. Proc Natl Acad Sci USA 97:12256-12259 (2000).
61.
Shaw G, Fenelon J, Sichlau M, Auchus RJ, Wilson JD, Renfree MB: Role of the alternate pathway of dihydrotestosterone formation in virilization of the Wolffian ducts of the tammar wallaby, Macropus eugenii. Endocrinology 147:2368-2373 (2006).
62.
Speiser PW, Azziz R, Baskin LS, Ghizzoni L, Hensle TW, et al: Congenital adrenal hyperplasia due to steroid 21-hydroxylase deficiency: an Endocrine Society clinical practice guideline. J Clin Endocrinol Metab 95:4133-4160 (2010).
63.
Suzuki K, Bachiller D, Chen YP, Kamikawa M, Ogi H, et al: Regulation of outgrowth and apoptosis for the terminal appendage: external genitalia development by concerted actions of BMP signaling. Development 130:6209-6220 (2003).
64.
Tong SY, Hutson JM, Watts LM: Does testosterone diffuse down the Wolffian duct during sexual differentiation? J Urol 155:2057-2059 (1996).
65.
Whitworth DJ: XX germ cells: the difference between an ovary and a testis. Trends Endocrinol Metab 9:2-6 (1998).
66.
Whitworth DJ, Shaw G, Renfree MB: Gonadal sex reversal of the developing marsupial ovary in vivo and in vitro. Development 122:4057-4063 (1996).
67.
Williams SC, Fletcher TP, Renfree MB: Puberty in the female tammar wallaby. Biol Reprod 58:1117-1122 (1998).
68.
Williamson P, Fletcher TP, Renfree MB: Testicular development and maturation of the hypothalamic-pituitary-testicular axis in the male tammar, Macropus eugenii. J Reprod Fertil 88:549-557 (1990).
69.
Wilson JD, George FW, Shaw G, Renfree MB: Virilization of the male pouch young of the tammar wallaby does not appear to be mediated by plasma testosterone or dihydrotestosterone. Biol Reprod 61:471-475 (1999).
70.
Wilson JD, Leihy MW, Shaw G, Renfree MB: Androgen physiology: unsolved problems at the millennium. Mol Cell Endocrinol 198:1-5 (2002a).
71.
Wilson JD, Shaw G, Leihy ML, Renfree MB: The marsupial model for male phenotypic development. Trends Endocrinol Metab 13:78-83 (2002b).
72.
Wilson JD, Auchus RJ, Leihy MW, Guryev OL, Estabrook RW, et al: 5α-androstane-3α,17β-diol is formed in tammar wallaby pouch young testes by a pathway involving 5α- pregnane-3α,17α-diol-20-one as a key intermediate. Endocrinology 144:575-580 (2003a).
73.
Wilson JD, Leihy MW, Shaw G, Renfree MB: Unsolved problems in male physiology: studies in a marsupial. Mol Cell Endocrinol 211:33-36 (2003b).
74.
Wilson JD, Shaw G, Renfree MB, Auchus RJ, Leihy MW, Eckery DC: Ontogeny and pathway of formation of 5α-androstane-3α,17β-diol in the testes of the immature brushtail possum Trichosurus vulpecula. Reprod Fertil Dev 17:603-609 (2005).
75.
Xie Q, Mackay S, Ullmann SL, Gilmore DP, Payne AP, Gray C: Postnatal development of Leydig cells in the opossum (Monodelphis domestica): an immunocytochemical and endocrinological study. Biol Reprod 58:664-669 (1998).
76.
Yamada G: Reproductive/urogenital organ development and molecular genetic cascades: glamorous developmental processes of bodies. J Biochem 137:665-669 (2005).
77.
Yamada G, Satoh Y, Baskin LS, Cunha GR: Cellular and molecular mechanisms of development of the external genitalia. Differentiation 71:445-460 (2003).
78.
Yamada G, Suzuki K, Haraguchi R, Miyagawa S, Satoh Y, et al: Molecular genetic cascades for external genitalia formation: an emerging organogenesis program. Dev Dyn 235:1738-1752 (2006).
Copyright / Drug Dosage / Disclaimer
Copyright: All rights reserved. No part of this publication may be translated into other languages, reproduced or utilized in any form or by any means, electronic or mechanical, including photocopying, recording, microcopying, or by any information storage and retrieval system, without permission in writing from the publisher.
Drug Dosage: The authors and the publisher have exerted every effort to ensure that drug selection and dosage set forth in this text are in accord with current recommendations and practice at the time of publication. However, in view of ongoing research, changes in government regulations, and the constant flow of information relating to drug therapy and drug reactions, the reader is urged to check the package insert for each drug for any changes in indications and dosage and for added warnings and precautions. This is particularly important when the recommended agent is a new and/or infrequently employed drug.
Disclaimer: The statements, opinions and data contained in this publication are solely those of the individual authors and contributors and not of the publishers and the editor(s). The appearance of advertisements or/and product references in the publication is not a warranty, endorsement, or approval of the products or services advertised or of their effectiveness, quality or safety. The publisher and the editor(s) disclaim responsibility for any injury to persons or property resulting from any ideas, methods, instructions or products referred to in the content or advertisements.