Instability of Sex-Determining Systems in Frogs

*Michihiko Ito*

## **Abstract**

All of the anuran amphibians examined so far have genetic sex-determining systems, which include female heterogametic ZZ/ZW and male heterogametic XX/ XY types. For example, the Japanese wrinkled frog *Glandirana rugosa* has both types. Most of frog species including the African clawed frog *Xenopus laevis* possess homomorphic sex chromosomes, while most mammalian and avian species have heteromorphic sex chromosomes. Thus, there should be a variety of sexdetermining genes and sex chromosomes in frogs, although only *X. laevis* W-linked gene *dm-W* has been reported as a sex-determining gene. Interestingly, estrogen or androgen can induce sex reversal in many frog species, suggesting a vital role of sex steroid hormones on sex identity. In other words, frogs in the same order are good examples for the understanding of diversity of sex-determining systems. In this chapter, I summarize the diversity of frog sex-determining systems and discuss why sex-determining genes and systems have been unstable in frogs.

**Keywords:** sex determination, sex chromosome, sex-determining gene, sex steroid, default sex, ectothermy

## **1. Introduction**

Sexual reproduction is the most common life cycle in animals and plants. Meiotic recombination mediated through sexual reproduction is believed to allow genetic variation for survival of some populations against environmental changes. Thus, sex systems are very important for life evolution and biodiversity. In vertebrates, female and male sexes could be mainly defined by the property of gonads, ovaries producing eggs and testes producing sperm, respectively. Importantly, undifferentiated gonads in most vertebrate species have potential to differentiate into ovaries and testes. Then sex determination could be defined as the decision of bipotential gonads to develop as either ovaries or testes in vertebrates.

There are a variety of sex-determining systems in organisms. In vertebrates, they could be classified roughly into two types: genetic and environmental types. Endothermic vertebrates exclusively have the former system, which includes female (ZW) and male (XY) heterogametic sex chromosomes. Most mammalian and avian species have the XX/XY and ZZ/ZW systems, respectively, while there are both ZZ/ ZW- and XX/XY-type systems in teleost fish, amphibians, and reptiles [1]. In addition, ectothermic vertebrates including reptiles and fish have not only the genetic sex-determining systems but also environmental sex-determining systems, such as temperature- and social-dependent types. Remarkably, all amphibian species possess the genetic systems, although they have ectothermic traits like reptiles and fish [1].

In the chapter, I introduce sex-determining systems, sex chromosomes, and sexdetermining genes in amphibian frogs and discuss the relationships among them.

## **2. Sex-determining systems and sex chromosomes in frogs**

As described in the above section, all anuran amphibians examined so far have the genetic sex-determining systems including the ZZ/ZW and XX/XY types (**Table 1**). For examples, the (African bullfrog) *Pyxicephalus adspersus*, African clawed frog *Xenopus laevis*, and the cane toad *Bufo marinus* have the ZZ/ZW type [2–5], while the African reed frog *Hyperolius viridiflavus* and the marsupial frog *Gastrotheca riobambae* adopt the XX/XY-type systems [6, 7]. Remarkably, the Japanese frog *Glandirana (Rana) rugosa* have five populations in Japan; their sexdetermining systems include two ZZ/ZW and three XX/XY types [8].


#### **Table 1.**

*Sex-determining systems, sex chromosomes, and sex-determining genes in frogs.*

#### **Figure 1.**

*A model for emergence and evolution of sex-determining genes and homomorphic and heteromorphic sex chromosomes in vertebrates. The model includes a proposal of "GENE-eat-GENE" model for changes of sex-determining genes in homomorphic sex chromosomes.*

**15**

*Instability of Sex-Determining Systems in Frogs DOI: http://dx.doi.org/10.5772/intechopen.89050*

**clawed frog**

most vertebrates as transcription factors [21].

evolved after allotetraploidization [24].

The XX/XY and ZZ/ZW systems in most mammals and all birds examined have been maintained for more than a 100 million years, which is greatly connected with the monophyletic and heteromorphic sex chromosomes among most species of therian mammals or avians: the monophyly of the Z or Y sex chromosomes is closely related to the maintainability of the sex-determining gene *Dmrt1* on the Z chromosome or *Sry* on the Y chromosome, respectively [1]. In contrast, more than 90% species of frogs including *X. laevis* have homomorphic sex chromosomes [9–11]. In fact, sex chromosome homomorphism is well conserved among many vertebrate species except for mammals and birds. In 2012, we proposed a hypothesis for the coevolution of sex chromosomes and sex-determining genes, in which homomorphic sex chromosomes easily allow changes of sex-determining genes, resulting in changes of sex chromosomes. On the contrary, highly differentiated heteromorphic sex chromosomes including mammalian XY and avian ZW chromosomes are easily maintained, resulting in a stable fixation of a particular sex-determining gene, because each sex chromosome has gained important functions except for sex determination ([12, 13]; **Figure 1**). This context could lead to the conclusion that there are a variety of sex-determining genes in frogs [1], although few amphibian sex-determining genes

except for *dm-W* we discovered in *X. laevis* [14] have been identified yet.

**3. Discovery of a female sex-determining gene** *dm-W* **in the African** 

In 1990, human *SRY* was discovered as a sex-determining gene, which was the first report among vertebrate species [15], followed by mouse *Sry* [16]. Now *Sry* is believed to be a sex-determining gene in many species of therian mammals. After about 10 years, the second vertebrate sex-determining gene named *dmy* (also known as *dmrt1bY*) was reported in the teleost fish medaka *Oryzias latipes* [17, 18]. Both the two genes function as Y-linked male-determining genes in the XX/ XY-type sex-determining systems. In 2008, we discovered a W-linked sex (female) determining gene *dm-W* from the frog *X. laevis* having a ZZ/ZW type [5]; *dm-W* was the first report as the sex-determining gene among amphibian species or ZZ/ ZW-type vertebrate species. Among sex-determining genes reported so far, the *dm-W* gene is unique in that the gene is female genome-specific (W-linked) and causes ovary formation [5, 19]. Both the *dmy* and *dm-W* genes emerged from the duplication of *dmrt1* independently during species diversity in genus *Oryzias* and *Xenopus*, respectively [12]. Next, Smith et al. (2010) reported that the Z-linked *dmrt1* gene is necessary for male sex determination in the chicken (*Gallus domesticus*) [20]. Here I should describe what protein is doublesex Mab-3-related transcription factor 1 (DMRT1). The protein including a DNA-binding domain, called "DM domain," functions in gonadal somatic cell masculinization and germ cell development in

*X. laevis* is an allotetraploid species, whose ancestor might emerge by hybridiza-

The DM domain of DM-W has about 90% amino acid sequence identity with those of DMRT1.L and DMRT1.S. However, the DM-W C-terminal region shares almost no similarity with those of DMRT1s. The last fourth exon of *dm-W* coding the C-terminal region emerged as a new exon [5]. We reported that DM-W and DMRT1 could cause primary ovarian and testicular formation in developing ZW

tion between two closely related *Xenopus* diploid species [22]. Therefore, there are two homoeologous L and S subgenome-derived genes in most of the genes in *X. laevis*. Partial duplication of S subgenome-derived *dmrt1* (*dmrt1.S*) leads to the emergence of *dm-W* [5, 23, 24]. In addition, we recently reported that *dm-W*

*Instability of Sex-Determining Systems in Frogs DOI: http://dx.doi.org/10.5772/intechopen.89050*

*Gene Expression and Phenotypic Traits*

**Species Sex-determining** 

*Xenopus laevis* (African clawed frog)

*Pyxicephalus adspersus* African bullfrog)

*Hyperolius viridiflavus* (African reed frog)

*Glandirana rugosa* (Japanese wrinkled

frog)

**Table 1.**

**type**

*Gastrotheca riobambae* XX/XY Heteromorphic

*Sex-determining systems, sex chromosomes, and sex-determining genes in frogs.*

In the chapter, I introduce sex-determining systems, sex chromosomes, and sexdetermining genes in amphibian frogs and discuss the relationships among them.

As described in the above section, all anuran amphibians examined so far have

**Morphology of sex chromosomes**

ZZ/ZW Homomorphic W-specific *dm-W*

**Sex-determining** 

**gene**

the genetic sex-determining systems including the ZZ/ZW and XX/XY types (**Table 1**). For examples, the (African bullfrog) *Pyxicephalus adspersus*, African clawed frog *Xenopus laevis*, and the cane toad *Bufo marinus* have the ZZ/ZW type [2–5], while the African reed frog *Hyperolius viridiflavus* and the marsupial frog *Gastrotheca riobambae* adopt the XX/XY-type systems [6, 7]. Remarkably, the Japanese frog *Glandirana (Rana) rugosa* have five populations in Japan; their sex-

ZZ/ZW Heteromorphic

XX/XY Homomorphic

ZZ/ZW or XX/XY Heteromorphic/homomorphic

**2. Sex-determining systems and sex chromosomes in frogs**

determining systems include two ZZ/ZW and three XX/XY types [8].

**14**

**Figure 1.**

*A model for emergence and evolution of sex-determining genes and homomorphic and heteromorphic sex chromosomes in vertebrates. The model includes a proposal of "GENE-eat-GENE" model for changes of* 

*sex-determining genes in homomorphic sex chromosomes.*

The XX/XY and ZZ/ZW systems in most mammals and all birds examined have been maintained for more than a 100 million years, which is greatly connected with the monophyletic and heteromorphic sex chromosomes among most species of therian mammals or avians: the monophyly of the Z or Y sex chromosomes is closely related to the maintainability of the sex-determining gene *Dmrt1* on the Z chromosome or *Sry* on the Y chromosome, respectively [1]. In contrast, more than 90% species of frogs including *X. laevis* have homomorphic sex chromosomes [9–11]. In fact, sex chromosome homomorphism is well conserved among many vertebrate species except for mammals and birds. In 2012, we proposed a hypothesis for the coevolution of sex chromosomes and sex-determining genes, in which homomorphic sex chromosomes easily allow changes of sex-determining genes, resulting in changes of sex chromosomes. On the contrary, highly differentiated heteromorphic sex chromosomes including mammalian XY and avian ZW chromosomes are easily maintained, resulting in a stable fixation of a particular sex-determining gene, because each sex chromosome has gained important functions except for sex determination ([12, 13]; **Figure 1**). This context could lead to the conclusion that there are a variety of sex-determining genes in frogs [1], although few amphibian sex-determining genes except for *dm-W* we discovered in *X. laevis* [14] have been identified yet.

## **3. Discovery of a female sex-determining gene** *dm-W* **in the African clawed frog**

In 1990, human *SRY* was discovered as a sex-determining gene, which was the first report among vertebrate species [15], followed by mouse *Sry* [16]. Now *Sry* is believed to be a sex-determining gene in many species of therian mammals. After about 10 years, the second vertebrate sex-determining gene named *dmy* (also known as *dmrt1bY*) was reported in the teleost fish medaka *Oryzias latipes* [17, 18]. Both the two genes function as Y-linked male-determining genes in the XX/ XY-type sex-determining systems. In 2008, we discovered a W-linked sex (female) determining gene *dm-W* from the frog *X. laevis* having a ZZ/ZW type [5]; *dm-W* was the first report as the sex-determining gene among amphibian species or ZZ/ ZW-type vertebrate species. Among sex-determining genes reported so far, the *dm-W* gene is unique in that the gene is female genome-specific (W-linked) and causes ovary formation [5, 19]. Both the *dmy* and *dm-W* genes emerged from the duplication of *dmrt1* independently during species diversity in genus *Oryzias* and *Xenopus*, respectively [12]. Next, Smith et al. (2010) reported that the Z-linked *dmrt1* gene is necessary for male sex determination in the chicken (*Gallus domesticus*) [20]. Here I should describe what protein is doublesex Mab-3-related transcription factor 1 (DMRT1). The protein including a DNA-binding domain, called "DM domain," functions in gonadal somatic cell masculinization and germ cell development in most vertebrates as transcription factors [21].

*X. laevis* is an allotetraploid species, whose ancestor might emerge by hybridization between two closely related *Xenopus* diploid species [22]. Therefore, there are two homoeologous L and S subgenome-derived genes in most of the genes in *X. laevis*. Partial duplication of S subgenome-derived *dmrt1* (*dmrt1.S*) leads to the emergence of *dm-W* [5, 23, 24]. In addition, we recently reported that *dm-W* evolved after allotetraploidization [24].

The DM domain of DM-W has about 90% amino acid sequence identity with those of DMRT1.L and DMRT1.S. However, the DM-W C-terminal region shares almost no similarity with those of DMRT1s. The last fourth exon of *dm-W* coding the C-terminal region emerged as a new exon [5]. We reported that DM-W and DMRT1 could cause primary ovarian and testicular formation in developing ZW

and ZZ gonads, respectively [19], and proposed a sex-determining model for the ZZ/ZW type that DM-W determines female sex by antagonizing DMRT1; *dm-W* evolved from a masculinizing gene *dmrt1* as a dominant negative-type gene [14].

## **4. Sex reversal and sex chromosome differentiation**

Although all frog species might genetically determine sex as mentioned above, most frog species could accept male-to-female or female-to-male sex reversals by treatment of sex steroids, estrogen, or androgen, respectively, during tadpole development [1]. Importantly, many frogs of them have homomorphic sex chromosomes. For example, *X. laevis* carries homomorphic W and Z sex chromosomes [5], and the estradiol-treated ZZ tadpoles developed to female adults [1]. In addition, we reported ZW female-to-male sex reversals in *X. laevis* transgenic tadpoles with *dm-W* knockdown or germline stem cell-specific knockdown of *dmrt1* and ZZ male-tofemale sex reversals in *X. laevis* transgenic tadpoles carrying the *dm-W* expression plasmid [5, 19, 21].

Moreover, we recently analyzed detail structures of the sex chromosomes on 2Lq32-33 in *X. laevis*, revealing 278 kb W-specific region including three W-specific genes, the sex-determining gene *dm-W*, *scanw*, and *ccdc69w*, and 83 kb Z-specific region including one Z-specific gene *capn5z* [24]. Importantly, both gynogenetic WW and estrogen-driven sex-reversed ZZ individuals could develop into normal fertile females [25, 26]. These findings suggest that the homomorphic W/Z sex chromosomes in *X. laevis* are now differentiating but not so differentiated yet. In other words, *X. laevis* sex chromosomes have the potential to accept sex reversal and a new sex-determining gene.

## **5. Conclusions and perspective**

All frogs examined possess genetic sex-determining systems, and most of them have homomorphic sex chromosomes. The genetic systems could be easy to change during species diversity, that is, the instability of the systems, maybe because of homomorphic sex chromosomes, which could have a potential to convert a sexdetermining gene into a new one on another chromosome, resulting in the change of sex chromosomes. Then I propose a "GENE-eat-GENE" model for turnover of sex-determining genes: there has been battles among the present sex-determining gene and candidates of new sex-determining genes for king/queen ship in some populations holding homomorphic sex chromosomes (**Figure 1**). Accordingly, I predict that there are great many sex-determining genes in frogs, although only one *dm-W* has been identified as sex-determining genes. Frogs belong to the order Anura, which collects several thousands of species. Therefore they could be good examples for studying the relationships between sex-determining systems and species diversity.

**17**

**Author details**

Department of Biosciences, School of Science, Kitasato University, Sagamihara,

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium,

\*Address all correspondence to: ito@sci.kitasato-u.ac.jp

provided the original work is properly cited.

Michihiko Ito

Japan

*Instability of Sex-Determining Systems in Frogs DOI: http://dx.doi.org/10.5772/intechopen.89050* *Instability of Sex-Determining Systems in Frogs DOI: http://dx.doi.org/10.5772/intechopen.89050*

*Gene Expression and Phenotypic Traits*

plasmid [5, 19, 21].

a new sex-determining gene.

**5. Conclusions and perspective**

and ZZ gonads, respectively [19], and proposed a sex-determining model for the ZZ/ZW type that DM-W determines female sex by antagonizing DMRT1; *dm-W* evolved from a masculinizing gene *dmrt1* as a dominant negative-type gene [14].

Although all frog species might genetically determine sex as mentioned above, most frog species could accept male-to-female or female-to-male sex reversals by treatment of sex steroids, estrogen, or androgen, respectively, during tadpole development [1]. Importantly, many frogs of them have homomorphic sex chromosomes. For example, *X. laevis* carries homomorphic W and Z sex chromosomes [5], and the estradiol-treated ZZ tadpoles developed to female adults [1]. In addition, we reported ZW female-to-male sex reversals in *X. laevis* transgenic tadpoles with *dm-W* knockdown or germline stem cell-specific knockdown of *dmrt1* and ZZ male-tofemale sex reversals in *X. laevis* transgenic tadpoles carrying the *dm-W* expression

Moreover, we recently analyzed detail structures of the sex chromosomes on 2Lq32-33 in *X. laevis*, revealing 278 kb W-specific region including three W-specific genes, the sex-determining gene *dm-W*, *scanw*, and *ccdc69w*, and 83 kb Z-specific region including one Z-specific gene *capn5z* [24]. Importantly, both gynogenetic WW and estrogen-driven sex-reversed ZZ individuals could develop into normal fertile females [25, 26]. These findings suggest that the homomorphic W/Z sex chromosomes in *X. laevis* are now differentiating but not so differentiated yet. In other words, *X. laevis* sex chromosomes have the potential to accept sex reversal and

All frogs examined possess genetic sex-determining systems, and most of them have homomorphic sex chromosomes. The genetic systems could be easy to change during species diversity, that is, the instability of the systems, maybe because of homomorphic sex chromosomes, which could have a potential to convert a sexdetermining gene into a new one on another chromosome, resulting in the change of sex chromosomes. Then I propose a "GENE-eat-GENE" model for turnover of sex-determining genes: there has been battles among the present sex-determining gene and candidates of new sex-determining genes for king/queen ship in some populations holding homomorphic sex chromosomes (**Figure 1**). Accordingly, I predict that there are great many sex-determining genes in frogs, although only one *dm-W* has been identified as sex-determining genes. Frogs belong to the order Anura, which collects several thousands of species. Therefore they could be good examples for studying the relationships between sex-determining systems and

**4. Sex reversal and sex chromosome differentiation**

**16**

species diversity.

## **Author details**

Michihiko Ito Department of Biosciences, School of Science, Kitasato University, Sagamihara, Japan

\*Address all correspondence to: ito@sci.kitasato-u.ac.jp

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

## **References**

[1] Ito M. Sex determination and differentiation in frogs. In: Kobayashi et al., editors. Reproductive and Developmental Strategies. Springer; 2018. pp. 349-366

[2] Abramyan J, Ezaz T, Graves JA, Koopman P. Z and W sex chromosomes in the cane toad (*Bufo marinus*). Chromosome Research. 2009;**17**:1015-1024

[3] Chang CY, Witschi E. Genic control and hormonal reversal of sex differentiation in *Xenopus*. Proceedings of the Society for Experimental Biology and Medicine. 1956;**93**:140-144

[4] Schmid M, Bachmann K. A frog with highly evolved sex chromosomes. Experientia. 1981;**37**:243-245

[5] Yoshimoto S, Okada E, Umemoto H, Tamura K, Uno Y, Nishida-Umehara C, et al. A W-linked DM-domain gene, DM-W, participates in primary ovary development in *Xenopus laevis*. Proceedings of the National Academy of Sciences of the United States of America. 2008;**105**:2469-2474

[6] De Almeida CG, Grafe TU, Guttenbach M, Schmid M. Karyotype and chromosome banding in the reed frog *Hyperolius viridiflavus ommatostictus* (Amphibia, *Anura*, *Hyperoliidae*). Experientia. 1990;**46**:509-511

[7] Schmid M, Haaf T, Geile B, Sims S. Chromosome banding in Amphibia. VIII. An unusual XY/XX-sex chromosome system in *Gastrotheca riobambae* (Anura, Hylidae). Chromosoma. 1983;**88**:69-82

[8] Miura I. An evolutionary witness: The frog rana rugosa underwent change of heterogametic sex from XY male to ZW female. Sexual Development. 2007;**1**:323-331

[9] Eggert C. Sex determination: The amphibian models. Reproduction,

Nutrition, Development. 2004;**44**: 539-549

[10] Malcom JW, Kudra RS, Malone JH. The sex chromosomes of frogs: Variability and tolerance offer clues to genome evolution and function. Journal of Genomics. 2014;**2**:68-76

[11] Schmid M, Steinlein C, Bogart JP, Feichtinger W, León P, La Marca E, et al. The chromosomes of terraranan frogs. Insights into vertebrate cytogenetics. Cytogenetic and Genome Research. 2010;**130-131**:1-568

[12] Ito M, Mawaribuchi S. Molecular evolution of genes involved in vertebrate sex determination. In: eLS. Chichester: John Wiley & Sons, Ltd; 2013

[13] Mawaribuchi S, Yoshimoto S, Ohashi S, Takamatsu N, Ito M. Molecular evolution of vertebrate sex-determining genes. Chromosome Research. 2012;**20**:139-151

[14] Yoshimoto S, Ito M. A ZZ/ZW-type sex determination in *Xenopus laevis*. The FEBS Journal. 2011;**278**:1020-1026

[15] Sinclair AH, Berta P, Palmer MS, Hawkins JR, Griffiths BL, Smith MJ, et al. A gene from the human sexdetermining region encodes a protein with homology to a conserved DNA-binding motif. Nature. 1990;**346**:240-244

[16] Koopman P, Gubbay J, Vivian N, Goodfellow P, Lovell-Badge R. Male development of chromosomally female mice transgenic for *Sry*. Nature. 1991;**351**:117-121

[17] Matsuda M, Nagahama Y, Shinomiya A, Sato T, Matsuda C, Kobayashi T, et al. DMY is a Y-specific DM-domain gene required for male development in the medaka fish. Nature. 2002;**417**:559-563

**19**

*Instability of Sex-Determining Systems in Frogs DOI: http://dx.doi.org/10.5772/intechopen.89050*

in *Xenopus laevis*. Molecular & General

[26] Villalpando I, Merchant-Larios H. Determination of the sensitive stages for gonadal sex-reversal in *Xenopus laevis* tadpoles. The International Journal of Developmental Biology.

Genetics. 1984;**194**:57-59

1990;**34**:281-285

[18] Nanda I, Kondo M, Hornung U, Asakawa S, Winkler C, Shimizu A, et al. A duplicated copy of DMRT1 in the sex-determining region of the Y chromosome of the medaka, *Oryzias latipes*. Proceedings of the National Academy of Sciences of the United States of America. 2002;**99**:11778-11783

[19] Yoshimoto S, Ikeda N, Izutsu Y, Shiba T, Takamatsu N, Ito M. Opposite roles of DMRT1 and its W-linked paralogue, DM-W, in sexual dimorphism of *Xenopus laevis*: Implications of a ZZ/ZW-type sexdetermining system. Development.

[20] Smith CA, Roeszler KN, Ohnesorg T, Cummins DM, Farlie PG, Doran TJ, et al. The avian Z-linked gene DMRT1 is required for male sex determination in the chicken. Nature. 2009;**461**:267-271

[21] Mawaribuchi S, Musashijima M, Wada M, Izutsu Y, Kurakata E, Park MK, et al. Molecular evolution of two distinct dmrt1 promoters for germ and somatic cells in vertebrate gonads. Molecular Biology and Evolution. 2017;**34**:724-733

[22] Session AM, Uno Y, Kwon T, Chapman JA, Toyoda A, Takahashi S, et al. Genome evolution in the allotetraploid frog *Xenopus laevis*.

Nature. 2016;**538**:336-343

[23] Bewick AJ, Anderson DW, Evans BJ. Evolution of the closely related, sex-related genes DM-W and DMRT1 in African clawed frogs (*Xenopus*). Evolution. 2011;**65**:698-712

[24] Mawaribuchi S, Takahashi S, Wada M, Uno Y, Matsuda Y, Kondo M, et al. Sex chromosome differentiation and the W- and Z-specific loci in *Xenopus laevis*. Developmental Biology.

[25] Colombelli B, Thiebaud CH, Muller WP. Production of WW superfemales by diploid gynogenesis

2017;**426**:393-400

2010;**137**:2519-2526

*Instability of Sex-Determining Systems in Frogs DOI: http://dx.doi.org/10.5772/intechopen.89050*

[18] Nanda I, Kondo M, Hornung U, Asakawa S, Winkler C, Shimizu A, et al. A duplicated copy of DMRT1 in the sex-determining region of the Y chromosome of the medaka, *Oryzias latipes*. Proceedings of the National Academy of Sciences of the United States of America. 2002;**99**:11778-11783

[19] Yoshimoto S, Ikeda N, Izutsu Y, Shiba T, Takamatsu N, Ito M. Opposite roles of DMRT1 and its W-linked paralogue, DM-W, in sexual dimorphism of *Xenopus laevis*: Implications of a ZZ/ZW-type sexdetermining system. Development. 2010;**137**:2519-2526

[20] Smith CA, Roeszler KN, Ohnesorg T, Cummins DM, Farlie PG, Doran TJ, et al. The avian Z-linked gene DMRT1 is required for male sex determination in the chicken. Nature. 2009;**461**:267-271

[21] Mawaribuchi S, Musashijima M, Wada M, Izutsu Y, Kurakata E, Park MK, et al. Molecular evolution of two distinct dmrt1 promoters for germ and somatic cells in vertebrate gonads. Molecular Biology and Evolution. 2017;**34**:724-733

[22] Session AM, Uno Y, Kwon T, Chapman JA, Toyoda A, Takahashi S, et al. Genome evolution in the allotetraploid frog *Xenopus laevis*. Nature. 2016;**538**:336-343

[23] Bewick AJ, Anderson DW, Evans BJ. Evolution of the closely related, sex-related genes DM-W and DMRT1 in African clawed frogs (*Xenopus*). Evolution. 2011;**65**:698-712

[24] Mawaribuchi S, Takahashi S, Wada M, Uno Y, Matsuda Y, Kondo M, et al. Sex chromosome differentiation and the W- and Z-specific loci in *Xenopus laevis*. Developmental Biology. 2017;**426**:393-400

[25] Colombelli B, Thiebaud CH, Muller WP. Production of WW superfemales by diploid gynogenesis in *Xenopus laevis*. Molecular & General Genetics. 1984;**194**:57-59

[26] Villalpando I, Merchant-Larios H. Determination of the sensitive stages for gonadal sex-reversal in *Xenopus laevis* tadpoles. The International Journal of Developmental Biology. 1990;**34**:281-285

**18**

2007;**1**:323-331

*Gene Expression and Phenotypic Traits*

[1] Ito M. Sex determination and differentiation in frogs. In: Kobayashi et al., editors. Reproductive and Developmental Strategies. Springer;

Nutrition, Development. 2004;**44**:

Malone JH. The sex chromosomes of frogs: Variability and tolerance offer clues to genome evolution and function. Journal of Genomics. 2014;**2**:68-76

[11] Schmid M, Steinlein C, Bogart JP, Feichtinger W, León P, La Marca E, et al. The chromosomes of terraranan frogs. Insights into vertebrate cytogenetics. Cytogenetic and Genome Research.

[12] Ito M, Mawaribuchi S. Molecular evolution of genes involved in vertebrate sex determination. In: eLS. Chichester:

John Wiley & Sons, Ltd; 2013

Research. 2012;**20**:139-151

1990;**346**:240-244

1991;**351**:117-121

[13] Mawaribuchi S, Yoshimoto S, Ohashi S, Takamatsu N, Ito M. Molecular evolution of vertebrate sex-determining genes. Chromosome

[14] Yoshimoto S, Ito M. A ZZ/ZW-type sex determination in *Xenopus laevis*. The FEBS Journal. 2011;**278**:1020-1026

[15] Sinclair AH, Berta P, Palmer MS, Hawkins JR, Griffiths BL, Smith MJ, et al. A gene from the human sexdetermining region encodes a protein with homology to a conserved DNA-binding motif. Nature.

[16] Koopman P, Gubbay J, Vivian N, Goodfellow P, Lovell-Badge R. Male development of chromosomally female mice transgenic for *Sry*. Nature.

[17] Matsuda M, Nagahama Y, Shinomiya A, Sato T, Matsuda C, Kobayashi T, et al. DMY is a Y-specific DM-domain gene required for male development in the medaka fish.

Nature. 2002;**417**:559-563

[10] Malcom JW, Kudra RS,

2010;**130-131**:1-568

539-549

[2] Abramyan J, Ezaz T, Graves JA, Koopman P. Z and W sex chromosomes in the cane toad (*Bufo marinus*).

[3] Chang CY, Witschi E. Genic control and hormonal reversal of sex differentiation in *Xenopus*. Proceedings of the Society for Experimental Biology

and Medicine. 1956;**93**:140-144

Experientia. 1981;**37**:243-245

[6] De Almeida CG, Grafe TU,

Guttenbach M, Schmid M. Karyotype and chromosome banding in the reed frog *Hyperolius viridiflavus ommatostictus* (Amphibia, *Anura*, *Hyperoliidae*). Experientia. 1990;**46**:509-511

[7] Schmid M, Haaf T, Geile B, Sims S. Chromosome banding in Amphibia. VIII. An unusual XY/XX-sex chromosome system in *Gastrotheca riobambae* (Anura, Hylidae). Chromosoma. 1983;**88**:69-82

[8] Miura I. An evolutionary witness: The frog rana rugosa underwent change of heterogametic sex from XY male to ZW female. Sexual Development.

[9] Eggert C. Sex determination: The amphibian models. Reproduction,

[4] Schmid M, Bachmann K. A frog with highly evolved sex chromosomes.

[5] Yoshimoto S, Okada E, Umemoto H, Tamura K, Uno Y, Nishida-Umehara C, et al. A W-linked DM-domain gene, DM-W, participates in primary ovary development in *Xenopus laevis*. Proceedings of the National Academy of Sciences of the United States of America. 2008;**105**:2469-2474

Chromosome Research. 2009;**17**:1015-1024

2018. pp. 349-366

**References**

**21**

**Chapter 3**

**Abstract**

sex determining genes

**1. Introduction**

Comparison of Sex Determination

The chapter is devoted to the consideration of sex determination in vertebrate groups of nonmammalians: fish, amphibians, reptiles, and birds. Attention is drawn to the fact that all these groups of animals, unlike mammals, are implemented hormonal control options for primary sex determination, and there is a possibility of sex reversion. Determination of gonadal development in vertebrates like testis or ovary was initially controlled mainly by sex hormones (fish and amphibians). Later, various sex determining genes were involved in this process. The system was quite plastic and was able to respond to changes in external conditions (reptiles). The appearance of heteromorphic sex chromosomes (birds) has led to the emergence of some specific W chromosomal signal, which provides estrogen control of the development of a heterogametic sex. In mammals, the control of the primary determination of sex (the appearance of the gonad) becomes purely genetic, and the role of sex hormones is reduced to the differentiation of testis or ovaries.

in Vertebrates (Nonmammals)

*Aleksandr F. Smirnov and Antonina V. Trukhina*

**Keywords:** sex determination, sex hormones, sex chromosomes,

Gender is a set of morphological and physiological characteristics of the organism, providing reproduction, the essence of which is to fertilization, i.e. the fusion of male and female germ cells (gametes) in zygote, which develops into a new organism. Differentiation of sex (its phenotypic manifestation) includes two successive stages: the primary determination of sex and the appearance of secondary (external) sexual characteristics (actual differentiation). It is believed that the concept of this process is conservative. Sex determination is both a genetic and ecological process, with the sex of the individual being determined by an alternative physiological solution. It is assumed that there are two main mechanisms for determining sex: genetic (GSD—genetic sex determination) and environmental (ESD—environmental sex determination). Genetic sex is determined at the time of conception and depends on genetic differences between males and females, and ecological sex depends on external conditions in the absence of significant genetic differences and is determined after fertilization in response to environmental conditions. For birds and mammals, only the GSD is characteristic, and for crocodiles—TSD (one of the forms of ESD). In addition, there are two varieties of the genetic sex determination system: with heterogametic males (XY, mammals) and heterogametic females (ZW, birds). It should be noted that amphibians have both

## **Chapter 3**

## Comparison of Sex Determination in Vertebrates (Nonmammals)

*Aleksandr F. Smirnov and Antonina V. Trukhina*

## **Abstract**

The chapter is devoted to the consideration of sex determination in vertebrate groups of nonmammalians: fish, amphibians, reptiles, and birds. Attention is drawn to the fact that all these groups of animals, unlike mammals, are implemented hormonal control options for primary sex determination, and there is a possibility of sex reversion. Determination of gonadal development in vertebrates like testis or ovary was initially controlled mainly by sex hormones (fish and amphibians). Later, various sex determining genes were involved in this process. The system was quite plastic and was able to respond to changes in external conditions (reptiles). The appearance of heteromorphic sex chromosomes (birds) has led to the emergence of some specific W chromosomal signal, which provides estrogen control of the development of a heterogametic sex. In mammals, the control of the primary determination of sex (the appearance of the gonad) becomes purely genetic, and the role of sex hormones is reduced to the differentiation of testis or ovaries.

**Keywords:** sex determination, sex hormones, sex chromosomes, sex determining genes

## **1. Introduction**

Gender is a set of morphological and physiological characteristics of the organism, providing reproduction, the essence of which is to fertilization, i.e. the fusion of male and female germ cells (gametes) in zygote, which develops into a new organism. Differentiation of sex (its phenotypic manifestation) includes two successive stages: the primary determination of sex and the appearance of secondary (external) sexual characteristics (actual differentiation). It is believed that the concept of this process is conservative. Sex determination is both a genetic and ecological process, with the sex of the individual being determined by an alternative physiological solution. It is assumed that there are two main mechanisms for determining sex: genetic (GSD—genetic sex determination) and environmental (ESD—environmental sex determination). Genetic sex is determined at the time of conception and depends on genetic differences between males and females, and ecological sex depends on external conditions in the absence of significant genetic differences and is determined after fertilization in response to environmental conditions. For birds and mammals, only the GSD is characteristic, and for crocodiles—TSD (one of the forms of ESD). In addition, there are two varieties of the genetic sex determination system: with heterogametic males (XY, mammals) and heterogametic females (ZW, birds). It should be noted that amphibians have both

genetic systems, and for lizards, snakes, turtles, and bony fish, all possible variants of sex determination are described [1–3].

Sex steroid hormones including androgens, estrogens, and progesterone are present in all vertebrates which play essential roles in modulating a variety of behavior and processes, such as embryonic development, sexual differentiation, growth, aggression, reproduction, learning, memory, social communication, and so on. Many signaling actions of these sex steroid hormones are mediated by their receptors that belong to the superfamily of steroid nuclear receptors. Once a sex steroid hormone ligand binds to its receptor, the receptor becomes phosphorylated and is translocated into the nucleus, where it binds to specific DNA sequences and activates gene transcription. Androgens have a critical physiological role in reproductive biology and sexual differentiation, particularly in the development of male secondary sex characteristics [4, 5].

It is assumed that sex determination is a combination of hormonal and genetic factors and is divided conditionally into appropriate stages. This phenomenon is reflected in the possibility of sex inversion—the possibility of its complete or partial hormonal alteration. For fishes and amphibians, there is the sensitivity of normal development of the gonads to androgens and estrogens. In reptiles, birds and marsupials, only estrogens are effective. The appearance of the gonads of placental mammals does not depend on sex hormones. This trend is associated with the stability of growing offspring or incubation of eggs [6].

The proposed chapter will consider the system of sex determination in fish, amphibians, reptiles, and birds in comparing the role of hormonal and genetic mechanisms, possibilities, and mechanisms of sex inversion.

## **2. Features of sex determination in fishes**

Fishes are perhaps the most complex group of animals in the mechanism of sex determination. Only bony fish include over 30,000 species. It is the largest group of vertebrates. They are divided into three groups in accordance with the laws of sex determination: (1) gonochoristic species whose sex is determined genetically or through environmental factors; (2) sequential hermaphrodites (about 2% of all existing species), changing the sex of males to females (protandrous), the sex of females to males (protogynous), or in both directions (serial) in the process of ontogenesis; (3) unisexual type of sex determination (characteristic only for Amazon mollies (*Poecilia formosa*)). Gonochoristic genetics of sex in fish is largely unclear. Functional hermaphroditism occurs in many different species of animals such as echinoderms, crustaceans, molluscs, and fish; however, it is lost in vertebrates during the transition from amphibians to mammals. From here, fishes provide a unique model for studying the mechanism of hermaphroditism in vertebrates. Unfortunately, only one species of fish (Japanese medaka—*Oryzias latipes*) was identified by a primary system of sex determination [7, 8].

The Japanese medaka (Oryzias latipes) and Maebashi medaka (*Oryzias curvinotus*)—species with heterogametic male sex with homomorphic sex chromosomes that are a very early stage of evolution, the recently described Y-chromosome plot, containing hypothetical gene *dmy*. This gene is specifically expressed in the gonads and is essential for embryo development in male type. Gene *dmrt1bY* (*dmy*) homologous (about 80%) of the *dmrt1* gene in other species of vertebrates represents the equivalent of *sry* gene in mammals. It is important that medaka *dmy*/*dmrt1bY* is a unique system. This species is described as ontology mammalian *sox9* gene, but in contrast to amniotes and amphibians, this does not play a role in determining the testes. Sex determination system of medaka is unstable. Medaka has interesting

**23**

**Figure 1.**

*Comparison of Sex Determination in Vertebrates (Nonmammals)*

significant genetic divergence: *dmy* gene is absent in some lines of the Japanese medaka (over 10%) and other types of fish of the genus *Oryzias.* In some laboratory lines, the proportion of homogametic males (XX) exceeds 20%. It is believed that gene *dmy* has occurred as a result of the *dmrt1* gene duplication and transposition of part of its copy size to 280 kbp about 10 million years ago. The products of these genes differ only in one amino acid replacement (Ser26/Thr), which may have led to such differences by gender. It has been shown that the rate of synonymous substitutions in the *dmy* is 1.78 times greater than that of *dmrt1* and this is consistent with the hypothesis of evolution through males (male-driven evolution hypothesis). In birds and salmon, it has the same orientation. The speed ratio of nonsynonymous substitutions (dN) to synonymous (dS) is also higher in comparison with *dmy*/*dmrt1*. Only two sex determining genes in vertebrates were described: *sry* and *dmy*. It is believed that the protein DMY performs two different functions in germ and somatic cells. In somatic cells surrounding germ ones, it affects the proliferation of the latter (for example, influencing a cascade of genes involved in the transmission of the estrogen signal). Another feature is the induction of development of pre-Sertoli cells (cells surrounding the primary germ cells (PGCs)) in the gonad heterogametic XY sex. In this case, there is an analogy with *sry*, which is involved in the activation of other genes that support the development of Sertoli cells. In medaka, there are other female-specific genes and male-specific genes (**Figure 1**). Moreover, the latter gene is located in autosomes. Some ideas of the diversity of sex

In this species, the sex determined region of the Y chromosome is only 260 kb (1% of the total length of the Y chromosome (59 Mbps)). In this area, there is suppression of recombination. In medaka, all XY individuals carry mutations in the gene *dmy* form ovaries. In individuals with altered *gsdf*-gene, sex inversion is also observed. It is believed that for medaka, the normal gene *dmrt1* (*dmy*) initiates the formation of the testes and controls their maintenance with *gsdf*. The study of sex chromosomes in six species of medaka from the group *celebensis* with XX/XY-sex determination showed that *O. marmoratus* and *O. profundicola* sex chromosomes homologous sex chromosomes *of O. latipes* from the LG10 linkage group. Four species *O. celebensis, O. matanensis, O. wolasi* and *O. woworae* marked homology

*A schematic diagram of sex determination and gonad (testis or ovary) differentiation in fish with XX/XY sex* 

*DOI: http://dx.doi.org/10.5772/intechopen.83831*

determining genes among medaka given.

*determination system (adopted from Mei and Gui [10]).*

## *Comparison of Sex Determination in Vertebrates (Nonmammals) DOI: http://dx.doi.org/10.5772/intechopen.83831*

*Gene Expression and Phenotypic Traits*

of sex determination are described [1–3].

secondary sex characteristics [4, 5].

stability of growing offspring or incubation of eggs [6].

**2. Features of sex determination in fishes**

mechanisms, possibilities, and mechanisms of sex inversion.

was identified by a primary system of sex determination [7, 8].

The Japanese medaka (Oryzias latipes) and Maebashi medaka (*Oryzias curvinotus*)—species with heterogametic male sex with homomorphic sex chromosomes that are a very early stage of evolution, the recently described Y-chromosome plot, containing hypothetical gene *dmy*. This gene is specifically expressed in the gonads and is essential for embryo development in male type. Gene *dmrt1bY* (*dmy*) homologous (about 80%) of the *dmrt1* gene in other species of vertebrates represents the equivalent of *sry* gene in mammals. It is important that medaka *dmy*/*dmrt1bY* is a unique system. This species is described as ontology mammalian *sox9* gene, but in contrast to amniotes and amphibians, this does not play a role in determining the testes. Sex determination system of medaka is unstable. Medaka has interesting

genetic systems, and for lizards, snakes, turtles, and bony fish, all possible variants

Sex steroid hormones including androgens, estrogens, and progesterone are present in all vertebrates which play essential roles in modulating a variety of behavior and processes, such as embryonic development, sexual differentiation, growth, aggression, reproduction, learning, memory, social communication, and so on. Many signaling actions of these sex steroid hormones are mediated by their receptors that belong to the superfamily of steroid nuclear receptors. Once a sex steroid hormone ligand binds to its receptor, the receptor becomes phosphorylated and is translocated into the nucleus, where it binds to specific DNA sequences and activates gene transcription. Androgens have a critical physiological role in reproductive biology and sexual differentiation, particularly in the development of male

It is assumed that sex determination is a combination of hormonal and genetic factors and is divided conditionally into appropriate stages. This phenomenon is reflected in the possibility of sex inversion—the possibility of its complete or partial hormonal alteration. For fishes and amphibians, there is the sensitivity of normal development of the gonads to androgens and estrogens. In reptiles, birds and marsupials, only estrogens are effective. The appearance of the gonads of placental mammals does not depend on sex hormones. This trend is associated with the

The proposed chapter will consider the system of sex determination in fish, amphibians, reptiles, and birds in comparing the role of hormonal and genetic

Fishes are perhaps the most complex group of animals in the mechanism of sex determination. Only bony fish include over 30,000 species. It is the largest group of vertebrates. They are divided into three groups in accordance with the laws of sex determination: (1) gonochoristic species whose sex is determined genetically or through environmental factors; (2) sequential hermaphrodites (about 2% of all existing species), changing the sex of males to females (protandrous), the sex of females to males (protogynous), or in both directions (serial) in the process of ontogenesis; (3) unisexual type of sex determination (characteristic only for Amazon mollies (*Poecilia formosa*)). Gonochoristic genetics of sex in fish is largely unclear. Functional hermaphroditism occurs in many different species of animals such as echinoderms, crustaceans, molluscs, and fish; however, it is lost in vertebrates during the transition from amphibians to mammals. From here, fishes provide a unique model for studying the mechanism of hermaphroditism in vertebrates. Unfortunately, only one species of fish (Japanese medaka—*Oryzias latipes*)

**22**

significant genetic divergence: *dmy* gene is absent in some lines of the Japanese medaka (over 10%) and other types of fish of the genus *Oryzias.* In some laboratory lines, the proportion of homogametic males (XX) exceeds 20%. It is believed that gene *dmy* has occurred as a result of the *dmrt1* gene duplication and transposition of part of its copy size to 280 kbp about 10 million years ago. The products of these genes differ only in one amino acid replacement (Ser26/Thr), which may have led to such differences by gender. It has been shown that the rate of synonymous substitutions in the *dmy* is 1.78 times greater than that of *dmrt1* and this is consistent with the hypothesis of evolution through males (male-driven evolution hypothesis). In birds and salmon, it has the same orientation. The speed ratio of nonsynonymous substitutions (dN) to synonymous (dS) is also higher in comparison with *dmy*/*dmrt1*.

Only two sex determining genes in vertebrates were described: *sry* and *dmy*. It is believed that the protein DMY performs two different functions in germ and somatic cells. In somatic cells surrounding germ ones, it affects the proliferation of the latter (for example, influencing a cascade of genes involved in the transmission of the estrogen signal). Another feature is the induction of development of pre-Sertoli cells (cells surrounding the primary germ cells (PGCs)) in the gonad heterogametic XY sex. In this case, there is an analogy with *sry*, which is involved in the activation of other genes that support the development of Sertoli cells. In medaka, there are other female-specific genes and male-specific genes (**Figure 1**). Moreover, the latter gene is located in autosomes. Some ideas of the diversity of sex determining genes among medaka given.

In this species, the sex determined region of the Y chromosome is only 260 kb (1% of the total length of the Y chromosome (59 Mbps)). In this area, there is suppression of recombination. In medaka, all XY individuals carry mutations in the gene *dmy* form ovaries. In individuals with altered *gsdf*-gene, sex inversion is also observed. It is believed that for medaka, the normal gene *dmrt1* (*dmy*) initiates the formation of the testes and controls their maintenance with *gsdf*. The study of sex chromosomes in six species of medaka from the group *celebensis* with XX/XY-sex determination showed that *O. marmoratus* and *O. profundicola* sex chromosomes homologous sex chromosomes *of O. latipes* from the LG10 linkage group. Four species *O. celebensis, O. matanensis, O. wolasi* and *O. woworae* marked homology

#### **Figure 1.**

*A schematic diagram of sex determination and gonad (testis or ovary) differentiation in fish with XX/XY sex determination system (adopted from Mei and Gui [10]).*

with the chromosomes of LG 24, which involves the transformation of chromosomes from *O. latipes* LG to 24 LG10 within this group. All six studied species share a common sex determined gene (SD). It is shown that genomic predecessor is the Y-chromosomal gene *sox3* and this process involves specific insertion (430 bp).

The zebrafish testes derived from *dmrt1* mutant fish fail to express the anti-Müllerian hormone (*amh*) gene, a key testis-expressed gene, and over-express the ovary-associated gene *foxl2*. Therefore, zebrafish *dmrt1* shares similar roles in male sexual development as other organisms in regulating sex determination and testis differentiation.

In other fishes, e.g., salmonids, there appear to be an early stage of differentiation of sex chromosomes. In rainbow trout (*Oncorhynchus mykiss*) with monofactorial XX/XY system of sex determination, a new gene *sdY* responsible for the development of testes is described. This gene is partially similar to the gene regulator of interferon 9. It has been found that highly conserved in *sdY* salmon is male Y-chromosomal gene for the majority of these species. It is assumed that it is the main testis determining gene for this group of fishes. For the two species of whitefish (subfamily *Coregoninae*), the *sdY* gene is found in both males and females. This implies that there is an alternative system of sex determination in this family. Among other candidate genes for sex determination, gene antimullerian hormone (*amh*) tilapia is discussed. Fishes with hermaphrodite sex determination (*Labridae*, fish-clowns—amphiprion (*Amphiprion),* and gobies—*Trimma okinawae*) have got bisexual gonads capable of restructuring with the participation of aromatase and gonadotropin receptors. For some species, such as blue tilapia (*Oreochromis aureus*), sex determined putative gene is located on the genetic map of a sex determining region consisting of more than 550 minisatellite markers [7, 9].

In vertebrates, until recently, only four sex determining genes were discovered: *sry* (in mammals), *dmrt1* (in domestic chicken), *dmy* (the Japanese medaka), and *dm-w* (the frog). Recently, four candidate genes were found for this role (and all fish): Patagonian aterin have *amhy*, Luzon ricefish (*Oryzias luzonensis*) have *gsdf*, and puffer (*Takifugu or Fugu*)—*amhr2* and rainbow trout—*sdy*. In the Nile tilapia (*Oreochromis niloticus)* gene *gdf*, (gonadal soma derived factor (*gsdf*)) also induces the development of the testes. Assume that the Atlantic salmon *sdY* gene product activates genes *gsdf* and *amh/mi*, thereby reducing the activity of aromatase (*cyp19a* gene), leading to the appearance of males. Where *sdY* is missing, aromatase is synthesized in quantities sufficient for the emergence of the females [8, 10].

Sex determining genes in fish are not conservative. It is believed that the reason for this is the more frequent variation of sex chromosomes in fish than other coldblooded animals and mammals (**Figure 1**).

These objects sex determination has a high plasticity and is, therefore, possible sex reversal, even in species with established regulatory genes. Striped Danio (*Danio rerio*) experimental data are in good agreement with polygenic sex determination (PSD) when the sex is determined by allelic combinations of several loci. Typically, these loci are dispersed throughout the genome, but some species of bony fish are placed in special sex chromosomes. In hermaphroditic fish, ovotestis develops first, and then secondary sex determination occurs. So, the black bass individuals (genus *Micropterus*) in the first 2 years of life are males, but in the third year, 50% of them are transformed into females. Sex determining male genes such as *dmrt1*, *amh*, and *amhr2* are activated during differentiation of the testis, and their expression is maintained at high level during the period of functioning as males. High dose estrogen E2 induces the development of ovarian and testicular tissue degradation [11, 12].

In fish, there are two systems of sex determination: XX/XY and ZW/ZZ. The most common one is the last. Exploring the flatfish *Cynoglossus semilaevis* as a model

**25**

**Figure 2.**

*Comparison of Sex Determination in Vertebrates (Nonmammals)*

species with genetic sex determination system of ZW-type and the simultaneous presence of ESD, it was found that about 14% of females at a temperature of 22°C become males (pseudomales). It is believed that there is *dmrt1* gene (double sex and mab-3 related transcription factor 1) which is the sex determining gene in this species. It was also shown that pseudomales change the level of methylation of a certain portion of the Z chromosome, resulting in the intensity of transcription in this area as in normal males. In females, on the contrary, the activity of the corresponding plot of W chromosome by methylation is suppressed. Unusual WXZ-system is described for the swordtail (*Xiphophorus helleri*). Not so many fish species had morphologically different sex chromosomes (about 10%) and in most species they are in the early stages of their differentiation. For many members of this class, sex is determined by the environment, and even changes under the influence of behav-

ioral factors. There are species with heterogametic male and female [13].

Fish is characterized by plasticity of germ and somatic cells. This plasticity is maintained throughout the life cycle. Furthermore, they have described the influence of factors on this process such as temperature, pH, density of population, etc. It should be noted that the temperature sensitivity of fish is different from that of reptiles, especially because these types of monosexual populations are rare, even under extreme conditions. TSD in fish is less common than previously thought. The effect of estrogens, acting via estrogen receptors (ER) and directly or indirectly regulating P450arom and AMH, is particularly noticeable. It is noted that the analysis of the differences between gonochoristic and hermaphroditic fish species will help to understand the mechanism of plasticity of sex determination in vertebrates. In addition, there is the idea that gender in fish depending

*The profiles of gonadal development in three different sexual phases in hermaphroditic Japanese black porgy, Acanthopagrus schlegelii. Maleness: the fishes are functional males in the first two spawning seasons. The testis exists at all stages of the reproductive cycles in maleness. Active femaleness: the fishes are functional females following the natural sex change that occurs in fish older than 2 years or when induced by the removal of the testis of the digonic gonad. The ovary could reach to the stage of vitellogenesis, vitellogenic, and mature oocytes. Passive femaleness: Long-term E2 (4–6 mg/kg feed) administration for 2–3 months results in the appearance of a dominant ovary (with the primary oocytes) with a regressed testis in fish younger than 2 years old, and no vitellogenic oocytes are observed in E2-induced sex-changing fish. A reversible sex change (from passive femaleness to maleness) exists after E2 administration has been withdrawn. Undiff. gonad, undifferentiated gonad; E2, estradiol-17β; PO, primary oocyte stage; and VO, vitellogenic oocyte stage (adopted from Wu and Chang [16]).*

*DOI: http://dx.doi.org/10.5772/intechopen.83831*

## *Comparison of Sex Determination in Vertebrates (Nonmammals) DOI: http://dx.doi.org/10.5772/intechopen.83831*

*Gene Expression and Phenotypic Traits*

differentiation.

with the chromosomes of LG 24, which involves the transformation of chromosomes from *O. latipes* LG to 24 LG10 within this group. All six studied species share a common sex determined gene (SD). It is shown that genomic predecessor is the Y-chromosomal gene *sox3* and this process involves specific insertion (430 bp). The zebrafish testes derived from *dmrt1* mutant fish fail to express the anti-Müllerian hormone (*amh*) gene, a key testis-expressed gene, and over-express the ovary-associated gene *foxl2*. Therefore, zebrafish *dmrt1* shares similar roles in male sexual development as other organisms in regulating sex determination and testis

In other fishes, e.g., salmonids, there appear to be an early stage of differentiation of sex chromosomes. In rainbow trout (*Oncorhynchus mykiss*) with monofactorial XX/XY system of sex determination, a new gene *sdY* responsible for the development of testes is described. This gene is partially similar to the gene regulator of interferon 9. It has been found that highly conserved in *sdY* salmon is male Y-chromosomal gene for the majority of these species. It is assumed that it is the main testis determining gene for this group of fishes. For the two species of whitefish (subfamily *Coregoninae*), the *sdY* gene is found in both males and females. This implies that there is an alternative system of sex determination in this family. Among other candidate genes for sex determination, gene antimullerian hormone (*amh*) tilapia is discussed. Fishes with hermaphrodite sex determination (*Labridae*, fish-clowns—amphiprion (*Amphiprion),* and gobies—*Trimma okinawae*) have got bisexual gonads capable of restructuring with the participation of aromatase and gonadotropin receptors. For some species, such as blue tilapia (*Oreochromis aureus*), sex determined putative gene is located on the genetic map of a sex determining

region consisting of more than 550 minisatellite markers [7, 9].

blooded animals and mammals (**Figure 1**).

In vertebrates, until recently, only four sex determining genes were discovered: *sry* (in mammals), *dmrt1* (in domestic chicken), *dmy* (the Japanese medaka), and *dm-w* (the frog). Recently, four candidate genes were found for this role (and all fish): Patagonian aterin have *amhy*, Luzon ricefish (*Oryzias luzonensis*) have *gsdf*, and puffer (*Takifugu or Fugu*)—*amhr2* and rainbow trout—*sdy*. In the Nile tilapia (*Oreochromis niloticus)* gene *gdf*, (gonadal soma derived factor (*gsdf*)) also induces the development of the testes. Assume that the Atlantic salmon *sdY* gene product activates genes *gsdf* and *amh/mi*, thereby reducing the activity of aromatase (*cyp19a* gene), leading to the appearance of males. Where *sdY* is missing, aromatase is synthesized in quantities sufficient for the emergence of the females [8, 10].

Sex determining genes in fish are not conservative. It is believed that the reason for this is the more frequent variation of sex chromosomes in fish than other cold-

These objects sex determination has a high plasticity and is, therefore, possible sex reversal, even in species with established regulatory genes. Striped Danio (*Danio rerio*) experimental data are in good agreement with polygenic sex determination (PSD) when the sex is determined by allelic combinations of several loci. Typically, these loci are dispersed throughout the genome, but some species of bony fish are placed in special sex chromosomes. In hermaphroditic fish, ovotestis develops first, and then secondary sex determination occurs. So, the black bass individuals (genus *Micropterus*) in the first 2 years of life are males, but in the third year, 50% of them are transformed into females. Sex determining male genes such as *dmrt1*, *amh*, and *amhr2* are activated during differentiation of the testis, and their expression is maintained at high level during the period of functioning as males. High dose estrogen E2 induces the development of ovarian and testicular tissue

In fish, there are two systems of sex determination: XX/XY and ZW/ZZ. The most common one is the last. Exploring the flatfish *Cynoglossus semilaevis* as a model

**24**

degradation [11, 12].

species with genetic sex determination system of ZW-type and the simultaneous presence of ESD, it was found that about 14% of females at a temperature of 22°C become males (pseudomales). It is believed that there is *dmrt1* gene (double sex and mab-3 related transcription factor 1) which is the sex determining gene in this species. It was also shown that pseudomales change the level of methylation of a certain portion of the Z chromosome, resulting in the intensity of transcription in this area as in normal males. In females, on the contrary, the activity of the corresponding plot of W chromosome by methylation is suppressed. Unusual WXZ-system is described for the swordtail (*Xiphophorus helleri*). Not so many fish species had morphologically different sex chromosomes (about 10%) and in most species they are in the early stages of their differentiation. For many members of this class, sex is determined by the environment, and even changes under the influence of behavioral factors. There are species with heterogametic male and female [13].

Fish is characterized by plasticity of germ and somatic cells. This plasticity is maintained throughout the life cycle. Furthermore, they have described the influence of factors on this process such as temperature, pH, density of population, etc. It should be noted that the temperature sensitivity of fish is different from that of reptiles, especially because these types of monosexual populations are rare, even under extreme conditions. TSD in fish is less common than previously thought. The effect of estrogens, acting via estrogen receptors (ER) and directly or indirectly regulating P450arom and AMH, is particularly noticeable. It is noted that the analysis of the differences between gonochoristic and hermaphroditic fish species will help to understand the mechanism of plasticity of sex determination in vertebrates. In addition, there is the idea that gender in fish depending

#### **Figure 2.**

*The profiles of gonadal development in three different sexual phases in hermaphroditic Japanese black porgy, Acanthopagrus schlegelii. Maleness: the fishes are functional males in the first two spawning seasons. The testis exists at all stages of the reproductive cycles in maleness. Active femaleness: the fishes are functional females following the natural sex change that occurs in fish older than 2 years or when induced by the removal of the testis of the digonic gonad. The ovary could reach to the stage of vitellogenesis, vitellogenic, and mature oocytes. Passive femaleness: Long-term E2 (4–6 mg/kg feed) administration for 2–3 months results in the appearance of a dominant ovary (with the primary oocytes) with a regressed testis in fish younger than 2 years old, and no vitellogenic oocytes are observed in E2-induced sex-changing fish. A reversible sex change (from passive femaleness to maleness) exists after E2 administration has been withdrawn. Undiff. gonad, undifferentiated gonad; E2, estradiol-17β; PO, primary oocyte stage; and VO, vitellogenic oocyte stage (adopted from Wu and Chang [16]).*

on species is a complex trait under the control of one or many genetic factors in addition to environmental effects [9, 14]. In the Chinese tongue sole (*Cynoglossus semilaevis)*, genetic ZZ females may change into pseudomales, thereby increasing aquaculture costs because of the lower growth rate of the males than that of the females. A new locus was identified to regulate sex reversal interactively with the SNPCyn\_Z\_6676874; the linkage between these two loci and the absence of W sperm for pseudomales clearly elucidate the genetic architecture of sex reversal in the tongue sole [15]. Sexual determination in zebrafish is unique in that laboratory strains lack a sex chromosome, and no sex determining gene has been identified. GPER (estrogen receptor) is not required for normal sex differentiation, gonad development, or gonad function in zebrafish [16]. Genetic studies suggest that gonadal sexual fate is not only established by competition for primacy between two sexes via antagonistic signaling pathways during embryonic development but also requires active maintenance to suppress the opposite sex during adulthood. Documented in about 2% of teleost species spanning over 20 families, functional sex change generally occurs in three ways: protogynous (female-to-male), protandrous (male-to-female), and sequentially bi-directional. Most sequentially hermaphroditic fish are protogynous. Sex change in all hermaphroditic species involves radical gonadal transformation, and follows diverse ontogenetic pathways in different lineages particularly where sequential hermaphroditism has independently evolved. Gonadal transition in sex-changing fish is accompanied by changes in plasma concentrations of gonadal steroids. These steroids control gonad differentiation and maintain sexual phenotypes in teleost fish, wherein 17β-estradiol (E2) and 11-ketotestosterone (11-KT) function as the major estrogen and androgen, respectively. The balance between estrogen and androgen production is expected to control sexual fate of the gonads during sex change. For example,

#### **Figure 3.**

*The potential mechanism for sexual fate decision through the Gnrhs—Gths—Dmrt1 axis (brain-pituitarytestis axis). The model shows that the male fate decision is controlled by gonadotropins through the Gnrhs— Gths—Dmrt1 axis. The testis may stimulate the epigenetic modification of the ovary by DNA methylation of the cyp19a1a promoter to suppress the cyp19a1a expression. Gnrhs, gonadotropin-releasing hormones; Gths, gonadotropins; and Gthrs, gonadotropin receptors (adopted from Wu and Chang [16]).*

**27**

*Comparison of Sex Determination in Vertebrates (Nonmammals)*

numerous factors that potentially regulate its expression [15].

*dm-w* participate in the sex determination [18, 19] (**Figure 4**).

novel sex determining mechanism are yet to be identified [17].

In the northern crested newt (*Triturus cristatus*), the proportion of males increases when the ambient temperature increases, and a decrease of temperature leads to an excess of females. Thus, in amphibians, an increase or decrease of the ambient temperature leads to a modification of the normal development of the gonads and sex determination. Here, sex-determining genes are not the decisive factor in determining sex. A number of experiments have shown that atrazine and some other pesticides that affect the endocrine system affect the formation of sex in frogs. As a result, males are changed to females. Exogenous steroids (introduced from the outside) are also changing the sex in amphibians [20]. The unexplainable mechanism of sex determination in the rice frog species was introduced. Amphibians bearing a

factors regulating *cyp19a1a* expression are strong candidates for the trigger that initiates gonadal sex change; *cyp19a1a* promoter regions contain binding motifs for

However, hermaphroditic fishes have a plastic sex, and a stable sex is difficult to maintain with sex steroids. The black porgy regulated the dynamic development of both sexes; only one sex can grow while the other sex exists in a rudimentary stage (**Figure 2**). The sexual fate of the digonic gonad is determined by the male fate maintenance and through the Gnrh—Gth—Dmrt1 signaling. Altogether, testicular *dmrt1* and ovarian *cyp19a1a* expression are critical to the sexual fate of a male phase

Amphibians have two sex determined systems: XX/XY and ZZ/ZW. Most tailed amphibians (order *Caudata*) have XX/XY-system. For 63 species of 1500, sex was determined and only 20 species have differing sex chromosomes. Males of some New Zealand frogs (*Leiopelma hamiltoni* and *L. hochstetteri*) have heterogametic sex. In most amphibians, sex chromosomes are homomorphic (undifferentiated) in both sexes and are characterized by frequent turnover. This is in sharp contrast to sex chromosomes in two major vertebrate groups, the mammals and birds, where they are heteromorphic in one sex and are highly conserved. Thus, amphibians are excellent research materials on the turnover of sex sensitive to a resistant state, indicating the relationship between sex chromosome turnover and sex ratio control. Models of sex differentiation in amphibians can be divided into three types: (1) a direct development of the undifferentiated gonads into testes or ovaries, (2) the development of the undifferentiated gonad into the ovary and subsequent development of the testis through the ovary, and (3) the development of the testes through the intersex phase (prodifferentiating type) [17]. For a long time, genes that determine sex could not be found in amphibians. Recently, for smooth clawed frog (*Xenopus laevis*), the candidate gene has been found suitable for such a role. It is believed that it is involved in the development of the ovary. African clawed frog has a ZZ/ZW system of sex determination. Its *dm-w* gene was described. It is localized in the X chromosome and possessed a DM-domain. The nucleotide sequence of gene encoding a DNA-binding domain has 89% identity with *dmrt1*, but there is not similarity in transactivational region *dm-w* and *dmrt1*: genes are expressed exclusively in the primordial gonads, and *dm-w* is expressed more actively than in the gonads of ZW-larvae. The gene *dmrt1* (*dmrt1α* and *dmrt1β*) is located in autosome and there are no differences in its expression in males and females. The product of this gene enhances the expression of *cyp19* and *foxl2* ones. A similar gene was not detected in other species of amphibians. It is assumed that in these frogs, homo- and heterodimer products of *dmrt1* and

*DOI: http://dx.doi.org/10.5772/intechopen.83831*

and female phase, respectively (**Figure 3**).

**3. Sex in amphibians**

*Comparison of Sex Determination in Vertebrates (Nonmammals) DOI: http://dx.doi.org/10.5772/intechopen.83831*

factors regulating *cyp19a1a* expression are strong candidates for the trigger that initiates gonadal sex change; *cyp19a1a* promoter regions contain binding motifs for numerous factors that potentially regulate its expression [15].

However, hermaphroditic fishes have a plastic sex, and a stable sex is difficult to maintain with sex steroids. The black porgy regulated the dynamic development of both sexes; only one sex can grow while the other sex exists in a rudimentary stage (**Figure 2**). The sexual fate of the digonic gonad is determined by the male fate maintenance and through the Gnrh—Gth—Dmrt1 signaling. Altogether, testicular *dmrt1* and ovarian *cyp19a1a* expression are critical to the sexual fate of a male phase and female phase, respectively (**Figure 3**).

## **3. Sex in amphibians**

*Gene Expression and Phenotypic Traits*

on species is a complex trait under the control of one or many genetic factors in addition to environmental effects [9, 14]. In the Chinese tongue sole (*Cynoglossus semilaevis)*, genetic ZZ females may change into pseudomales, thereby increasing aquaculture costs because of the lower growth rate of the males than that of the females. A new locus was identified to regulate sex reversal interactively with the SNPCyn\_Z\_6676874; the linkage between these two loci and the absence of W sperm for pseudomales clearly elucidate the genetic architecture of sex reversal in the tongue sole [15]. Sexual determination in zebrafish is unique in that laboratory strains lack a sex chromosome, and no sex determining gene has been identified. GPER (estrogen receptor) is not required for normal sex differentiation, gonad development, or gonad function in zebrafish [16]. Genetic studies suggest that gonadal sexual fate is not only established by competition for primacy between two sexes via antagonistic signaling pathways during embryonic development but also requires active maintenance to suppress the opposite sex during adulthood. Documented in about 2% of teleost species spanning over 20 families, functional sex change generally occurs in three ways: protogynous (female-to-male), protandrous (male-to-female), and sequentially bi-directional. Most sequentially hermaphroditic fish are protogynous. Sex change in all hermaphroditic species involves radical gonadal transformation, and follows diverse ontogenetic pathways in different lineages particularly where sequential hermaphroditism has independently evolved. Gonadal transition in sex-changing fish is accompanied by changes in plasma concentrations of gonadal steroids. These steroids control gonad differentiation and maintain sexual phenotypes in teleost fish, wherein 17β-estradiol

(E2) and 11-ketotestosterone (11-KT) function as the major estrogen and androgen, respectively. The balance between estrogen and androgen production is expected to control sexual fate of the gonads during sex change. For example,

*The potential mechanism for sexual fate decision through the Gnrhs—Gths—Dmrt1 axis (brain-pituitarytestis axis). The model shows that the male fate decision is controlled by gonadotropins through the Gnrhs— Gths—Dmrt1 axis. The testis may stimulate the epigenetic modification of the ovary by DNA methylation of the cyp19a1a promoter to suppress the cyp19a1a expression. Gnrhs, gonadotropin-releasing hormones; Gths,* 

*gonadotropins; and Gthrs, gonadotropin receptors (adopted from Wu and Chang [16]).*

**26**

**Figure 3.**

Amphibians have two sex determined systems: XX/XY and ZZ/ZW. Most tailed amphibians (order *Caudata*) have XX/XY-system. For 63 species of 1500, sex was determined and only 20 species have differing sex chromosomes. Males of some New Zealand frogs (*Leiopelma hamiltoni* and *L. hochstetteri*) have heterogametic sex. In most amphibians, sex chromosomes are homomorphic (undifferentiated) in both sexes and are characterized by frequent turnover. This is in sharp contrast to sex chromosomes in two major vertebrate groups, the mammals and birds, where they are heteromorphic in one sex and are highly conserved. Thus, amphibians are excellent research materials on the turnover of sex sensitive to a resistant state, indicating the relationship between sex chromosome turnover and sex ratio control.

Models of sex differentiation in amphibians can be divided into three types: (1) a direct development of the undifferentiated gonads into testes or ovaries, (2) the development of the undifferentiated gonad into the ovary and subsequent development of the testis through the ovary, and (3) the development of the testes through the intersex phase (prodifferentiating type) [17]. For a long time, genes that determine sex could not be found in amphibians. Recently, for smooth clawed frog (*Xenopus laevis*), the candidate gene has been found suitable for such a role. It is believed that it is involved in the development of the ovary. African clawed frog has a ZZ/ZW system of sex determination. Its *dm-w* gene was described. It is localized in the X chromosome and possessed a DM-domain. The nucleotide sequence of gene encoding a DNA-binding domain has 89% identity with *dmrt1*, but there is not similarity in transactivational region *dm-w* and *dmrt1*: genes are expressed exclusively in the primordial gonads, and *dm-w* is expressed more actively than in the gonads of ZW-larvae. The gene *dmrt1* (*dmrt1α* and *dmrt1β*) is located in autosome and there are no differences in its expression in males and females. The product of this gene enhances the expression of *cyp19* and *foxl2* ones. A similar gene was not detected in other species of amphibians. It is assumed that in these frogs, homo- and heterodimer products of *dmrt1* and *dm-w* participate in the sex determination [18, 19] (**Figure 4**).

In the northern crested newt (*Triturus cristatus*), the proportion of males increases when the ambient temperature increases, and a decrease of temperature leads to an excess of females. Thus, in amphibians, an increase or decrease of the ambient temperature leads to a modification of the normal development of the gonads and sex determination. Here, sex-determining genes are not the decisive factor in determining sex. A number of experiments have shown that atrazine and some other pesticides that affect the endocrine system affect the formation of sex in frogs. As a result, males are changed to females. Exogenous steroids (introduced from the outside) are also changing the sex in amphibians [20]. The unexplainable mechanism of sex determination in the rice frog species was introduced. Amphibians bearing a novel sex determining mechanism are yet to be identified [17].

**Figure 4.**

*Model of ZZ/ZW-sex determined system and the formation of the ovary from Xenopus laevis (adopted from Liu et al. [18]).*

#### **Figure 5.**

*The role of steroid hormones in sex determination from Rana rugosa. At the stage of sex determination in the undifferentiated gonads of males, testosterone is synthesized at the same time females synthesize estradiol-17β. Letters ZZ, ZW indicate sex chromosomes. AR-T and ER-E2 represent complex androgen receptor (AR) to testosterone, and respectively, estrogen receptor and estradiol-17β (adopted from Nakamura [23]).*

The dominant hypothesis of sex determination for amphibians is proposed in relation to the *Rana rugosa.* In the Japanese wrinkled frog (Rana rugosa), four populations are described, in one of which (the northern population) females are heterogametic. Assume that sex determining genes really do not need to determine the sex of

**29**

*Comparison of Sex Determination in Vertebrates (Nonmammals)*

male sex-determinant in an amphibian species [22, 23].

**temperature**

called "window" of vulnerability [24].

amphibians [25] (**Figure 6**).

aromatase enzyme.

**4. Sex in reptiles: determination of sex under the influence of** 

Sex determination by environmental factors is mainly known in reptiles. The most well studied temperature sex determination (TSD) is occurring in three of the five main taxonomic groups of reptiles: turtles, crocodiles, and lizards, but it is not found in snakes. The adaptive significance of such sex determination mechanism is shown. During early embryonic development of gonad, epithelial cells are divided and unite in the epidermal strip of mesonephros mesenchyme. Further, during the so-called temperature-dependent period under the level of endogenous estrogen, such strip forms seminiferous tubules with Sertoli cell epithelium or gaps with squamous epithelium. The mechanism of this sex determination is poorly understood. Obviously, it is found in species with undifferentiated Y chromosome. The transition from the female promoting temperature (FPT) to male promoting temperature (MPT) is carried out in a temperature-period (TSP), during the so-

In some species of reptiles, GSD is not fixed for life, and the original gender may change during development without changing the genotype. This phenomenon is known as environmental sex reversal (ESR) and observed also in insects, fish, and

In reptiles, there is an "open" sex determination program that is different from a "closed" program, characteristic of birds and mammals. It is believed that in this case, the gender depends on the ratio of estrogens and androgens during sexual differentiation of the gonads. The temperature of incubation may change the activity of genes encoding aromatase, estrogen receptor, and reductase. It is not excluded that different taxonomic groups of animals with TSD have different mechanisms of regulation of sex. There may be temperature-sensitive genes *sox9* and *dax1* (freshwater turtles—*Emydidae*) and genes *sox9*, *sf1*, and *wt1* (*Testudinidae*). In mammals, this mechanism is not valid, because the Y chromosome has genes that inhibit the

For Mississippi alligator (*Alligator mississippiensis)*, pond slider turtle (*Trachemys scripta*), and olive ridley (*Lepidochelys olivacea*, from the family of sea turtles), the expression level of the gene *dmrt1* was higher during the incubation of embryos at a temperature that contributed to the emergence of males. In reptiles and in particular

amphibians, as well as the presence of the transcription factor, localized in the X or W chromosomes, influencing the feminization of vertebrates with TSD or GSD systems of sex determination. In males, if there is a specific mechanism of sex determination, it is likely that it supports the regulation of steroid hormones in undifferentiated gonads through the inhibition of *cyp19* gene transcription for the formation of the ovaries. In the scheme of **Figure 5**, a possible role of steroid hormones in sex determination is shown for *Rana rugosa* [21]. According to the next experimental data, complete female-to-male sex reversal occurred in the AR-Tg-transgenic ZW female frogs when a low dosage of T was supplied in the rearing water of tadpoles. In the sex reversed testes, the expression of *dmrt1*, *ar*, and *cyp17* genes required for masculinization was significantly upregulated. Next, AR-knockdown (KD) ZW female frogs were produced by the CRISPR/Cas9 system. Interestingly, no sex-reversal was observed in AR-KD ZW female frogs when the gonads were treated with dosages of T high enough to induce complete female-to-male sex-reversal, even in wild type frogs. In the AR-KD ZW female gonads, the expression of genes required for masculinization was not up-regulated. These results indicate that AR together with androgens can be a

*DOI: http://dx.doi.org/10.5772/intechopen.83831*

*Comparison of Sex Determination in Vertebrates (Nonmammals) DOI: http://dx.doi.org/10.5772/intechopen.83831*

*Gene Expression and Phenotypic Traits*

**28**

**Figure 5.**

**Figure 4.**

*Liu et al. [18]).*

*The role of steroid hormones in sex determination from Rana rugosa. At the stage of sex determination in the undifferentiated gonads of males, testosterone is synthesized at the same time females synthesize estradiol-17β. Letters ZZ, ZW indicate sex chromosomes. AR-T and ER-E2 represent complex androgen receptor (AR) to testosterone, and respectively, estrogen receptor and estradiol-17β (adopted from Nakamura [23]).*

*Model of ZZ/ZW-sex determined system and the formation of the ovary from Xenopus laevis (adopted from* 

The dominant hypothesis of sex determination for amphibians is proposed in relation to the *Rana rugosa.* In the Japanese wrinkled frog (Rana rugosa), four populations are described, in one of which (the northern population) females are heterogametic. Assume that sex determining genes really do not need to determine the sex of

amphibians, as well as the presence of the transcription factor, localized in the X or W chromosomes, influencing the feminization of vertebrates with TSD or GSD systems of sex determination. In males, if there is a specific mechanism of sex determination, it is likely that it supports the regulation of steroid hormones in undifferentiated gonads through the inhibition of *cyp19* gene transcription for the formation of the ovaries. In the scheme of **Figure 5**, a possible role of steroid hormones in sex determination is shown for *Rana rugosa* [21]. According to the next experimental data, complete female-to-male sex reversal occurred in the AR-Tg-transgenic ZW female frogs when a low dosage of T was supplied in the rearing water of tadpoles. In the sex reversed testes, the expression of *dmrt1*, *ar*, and *cyp17* genes required for masculinization was significantly upregulated. Next, AR-knockdown (KD) ZW female frogs were produced by the CRISPR/Cas9 system. Interestingly, no sex-reversal was observed in AR-KD ZW female frogs when the gonads were treated with dosages of T high enough to induce complete female-to-male sex-reversal, even in wild type frogs. In the AR-KD ZW female gonads, the expression of genes required for masculinization was not up-regulated. These results indicate that AR together with androgens can be a male sex-determinant in an amphibian species [22, 23].

## **4. Sex in reptiles: determination of sex under the influence of temperature**

Sex determination by environmental factors is mainly known in reptiles. The most well studied temperature sex determination (TSD) is occurring in three of the five main taxonomic groups of reptiles: turtles, crocodiles, and lizards, but it is not found in snakes. The adaptive significance of such sex determination mechanism is shown. During early embryonic development of gonad, epithelial cells are divided and unite in the epidermal strip of mesonephros mesenchyme. Further, during the so-called temperature-dependent period under the level of endogenous estrogen, such strip forms seminiferous tubules with Sertoli cell epithelium or gaps with squamous epithelium. The mechanism of this sex determination is poorly understood. Obviously, it is found in species with undifferentiated Y chromosome. The transition from the female promoting temperature (FPT) to male promoting temperature (MPT) is carried out in a temperature-period (TSP), during the socalled "window" of vulnerability [24].

In some species of reptiles, GSD is not fixed for life, and the original gender may change during development without changing the genotype. This phenomenon is known as environmental sex reversal (ESR) and observed also in insects, fish, and amphibians [25] (**Figure 6**).

In reptiles, there is an "open" sex determination program that is different from a "closed" program, characteristic of birds and mammals. It is believed that in this case, the gender depends on the ratio of estrogens and androgens during sexual differentiation of the gonads. The temperature of incubation may change the activity of genes encoding aromatase, estrogen receptor, and reductase. It is not excluded that different taxonomic groups of animals with TSD have different mechanisms of regulation of sex. There may be temperature-sensitive genes *sox9* and *dax1* (freshwater turtles—*Emydidae*) and genes *sox9*, *sf1*, and *wt1* (*Testudinidae*). In mammals, this mechanism is not valid, because the Y chromosome has genes that inhibit the aromatase enzyme.

For Mississippi alligator (*Alligator mississippiensis)*, pond slider turtle (*Trachemys scripta*), and olive ridley (*Lepidochelys olivacea*, from the family of sea turtles), the expression level of the gene *dmrt1* was higher during the incubation of embryos at a temperature that contributed to the emergence of males. In reptiles and in particular

### **Figure 6.**

*The continuum of sex determination. Distribution mechanisms from GSD to ESD, including intermediate system to overcome genetic sex determination with environmental factors (GSD + EE) (adopted from Valenzuela et al. [26]).*

*Trachemys scripta elegans*, a large amount of the KDM6B product is observed at a temperature favorable for males (MPT) and activates the expression of the *dmrt1* gene, and its reduction represses the expression of *dmrt1* and promotes the appearance of females. The latter is associated with H3K27 trimethylation. KDM6B is a member of the Jumonji gene family. It is believed that such genes are somehow regulated. One such regulator—*cirbp* (cold-inducible RNA binding protein)—has recently been described in the turtle *Chelydra serpentina* [27]. It managed to detect differences in the structure of *dmrt1*-gene in 34 species of reptiles with temperature and genetic mechanisms of sex determination, affecting sequence in exon 2 near DM-binding domain. In species with TSD, threonine occurs at position 54 (T54) and serine at position 57 (S57), while in species with a genetic sex determination mechanism, serine is observed in the S54-S57 position. This is obviously only the discovery of the molecular differences in sex determining gene when changing the mechanism of sex determination [28]. The discovery of the triploid male (ZZW) in the colubrid snake testifies to the absence of a particular role of the B chromosome in the determination of sex in this species [29].

Sex reversal has not yet been demonstrated in nature for any amniote, although it occurs in fish and rarely in amphibians. There is only one report about sex change in reptiles in the wild (Australian bearded dragon (*Pogona vitticeps*)) and the use of animals with inverse sex in order to experimentally induce a rapid transition from GSD to ESD. Controlled mating of normal males to sex-reversed females produces a viable and fertile offspring whose phenotypic sex is determined solely by temperature (temperature-dependent sex determination). The W sex chromosome is eliminated from this lineage in the first generation which indicates its specific role in genetic sex [30].

## **5. Sex determination in bird**

In birds, estrogens play an important role in sex determination. They regulate expression of key sex determining genes during the first 3 days of embryonic

**31**

**Figure 7.**

*Comparison of Sex Determination in Vertebrates (Nonmammals)*

development and further. At the same time, the set of sex chromosomes is equally important. Embryos with two Z chromosomes in birds develop as males, and those with ZW chromosomes develop as females. At present, two hypotheses on sex determination in birds compete. One of these hypotheses considers the number of Z chromosomes as a key sex determining factor, while the other hypothesis supposes the presence in W chromosome of the key gene controlling ovarian development or suppressing the appearance of testes. The presence in Z chromosome of a strong candidate gene for sex determination (DMRT1 gene) supports the dose scheme. **Figure 7** presents a hypothetical scheme of genetic control of primary sex differentiation in *Gallus gallus.* The gonad appears on the 3.5th day (stage 22) as thickening on the surface of mesonephros. It consists of the epithelial layer of somatic and germ cells and medullary cordate layer (epithelial cords), which is mixed with mesenchymal cells. On the 6.5th day (30th stage), the first sex determining genes are activated. In the modern scheme of the genetic control of sex determination in birds (practically within the dose scheme), an epigenetic mechanism for switching off the single allele of avian key sex determining *dmrt1* gene in females through hypermethylation and using noncoding MHM RNA came into sharp focus (**Figure 8**) [30–34]. Synthetic aromatase inhibitors (an enzyme catalyzing the synthesis of estrogens) can induce steady female → male sex inversion. In this case, the left gonad becomes an ovotestis, or a testis, and the right gonad becomes a testis. Injection of aromatase inhibitors *in ovo* in most experiments was carried out on the third or fourth day of incubation. At the same time, in experimental males, injection of estradiol results in reversible feminization of the gonads [35, 36]. Unfortunately, the genetic and

*Possible models of primary sex determination in birds by the example of Gallus gallus (adopted from Kuroiwa [33]).*

*DOI: http://dx.doi.org/10.5772/intechopen.83831*

*Comparison of Sex Determination in Vertebrates (Nonmammals) DOI: http://dx.doi.org/10.5772/intechopen.83831*

*Gene Expression and Phenotypic Traits*

*Trachemys scripta elegans*, a large amount of the KDM6B product is observed at a temperature favorable for males (MPT) and activates the expression of the *dmrt1* gene, and its reduction represses the expression of *dmrt1* and promotes the appearance of females. The latter is associated with H3K27 trimethylation. KDM6B is a member of the Jumonji gene family. It is believed that such genes are somehow regulated. One such regulator—*cirbp* (cold-inducible RNA binding protein)—has recently been described in the turtle *Chelydra serpentina* [27]. It managed to detect differences in the structure of *dmrt1*-gene in 34 species of reptiles with temperature and genetic mechanisms of sex determination, affecting sequence in exon 2 near DM-binding domain. In species with TSD, threonine occurs at position 54 (T54) and serine at position 57 (S57), while in species with a genetic sex determination mechanism, serine is observed in the S54-S57 position. This is obviously only the discovery of the molecular differences in sex determining gene when changing the mechanism of sex determination [28]. The discovery of the triploid male (ZZW) in the colubrid snake testifies to the absence of a particular role of the B chromosome in the determination of sex in this

*The continuum of sex determination. Distribution mechanisms from GSD to ESD, including intermediate system to overcome genetic sex determination with environmental factors (GSD + EE) (adopted from* 

Sex reversal has not yet been demonstrated in nature for any amniote, although it occurs in fish and rarely in amphibians. There is only one report about sex change in reptiles in the wild (Australian bearded dragon (*Pogona vitticeps*)) and the use of animals with inverse sex in order to experimentally induce a rapid transition from GSD to ESD. Controlled mating of normal males to sex-reversed females produces a viable and fertile offspring whose phenotypic sex is determined solely by temperature (temperature-dependent sex determination). The W sex chromosome is eliminated from this lineage in the first generation which indicates its specific role

In birds, estrogens play an important role in sex determination. They regulate expression of key sex determining genes during the first 3 days of embryonic

**30**

species [29].

**Figure 6.**

*Valenzuela et al. [26]).*

in genetic sex [30].

**5. Sex determination in bird**

development and further. At the same time, the set of sex chromosomes is equally important. Embryos with two Z chromosomes in birds develop as males, and those with ZW chromosomes develop as females. At present, two hypotheses on sex determination in birds compete. One of these hypotheses considers the number of Z chromosomes as a key sex determining factor, while the other hypothesis supposes the presence in W chromosome of the key gene controlling ovarian development or suppressing the appearance of testes. The presence in Z chromosome of a strong candidate gene for sex determination (DMRT1 gene) supports the dose scheme. **Figure 7** presents a hypothetical scheme of genetic control of primary sex differentiation in *Gallus gallus.* The gonad appears on the 3.5th day (stage 22) as thickening on the surface of mesonephros. It consists of the epithelial layer of somatic and germ cells and medullary cordate layer (epithelial cords), which is mixed with mesenchymal cells. On the 6.5th day (30th stage), the first sex determining genes are activated. In the modern scheme of the genetic control of sex determination in birds (practically within the dose scheme), an epigenetic mechanism for switching off the single allele of avian key sex determining *dmrt1* gene in females through hypermethylation and using noncoding MHM RNA came into sharp focus (**Figure 8**) [30–34]. Synthetic aromatase inhibitors (an enzyme catalyzing the synthesis of estrogens) can induce steady female → male sex inversion. In this case, the left gonad becomes an ovotestis, or a testis, and the right gonad becomes a testis. Injection of aromatase inhibitors *in ovo* in most experiments was carried out on the third or fourth day of incubation. At the same time, in experimental males, injection of estradiol results in reversible feminization of the gonads [35, 36]. Unfortunately, the genetic and

**Figure 7.**

*Possible models of primary sex determination in birds by the example of Gallus gallus (adopted from Kuroiwa [33]).*

#### **Figure 8.**

*The large Z chromosome (82.3 Mb) is drawn to scale next to the degenerate W chromosome (7 Mb). (A) Male (ZZ) have two copies of DMRT1 and HEMGN, while the female (ZW) only has one. The MHM locus is transcribed from the single Z in the female and may play a role in local dosage and epigenetic regulation of DMRT1 in the female. (B) Location and orientation of the 28 protein coding genes that are located on the W chromosome (adopted from Hirst et al. [34]).*

hormonal status of individuals with sex inversion was not investigated. The two enzymes required for the synthesis of estrogen, aromatase, and 17-beta-hydroxysteroid dehydrogenase (17β-HSD) are synthesized only in ovarian medullary cords at the onset of morphological differentiation. It is suggested that the earliest expression of aromatase in birds is detected only on the fifth day of embryonic development. It is worth mentioning that the appearance of aromatase was recently demonstrated as early as in the maternal body, upon oogenesis in the theca layer of early follicles [37]. The data obtained make it possible to suggest earlier appearance of aromatase and estrogens in female gonadogenesis than that follows from the classical scheme of primary sex determination in *Gallus gallus* [38].

In birds, sex determination depends on sex hormones and sex-hormone-specific receptors. Estrogen receptors are also important in this process. In a recent study, the gonads and endocrine profile of a gynadromorphic chicken were described. It had male features on the right and female features on the left. At sexual maturity, the gonads of this bird were largely testicular. The right gonad was a testis, with SOX9+ Sertoli cells, DMRT1+ germ cells, and active spermatogenesis. According to histology, the left gonad was primarily testicular, but with a few number of peripheral aromatase follicles. The gynandromorph had low levels of serum 17β-estradiol (39 pmol/L). In contrast, the gynandromorph had very elevated levels of serum testosterone (41.3 nmol/L). Despite the elevated testosterone, the bird was female on one side of the body. The right male side was almost entirely ZZ (96%), whereas those from the left female side were a mixture of male (77% ZZ) and female (23% ZW) cells. It had a low percentage of ZW cells on the female side, but still

**33**

*Comparison of Sex Determination in Vertebrates (Nonmammals)*

somes, GRS) found only in the germ cells of songbirds.

hormones is reduced to the differentiation of testis or ovaries.

term for nonmammal's sex change [43].

**Acknowledgements**

potential conflict of interest.

had female sex-linked feathering, smaller muscle mass, smaller leg and spur, and smaller wattle. This indicates that sexually dimorphic structures such as the wattle, spur, and feathering must be at least partly independent of sex steroid effects. Even a small percentage of ZW cells appear sufficient to support female-type sexual differentiation [39–41]. Studies of chimeric embryos also support the hypothesis that avian sexual differentiation is largely, or partly, cell autonomous, involving direct

So, estrogens and androgens play important roles in sexual differentiation and reproduction, particularly in the development and expression of male and female sexual characteristics. These effects are principally mediated by the estrogen and androgen receptors (ESRs and ARs), which belong to superfamily of the nuclear receptors [42]. The nature of the relationship between sex hormones and gender determining genes and the patterns of their interaction remains unclear. For some amphibians, the absence of appropriate genes and the replacement by control factors of steroid hormones and receptors are postulated. For birds, we can assume a special role of heteromorphism of sex chromosome and the presence of a specific interaction of the W and Z chromosomes. In this regard, we should mention the phenomenon of detection of specific chromosomes (germ line restricted chromo-

In mammals, aromatase is expressed later in embryonic development and the gonadal sex is formed independently of sex hormones and differentiation can occur in the absence of steroidogenesis. For mammals, two-step primary sex determination is typical. At the first stage, its determination is carried out by the *sry* gene. At the second one, sex hormones are synthesized in gonads and genetic endocrine regulation of sex development is maintained. It raises questions about the sensitivity to androgens and estrogens of sex determination in fish, amphibians, reptiles, and birds. The functional role of the emerging chromosome heteromorphism is not clear. It is believed that the realization of the phenomenon of sex reversal is different in nonmammal vertebrates and mammals. It is intended to introduce a special

So, determination of gonadal development in vertebrates like testis or ovary was initially controlled mainly by sex hormones (fish and amphibians). Later, various sex determining genes were involved in this process. The system was quite plastic and was able to respond to changes in external conditions (reptiles). The appearance of heteromorphic sex chromosomes (birds) has led to the emergence of some specific W chromosomal signal, which provides estrogen control of the development of a heterogametic sex. In mammals, the control of the primary determination of sex (the appearance of the gonad) becomes purely genetic, and the role of sex

This research was supported by a grant 17-04-01321A Russian Foundation for Basic Research (RFBR). The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be considered as a

*DOI: http://dx.doi.org/10.5772/intechopen.83831*

genetic factors and steroid hormones.

**6. Conclusion**

had female sex-linked feathering, smaller muscle mass, smaller leg and spur, and smaller wattle. This indicates that sexually dimorphic structures such as the wattle, spur, and feathering must be at least partly independent of sex steroid effects. Even a small percentage of ZW cells appear sufficient to support female-type sexual differentiation [39–41]. Studies of chimeric embryos also support the hypothesis that avian sexual differentiation is largely, or partly, cell autonomous, involving direct genetic factors and steroid hormones.

## **6. Conclusion**

*Gene Expression and Phenotypic Traits*

hormonal status of individuals with sex inversion was not investigated. The two enzymes required for the synthesis of estrogen, aromatase, and 17-beta-hydroxysteroid dehydrogenase (17β-HSD) are synthesized only in ovarian medullary cords at the onset of morphological differentiation. It is suggested that the earliest expression of aromatase in birds is detected only on the fifth day of embryonic development. It is worth mentioning that the appearance of aromatase was recently demonstrated as early as in the maternal body, upon oogenesis in the theca layer of early follicles [37]. The data obtained make it possible to suggest earlier appearance of aromatase and estrogens in female gonadogenesis than that follows from the

*The large Z chromosome (82.3 Mb) is drawn to scale next to the degenerate W chromosome (7 Mb). (A) Male (ZZ) have two copies of DMRT1 and HEMGN, while the female (ZW) only has one. The MHM locus is transcribed from the single Z in the female and may play a role in local dosage and epigenetic regulation of DMRT1 in the female. (B) Location and orientation of the 28 protein coding genes that are located on the W* 

In birds, sex determination depends on sex hormones and sex-hormone-specific receptors. Estrogen receptors are also important in this process. In a recent study, the gonads and endocrine profile of a gynadromorphic chicken were described. It had male features on the right and female features on the left. At sexual maturity, the gonads of this bird were largely testicular. The right gonad was a testis, with

histology, the left gonad was primarily testicular, but with a few number of peripheral aromatase follicles. The gynandromorph had low levels of serum 17β-estradiol (39 pmol/L). In contrast, the gynandromorph had very elevated levels of serum testosterone (41.3 nmol/L). Despite the elevated testosterone, the bird was female on one side of the body. The right male side was almost entirely ZZ (96%), whereas those from the left female side were a mixture of male (77% ZZ) and female (23% ZW) cells. It had a low percentage of ZW cells on the female side, but still

germ cells, and active spermatogenesis. According to

classical scheme of primary sex determination in *Gallus gallus* [38].

**32**

SOX9+

**Figure 8.**

Sertoli cells, DMRT1+

*chromosome (adopted from Hirst et al. [34]).*

So, estrogens and androgens play important roles in sexual differentiation and reproduction, particularly in the development and expression of male and female sexual characteristics. These effects are principally mediated by the estrogen and androgen receptors (ESRs and ARs), which belong to superfamily of the nuclear receptors [42]. The nature of the relationship between sex hormones and gender determining genes and the patterns of their interaction remains unclear. For some amphibians, the absence of appropriate genes and the replacement by control factors of steroid hormones and receptors are postulated. For birds, we can assume a special role of heteromorphism of sex chromosome and the presence of a specific interaction of the W and Z chromosomes. In this regard, we should mention the phenomenon of detection of specific chromosomes (germ line restricted chromosomes, GRS) found only in the germ cells of songbirds.

In mammals, aromatase is expressed later in embryonic development and the gonadal sex is formed independently of sex hormones and differentiation can occur in the absence of steroidogenesis. For mammals, two-step primary sex determination is typical. At the first stage, its determination is carried out by the *sry* gene. At the second one, sex hormones are synthesized in gonads and genetic endocrine regulation of sex development is maintained. It raises questions about the sensitivity to androgens and estrogens of sex determination in fish, amphibians, reptiles, and birds. The functional role of the emerging chromosome heteromorphism is not clear. It is believed that the realization of the phenomenon of sex reversal is different in nonmammal vertebrates and mammals. It is intended to introduce a special term for nonmammal's sex change [43].

So, determination of gonadal development in vertebrates like testis or ovary was initially controlled mainly by sex hormones (fish and amphibians). Later, various sex determining genes were involved in this process. The system was quite plastic and was able to respond to changes in external conditions (reptiles). The appearance of heteromorphic sex chromosomes (birds) has led to the emergence of some specific W chromosomal signal, which provides estrogen control of the development of a heterogametic sex. In mammals, the control of the primary determination of sex (the appearance of the gonad) becomes purely genetic, and the role of sex hormones is reduced to the differentiation of testis or ovaries.

## **Acknowledgements**

This research was supported by a grant 17-04-01321A Russian Foundation for Basic Research (RFBR). The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be considered as a potential conflict of interest.

*Gene Expression and Phenotypic Traits*

## **Author details**

Aleksandr F. Smirnov and Antonina V. Trukhina\* Department of Genetics and Biotechnology, Saint Petersburg State University, Saint Petersburg, Russia

\*Address all correspondence to: a.trukhina@spbu.ru

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

**35**

*Comparison of Sex Determination in Vertebrates (Nonmammals)*

[9] Martínez P, Viñas AM, Sánchez L, et al. Genetic architecture of sex determination in fish: Applications to sex ratio control in aquaculture. Frontiers in Genetics. 2014;**5**:340. DOI:

[10] Mei J, Gui JF. Genetic basis and biotechnological manipulation of sexual dimorphism and sex determination in fish. Science China. Life Sciences. 2015;**58**(2):124-136. DOI: 10.1007/

[11] Kikuchi K, Hamaguchi S. Novel sex-determining genes in fish and sex chromosome evolution. Developmental Dynamics. 2013;**242**(4):339-353. DOI:

[12] Kobayashi Y, Nagahama Y,

[13] Shao C, Li Q , Chen S, et al.

[15] Crowder CM, Romano SN, Gorelick DA. G protein-coupled estrogen receptor is not required for sex determination or ovary function in zebrafish. Endocrinology. 2018;**159**(10):3515-3523. DOI: 10.1210/

[16] Wu GC, Chang CF. Primary males guide the femaleness through the regulation of testicular *dmrt1* and ovarian *cyp19a1a* in protandrous

en.2018-00685

10.1101/gr.162172.113

Nakamura M. Diversity and plasticity of sex determination and differentiation in fishes. Sexual Development. 2013;**7** (1-3):115-125. DOI: 10.1159/000342009

Epigenetic modification and inheritance in sexual reversal of fish. Genome Research. 2014;**24**(4):604-615. DOI:

[14] Shen Z, Wang H. Molecular players involved in temperature-dependent sex determination and sex differentiation in Teleost fish. Genetics, Selection, Evolution. 2014;**46**(1):26. DOI: 10.1186/1297-9686-46-26

10.3389/fgene.2014.00340

s11427-014-4797-9

10.1002/dvdy.23927

*DOI: http://dx.doi.org/10.5772/intechopen.83831*

[1] Johnson Pokorná M, Kratochvíl L. What was the ancestral sex-determining mechanism in amniote vertebrates? Biological Reviews of the Cambridge Philosophical Society. 2016;**91**(1):1-12.

**References**

DOI: 10.1111/brv.12156

[3] Smirnov AF, Trukhina

978-1-61896-390-1

bbagrm.2014.05.020

10.1159/000342272

s11160-009-9123-4

[2] Schärer L. The varied ways of being male and female. Molecular Reproduction and Development.

2017;**84**:94-104. DOI: 10.1002/mrd.22775

[4] Murashima A, Kishigami S, Thomson A, et al. Androgens and mammalian male reproductive tract development. Biochimica et Biophysica Acta. 2015;**1849**(2):163-170. DOI: 10.1016/j.

[5] Morohashi K, Baba T, Tanaka M. Steroid hormones and the development

Development. 2013;**7**(1-3):61-79. DOI:

[6] Tagirov MT. Sex determination and control mechanisms in birds. Biotechnologia Acta. 2013;**6**(1):62-72. DOI: 10.15407/biotech6.01.062

[7] Guerrero-Estévez S, Moreno-Mendoza N. Sexual determination and differentiation in teleost fish. Reviews in Fish Biology and Fisheries. 2009;**20**(1):101-121. DOI: 10.1007/

[8] Hattori RS, Strüssmann CA, Fernandino JI, et al. Genotypic sex determination in teleosts: Insights from the testis-determining *amhy* gene. General and Comparative Endocrinology. 2013;**192**:55-59. DOI:

10.1016/j.ygcen.2013.03.019

of reproductive organs. Sexual

AV. Molecular-Genetic Mechanisms of Sex Determination in Animals. Scientific Research Publishing: An Academic Publisher; 2017. p. 118. ISBN: *Comparison of Sex Determination in Vertebrates (Nonmammals) DOI: http://dx.doi.org/10.5772/intechopen.83831*

## **References**

*Gene Expression and Phenotypic Traits*

**34**

**Author details**

Saint Petersburg, Russia

provided the original work is properly cited.

Aleksandr F. Smirnov and Antonina V. Trukhina\*

\*Address all correspondence to: a.trukhina@spbu.ru

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium,

Department of Genetics and Biotechnology, Saint Petersburg State University,

[1] Johnson Pokorná M, Kratochvíl L. What was the ancestral sex-determining mechanism in amniote vertebrates? Biological Reviews of the Cambridge Philosophical Society. 2016;**91**(1):1-12. DOI: 10.1111/brv.12156

[2] Schärer L. The varied ways of being male and female. Molecular Reproduction and Development. 2017;**84**:94-104. DOI: 10.1002/mrd.22775

[3] Smirnov AF, Trukhina AV. Molecular-Genetic Mechanisms of Sex Determination in Animals. Scientific Research Publishing: An Academic Publisher; 2017. p. 118. ISBN: 978-1-61896-390-1

[4] Murashima A, Kishigami S, Thomson A, et al. Androgens and mammalian male reproductive tract development. Biochimica et Biophysica Acta. 2015;**1849**(2):163-170. DOI: 10.1016/j. bbagrm.2014.05.020

[5] Morohashi K, Baba T, Tanaka M. Steroid hormones and the development of reproductive organs. Sexual Development. 2013;**7**(1-3):61-79. DOI: 10.1159/000342272

[6] Tagirov MT. Sex determination and control mechanisms in birds. Biotechnologia Acta. 2013;**6**(1):62-72. DOI: 10.15407/biotech6.01.062

[7] Guerrero-Estévez S, Moreno-Mendoza N. Sexual determination and differentiation in teleost fish. Reviews in Fish Biology and Fisheries. 2009;**20**(1):101-121. DOI: 10.1007/ s11160-009-9123-4

[8] Hattori RS, Strüssmann CA, Fernandino JI, et al. Genotypic sex determination in teleosts: Insights from the testis-determining *amhy* gene. General and Comparative Endocrinology. 2013;**192**:55-59. DOI: 10.1016/j.ygcen.2013.03.019

[9] Martínez P, Viñas AM, Sánchez L, et al. Genetic architecture of sex determination in fish: Applications to sex ratio control in aquaculture. Frontiers in Genetics. 2014;**5**:340. DOI: 10.3389/fgene.2014.00340

[10] Mei J, Gui JF. Genetic basis and biotechnological manipulation of sexual dimorphism and sex determination in fish. Science China. Life Sciences. 2015;**58**(2):124-136. DOI: 10.1007/ s11427-014-4797-9

[11] Kikuchi K, Hamaguchi S. Novel sex-determining genes in fish and sex chromosome evolution. Developmental Dynamics. 2013;**242**(4):339-353. DOI: 10.1002/dvdy.23927

[12] Kobayashi Y, Nagahama Y, Nakamura M. Diversity and plasticity of sex determination and differentiation in fishes. Sexual Development. 2013;**7** (1-3):115-125. DOI: 10.1159/000342009

[13] Shao C, Li Q , Chen S, et al. Epigenetic modification and inheritance in sexual reversal of fish. Genome Research. 2014;**24**(4):604-615. DOI: 10.1101/gr.162172.113

[14] Shen Z, Wang H. Molecular players involved in temperature-dependent sex determination and sex differentiation in Teleost fish. Genetics, Selection, Evolution. 2014;**46**(1):26. DOI: 10.1186/1297-9686-46-26

[15] Crowder CM, Romano SN, Gorelick DA. G protein-coupled estrogen receptor is not required for sex determination or ovary function in zebrafish. Endocrinology. 2018;**159**(10):3515-3523. DOI: 10.1210/ en.2018-00685

[16] Wu GC, Chang CF. Primary males guide the femaleness through the regulation of testicular *dmrt1* and ovarian *cyp19a1a* in protandrous black porgy. General and Comparative Endocrinology. 2018;**261**:198-202. DOI: 10.1016/j.ygcen.2017.01.033

[17] Cui Y, Wang W, Ma L, et al. New locus reveals the genetic architecture of sex reversal in the Chinese tongue sole (*Cynoglossus semilaevis*). Heredity. 2018;**121**(4):319-326. DOI: 10.1038/ s41437-018-0126-6

[18] Liu H, Todd EV, Lokman PM, et al. Sexual plasticity: A fishy tale. Molecular Reproduction and Development. 2017;**84**(2):171-194. DOI: 10.1002/ mrd.22691

[19] Miura I. Sex determination and sex chromosomes in amphibia. Sexual Development. 2017;**11**(5-6):298-306. DOI: 10.1159/000485270

[20] Yoshimoto S, Ito M. A ZZ/ZW-type sex determination in *Xenopus laevis*. FEBS Journal. 2011;**278**(7):1020-1026. DOI: 10.1111/j.1742-4658.2011.08031.x

[21] Piprek RP, Damulewicz M, Kloc M, Kubiak JZ. Transcriptome analysis identifies genes involved in sex determination and development of *Xenopus laevis* gonads. Differentiation. 2018;**100**:46-56. DOI: 10.1016/j. diff.2018.02.004

[22] Flament S. Sex reversal in amphibians. Sexual Development. 2016;**10**(5-6):267-278. DOI: 10.1159/000448797

[23] Nakamura M. Is a sex-determining gene(s) necessary for sex-determination in amphibians? Steroid hormones may be the key factor. Sexual Development. 2013;**7**(1-3):104-114. DOI: 10.1159/000339661

[24] Oike A, Kodama M, Yasumasu S, et al. Participation of androgen and its receptor in sex determination of an amphibian species. PLoS One. 2017;**12**(6):e0178067. DOI: 10.1371/ journal.pone.0178067

[25] Merchant-Larios H, Díaz-Hernández V. Environmental sex determination mechanisms in reptiles. Sexual Development. 2013;**7**(1-3): 95-103. DOI: 10.1159/000341936

[26] Valenzuela N, Badenhorst D, Montiel EE, Literman R. Molecular cytogenetic search for cryptic sex chromosomes in painted turtles *Chrysemys picta*. Cytogenetic and Genome Research. 2014;**144**(1):39-46. DOI: 10.1159/000366076

[27] Pieau C, Dorizzi M, Richard-Mercier N. Temperature-dependent sex determination and gonadal differentiation in reptiles. In: Scherer G, Schmid M, editors. Genes and Mechanisms in Vertebrate Sex Determination. Switzerland: Birkhäuser Verlag Basel; 2001. pp. 117-141

[28] Georges A, Holleley CE. How does temperature determine sex? Science. 2018;**360**(6389):601-602. DOI: 10.1126/ science.aat5993

[29] Janes DE, Organ CL, Stiglec R, et al. Molecular evolution of *dmrt1* accompanies change of sex-determining mechanisms in reptilia. Biology Letters. 2014;**10**(12):20140809. DOI: 10.1098/ rsbl.2014.0809

[30] Rovatsos M, Augstenová B, Altmanová M, et al. Triploid colubrid snake provides insight into the mechanism of sex determination in advanced snakes. Sexual Development. 2018;**12**:251-255. DOI: 10.1159/000490124

[31] Holleley CE, O'Meally D, Sarre SD, et al. Sex reversal triggers the rapid transition from genetic to temperature-dependent sex. Nature. 2015;**523**(7558):79-82. DOI: 10.1038/ nature14574

[32] Schmid M, Smith J, Burt DW, et al. Third report on chicken genes and chromosomes 2015. Cytogenetic and

**37**

*Comparison of Sex Determination in Vertebrates (Nonmammals)*

[40] Trukhina AV, Lukina NA, Smirnov AF. Hormonal sex inversion and some aspects of its genetic determination in chicken. Russian Journal of Genetics. 2018;**54**(9):1069-1077. DOI: 10.1134/

[41] Morris KR, Hirst CE, Major AT, et al. Gonadal and endocrine analysis of a gynandromorphic chicken.

DOI: 10.1210/en.2018-00553

Endocrinology. 2018;**159**(10):3492-3502.

[42] Ogino Y, Tohyama S, Kohno S, et al. Functional distinctions associated with the diversity of sex steroid hormone receptors ESR and AR. The Journal of Steroid Biochemistry and Molecular Biology. 2018;**184**:38-46. DOI: 10.1016/j.

[43] Weber C, Capel B. Sex reversal. Current Biology. 2018;**28**:R1234-R1236

S1022795418090144

jsbmb.2018.06.002

*DOI: http://dx.doi.org/10.5772/intechopen.83831*

Genome Research. 2015;**145**(2):78-179.

[33] Kuroiwa A. Sex-determining mechanism in avians. Advances in Experimental Medicine and Biology. 2017;**1001**:19-31. DOI: 10.1007/978-981-10-3975-1\_2

[34] Hirst CE, Major AT, Smith CA. Sex determination and gonadal sex differentiation in the chicken model. The International Journal of Developmental Biology. 2018;**62**(1-3):153-166. DOI:

[35] Caetano LC, Gennaro FG, Coelho K, et al. Differential expression of the MHM region and of sex-determining-

[36] Sánchez L, Chaouiya C. Logical modelling uncovers developmental

[37] Elbrecht A, Smith RG. Aromatase

[38] Vaillant S, Guémené D, Dorizzi M, et al. Degree of sex reversal as related to plasma steroid levels in genetic female chickens (*Gallus domesticus*) treated with Fadrozole. Molecular Reproduction and Development. 2003;**65**(4):420-428.

determination in chickens. Science. 1992;**255**(5043):467-470. DOI: 10.1126/

related genes during gonadal development in chicken embryos. Genetics and Molecular Research. 2014;**13**(1):838-849. DOI: 10.4238/2014.

constraints for primary sex determination of chicken gonads. Journal of The Royal Society Interface. 2018;**15**(142):20180165. DOI: 10.1098/

enzyme activity and sex

DOI: 10.1002/mrd.10318

[39] Wang J, Gong Y. Transcription of CYP19A1 is directly regulated by SF-1 in the theca cells of ovary follicles in chicken. General and Comparative Endocrinology. 2017;**247**:1-7. DOI: 10.1016/j.ygcen.2017.03.013

DOI: 10.1159/000430927

10.1387/ijdb.170319cs

February.13.2

rsif.2018.0165

science.1734525

*Comparison of Sex Determination in Vertebrates (Nonmammals) DOI: http://dx.doi.org/10.5772/intechopen.83831*

Genome Research. 2015;**145**(2):78-179. DOI: 10.1159/000430927

[33] Kuroiwa A. Sex-determining mechanism in avians. Advances in Experimental Medicine and Biology. 2017;**1001**:19-31. DOI: 10.1007/978-981-10-3975-1\_2

*Gene Expression and Phenotypic Traits*

10.1016/j.ygcen.2017.01.033

s41437-018-0126-6

mrd.22691

black porgy. General and Comparative Endocrinology. 2018;**261**:198-202. DOI: [25] Merchant-Larios H, Díaz-Hernández V. Environmental sex determination mechanisms in reptiles. Sexual Development. 2013;**7**(1-3): 95-103. DOI: 10.1159/000341936

[26] Valenzuela N, Badenhorst D, Montiel EE, Literman R. Molecular cytogenetic search for cryptic sex chromosomes in painted turtles *Chrysemys picta*. Cytogenetic and Genome Research. 2014;**144**(1):39-46.

[27] Pieau C, Dorizzi M, Richard-Mercier N. Temperature-dependent sex determination and gonadal differentiation in reptiles. In: Scherer G, Schmid M, editors. Genes and Mechanisms in Vertebrate Sex

Verlag Basel; 2001. pp. 117-141

science.aat5993

rsbl.2014.0809

10.1159/000490124

nature14574

Determination. Switzerland: Birkhäuser

[28] Georges A, Holleley CE. How does temperature determine sex? Science. 2018;**360**(6389):601-602. DOI: 10.1126/

[29] Janes DE, Organ CL, Stiglec R, et al. Molecular evolution of *dmrt1* accompanies change of sex-determining mechanisms in reptilia. Biology Letters. 2014;**10**(12):20140809. DOI: 10.1098/

[30] Rovatsos M, Augstenová B, Altmanová M, et al. Triploid colubrid snake provides insight into the mechanism of sex determination in advanced snakes. Sexual

Development. 2018;**12**:251-255. DOI:

[31] Holleley CE, O'Meally D, Sarre SD, et al. Sex reversal triggers the rapid transition from genetic to temperature-dependent sex. Nature. 2015;**523**(7558):79-82. DOI: 10.1038/

[32] Schmid M, Smith J, Burt DW, et al. Third report on chicken genes and chromosomes 2015. Cytogenetic and

DOI: 10.1159/000366076

[17] Cui Y, Wang W, Ma L, et al. New locus reveals the genetic architecture of sex reversal in the Chinese tongue sole (*Cynoglossus semilaevis*). Heredity. 2018;**121**(4):319-326. DOI: 10.1038/

[18] Liu H, Todd EV, Lokman PM, et al. Sexual plasticity: A fishy tale. Molecular Reproduction and Development. 2017;**84**(2):171-194. DOI: 10.1002/

[19] Miura I. Sex determination and sex chromosomes in amphibia. Sexual Development. 2017;**11**(5-6):298-306.

[20] Yoshimoto S, Ito M. A ZZ/ZW-type sex determination in *Xenopus laevis*. FEBS Journal. 2011;**278**(7):1020-1026. DOI: 10.1111/j.1742-4658.2011.08031.x

[21] Piprek RP, Damulewicz M, Kloc M, Kubiak JZ. Transcriptome analysis identifies genes involved in sex determination and development of *Xenopus laevis* gonads. Differentiation.

2018;**100**:46-56. DOI: 10.1016/j.

[22] Flament S. Sex reversal in amphibians. Sexual Development. 2016;**10**(5-6):267-278. DOI:

[23] Nakamura M. Is a sex-determining gene(s) necessary for sex-determination in amphibians? Steroid hormones may be the key factor. Sexual Development. 2013;**7**(1-3):104-114.

[24] Oike A, Kodama M, Yasumasu S, et al. Participation of androgen and its receptor in sex determination of an amphibian species. PLoS One. 2017;**12**(6):e0178067. DOI: 10.1371/

diff.2018.02.004

10.1159/000448797

DOI: 10.1159/000339661

journal.pone.0178067

DOI: 10.1159/000485270

**36**

[34] Hirst CE, Major AT, Smith CA. Sex determination and gonadal sex differentiation in the chicken model. The International Journal of Developmental Biology. 2018;**62**(1-3):153-166. DOI: 10.1387/ijdb.170319cs

[35] Caetano LC, Gennaro FG, Coelho K, et al. Differential expression of the MHM region and of sex-determiningrelated genes during gonadal development in chicken embryos. Genetics and Molecular Research. 2014;**13**(1):838-849. DOI: 10.4238/2014. February.13.2

[36] Sánchez L, Chaouiya C. Logical modelling uncovers developmental constraints for primary sex determination of chicken gonads. Journal of The Royal Society Interface. 2018;**15**(142):20180165. DOI: 10.1098/ rsif.2018.0165

[37] Elbrecht A, Smith RG. Aromatase enzyme activity and sex determination in chickens. Science. 1992;**255**(5043):467-470. DOI: 10.1126/ science.1734525

[38] Vaillant S, Guémené D, Dorizzi M, et al. Degree of sex reversal as related to plasma steroid levels in genetic female chickens (*Gallus domesticus*) treated with Fadrozole. Molecular Reproduction and Development. 2003;**65**(4):420-428. DOI: 10.1002/mrd.10318

[39] Wang J, Gong Y. Transcription of CYP19A1 is directly regulated by SF-1 in the theca cells of ovary follicles in chicken. General and Comparative Endocrinology. 2017;**247**:1-7. DOI: 10.1016/j.ygcen.2017.03.013

[40] Trukhina AV, Lukina NA, Smirnov AF. Hormonal sex inversion and some aspects of its genetic determination in chicken. Russian Journal of Genetics. 2018;**54**(9):1069-1077. DOI: 10.1134/ S1022795418090144

[41] Morris KR, Hirst CE, Major AT, et al. Gonadal and endocrine analysis of a gynandromorphic chicken. Endocrinology. 2018;**159**(10):3492-3502. DOI: 10.1210/en.2018-00553

[42] Ogino Y, Tohyama S, Kohno S, et al. Functional distinctions associated with the diversity of sex steroid hormone receptors ESR and AR. The Journal of Steroid Biochemistry and Molecular Biology. 2018;**184**:38-46. DOI: 10.1016/j. jsbmb.2018.06.002

[43] Weber C, Capel B. Sex reversal. Current Biology. 2018;**28**:R1234-R1236

**39**

**Chapter 4**

**Abstract**

domesticus

role in bird sex determination.

**1. Introduction**

genes, bird, Gallus *g*allus domesticus

*Aleksandr Fedorovich Smirnov* 

*and Antonina Vladimirovna Trukhina*

Specific Features of Sex

Determination in Birds on

the Example of *Gallus gallus*

The chapter is devoted to the consideration of sex determination in birds. The appearance of heteromorphic sex chromosomes (birds) has led to the emergence of some specific W chromosomal signal, which provides estrogen control of the development of a heterogametic sex. At present, two hypotheses about sex determination in birds compete. One of these hypotheses considers the number of Z chromosomes as a key sex-determining factor, while the other hypothesis supposes the presence in W chromosome of the key gene controlling ovarian development or suppressing the appearance of testes. Into the modern scheme of the genetic control of sex determination in birds (practically within the hypothesis of dose compensation), an epigenetic mechanism was added. The appearance of gonads in birds is most likely determined by sex hormones and to the greatest extent by estrogen under the control of W chromosome. It is desirable to pay attention to noncoding RNAs, their connection with the W chromosome and their

**Keywords:** sex determination, sex hormones, sex chromosomes, sex-determining

Sex is characterized by a set of features that ensure sexual reproduction. We distinguish the primary definition of sex—the emergence of one of two types of gonads (organs somatic of nature), their sexual differentiation into final system, and the development of two types germ cells. In different groups of vertebrates, different mechanisms of sex determination are realized. We consider hypothetical schemes of such a process in birds using the example of *Gallus gallus domesticus* [1]. On the one hand, chicken is an important model object of fundamental genetics, especially embryogenetics [2], and on the other hand, it has significant practical importance for humans: 210 million tons of meat and 1482 billion eggs per year [2, 3]. Both males and females are fattened in broiler production. There is currently no economically worthwhile use of the male of egg breeds. Therefore, the 1-day cockerels are destroyed, and this applies to 330 million chickens annually in

## **Chapter 4**
