Tissue Specific Gene Expression in Chelydra Serpentina

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Theses and Dissertations Theses, Dissertations, and Senior Projects

January 2019

Tissue Specific Gene Expression In Chelydra Serpentina

Jasmine Carly Kasanke

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Recommended Citation Kasanke, Jasmine Carly, "Tissue Specific Gene Expression In Chelydra Serpentina" (2019). Theses and Dissertations. 2856. https://commons.und.edu/theses/2856

This Thesis is brought to you for free and open access by the Theses, Dissertations, and Senior Projects at UND Scholarly Commons. It has been accepted for inclusion in Theses and Dissertations by an authorized administrator of UND Scholarly Commons. For more information, please contact [email protected]. TISSUE SPECIFIC GENE EXPRESSION IN CHELYDRA SERPENTINA

Jasmine Carly Kasanke Bachelor of Biology, University of North Dakota, 2016

A Thesis

Submitted to the Graduate Faculty of the University of North Dakota in partial fulfillment of the requirements

for the degree of Master of Science

Grand Forks, North Dakota

December 2019

PERMISSION

Title Tissue Specific Gene Expression in Chelydra serpentina

Department Biology

Degree Master of Science

In presenting this thesis in partial fulfillment of the requirements for a graduate degree from the University of North Dakota, I agree that the library of this University shall make it freely available for inspection. I further agree that permission for extensive copying for scholarly purposes may be granted by the professor who supervised my thesis work or, in his absence, by the Chairperson of the department or the dean of the School of Graduate Studies. It is understood that any copying or publication or other use of this thesis or part thereof for financial gain shall not be allowed without my written permission. It is also understood that due recognition shall be given to me and to the University of North Dakota in any scholarly use which may be made of any material in my thesis.

Jasmine Carly Kasanke 28 October 2019

TABLE OF CONTENTS LIST OF FIGURES ...... vii LIST OF TABLES ...... viii ACKNOWLEDGEMENTS ...... ix ABSTRACT ...... x CHAPTER 1 ...... 1 TISSUE SPECIFIC GENE EXPRESSION ...... 1 Abstract ...... 1 Embryonic Development ...... 2 Gonad Development ...... 3 Sex Determination ...... 5 Tissue Specific Gene Expression ...... 7 Anti-Müllerian hormone ...... 8 Dmrt1 ...... 10 Sox9 ...... 12 FoxL2 ...... 15 Somatostatin ...... 17 Growth Hormone ...... 18 Cdx1 ...... 19 Slc12a1 ...... 20 Gene Regulation...... 21 DNA Methylation ...... 22 Histone Modifications ...... 22 Conclusions ...... 23 CHAPTER 2 ...... 24 TISSUE SPECIFIC EXPRESSION IN CHELYDRA SERPENTINA .....24 Abstract ...... 24 Introduction ...... 25 Materials and Methods ...... 27

Acquisition of Tissues...... 27 RNA Extractions ...... 28 Reverse Transcription ...... 29 Quantitative PCR ...... 30 Statistical Analysis ...... 31 Results ...... 32 Discussion ...... 40 Future Work ...... 51 REFERENCES ...... 53

LIST OF FIGURES

Figure Page

Plot of Means for Tissue Specific Expression of Amh ...... 33

Plot of Means for Tissue Specific Expression of Dmrt1 ...... 34

Plot of Means for Tissue Specific Expression of Sox9...... 35

Plot of Means for Tissue Specific Expression of FoxL2 ...... 36

Plot of Means for Tissue Specific Expression of Sst ...... 37

Plot of Means for Tissue Specific Expression of Gh1 ...... 38

Plot of Means for Tissue Specific Expression of Cdx1 ...... 39

Plot of Means for Tissue Specific Expression of Slc12a1 ...... 40

vii

LIST OF TABLES

Table Page

Primer Sequences for qPCR ...... 30

ANCOVA Results for Tissue Specific Expression ...... 32

Summary of qPCR Results for tissue specific expression in Chelydra serpentina in comparison to mammals...... 52

viii

ACKNOWLEDGEMENTS

I wish to express my sincere appreciation to my graduate advisor Turk Rhen for giving me the opportunity to work in his lab, and for his guidance and support. I would also like to thank my graduate advisory committee, Dr. Tristan Darland and Dr. Peter

Meberg, for their guidance, support and encouragement during my time in the master’s program at the University of North Dakota. Thank you to the members of Dr. Rhen’s lab for your assistance and contribution to this work. I would especially like to thank Sunil

Singh for his tutelage and guidance, Debojyoti Das for his help with R, and finally Jacob

Bierstedt for his help with the lab work. I also wish to thank the faculty members and staff of the graduate school and Biology Department of the University of North Dakota, and finally the NSF for funding this research.

ABSTRACT

Epigenetic regulation has been studied in mammals, but research in reptiles is lacking. Knowing where specific genes are expressed is the first step to examining epigenetic regulation of tissue specific and cell-type specific gene expression.

Differences in gene expression allude to differences in gene regulation and gene function and will require further study. In order to address the question of tissue specific gene expression, we review literature on embryonic development, sex determination, and specific genes that display tissue specific or cell-type specific expression patterns. Next, we review what is known about regulation of gene expression in reptiles. We then outline the protocol used to examine expression patterns of Anti-Müllerian hormone, Dmrt1,

Sox9, FoxL2, Somatostatin, Growth hormone, Cdx1, and Slc12a1 in the common snapping turtle. Finally, we compare these expression patterns to those found in mammals and discuss any observed differences.

CHAPTER 1

TISSUE SPECIFIC GENE EXPRESSION

Abstract

Each vertebrate begins as a single cell. This cell contains all of the information necessary to direct growth and development into a mature adult. Even though the genetic code is identical in all cells (excluding gametes and B cells), the organism has a variety of tissues and cell types that perform different functions. Differences among cell types comes from differential expression of specific genes. These genes are regulated by epigenetic modifications. Epigenetic regulation has been studied in mammals, but research in reptiles is lacking. Of particular interest is the study of reptiles with regard to sex determination. The overarching goal of this research is to study the genes involved in temperature dependent sex determination, and how they are regulated. Therefore, many of the genes and tissues chosen for this research, aside from the controls, have some relation to sex determination. Knowing where specific genes are expressed is the first step to examining epigenetic regulation of tissue specific and cell type specific gene expression. This chapter begins with a brief explanation of embryonic development, followed by bipotential gonad development, and male and female reproductive tract differentiation. Next is a review of information on several genes that exhibit tissue specific expression. This is followed by a discussion of what is known about DNA methylation and histone modifications in reptiles.

Embryonic Development

Every vertebrate starts as a single cell. Soon after fertilization, the single celled zygote begins to divide and cells multiply, forming the blastodisc. In egg laying species like birds and turtles, blastodisc development occurs inside of an amniotic egg. Extensive research has been performed on chicken development since the 1800s, and thus it is the model that is described. Bird and reptilian cells multiply by discoidal meroblastic cleavage that forms the blastoderm. Eventually, a subgerminal cavity is formed when fluid is secreted between these cells and the yolk. Cell division continues to lead to development of the marginal zone, in which some cells are critical for formation of the primitive streak. The primary and secondary hypoblasts develop and contain cells that provide signals to aid in directing the migration of epiblast cells. After this, the primitive streak is formed. Epiblast cells later develop via the process of gastrulation to form the three germ layers from which all parts of the organism develop: the endoderm, ectoderm, and mesoderm.

Also coordinated with the formation and development of the primitive streak is the process of neurulation in which the notochord forms from mesoderm and the neural plate folds to form the neural tube. The neural tube later develops into the brain and spinal cord. Cells then move to cover the neural tube and will later become the epidermis.

Neurulation is the first step in the process of organogenesis, the formation of the organs.

The formation of the notochord leads to changes in nearby cells that cause the mesoderm to form somites. Somites form as discrete divisions of the mesoderm of an embryo and later become the spine, muscle, and dermis. A split along the somite axis forms the coelom. The three germ layers form various organs by folding, splitting, and condensing.

The endoderm or inner layer develops into the lining of the gastrointestinal tract, lungs, liver, pancreas and other glands. The ectoderm, or outer layer develops into the skin, and the central and peripheral nervous systems. The mesoderm or middle layer develops into the dermis, heart, muscular system, urogenital system, and skeletal system. The mesoderm layer also gives rise to the gonads (Gilbert, 2014).

Gonad Development

Gonad development provides a specific example of the process and mechanisms involved in organogenesis. There are three steps to the formation of the gonads. The first step is development of the bipotential gonads, also known as gonadogenesis (Swain &

Lovell-Badge, 1999; McLaren, 2000). In the second step, the fate of the bipotential organs is determined and there is irreversible commitment to develop into testes or ovaries, the male or female reproductive organs, depending on the activation and silencing of certain genes (Lillie, 1917; Fisher & Merkenschlager, 2002; Loo & Rine,

1995). In the third step, gonadal cells differentiate and reorganize to produce morphologically distinct testes and ovaries.

Initial formation of the bipotential gonad begins with proliferation of coelomic epithelial cells on the ventromedial surface of the mesonephros (Ginsburg, Snow, &

McLaren, 1990). The paired mesonephroi are each comprised of three regions: the pronephros, mesonephros, and metanephroi, from anterior to posterior. The pronephros give rise to adrenal tissue, the mesonephros is the area in which the gonad forms, and the metanephros develops into the kidney (Wilhelm, Palmer, & Koopman, 2007).

Primordial germ cells migrate from the yolk sac along the gut tube to the site of gonad formation, the mesonephros, at about 10.5 days post coitum (dpc) in mice (Swain

& Lovell-Badge, 1999; McLaren, 2000), which is when the gonads emerge as a ridge of tissue on the mesonephros (Karl, et al., 1995). Migrating primordial germ cells (PGC’s) contribute to formation of the gonads, as these cells are undifferentiated precursors to the gametes (Molyneaux & Wylie, 2004). PGC’s are first found at the allantois, the fetal membrane below the chorion in vertebrates (Ginsburg & Snow, 1990). They migrate along the hindgut into the developing urogenital ridge. During migration, the PGC’s are also proliferating (Bendel-Stenzel, Anderson, Heasman, & Wylie, 1998).

The second step in gonad development is sex determination when cell fate and gonad fate is determined: i.e., an irreversible developmental decision is made for the gonads to become testes or ovaries (Swain & Lovell-Badge, 1999; McLaren, Germ and

Somatic Cell Lineages in the Developing Gonad, 2000). The bipotential gonad consists of an outer cortex surrounding an inner medulla. If the cells in the medulla begin to divide, differentiate, and reorganize to form testicular cords, the bipotential gonad morphs into a testis. However, if cells in the cortex proliferate, differentiate, and reorganize to form ovarian follicles, the bipotential gonad forms an ovary (Nef & Vasalli, 2009; McLaren,

1998). There is also movement of cells between these compartments and migration of cells from underlying mesonephros into the gonads.

The third step involves further differentiation of testicular and ovarian cell types and completion of morphogenesis. This whole cascade of events begins with determination and differentiation of supporting cell precursors into male or female specific cell types: Sertoli cells in testes or granulosa cells in ovaries (Kim & Capel,

2006). Sertoli and granulosa cells support and provide nutrients to germ cells to help sustain them and guide their development into mature sperm and eggs, respectively

(Merchant-Larios, Moreno-Mendoza, & Buehr, 1993). Sertoli cells are also thought to induce the differentiation of other cell lineages during testis development (Kim & Capel,

2006). Leydig cells in the testis and theca cells in the ovary both secrete steroids and are thought to derive from the same precursor cells (Wilhelm, Palmer, & Koopman, 2007).

Evidence from previous studies suggests that cell fate in the gonad is dependent upon the local environment of the progenitor cells (Racine, et al., 1998; Habert, Lejuene, & Saez,

2001; Yao, Whoriskey, & Capel, 2002; Kim & Capel, 2006).

Sex Determination

In mammals, testis differentiation is caused by expression of sex-determining region Y protein (Sry), which is a DNA-binding transcription factor that induces somatic cells to become Sertoli cells (Wilhelm, Palmer, & Koopman, 2007). The Sry gene is not present in reptiles, and it is unknown whether a gene with similar function exists, or if multiple genes have a similar overall effect (Griffiths, 1991; Coriat, Muller, Harry,

Uwanogho, & Sharpe, 1993; Foster & Graves, 1994). Studies generally agree, however, that gonad development in reptiles and mammals employs similar molecular pathways, except for the Sry gene (Zarkower, 2001; Raymond, et al., 1998; Marin & Baker, 1998).

Although many of the genes involved in testis and ovary development appear to be evolutionarily conserved, the relative timing of developmental events may differ.

The mechanism that triggers the bipotential gonads to become testes or ovaries also varies among vertebrates. There are five categories of sex determination found in animals, XY/XX sex chromosome system, haplodiploidy, a ZW/ZZ sex chromosome

5 system, a polygenic system, and environmental sex determination (Graves & Shetty,

2001). In placental mammals, for instance, sex is determined by the X and Y chromosomes. If the zygote has an XY karyotype and the Sry gene, it will develop testes and become a male. Haplodiploidy is a system in which females have two X chromosomes and males have one X chromosome, or XO. Fertilized eggs become females, whereas unfertilized eggs develop as males. This form of sex determination is often found among insects (Alpedrinha, Gardner, & West, 2014). Birds, snakes, and some turtles, lizards and fish have evolved a ZZ/ZW system of sex determination. In this mode of sex determination, homozygotes for the Z chromosome develop testes and become male, while heterozygotes for the Z and W chromosomes develop ovaries and become female. A gene mapping study has revealed that XY and ZW sex chromosomes in amniotic vertebrates share no homology, and thus evolved from different pairs of autosomal chromosomes (Matsubara, et al., 2006). Finally, it has long been known that turtles, and many other reptiles, have a form of environmental sex determination called temperature-dependent sex determination (TSD).

TSD is unique in that the environment dictates what sex the embryo will become.

There are at least two species of lizard that have genotypic sex determination (GSD), but genetic sex can be overridden by temperature during development (Quinn, et al., 2007;

Radder, Quinn, Georges, Sarre, & Shine, 2008; Ezaz, Sarre, O'Meally, Graves, &

Georges, 2010). Evolutionary transitions between TSD and GSD have occurred more than once in reptiles (Prkana & Kratochvil, 2009). There are three distinct versions of

TSD. In one type, males develop at intermediate temperatures while females develop at warmer and cooler temperatures (FMF pattern), which is the pattern exhibited in our

6 focal population of snapping turtles. In another type, females develop at cooler temperatures and males develop at warmer temperatures (FM pattern). Finally, in the third type, males develop at cooler temperatures and females at warmer temperatures

(MF pattern). Sex is determined during a temperature sensitive period (TSP) that occurs when the bipotential gonads are forming (Shen & Wang, 2014; Rhen, Fagerlie,

Schroecer, Crossley II, & Lang, 2015). Once gonadal fate is determined, the differentiation of the gonads into testes or ovaries begins.

Tissue Specific Gene Expression

Development and differentiation of the gonads and other organs occurs through cell-type specific and tissue specific expression of certain genes. These genes can be divided into two functional categories. One category of genes determines cell fate while the second category includes downstream effector genes that give cells their unique phenotype (Barsi, Tu, & Davidson, 2014). Here we review the function of several genes that play a role in cell fate determination and/or cellular differentiation. Sry-related high mobility group 9 (Sox9), Anti-Müllerian hormone (Amh), doublesex and mab-3 related transcription factor 1 (Dmrt1), and pituitary Forkhead factor (FoxL2), are expressed in the male and female gonads. The gonads, together with the hypothalamus and pituitary gland form the hypothalamic-pituitary-gonadal (HPG) axis. The hypothalamus is known to express the Somatostatin (Sst) gene while the pituitary expresses the growth hormone

(Gh1) gene (Spiess, Villarreal, & Vale, 1981; Gahete, et al., 2009). These tissues and their hormones are essential for reproduction and somatic growth in vertebrates. The next tissues chosen were kidney and intestine samples because of their proximity to the gonads. The embryonic kidneys share precursor cells with the developing gonads

(Wilhelm, Palmer, & Koopman, 2007) and are known to express Slc12a1 (Greger &

Schlatter, 1981). The developing intestine also interacts with cells that later reside in the gonads. Primordial germ cells migrate along the gut tube to the developing gonads

(Swain & Lovell-Badge, 1999; McLaren, 2000). Cdx1 is expressed in the intestine

(Silberg, Swain, Suh, & Traber, 2000). Below we discuss the molecular function of these genes and their role in development and differentiation of the gonads, hypothalamus, pituitary, kidney and intestine. We begin with evolutionarily conserved genes that play important roles in testis formation including: Amh, Dmrt1, and Sox9 (Munsterberg &

Lovell-Badge, 1991; Hanley, et al., 2000).

Anti-Müllerian hormone

Anti-Müllerian hormone (Amh), also known as Müllerian inhibiting substance

(Mis), is a member of the transforming growth factor-β (Tgf-β) family of growth and differentiation factors (Mullen et al., 2019). Amh is thought to be activated by the transcription factors Sox9 and Sf1 in mammals (Santa Barbara, et al., 1998; Arango &

Lovell-Badge, 1999). This gene is critical for development along the male pathway as it is expressed in Sertoli cells (Behringer, Finegold, & Cate, 1994). The source of Amh production in male gonads was determined by radiation treatment to kill germ cells, which are more sensitive to radiation than other cell types in the gonads (Liu G. et al.,

2006; Liu G. et al., 2007). Because Amh expression did not decrease with germ cell mortality, the authors concluded that Amh is likely produced by the neighboring Sertoli cells. This finding was later supported with the cloning of cDNA and in situ hybridization to directly detect Amh mRNA expression in Sertoli cells (Josso, 1974; Munsterberg &

Lovell-Badge, 1991). Although embryonic ovaries produce very little or no Amh,

8 significant expression of Amh in adult female gonads is found in granulosa cells of the

Graafian follicle. Primary follicles, those that were growing, and the corpus luteum were found to lack Amh expression, as did the Theca cells (Takahashi, Hayashi, Manganaro, &

Donahoe, 1986). The finding that Sertoli cells and Granulosa cells are the sources of

Amh in both sexes further supports the hypothesis that these cell types share a common origin during embryonic development. (Albrecht & Eicher, 2001).

Amh is involved in Müllerian duct regression during male testis development

(MacLaughlin, et al., 1992) and is an early indicator of testis determination (Takada,

DiNapoli, Capel, & Koopman, 2004). Unlike Sox9, it has greater variation among species at the amino acid level (Carre Eusebe, et al., 1996; Neeper, et al., 1996; Western, Harry,

Marshall Graves, & Sinclair, 1999; Miura, Miura, Konda, & Yamauchi, 2002). Perhaps this is because the function of Amh appears to have diversified during evolution. Data suggests that the original function of Amh is regulation of germ cell proliferation (Skaar, et al., 2011; Nobraga, et al., 2015; Safian, Bogerd, & Schulz, 2018; Miura, Miura, Konda,

& Yamauchi, 2002). This is further supported by a study in mice with a mutant, nonfunctional version of Amh. Female mice homozygous for this mutation had earlier depletion of primordial follicles when compared to wild type females (Durlinger et al.,

1999). In humans, females with mutations in Amh or its receptor display premature ovarian insufficiency (Mercadal et al., 2015). In addition to a male reproductive system, male mice homozygous for the same Amh mutation developed uteri, oviducts, and vaginal tissue. These tissues are derived from the Müllerian ducts (Mullen & Behringer,

2014). Males had testes that were morphologically normal, but only 13 percent were able to produce offspring (Mullen, et al., 2019, in press). The absence of functional Amh or its

9 receptor AmhR2 in humans results in Persistent Müllerian Duct Syndrome, a rare disorder in which the female reproductive tract is found in males (Josso, et al., 1993).

Dmrt1

Another gene that is critical for male development is doublesex and mab-3 related transcription factor 1 (Dmrt1). Unlike Amh, this gene is not known to play a role in females. Dmrt1 is involved in sex determination in fish, mammals, and reptiles (Matson

& Zarkower, 2012). Dmrt1 expression is higher in the developing gonads of males compared to females (Wilhelm, Palmer, & Koopman, 2007; De Grandi, et al., 2000;

Kettlewell, Raymond, & Zarkower, 2000; Moniot, Berta, Scherer, Sudbeck, & Poulat,

2000; Raymond, Kettlewell, Hirsch, Bardwell, & Zarkower, 1999; Smith, McClive,

Western, Reed, & Sinclair, 1999). There are other Dmrt proteins that may also play a role in sex determination, including Dmrt3, Dmrt5, and Dmrt7. These genes are of potential interest because of their spatial and temporal pattern of expression is similar to Dmrt1

(Matsuda, et al., 2002; Nanda, et al., 2002; Kim, Kettlewell, Anderson, Bardwell, &

Zarkower, 2003).

Sun et al. (2017) performed studies with the turtle Pelodiscus sinensis, finding that Dmrt1 expression was induced in ZW females treated with an aromatase inhibitor, leading to masculinized females. RNA interference knockdown of Dmrt1 induced male to female sex reversal. Downregulation of Amh and Sox9, and upregulation of Cyp19a1 and

FoxL2 was also observed. This data indicates that Dmrt1 likely acts as an upstream initiator of male development and is needed for proper development of male gonads (Sun et al., 2017). The red-eared slider turtle and snapping turtle exhibit upregulation of Dmrt1 during the sex determining period before gonad differentiation (Shoemaker, Queen, &

Crews, 2007; Kettlewell, Raymond, & Zarkower, 2000; Rhen T. , Metzger, Shroeder, &

Woodward, 2007). Expression of Dmrt1 was seen during the period of differentiation in pre-Sertoli cell nuclei (Sun, et al., 2017).

Webster et al. (2017) examined zebrafish with Dmrt1 mutations. Most of the mutant fish developed as females despite their male genotype. Embryos that developed as males were unable to reproduce due to testicular dysgenesis. These males displayed abnormal gonadal development including incomplete apoptosis of oocytes, intersex gonads, and no spermatogenesis. Dmrt1 is needed for regulation of Amh and FoxL2.

Female zebrafish with mutated Dmrt1 developed normally (Webster, et al., 2017).

Dmrt1 is also known to show sexually dimorphic expression in mammals

(Kettlewell, Raymond, & Zarkower, 2000). Dmrt1 contains a DM domain, which is a

DNA-binding motif containing a zinc finger (Erdman & Burtis, 1993; Raymond, et al.,

1998). There is some evidence to suggest that Dmrt1 is related to the dsx gene in

Drosophilia, and the mab-3 gene in Caenorhabditis elegans (Baker & Ridge, 1980; Shen

& Hodgkin, 1988; Raymond, et al., 1998). Interestingly, when the cloned male isoform of dsx was injected in C. elegans in vivo, it was able to function in the place of knocked out mab-3, but the female-specific isoform did not function in place of the mab-3 protein

(Raymond, et al., 1998). Further research is needed to determine if these genes are in fact related, or simply have similar functions through convergent evolution.

Dmrt1, shows a high degree of similarity between vertebrate species. In the pond slider turtle, it has 94% identity to alligator, and 78% identity to chicken sequences and its expression was found in pre-Sertoli and germ cells (Kettlewell, Raymond, &

Zarkower, 2000). Mutations in the short arm of chromosome 9 have been linked to XY

11 sex reversal in humans (Bennett, et al., 1993; Veitia, et al., 1998; Flejter, Fergestad,

Gorski, Varvill, & Chandrasekharppa, 1998). It is now known that this chromosome contains Dmrt1 and Dmrt2 genes, and thus explains sex reversal. Mutations in one or both copies of Dmrt1 affect male sexual development in humans (Raymond, et al., 1999).

Interestingly, Dmrt1 is found on the Z chromosome in birds (Nanda, et al., 2000) and it is thought that two doses of this gene in ZZ individuals are required for proper male development (Smith, et al., 2009).

Sox9

Vertebrates have an autosomal, evolutionarily conserved gene called Sry-related high mobility group 9 (Sox9) (Koopman 1999). The Sox9 protein is first expressed in the bipotential gonad at low levels. Sry, a transcription factor, upregulates Sox9 expression in pre-Sertoli cells in males (Wilhelm, Palmer, & Koopman, 2007). Sox9 contains a high mobility group DNA-binding domain and two transactivation domains (Harley, Clarkson,

& Argentaro, 2003; Sudbeck, Schmitz, Baeuerle, & Scherer, 1996). However, non- mammalian vertebrates have no Sry gene so some other transcription factor must regulate

Sox9 expression in these groups (Griffiths, 1991; Coriat, Muller, Harry, Uwanogho, &

Sharpe, 1993; Foster & Graves, 1994). The fact that Sox9 is found in mammals, birds, and reptiles and is expressed at a higher level in developing testes than in ovaries leads scientists to believe that it is an important testis-determining factor that is paramount in all vertebrates (Shoemaker, Ramsey, Queen, & Crews, 2007; Kent, Wheatley, Andrews,

Sinclair, & Koopman, 1996; Silva, et al., 1996). In mammals, Sry expression in the developing male gonad initiates a cascade of events that leads to phosphorylation of

Sox9, which is followed by shuttling of Sox9 protein into the nucleus (Koopman,

Bullejos, & Bowles, 2001). This gene is necessary for differentiation and maintenance of

Sertoli cells and thus the development of male phenotype. Contrary to its expression in males, Sox9 is expressed at a lower level in females and does not translocate to the nucleus. In males, Sox9, regulates expression of Amh and Vanin-1. These genes are also required for proper development of male phenotype (Arango & Lovell-Badge, 1999;

Santa Barbara, et al., 1998; Wilson, Jeyasuria, Parker, & Koopman, 2005).

As described above, Sox9 and Sry share the HMG box domain (Jager, Anvret,

Hall, & Scherer, 1990; Koopman, Munsterberg, Capel, Vivian, & Lovell-Badge, 1990;

Berta, et al., 1990; Denny, et al., 1992; Wright, Snopek, & Koopman, 1993). Like its paralog, Sox9 is a transcription factor (Santa Barbara, et al., 1998; Bell, et al., 1997;

Akiyama, 2008). It is expressed in developing kidney, testes, the central nervous system, and tissue that later forms the skeleton (Wright, et al., 1995; Kent, Wheatley, Andrews,

Sinclair, & Koopman, 1996).

From its expression pattern it is apparent that Sox9 is involved in many processes, including chondrogenesis and skeletal development (Lefebvre, Behringer, &

Crombrugghe, 2001; Wright, et al., 1995; Bi, Deng, Zhang, Behringer, & Crombrugghe,

1999; Bi, et al., 2001). The authors used Alcian blue to visualize developing cartilage and

Sox9 expression in mouse embryos and found that Sox9 expression preceded sites of chondrogenesis (Wright, et al., 1995).

Further evidence that Sox9 is involved in chondrogenesis is seen in its link to campomelic dysplasia, which is a disease that is characterized by skeletal deformities including underdeveloped shoulder blades and eleven pairs of ribs. These deformities often lead to respiratory distress and thus the mutation is usually lethal with death

13 occurring soon after birth (Mansour, Hall, Pembrey, & Young, 1995). Most genetically male patients with this disease also exhibit XY sex reversal, which demonstrates the critical role for Sox9 in testis development (Mansour, Hall, Pembrey, & Young, 1995).

Another clue to the cause of this disease is that most patients had chromosomal rearrangements associated with chromosome 17q, which is where the Sox9 gene is located (Young, Zuccollo, Maltby, & Broderick, 1992; Tommerup, et al., 1993). Finally, in 1995, it was discovered that mutations within the open reading frame of Sox9 caused the disease (Kwok, et al., 1995).

Sox9 expression has been found to be conserved in testes of vertebrates including mammals, birds, and reptiles, indicating that it is important in animals that have genotypic sex determination and those with temperature dependent sex determination

(Shoemaker, Ramsey, Queen, & Crews, 2007; Western, Harry, Marshall Graves, &

Sinclair, 1999; Kent, Wheatley, Andrews, Sinclair, & Koopman, 1996). Silva et al.

(1996) studied Sox9 expression in developing mouse and chick gonads using whole mount in situ hybridization, RNase protein assays, and immunohistochemistry. They found that Sox9 expression was similar in developing mouse and chick testes and its expression was mostly localized to the nuclei of Sertoli cells. Kent et al. (1996) also examined Sox9 expression in developing mouse and chicken embryos and wrote “The conservation of sexually dimorphic expression in two vertebrate classes which have significant differences in their sex determination mechanisms, points to a fundamental role for Sox9 in testis determination in vertebrates”. In turtles, Sox9 is expressed equally in the bipotential gonads of embryos incubated at male and female producing

14 temperatures but expression begins to differentiate at stage 20 and becomes male specific

(Spotila, Spotila, & Hall, 1998).

Sox9 is a transcription factor that regulates expression of Amh in mice and humans (Arango & Lovell-Badge, 1999; Chaboissier, et al., 2004; Santa Barbara, et al.,

1998). However, the relationship between Sox9 and Amh is different for birds and reptiles. In these vertebrate classes, Amh expression precedes Sox9 expression (Western,

Harry, Marshall Graves, & Sinclair, 1999; Oreal, et al., 1998).

FoxL2

Amh has recently been found to increase FoxL2 mRNA and protein expression in human granulosa cells in vitro (Sacchi, et al., 2017). Conversely, FoxL2 has been shown to activate Amh expression. Amh knockdown leads to accelerated follicle growth.

However, normal Amh expression, combined with FoxL2, was found to result in normal follicle growth in mouse ovaries (Park, Suh, Lee, & Bae, 2014). Based on this information, it is hypothesized that Amh and FoxL2 are part of a positive feedback loop that is critical for proper follicle development (Sacchi, et al., 2017; Park, Suh, Lee, &

Bae, 2014).

FoxL2 is a member of the winged-helix, or Forkhead, transcription factor family.

It was initially discovered in the developing pituitary gland, and thus named pituitary

Forkhead factor (P-Frk) (Treier, et al., 1998). FoxL2 is a transcription factor that has both repressor and activator functions (Crisponi, et al., 2001; Treier, et al., 1998). FoxL2 is expressed in fetal and adult ovaries, the pituitary gland, and also plays a role in eyelid development (Crisponi, et al., 2001; Treier, et al., 1998).

FoxL2 was found in fetal female gonads with little or no expression in developing males (Cocquet, et al., 2002). It is thought that FoxL2 plays a role in maintaining and sustaining development of the ovarian follicles (Crisponi, et al., 2001). FoxL2 is known to induce apoptosis in granulosa cells (Kim, et al., 2011), and is mainly expressed in undifferentiated granulosa cells in the ovary (Crisponi, et al., 2001; Pisarska, Bae, Klein,

& Hsueh, 2004; Loffler, Zarkower, & Koopman, 2003). FoxL2 was expressed in granulosa cells of small, medium, and a few later follicles of infants and juveniles (Uda, et al., 2004).

FoxL2 binds to and represses the gene that encodes steroidogenic acute regulatory protein (StAR) (Pisarska, Bae, Klein, & Hsueh, 2004). StAR is a marker of granulosa cell differentiation (Pisarska, Bae, Klein, & Hsueh, 2004) and controls the rate limiting step for steroidogenesis (Clark, Wells, King, & Stocco, 1994; Lin, et al., 1995). Research shows that FoxL2 is expressed in somatic cells that can develop into granulosa, theca, or stromal cells at 17.5 days post coitum (dpc) in fetal gonads. FoxL2 was found in granulosa cells of follicles at 13 days of age, but the oocyte did not have any transcripts detected by real time PCR, neither did antral follicles or the corpus luteum (Pisarska,

Bae, Klein, & Hsueh, 2004). Another function is in eyelid formation. FoxL2 is expressed during development around the optic cup and in the ridges of the developing eyelids

(Crisponi, et al., 2001).

Due to its critical roles in eyelid and reproductive development, FoxL2 is highly conserved in its amino acid sequence and timing and pattern of expression across different vertebrate species (Loffler, Zarkower, & Koopman, 2003) and has been found to be under purifying selection (Cocquet et al., 2002). Purifying selection is when a gene

16 critical for survival and production of future generations is purged of deleterious or negative genetic polymorphisms that are derived from random mutations.

Mutations of FoxL2 disable its repressive function, which is thought to increase levels of StAR transcription and steroidogenesis as well as accelerate follicle development. Accelerated follicle development is the cause of premature ovarian failure

(Pisarska, Bae, Klein, & Hsueh, 2004). Premature ovarian failure with malformation of the eyelids is characteristic of a disease called blepharophimosis-ptosis-epicanthus inversus syndrome (Crisponi, et al., 2001; De Baere, et al., 2001). With so many phenotypes caused by a faulty FoxL2 there is clearly more to learn about this gene and how it functions.

Somatostatin

Moving to another component of the HPG axis, the hypothalamus expresses Sst

(Spiess, Villarreal, & Vale, 1981). The somatostatin gene encodes a larger precursor protein that is post-translationally processed to produce the mature hormone of 28 amino acids (Schally, et al., 1980). Human, pig and sheep Sst-28 are identical in sequence.

Comparisons to anglerfish pre-pro-somatostatin sequences revealed that the carboxy- terminal region is conserved, along with the adjacent six amino acids. The remainder of the protein is more diverse, but with many conservative amino acid substitutions (Shen,

Pictet, & Rutter, 1982). Sst is expressed in the hypothalamus of all vertebrate classes examined to date (Spiess, Villarreal, & Vale, 1981; Tostivint, Joly, Lihrmann, Ekker, &

Vaudry, 2004) and is part of a feedback loop that inhibits growth hormone synthesis and secretion from the anterior pituitary (Brazeau, et al., 1973). The hormone is also

17 expressed in the digestive system and plays a part in regulating secretion, absorption, and motility of the gut (Corleto, 2010).

Literature on Sst mutation is lacking. However, there is information on Sst receptors of which there are five (Hejna, Schmidinger, & Raderer, 2002). Somatostatin receptors 1, 2, and 5 have been linked to inhibition of cell growth (Buscail, et al., 1994;

Buscail, et al., 1995; Kreienkamp, Akgun, Baumeister, Meyerhof, & Richter, 1999). A mutation in Sst receptor 5 (SSTR5) in a patient with acromegaly caused the patient to be resistant to a Sst analog that is used to treat the disease (Ballare, et al., 2001).

Growth Hormone

Somatostatin produced by the hypothalamus and secreted into the hypophysial portal system inhibits growth hormone (Gh1) synthesis and secretion from somatotropes in the anterior pituitary gland (Brazeau, et al., 1973) while growth hormone releasing hormone stimulates Gh1 synthesis and secretion (Goldenberg & Barkan, 2007; Gahete, et al., 2009). The growth hormone gene encodes a protein that is critical for regulating postnatal growth in vertebrates (Lupu, Terwilliger, Lee, Segre, & Efstratiadis, 2001). Gh1 binds to receptors in the liver and stimulates somatomedin (Igf1) production (Marques,

Silva, & Turyn, 1979; Salvatori, 2004).

Studies using immunochemical methods and radioimmunoassay demonstrate that

Gh1 is highly conserved among the common snapping turtle, rat, fish, monkey, ant eater, and possum (Hayashida, Farmer, & Papkoff, 1975). However, Gh1 from non-primates has no effect on humans indicating that functional interactions between Gh1 and its receptor have evolved (Forsynth & Wallis, 2002). Mutations in the sequence of Gh1 lead to isolated growth hormone deficiency (IGHD) (Keselman, et al., 2012), which is a

18 disorder characterized by severe growth retardation at birth, postnatal hypoglycemia, and dwarfism (Giordano, 2016).

Cdx1

We next move caudally to the intestine, which is known to express Cdx1 (Silberg,

Swain, Suh, & Traber, 2000). Like several of the genes discussed above, Cdx1 is a transcription factor. It is a member of the caudal-related homeobox gene family (Mallo, et al., 1997; Duprey, et al., 1988). The first member of the Cdx gene family was found in

Drosophilia and was named caudal (cad) (Mlodzik & Gehring, 1987; Ghering, et al.,

1990). Later, a cDNA probe from Drosophilia cad was used to isolate Cdx1 from in the mouse (Duprey, et al., 1988). Human and mouse Cdx1 sequences are 85% identical

(Mallo, et al., 1997). Like the cad gene, Cdx genes in vertebrates are thought to play a role in specifying development of the caudal region of embryos and anteroposterior patterning (Silberg, Swain, Suh, & Traber, 2000). There is a gradient of expression of

Cdx1 and Cdx2 that is thought to direct development of the intestine and maintain different cell types of the intestine and colon (Silberg, Swain, Suh, & Traber, 2000)

(Meyer & Gruss, 1993). This protein gradient is also present in Drosophilia and is initiated by the cad gene (Mlodzik & Gehring, 1987).

There are two different stages of expression for Cdx1. Transcripts are first detected on day 7.5 and are expressed until 12 days post coitum in embryonic mice

(Silberg, Swain, Suh, & Traber, 2000) (Meyer & Gruss, 1993). Expression is found in the primitive streak, and later in the developing limbs, neuroectoderm, and somites. In the second phase, Cdx1 expression is restricted to the intestine and colon. Intestinal

19 expression continues into adulthood and is critical for intestinal maintenance (Silberg,

Swain, Suh, & Traber, 2000).

Studies have shown that early expression of Cdx1 is initiated by two Wnt/beta- catenin responsive elements, while a retinoic acid (RA) response element is critical for maintaining its expression (Lickert & Kemler, 2002; Lickert, et al., 2000). Research suggests that Cdx1 regulates Hox gene expression during embryonic development

(Subramanian & Meyer, 1995).

Interestingly, mutations or knock out of Cdx1 in mice also leads to skeletal abnormalities, including the occipital bones of the skull, and transformations involving all seven of the cervical vertebrae and thoracic vertebra 8 through 10. Lumbar vertebrae and hindlimbs were normal. There were no abnormalities seen past thoracic vertebra 9

(Subramanian & Meyer, 1995).

Slc12a1

The final gene examined in the review Slc12a1, is expressed in the kidneys

(Greger & Schlatter, 1981; Kaplan, Plotkin, Brown, Hebert, & Delpire, 1996). This tissue was chosen due to its proximity to the gonads. During gonad development, cells migrate from the mesonephros to the site of gonad formation (Wilhelm, Palmer, & Koopman,

2007). Slc12a1 codes for an electroneutral transporter for Na+, K+, and Cl- (Gamba, et al.,

1994). The first physiological evidence for this type of gene was found in Ehrlich ascites tumor cells. While researchers knew that a cotransport system functioned in relation to cell volume, they were unable to determine whether the process involved primary or secondary transport (Geck, Pietrzyk, Pfeifferl, & Heinz, 1980). It was later found that this cotransporter works by secondary active transport and is powered by ion gradients

20 generated by the Na+/K+-ATPase. This gene is in the family of solute carrier 12 family of transporters (Markadieu & Delpire, 2014). It is expressed in the thick ascending limb of the loop of Henle in the kidney (Greger & Schlatter, 1981; Kaplan, Plotkin, Brown,

Hebert, & Delpire, 1996) where it reabsorbs NaCl that had been filtered by the glomerulus (Wang, White, Geibel, & Giebisch, 1990).

Mutations of Slc12a1 have been linked to Bartter’s syndrome (Simon, et al.,

1996), which is associated with dehydration, low blood pressure, seizures, tetany, muscle weakness, paresthesia, and joint pain (Markadieu & Delpire, 2014).

Gene Regulation

These are only a handful of the genes that display cell-type specific and/or tissue specific expression. Given that the DNA in all cell types is identical, excluding germ cells going through meiosis and B cells that produce variable antibodies, something must be regulating differential gene expression (Sonawane, et al., 2017).

There are many ways gene expression can be modified, in addition to epigenetic regulation. For this discussion, the focus will be on two of the most studied epigenetic mechanisms: DNA methylation and histone methylation. Even though regulation of gene expression is not exclusively performed through chromatin remodeling, differences in its chromatin state are associated with a more permanent state of tissue specific expression

(Mikkelsen, et al., 2007). An epigenetic mark is one that is stably inherited from mother to daughter cells during mitosis within a developing organism. Inherited patterns of gene expression result from changes in chromatin structure, rather than alterations to the DNA sequence itself (Berger, Kouzarides, Shiekhattar, & Shilatifard, 2009). Very little research has been performed regarding epigenetic modifications in reptiles.

DNA Methylation

DNA methylation is known to silence gene expression. One study looked at DNA methylation and its relation to body temperature over extended periods of time. Varriale and Bernardi (2006) found that DNA methylation was highly variable in relation to body temperature. The study looked at nucleotide composition, band asymmetry in a cesium chloride gradient, and compositional heterogeneity. The researchers found that methylation of reptilian genomes was heterogenous in comparison with other vertebrates.

This study only examined one crocodile, two turtles, one varanid, and three snakes

(Varriale & Bernardi, 2006). DNA methylation has been examined at a specific sex- determining locus in the red-eared slider turtle, a species with TSD. The aromatase gene is expressed at a female producing temperature, but not at a male producing temperature

(Crews & Bergeron, 1994). DNA methylation in the aromatase promoter was lower at a female producing temperature compared to a male producing temperature, suggesting that DNA methylation may inhibit transcription (Matsumoto, Buemio, Chu, Vafaee, &

Crews, 2016).

Histone Modifications

Histone modifications can also alter gene expression. Histone methylation can silence or activate gene expression depending on the residue that is methylated.

Trimethylation of histone H3 on lysine 27 (H3K27me3) is associated with gene silencing

(Cedar & Bergman, 2009), while trimethylation on lysine 4 (H3K4me3) is associated with gene activation (Howe, Fischl, Murray, & Mellor, 2017). In the red-eared slider turtle, H3K4me3 histone modification was found near the aromatase gene promoter at a female producing temperature. H3K27me3 was observed at both male and female

22 producing temperatures but was later demethylated at both temperatures (Matsumoto,

Buemio, Chu, Vafaee, & Crews, 2016).

Conclusions

There are very few studies that link tissue specific expression of genes to epigenetic modifications in reptiles. Some correlational evidence does suggest that regulatory patterns could be similar to mammals. For instance, bivalent chromatin modifications may regulate expression of sex determining genes in the red-eared slider turtle (Matsumoto, Hannigan, & Crews , 2016). However, more experimental work is needed to determine if epigenetic mechanisms actually regulate gene expression. The final goal of this research is to study tissue specific gene regulation in relation to temperature-dependent sex determination and determine if expression of sex-determining genes is regulated epigenetically as it is in mammals. The first step to studying these mechanisms is to identify genes that display tissue-specific expression patterns. The next step will be to test whether epigenetic marks at specific genes are correlated with their tissue specific patterns of expression. Finally, experimental disruption of these marks will be required to test whether the marks do indeed play a role in regulation of gene transcription.

CHAPTER 2

TISSUE SPECIFIC GENE EXPRESSION IN CHELYDRA SERPENTINA

Abstract

Many studies have been conducted on gene expression, patterning, and function in mammals. However, there have been few studies performed on reptiles with the exception of sex-determining genes. Studies of tissue specific gene expression are the first step to gaining more information on gene patterning and function in reptiles, and to help determine whether they are similar, or drastically different from mammals particularly with regard to sex determination. Any differences in gene expression could allude to differences in gene regulation and gene function and will require further study.

Hatchling turtles were dissected at 30 days of age, and tissues stored in RNA later. RNA was isolated, converted to cDNA, and was used in quantitative real time PCR to measure gene expression. Gene expression state was examined in hypothalamus, pituitary, testis, ovary, intestine, and kidney for Anti-Müllerian hormone (Amh), Sox9, FoxL2, Dmrt1, growth hormone 1 (GH1), Somatostatin (Sst), Cdx1, and Slc12a1. Expression patterns were similar to patterns found in mammals with several interesting differences. Sox9 was found to be expressed in the hypothalamus and pituitary glands in addition to male gonads. Amh levels were higher in male gonads compared to female. FoxL2 expression was high in the hypothalamus, and there was also expression in testes. CDX1 was found in gonads in addition to the intestine and was higher in male compared to female gonads.

Slc12a1 was found in the kidney as expected, but also in the gonads of both sexes.

Introduction

Differentiation of cells is crucial for the formation of the different tissues and organs in the body of multicellular organisms like animals. While all cells within the body have the same DNA content and sequence, different genes in those cells are silenced and activated to produce cells with different phenotypic characteristics. One example of this process of cell, tissue, and organ differentiation is gonad development.

There are three steps to the formation of the gonads. The first step is development of the bipotential gonads, also known as gonadogenesis (Swain & Lovell-Badge, 1999;

McLaren, 2000). These bipotential organs can either develop into testes or ovaries, the male or female reproductive organs (Lillie, 1917; Fisher & Merkenschlager, 2002; Loo &

Rine, 1995). The second step is sex determination when the developmental fate of cells and gonads is determined: i.e., an irreversible developmental decision is made for the gonads to become testes or ovaries (Swain & Lovell-Badge, 1999; McLaren, Germ and

Somatic Cell Lineages in the Developing Gonad, 2000). The third step involves phenotypic differentiation of various testicular and ovarian cell types and reorganization of these cells during morphogenesis of testes and ovaries. This whole cascade of events begins with determination and differentiation of precursor support cells, which later become Granulosa or Sertoli cells in females and males respectively (Kim & Capel,

2006).

There are several ways genes can be regulated during development to allow for tight control of gene expression and thus initiate the cascade of cell fate determination and cellular differentiation. Two interrelated mechanisms are tissue-specific transcription

25 factors and epigenetic mechanisms. The definition of an epigenetic trait is one that is stably inherited from mother cell to daughter cells during mitosis. Inherited patterns of gene expression result from changes in chromosome structure, rather than alteration of the DNA sequence itself (Berger, Kouzarides, Shiekhattar, & Shilatifard, 2009). This is still a relatively new field of study. The most studied forms of epigenetic modifications are DNA methylation as well as acetylation and methylation of the histone proteins around which the DNA is wrapped (Bird, 1992; Cedar & Bergman, 2009). Very little is known about epigenetic mechanisms in reptiles as research in this class of vertebrates is lacking. Knowledge on which genes are expressed in specific tissues is the first step to studying gene regulatory mechanisms in reptiles.

For this reason, we have chosen several tissues to study gene expression in

Chelydra serpentina. For this study, we focus on five tissues from males and include a comparison of gonads between males and females (i.e., testes versus ovaries). The gonads, along with the hypothalamus and pituitary gland forms the HPG axis. These tissues, together with the hormones they produce, are essential for reproduction. The kidneys and intestine were chosen because of their location near the gonads. The embryonic kidneys also have precursor cells that later contribute to formation of the gonads (Wilhelm, Palmer, & Koopman, 2007). While the gonads are forming, the primordial germ cells (PGCs) migrate along the gut tube, which later becomes part of the intestine (Swain & Lovell-Badge, 1999; McLaren, 2000). Research conducted on mammals suggests that we should see expression of Sox9, Amh, Dmrt1, and FoxL2 in the gonads, with Sox9, Amh, and Dmrt1 higher in testes, and FoxL2 higher in ovaries

(De Grandi, et al., 2000; Kettlewell, Raymond, & Zarkower, 2000; Spotila, Spotila, &

Hall, 1998; Park, Suh, Lee, & Bae, 2014). Somatostatin (Sst) is produced by the hypothalamus while growth hormone (Gh1) is active in the pituitary (Spiess, Villarreal,

& Vale, 1981; Gahete, et al., 2009). Cdx1 is specific to the intestine while Slc12a1 is produced in the kidney (Silberg, Swain, Suh, & Traber, 2000; Greger & Schlatter, 1981).

Materials and Methods:

All protocols and methods for the use of animals were approved by the University of North Dakota Institutional Animal Care and Use Committee (IACUC).

Acquisition of Tissues

Eggs were produced by the breeding colony of adult snapping turtles at the

University of North Dakota in June of 2017. Turtle eggs were collected on the same day that they were laid. Eggs were washed in tepid water, marked with a permanent Sharpie marker with a unique clutch and egg number, and placed in boxes filled with moist vermiculite (dry vermiculite and water were mixed in a 1:1 ratio by mass). Eggs from each clutch were evenly split between incubators set to 22.5° C, 26° C, 27° C, or 28° C for the entire embryonic period. Eggs were checked on a daily basis near the end of incubation to determine when turtles pipped or ruptured the egg shell prior to fully emerging from the egg.

Turtle hatchlings were rapidly decapitated and dissected 30 days following their pip date. All tissues for this study, excluding ovaries, were collected from males. All tissues, including testes, hypothalamus, pituitary, intestine section, and kidneys, were collected from each male. The only tissues collected from females were ovaries. Tissues were removed and immediately placed in RNAlater. Tissues were stored at -20° C until used for RNA extraction. Hatchlings were selected from three clutches with the majority

27 being from the same clutch. This clutch had a significant number of hatchlings that were female despite being incubated at 26° Celsius, which usually produces males. All of the samples were from embryos incubated at one temperature to avoid any temperature effects, which have been shown to affect growth rates (Rhen & Lang, Phenotypic

Plasticity for Growth in the Common Snapping Turtle: Effects of Incubation

Temperature, Clutch, and Their Interaction, 1995). Most hatchlings came from just two clutches to minimize any differences in gene expression due to genetic variation. For the ovary, testis, pituitary, and hypothalamus samples, the whole organ was used. For intestine and kidney, a 1mm tissue punch was used, and the samples were obtained from the same part of the organ each time. One punch was adequate for intestine samples, while 2 punches were needed for kidney samples. Intestine samples were from the ileum, just before the start of the large intestine while kidney samples were from the caudal region just below the adrenal glands.

RNA Extractions

RNA extractions were performed using the PicoPure RNA Isolation kit (catalog #

KIT0202, KIT0204). There were ten replicates for hypothalamus, ovary, and kidney tissues. Pituitary and intestine tissues had 9 replicates and testes had 13 replicates. Tissue was removed from RNAlater, blotted of excess RNAlater, placed in 100µl of extraction buffer, and homogenized with a Pro Scientific Bio-Gen Pro200 Handheld Homogenizer.

The CapSure Macro LCM Caps protocol was followed with the DNase (Qiagen) step included. Samples were then checked for RNA concentration using a Nanodrop OneC spectrophotometer and a Qubit 3.0 Fluorometer. Samples were diluted to the same

28 concentration of RNA (48ng of total RNA in 20µl of ultrapure water) in a 96 well plate.

When not in use, the plate containing RNA was sealed and stored at -80° Celsius.

Reverse Transcription

We first used quantitative PCR to test for contaminating genomic DNA in the total RNA with primers for Dmrt1. A positive control using 1ul of genomic DNA at a concentration of 144ng/ul was also included. Genomic DNA amplified at a Ct of 22.90.

Most RNA samples did not amplify at all while the earliest cycle threshold (Ct) value for amplification was 33.87 (11 orders of magnitude lower than the genomic DNA positive control). This indicates that there was very little to no contamination of genomic DNA and that RNA was pure.

We used the High Capacity cDNA Reverse Transcription Kit to synthesize cDNA according to the manufacturer’s instructions (Applied Biosystems). For each sample, 2ul of 10x RT Buffer, 0.8ul of 25x DNTPs, 2ul of 10x Random Primers, 1ul of Multiscribe

RT, and 4.2ul of Milli-Q water were combined to make a 2x mastermix. Normalized

RNA was removed from -80° C and allowed to thaw. The sealed plate was then placed in the thermocycler at 65° C for five minutes to denature RNA secondary structure, followed by a five-minute incubation on ice, after which 10uL of the cDNA mastermix was added to each sample well. The plate was then sealed and placed in the thermocycler following the recommended thermal profile: 25° C for 10 minutes, followed by 37° C for

120 minutes, then 85° C for 5 minutes, and finally 4° C for a holding temperature. After reverse transcription, cDNA was stored at -80° C until ready for use as template in quantitative PCR reactions.

Quantitative PCR

All qPCR reactions were set up using a Biomek FXP liquid handling robot.

Quantitative PCR was performed using the Bio-Rad CFX 384 ™ Real-Time System to assess gene expression. Master mix was made using SSoFast ™ EvaGreen ® Supermix from Bio-Rad, and primers were designed using Primer Express Software, and ordered from Integrated DNA Technologies (IDT). We combined 5uL of SSoFast TM Evagreen ®

Supermix, 0.6uL of each forward and reverse primers (10mM stock), and 1.8ul of ultrapure water for each sample and standard. Two microliters of each sample or standard was added for each reaction with 8ul of master mix. A standard curve was generated for each gene using known amounts of PCR product that were made via serial dilution. The thermal profile for qPCR was as follows: an initial polymerase activation step of 30 seconds of 95˚ C, which was followed by 40 rounds of two-step PCR with 95˚ C for 5 seconds and 60˚ C for 5 seconds. Standard curves were used to convert Ct values to absolute amounts of cDNA for each sample.

Primer Sequences Gene Type Primer Sequence 5' → 3' Sox9 Forward GAC ACA CAT CAA GAC GGA GCA Sox9 Reverse AAT GGA AGG CAC TCC GGT AGT Amh Forward CAG CTG CTA CTC ATC TCC TCT TGA Amh Reverse GGG CAT TTT GCG CCT TT FoxL2 Forward CGC GTT GCT GCC TCT TG FoxL2 Reverse CCC TTA GGA GCT GGT AAA GTT TCT C Dmrt1 Forward TCA GAG ACG ATG CCC AAT GA Dmrt1 Reverse GGC ATG ATG GTC CGA AGA AG Cdx1 Forward CTG AAC CTC AAT CCC CAG AAC T Cdx1 Reverse CCT GGC ACG TGA TGG TAG CT Slc12a1 Forward CCC AAG AAG ATG CCC ATC CT Slc12a1 Reverse TGA TCC TGT GTC TGC CAT GTG Sst Forward TGC AGG GAA CCG ACC TTA AC Sst Reverse GCT GGT TTC GCC TGA AAG C Gh1 Forward GGG CCT CAG GTC TGA TCC A Gh1 Reverse GAT AAC CTC CCT GTT GCT TAG GAA

Table 1: Primer sequences for qPCR were downstream from the transcription start site only. Primers were designed using Primer Express software and were ordered from IDT.

Statistical Analysis

Quantitative qPCR results were analyzed using JMP software version 14.0.0. Data was transformed to reduce heteroskedasticity. The natural log transformation was used for all genes with the exception of Slc12a1, which was transformed using Log 10.

Residuals were examined to ensure that the transformed gene expression values met the assumptions of Analysis of Covariance (ANCOVA). The quantiles function was used to detect and exclude outliers. There were three or fewer outliers for all genes except Sox9, which had 4 outliers. The fit model function was used to run (ANCOVA) with tissue type as the independent variable and expression of 18S rRNA as the covariate. We used Ct values for 18S rRNA as a covariate to control for random variation among samples due to

31 factors like variation in the quality of input RNA, variation in the efficiency of reverse transcription reactions, and pipetting errors. We measured absolute expression rather than relative expression because normalization via the delta-delta Ct method induces spurious differences when “housekeeping genes” vary in expression among groups of interest.

Moreover, measurement of absolute expression allows more direct and quantitative analysis of differences in gene expression. When the effect of tissue type was statistically significant, we used Tukey’s HSD post hoc test to compare gene expression levels among all tissues.

Plots of means were created in RStudio using the ggplot2 package. Outliers have been excluded in these plots as well. The means of the transformed data were calculated, plotted, and the bars on either side indicate one standard error.

Results

Overall, the ANCOVA revealed highly significant differences in expression levels for all eight genes among tissues (Table 1). The 18S rRNA covariate was also significant for some genes but not others (Table 1). We present tissue specific expression patterns for each gene in the following paragraphs. First are genes expected to be expressed at a higher level in testis (Amh, Dmrt1, and Sox9), then ovary (FoxL2), hypothalamus (Sst), pituitary (Gh1), intestine (Cdx1), and kidney (Slc12a1).

Amh Dmrt1 Sox9 FoxL2 F F F F [5,45] P [5,47] [5,49] [5,41] Effect ratio ratio P ratio P ratio P Tissue 118.3197 <.0001* 146.7146 <.0001* 155.5678 <.0001* 299.8967 <.0001* 18S rRNA 3.8956 0.0546 1.9878 0.1652 71.3242 <.0001* 5.2414 0.0273*

Sst Gh1 Cdx1 Slc12a1 F F F Effect [5,47] P [5,45] P [5,45] P F ratio P ratio ratio ratio [5,49] Tissue 143.5192 <.0001* 104.503 <.0001* 207.4654 <.0001* 38.9819 <.0001* 18S 1.192 0.2805 0.4864 0.4864 1.412 0.241 11.0803 0.0016* rRNA

Table 2: ANCOVA results for tissue specific expression. Asterisks indicate significance. Data was analyzed using JMP software version 14.0.0 The numbers following the F ratio indicate degrees of freedom for the variance between groups, and within groups respectively.

Anti-Müllerian Hormone (Amh) expression varied significantly among the different tissue types (Table 1). The 18S rRNA covariate was not statistically significant

(Table 1). Amh expression was highest in testes, followed by ovaries, and then the hypothalamus (Figure 1). Expression of Amh was lowest in the pituitary, kidney, and intestine and were not significantly different from each other (Figure 1).

Figure 1: Tissue specific expression of Amh. Plot of means was created using the ggplot2 package in RStudio and was generated from transformed data. Error bars indicate the standard error. The letters indicate the statistical differences via Tukey’s Honest Significance Test. All statistics were performed using JMP software version 14.0.0.

Expression of Dmrt1 varied significantly among tissue types (Table 1). Again, the

18S rRNA was not significant (Table 1). Dmrt1 expression was highest in testes, followed by ovaries, and then the pituitary gland (Figure 2). Dmrt1 expression was lowest in kidney, intestine, and the hypothalamus, with no differences among these tissues (Figure 2).

Figure 2: Tissue specific expression of Dmrt1. Plot of means was created using the ggplot2 package in RStudio and was generated from transformed data. Error bars indicate the standard error. The letters indicate the statistical differences via Tukey’s Honest Significance Test. All statistics were performed using JMP software version 14.0.0.

The ANCOVA revealed highly significant differences in Sox9 expression levels among tissues (Table 1). The 18S rRNA covariate was significant (Table 1), and there is more variation within groups in this gene compared to the others. This could indicate our primers are amplifying DNA other than Sox9. Sox9 was expressed in the testis and hypothalamus at roughly equal levels (Figure 3). Expression in the intestine and pituitary was lower than in hypothalamus and testes, but significantly higher than in the ovaries and kidneys (Figure 3).

Figure 3: Tissue specific expression of Sox9. Plot of means was created using the ggplot2 package in RStudio and was generated from transformed data. Error bars indicate the standard error. The letters indicate the statistical differences via Tukey’s Honest Significance Test. All statistics were performed using JMP software version 14.0.0.

FoxL2 expression levels varied significantly among the different tissue types

(Table 1). The 18S rRNA covariate was statistically significant (Table 1). Expression of

FoxL2 was highest in the pituitary and ovaries but did not differ between these two tissues (Figure 4). Expression levels were significantly lower in hypothalamus and then testes (Figure 4). Expression levels were lowest in the kidney and intestine (Figure 4).

Figure 4: Tissue specific expression of FoxL2. Plot of means was created using the ggplot2 package in RStudio and was generated from transformed data. Error bars indicate the standard error. The letters indicate the statistical differences via Tukey’s Honest Significance Test. All statistics were performed using JMP software version 14.0.0.

Somatostatin (Sst) levels varied dramatically among tissues (Table 1). The 18S rRNA covariate was not significant (Table 1). Hypothalamus showed the highest Sst expression (Figure 5). Expression of Sst was significantly lower in the intestine followed by the lowest levels in the pituitary, testis, ovary, and kidney samples (Figure 5).

Figure 5: Tissue specific expression of Sst. Plot of means was created using the ggplot2 package in RStudio and was generated from transformed data. Error bars indicate the standard error. The letters indicate the statistical differences via Tukey’s Honest Significance Test. All statistics were performed using JMP software version 14.0.0.

Growth hormone 1 (Gh1) expression differed significantly among tissue types

(Table 1), but the 18S rRNA covariate was not significant. The highest expression of Gh1 was observed in the pituitary gland (Figure 6). Expression was significantly lower in testis and ovary but did not differ between these two tissues (Figure 6). The lowest Gh1 expression was observed in hypothalamus, intestine and kidney (Figure 6).

Figure 6: Tissue specific expression of Gh1. Plot of means was created using the ggplot2 package in RStudio and was generated from transformed data. Error bars indicate the standard error. The letters indicate the statistical differences via Tukey’s Honest Significance Test. All statistics were performed using JMP software version 14.0.0.

Cdx1 expression levels were significantly different among tissues (Table 1). The

18S rRNA covariate was not significant (Table 1). The highest Cdx1 expression was seen in the intestine (Figure 7), with significantly lower levels found in the testis and ovary

(Figure 7). Next was the pituitary, followed by kidney and hypothalamus (Figure 7).

Figure7: Tissue specific expression of Cdx1. Plot of means was created using the ggplot2 package in RStudio and was generated from transformed data. Error bars indicate the standard error. The letters indicate the statistical differences via Tukey’s Honest Significance Test. All statistics were performed using JMP software version 14.0.0.

Finally, Slc12a1 expression differed among tissue types (Table 1). The 18S rRNA was a significant covariate (Table 1). This could be due to pipetting errors, or variation in the efficiency of the reverse transcriptase. The highest expression was observed in the kidney (Figure 8). Expression of Slc12a1 was significantly lower in all other tissues with lowest expression in the intestine (Figure 8).

Figure 8: Tissue specific expression of Slc12a1. Plot of means was created using the ggplot2 package in RStudio and was generated from transformed data. Error bars indicate the standard error. The letters indicate the statistical differences via Tukey’s Honest Significance Test. All statistics were performed using JMP software version 14.0.0.

Discussion

Results of this study were consistent with previous literature but with several interesting differences. Sox9 expression was found to be expressed in the hypothalamus and pituitary gland in addition to male gonads. Amh levels were higher in male gonads compared to female gonads. FoxL2 expression was high in the hypothalamus, and there was also some expression in testes. Cdx1 was found in gonads in addition to the intestine and was higher in male compared to female gonads. Slc12a1 was found in the kidney as expected, but also in the gonads of both sexes.

Sox9 expression in the intestine is not surprising or unusual upon reviewing the literature. Sox9 knockdown in mice reveals that it is essential for Paneth cell

41 differentiation in the intestines. Knockdown using a line of mice with a conditional Sox9 gene combined with a Villin-Cre transgene revealed an absence of Paneth cells in the intestines of these mice, with widening crypts, and replacement of Paneth cells by epithelial tissue (Mori-Akiyama, et al., 2007). There are no prior studies reporting Sox9 expression in reptile intestines. Given that Sox9 is involved in development of Paneth cells in mammals, this may explain why it is higher in the intestine compared to the ovary and kidney samples.

No studies could be found on Sox9 expression in the hypothalamus, however it is known that Sox9 is expressed in brain tissues in mammals (Pompolo & Harley, 2001).

Evidence has been found of Sox9 expression in the brains of the Red-Eared Slider Turtle, and the Orange-Spotted Grouper, while in the Japanese rice fish one of the two Sox9 variants found in this species was expressed in brain tissue (Spotila, Spotila, & Hall,

1998; Luo, Hu, Liu, Lin, & Zhu, 2010; Kluver, Kondo, Herpin, Mitani, & Schartl, 2005).

Sox9 is expressed mostly in astrocytes in the forebrain of adult rats and mice, which contains the hypothalamus. Sox9 protein was detected in white and gray matter using a monoclonal antibody and immunofluorescence. Sox9 was found in the nuclei of cells in male and female brains (Pompolo & Harley, 2001). Another study looked at Sox9 expression and the development of neural stem cells. The researchers found that Sox9 is involved in the neurosphere formation which is from neuroepithelial cells, and that Sox9 is needed for the maintenance of multipotentiality of neural stem cells (Scott, et al.,

2010).

Sox9 expression is also necessary for proper testis development (Chaboissier, et al., 2004; Arango & Lovell-Badge, 1999). In one study, homozygous knockout of the

42 gene in mice leads to a high proportion of animals with abnormal sex cord formation. The organization of germ cells was normal, however, there were fewer seminiferous cords with an irregular outline. Amh expression was also lower in mice with lower Sox9 expression, as was Sox8 expression. Homozygous mice for deletion of Sox9 died before gonad differentiation, so the gonads were removed and cultured, and did not form sex cords, or express male specific genes such as Amh, or Cyp11a1, which is a Leydig cell- specific marker. Instead, the gonads showed expression of Bmp2 and Follistatin, which are required for development of ovaries (Chaboissier, et al., 2004). Deletion of an upstream enhancer of Sox9 leads to decreased levels of expression consistent with female gonads and XY sex reversal (Gonen, et al., 2018). Without proper expression of Sox9,

Amh expression is also disrupted which can result in pseudohermaphrodites (Arango &

Lovell-Badge, 1999). Conversely, ectopic expression of Sox9 is sufficient to induce testis development in XX female mice (Vidal, Chaboissier, de Rooij, & Schedl, 2001).

Expression of Sox9 is influenced by incubation temperature with increasing levels at male producing temperatures during the temperature sensitive period of development in the common snapping turtle (Rhen T. , Metzger, Shroeder, & Woodward, 2007). When eggs were shifted from a male producing temperature to a female producing temperature, expression levels of Sox9 decreased. Males had nine times the amount of Sox9 after hatching compared to females (Rhen T. , Metzger, Schroeder, & Woodward, 2007).

Expression of Sox9 was the lowest in ovaries and kidneys. Expression of Sox9 is at very low levels in the gonadal ridge during embryonic development in mammals.

Expression is maintained at this low level in ovaries through development in humans and in reptiles (Hanley, et al., 2000; Western, Harry, Marshall Graves, & Sinclair, 1999).

Although expression in the kidney was low in hatchling tissues, the opposite may be true for embryonic kidneys. Sox9 is briefly expressed in the mesonephric tubules in chicks and the metanephros of mouse embryos (Silva, et al., 1996). Sox9 expression has not previously been studied in kidneys of reptiles, although there is information on expression in adrenal kidney gonad complexes (AKGs) (Spotila, Spotila, & Hall, 1998;

Shoemaker, Ramsey, Queen, & Crews, 2007).

Sox9 is expressed in the pituitary gland most likely because it is a transcription factor associated with stem/progenitor cells in the pituitary (Chen, et al., 2009; Rizzoti,

Akiyama, & Lovell-Badge, 2013). Pituitary progenitor cells were found to express Sox2 when proliferating, followed by Sox9, which is thought to stop proliferation and revert cells to a non-proliferative state (Rizzoti, Akiyama, & Lovell-Badge, 2013).

Unlike Sox9, Amh expression level and location is initially the same in the bipotential gonads of a TSD species (Shoemaker, Ramsey, Queen, & Crews, 2007;

Takada, DiNapoli, Capel, & Koopman, 2004). Müllerian ducts form as a separate set of ducts that are connected to the bipotential gonads and become the oviducts, uterus, and upper vagina in mammals if the gonads develop along the female pathway (Munsterberg

& Lovell-Badge, 1991). In males, these ducts are not needed, and thus Amh is produced to cause their regression (Munsterberg & Lovell-Badge, 1991). Expression of Amh occurs soon after testis differentiation begins in the red-eared slider turtle, a turtle with

TSD (Wibbels, Wilson, & Crews, 1999). In accord with this finding, Amh levels were found to be highest in testes from 30-day old snapping turtles. This contrasts with a study done in mice, which found there is no postnatal expression of Amh in males

(Munsterberg & Lovell-Badge, 1991). This suggests differences between mammals and

44 reptiles in the timing of Müllerian duct regression. However, in male red-eared slider turtles, Müllerian duct regression was found to occur at stages 22-23 of embryonic development and were absent by stage 26 (hatching) That study did not describe Amh levels, as the Amh gene had not been cloned in reptiles at the time (Wibbels, Wilson, &

Crews, 1999). In relation to TSD species, most research is focused on gene expression levels during the temperature sensitive period (Durando, et al., 2013; Wibbels, Wilson, &

Crews, 1999; Takada, DiNapoli, Capel, & Koopman, 2004). In humans, it has been found that Amh levels are elevated from the formation of the seminiferous cords until puberty, when Amh levels decrease and testosterone levels increase (Rey, et al., 1993).

Expression of Amh is required for proper postnatal development of the ovaries, after gonad fate has been established. Amh has been found to play a role in regulating germ cell proliferation (Durlinger, et al., 1999; Skaar, et al., 2011; Nobraga, et al., 2015;

Safian, Bogerd, & Schulz, 2018; Miura, Miura, Konda, & Yamauchi, 2002) and Amh is expressed in granulosa cells in sexually mature females (Takahashi, Hayashi, Manganaro,

& Donahoe, 1986). Future studies could test whether Amh expression in female turtles increases at reproductive maturity when granulosa cells and follicles would be growing.

However, female mice have detectable Amh levels in granulosa cells six days after birth

(Munsterberg & Lovell-Badge, 1991). Had embryonic ovaries been used, likely no Amh expression would be seen (Munsterberg & Lovell-Badge, 1991).

Some Amh expression was observed in the hypothalamus of hatchling turtles.

Amh expression in the hypothalamus correlates with a study conducted on zebrafish that found Amh is expressed at a higher level in male brains compared to female brains early in development, before sexual differentiation of the gonads. This indicates that sexual

45 differentiation is likely occurring in the brain before the gonads (Poonlaphdecha, et al.,

2010). That study was conducted in embryonic brains, whereas the current study was conducted in hatchling turtles that were thirty days old. Without expression data in hypothalamic tissues from female turtles it is impossible to know if Amh expression is also sexually dimorphic in turtles, or if it is part of normal regulatory processes of the brain and endocrine system.

Our results for Dmrt1in snapping turtle tissues support previous research. The gene is highly expressed in the testes, followed by ovaries, and finally some expression in the male pituitary gland. It is known that this gene is expressed at greater levels in male gonads compared to female gonads during the thermosensitive period of sex determination in a turtle species that has temperature-dependent sex determination (Rhen

T. , Metzger, Schroeder, & Woodward, 2007). This is also the gene known to show the earliest sexually dimorphic expression in reptiles and is therefore an early marker of testis fate (Kettlewell, Raymond, & Zarkower, 2000). In mouse and chicken, Dmrt1 was expressed in the genital ridge before sex differentiation (Raymond, Kettlewell, Hirsch,

Bardwell, & Zarkower, 1999). Dmrt1 may play a different role or perhaps be expressed at a different point in the chain of events during sex determination in reptiles and birds versus mammals. While Dmrt1 is expressed before sex differentiation in reptiles

(Kettlewell, Raymond, & Zarkower, 2000), in mice, Dmrt1 is initially expressed in bipotential gonads and later becomes male specific (Raymond, Kettlewell, Hirsch,

Bardwell, & Zarkower, 1999). A study conducted on embryonic gonads shows that embryos incubated at male producing temperatures had steadily increasing levels of

Dmrt1. Upon hatching, testes had 150-fold higher levels of Dmrt1 compared to ovaries

(Rhen T. , Metzger, Shroeder, & Woodward, 2007). Altogether, this information about

Dmrt1 suggests that it is an important, and conserved gene that is critical for development of testes. In addition to studying three genes (Sox9, Amh, and Dmrt1) that are classically associated with testis determination and sexual differentiation of the internal reproductive ducts, we examined a gene (FoxL2) that is typically linked to ovary determination.

FoxL2 expression was observed at equal levels in the pituitary and ovary and then in the hypothalamus and testes in decreasing order, respectively. This gene was originally discovered in mice and it exhibits a restricted expression pattern during the formation of the Rathke’s pouch which develops into the anterior pituitary (Treier, et al., 1998).

Further studies show FoxL2 is critical for normal development of the pituitary gland.

FoxL2 is expressed from just before mid-embryogenesis, stage 11.5, into adulthood in mice, which is suggestive of a maintenance role. Furthermore, there is evidence that

FoxL2 is involved in promoting differentiation in dividing cells (Ellsworth, et al., 2006).

Another study in which FoxL2 was selectively ablated by the Cre/lox approach in the anterior pituitary found that adult male spermatogenesis was impaired. In addition, adult females had smaller ovaries and fewer ovulated oocytes than controls. Both sexes were deficient for follicle stimulating hormone (FSH), caused by a deficiency of FSH subunit beta. This indicates that FoxL2 is required for gonadotrope development in the anterior pituitary (Tran, et al., 2013). Again, it would be curious to study male and female pituitary to determine whether there are sex differences in FoxL2 expression.

It is not surprising that FoxL2 expression was high in the ovaries. Studies in other animal models have shown that this gene is expressed in ovaries once ovarian fate has been determined (Loffler, Zarkower, & Koopman, 2003; Crisponi, et al., 2001; Kim, et

47 al., 2011). FoxL2 is also involved with ovarian maintenance and function in adulthood

(Pisarska, Bae, Klein, & Hsueh, 2004). No expression of FoxL2 was seen until morphological differentiation in mouse ovaries, while no FoxL2 was seen in developing male gonads at all. Expression in ovaries continued through embryonic development and past birth (Loffler, Zarkower, & Koopman, 2003). In fetal and mature mice, FoxL2 was found to be expressed in the germ cells of the ovary (Loffler, Zarkower, & Koopman,

2003). Our data is in agreement with research in mice. There is evidence of strong selective pressure on FoxL2 that keeps the gene similar in sequence across species

(Cocquet, et al., 2002).

High expression levels of FoxL2 in the hypothalamus was surprising and could be due to contamination from some pituitary gland tissues. However, there is little to no research on FoxL2 expression in the hypothalamus in other animal species.

Low expression of FoxL2 was seen in testes after 13.5 dpc and in adult testes in mice (Loffler, Zarkower, & Koopman, 2003). In humans and goats, FoxL2 protein was not observed at any stages of development in male gonads (Cocquet, et al., 2002). This is inconsistent with the data from the current study, as expression in male gonads was relatively high compared to tissues that do not express FoxL2 (kidney and intestine). This could be due to a possible difference in the rate of development of the testes in male mice versus male turtles. However, there is little information on FoxL2 expression in male gonads. FoxL2 mRNA expression was found to be influenced by incubation temperature in the common snapping turtle. Female producing temperatures had higher levels of

FoxL2 than male producing temperatures. Ovaries had 25-fold higher levels of FoxL2 mRNA compared to testes (Rhen T. , Metzger, Shroeder, & Woodward, 2007). FoxL2 is

48 a known marker for granulosa cell differentiation (Pisarska, Bae, Klein, & Hsueh, 2004), so it would be highly informative to understand its function in the male and female gonads of the common snapping turtle.

Finally, FoxL2 regulates follistatin transcription, which is expressed in many tissues of higher animals (Blount, et al., 2009). The snapping turtle Chelydra serpentina does in fact contain follistatin in its genome (T. Rhen, unpublished data), and further studies could reveal whether it is regulated by FoxL2. FoxL2, like Sox9, has also been found to play a role in skeletal development and cartilage formation in mice (Marongiu, et al., 2015).

It is known that Sst is expressed in the hypothalamus and that it plays a key role in regulating expression and secretion of growth hormone (Gh1) (Spiess, Villarreal, &

Vale, 1981; Schally, et al., 1980; Koerker, et al., 1974). Therefore, expression in the snapping turtle hypothalamus is expected. Expression in the enteric nervous system has also been reported in mammals (Corleto, 2010), which explains why it was found to be second highest in the intestine samples. It would be fascinating to disrupt Sst or its receptors in the common snapping turtle to see how it would affect intestine development or function.

As stated previously, Gh1 is produced in the somatotropes of the anterior pituitary

(Gahete, et al., 2009). Therefore, it is not surprising that expression of Gh1 was highest in the pituitary. Studies have shown that expression of Gh1 in humans and rat has a sexually dimorphic pattern. Male rats show discrete pulses, while females have fewer pulses, but higher levels between the peaks (Giustina & Veldhuis, 1998). It would be curious to study the expression pattern of growth hormone in turtles to see if there are differences

49 between the sexes. There is evidence that Gh1 has no affect in embryonic development

(Garcia-Aragon, et al., 1992), but there is also a study that counters this and found mRNA transcripts for Gh1 in the developing mouse embryo at the morula stage

(Pantaleon, et al., 1997). It would be interesting to determine if Gh1 is expressed embryonically in the common snapping turtle.

Intestine samples in this study were taken from the ileum, just before the start of the large intestine. Cdx1 has been shown to be localized to the epithelial nuclei of intestinal crypts in the adult, suggestive that it is associated with proliferating cells.

Staining was found to be strongest in the duodenum, and it was clear that a cephalocaudal gradient was established (Subramanian & Meyer, 1998). Due to the lack of research in reptiles, Cdx1 expression in this study has been compared to expression in mice. In normal adult mice, expression of Cdx1 is detected via Northern blotting and in situ hybridization in the intestine but not in any other tissues (Duprey, et al., 1988). This is inconsistent with expression patterns in the snapping turtle. However, qPCR is much more sensitive than nucleic acid hybridization-based methods, and that could explain the presence of Cdx1 in gonads in turtles, and not in mice. Or perhaps, expression in turtles is due to continued development and maturation of the gonads. Research in this area is again lacking.

It is important to note that in mammals, the intestine is not fully developed until animals are weaned. Perhaps this is also true in reptiles, however instead of suckling, hatchling turtles survive on residual yolk (Hewavisenthi & Parmenter, 2002). The hatchlings studied had not been fed and were thus surviving on nutrients from the internalized yolk. Perhaps Cdx1 expression would be different in fully mature versus

50 hatchling intestine. Knowing this gene is in fact expressed in reptiles is an excellent starting point to compare expression in mammals. The next step would be to measure expression in the intestine during development and in adulthood to see if the patterns are similar, and if Cdx1 plays a role in dictating cell differentiation and proliferation in the intestine. Any differences in expression could provide insight to the function of Cdx1 in reptiles. It would be beneficial to study Cdx1 localization throughout the reptilian intestine to see if expression occurs in specific cell types (i.e., Paneth cells) or in specific regions of the intestine along the anterior-posterior axis.

High Slc12a1 expression in the kidney was expected, as this gene is expressed in the loop of Henle in mammals (Greger & Schlatter, 1981). Unfortunately, no literature could be found about Slc12a1 expression in the gonads or other tissues. However, it is thought that the protein encoded by the Slc12a1 gene may influence blood pressure

(Lifton, Gharavi, & Geller, 2001). Perhaps it is used to aid in regulation of salt balance in other tissues as well. Expression was lowest in the intestine. Had a part of the large intestine been isolated instead, expression of Slc12a1 may have been higher, as this part of the intestine is associated with absorbing water and salts (Levitan, Fordtran, Burrows,

& Ingelfinger, 1962).

In summary gene expression was similar to mammals with a few exceptions.

Sox9 expression was found in the hypothalamus and pituitary glands in addition to male gonads. Amh expression was highest in male gonads, but there was also expression in female gonads. FoxL2 was found in the hypothalamus at similar levels to expression in the ovaries and was found at lower levels in the testes. Cdx1 was found at high levels in the intestine as expected, but had lower expression in the gonads, and was slightly higher

51 in testes compared to ovaries. Slc12a1 was high in the kidneys and was also found in the gonads at low levels.

Gene Pituitary Hypothalamus Testis Ovary Intestine Kidney Amh Off Low High Low Off Off Dmrt1 Low Off High Low Off Off Sox9 Medium High High Off Medium Off FoxL2 High Medium Low High Off Off Sst Off High Off Off Low Off Gh1 High Off Off Off Off Off Cdx1 Off Off Low Low High Off Slc12a1 Off Off Low Low Off High

Table 3: Summary of qPCR results for tissue specific expression in Chelydra serpentine in comparison to mammals. Bolded expression values indicate differences in expression compared to mammalian literature

Future work:

This research provides the foundation for studies of the epigenetic mechanisms underlying tissue specific expression in reptiles, especially during temperature-dependent sex determination. Future experiments to understand how tissue specific expression is regulated would involve DNase/MNase Foot Printing, Assay for Transposase-Accessible

Chromatin using Sequencing (ATAC-Seq), or Chromatin Immunoprecipitation followed by sequencing (ChIP-seq) to assess the role of chromatin structure. The first two methods examine whether the chromatin is tightly packed, or looser around the genes of interest to inhibit or allow transcription (Song & Crawford, 2010; Buenrostro, Giresi, Zaba, Chang,

& Greenleaf, 2013). ChIP-seq uses antibodies that target a specific histone modification

(Thurman, et al., 2012). For example, histone 3 on lysine 27 (H3K27) is associated with gene silencing when it is trimethylated (Lund & Lohuizen, 2004), while histone 3 lysine

4 (H3K4) trimethylation is associated with relaxed chromatin and gene transcription

(Barski, et al., 2007). Understanding how genes are silenced or activated in certain tissues can provide insight into basic mechanisms of cell differentiation and development. With increased knowledge of the mechanics of this process, developmental disorders could be better treated or perhaps even prevented.

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