Evolution of Mammalian Sex Chromosomes and Sex
Determination Genes: Insights from Monotremes
A thesis submitted for the degree of Doctor of Philosophy
Deborah Fernanda Toledo-Flores
School of Molecular and Biomedical Science
Discipline of Genetics
December 2014
Table of Contents
Table of Contents
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Abstract
Genetic sex determination systems are generally based on the presence of differentiated
sex chromosomes. Birds have a ZZ/ZW sex chromosome system in which males are ZZ
and females ZW, whereas mammals have an XX/XY system with males being XY and
females XX. Monotremes have an extraordinary sex chromosome system that consists
of multiple sex chromosomes: 5X5Y in platypus and 5X4Y in echidna. Intriguingly, the
monotreme sex chromosomes show extensive homology to the bird ZW and not to the
therian XY. However, sex determination in monotremes is still a mystery; the Yspecific Sry gene that triggers male sex determination in therian mammals is absent and
so far very few genes have been identified on Y chromosomes in monotremes. To gain
more insights into the gene content of Y-chromosomes and to identify potential sex
determination genes in the platypus a collaborative large scale transcriptomic approach
led to the identification of new male specific genes including the anti-Muellerian
hormone AMH that I mapped to Y5, this makes Amhy an exciting new candidate for sex
determination in monotremes.
Platypus chromosome 6 is largely homologous to the therian X and therefore it
represents the therian proto sex chromosome. In addition, this autosome features a large
heteromorphic nucleolus organizer region (NOR) and associates with the sex
chromosomes during male meiosis (Casey and Daish personal communication). I
investigated chromosome 6 heteromorphism in both sexes and found a number of sexspecific characteristics related to the extent of the NOR heteromorphism, DNA
methylation, silver staining patterns and interestingly, meiotic segregation bias. This
1
raises the possibility that chromosome 6 may have commenced differentiation prior to
monotreme therian divergence.
These results led me to investigate the chromosome 6 borne gene Sox3, from which Sry
evolved in therian mammals. This revealed a platypus male-specific Sox3 allele, which
differs from the alleles observed also in females on the length of one of the Sox3
polyalanine tracts. This raises the possibility that Sox3 may be working differently in
males and females.
We have used our unique knowledge of monotreme sex chromosomes to determine the
sex of captively bred echidnas. I used a PCR based genetic sexing technique that
utilizes DNA from small hair samples and primers that amplify male-specific genes.
Interestingly, I found that seven out of eight echidnas born in captivity were females.
Furthermore, I found a Sox3 deletion in the only male echidna born in captivity. This
gives us the unique opportunity to investigate the sexual development of an animal in
which this gene is naturally deleted providing an exceptional situation in which to study
monotreme sex determination. Furthermore, this sexing technique has the potential of
being applied in the wild to investigate sex ratio in natural populations of monotremes,
including the critically endangered long-beaked echidna.
2
Thesis Declaration
I certify that this work contains no material which has been accepted for the award of
any other degree or diploma in my name, in any university or other tertiary institution
and, to the best of my knowledge and belief, contains no material previously published
or written by another person, except where due reference has been made in the text. In
addition, I certify that no part of this work will, in the future, be used in a submission in
my name, for any other degree or diploma in any university or other tertiary institution
without the prior approval of the University of Adelaide and where applicable, any
partner institution responsible for the joint-award of this degree.
I give consent to this copy of my thesis when deposited in the University Library, being
made available for loan and photocopying, subject to the provisions of the Copyright
Act 1968.
The author acknowledges that copyright of published works contained within this thesis
resides with the copyright holder(s) of those works.
I also give permission for the digital version of my thesis to be made available on the
web, via the University’s digital research repository, the Library Search and also
through web search engines, unless permission has been granted by the University to
restrict access for a period of time.
Deborah Fernanda Toledo-Flores
20 – Dec – 2014
3
Acknowledgments
I would like to thank all the people that have supported me and guided me throughout
my Ph.D. studies. Thanks to my supervisor Prof. Frank Grützner for his constant advice
and support and to all the current and former members of the Grützner’s lab for being
excellent colleagues and friends. Thanks to our collaborator Prof. Henrik Kaessmann
and his group at the University of Lausanne, Switzerland, and to Prof. Vincent Harley
and his group at the Prince Henry’s Institute, Melbourne. Thanks to Dr. Dan Kortschak,
Prof. Jeremy Timmis and Prof. Jenny Graves for constructive discussion and feedback.
I would like to acknowledge the support of the Mexican National Council for Science
and Technology (CONACYT) for sponsoring me during my Ph.D.
Thanks to all the people that made this journey so enriching and enjoyable; the School
of Molecular and Biomedical Science students and staff, my friends and my boyfriend. I
want to particularly thank Nicole and Reuben for their support, feedback and for so
many good chats and lunch sessions. Also, thanks to Nicole for keeping my cookies in
her drawer!
Finally, I would like to specially thank my family because without them this would not
have been possible. Thanks to my mom, dad and my little brother for always being
there to support me, advice me, encourage me and inspire me. Thank you. – Finalmente,
quisiera agradecer muy especialmente a mi familia porque sin ellos, esto no hubiera
sido posible. Gracias a mi mamá, papá y a mi hermanito por siempre estar ahí para
apoyarme, aconsejarme, motivarme e inspirarme. Gracias. ¡Soy hija de mi apá y de mi
amá! ¡Y hermanita de mi hermanito!
4
CHAPTER 1: Introduction
This chapter consists of a conventional thesis introduction.
Chapter overview
In this chapter I present an overview of the evolution and function of sex determination
and sex chromosome systems in vertebrates, which is relevant for the work presented in
this thesis. I will discuss the complexity of the monotreme sex chromosomes and their
homology to the ZW-system in birds. In addition, I will discuss how the absence of Sry
in monotremes together with the lack of information on Y-specific genes leads to a gap
in the knowledge of sex determination in these animals. This will allow a better
perspective of how this work on monotremes contributes to our understanding of the
evolution of sex determination and sex chromosomes. In addition, I will briefly discuss
the relevance of these studies in monotreme conservation.
5
Introduction
This research project is based on the study of monotermes in order to obtain insights
onto the evolution of sex determination in the mammalian clade. This basal group of
mammals have amazed and intrigued scientists for more than 200 years due to their
fascinating appearance and extraordinary biology. Monotremes consists of the platypus
(Ornithorhynchus anatinus) and four extant species of echidnas (Tachyglossus
aculeatus, Zaglossus bruijni, Zaglossus attenboroughi and Zaglossus bartoni). They
diverged from therian mammals about ~200 million years ago [1], whereas mammals
diverged from birds and reptilians 315 million years ago (Fig. 1) [2]. Monotremes
possess several traits that establish them as mammals such as fur and milk-producing
mammary glands to feed their young [3], but they have also retained some features seen
in birds and reptiles such as oviparous reproduction and a single opening for the
intestinal, genital and urinary tracts [3].
Interestingly, this mixture of characteristics is also observed at the molecular level.
Similar to therian mammals monotremes have a XY male heterogametic system, which
is comprised by multiple sex chromosomes (male platypus have 5X5Y and male
echidna 5X4Y) [4, 5] that share homology to the bird sex chromosomes [4-6] and no
homology to the therian sex chromosomes. With such a complex system, it is unclear
what are the genes involved in triggering male sex determination in monotremes
representing an important gap in the knowledge of the evolution of mammalian sex
determination.
6
Figure 1. Variation in sex determination genes and sex chromosome systems among
amniotes. Divergence times between clades are represented in millions of years (Mya).
Homology relationships are represented in blue, green and orange. TSD: Temperature
sex determination. Croc: Crocodilians. Figure adapted from [7].
7
Sex chromosome evolution in vertebrates
Sexual reproduction is the predominant mode of reproduction in vertebrate species. It is
achieved through the combination of the genetic content of two gametes. Usually,
females are defined as the sex that produces the larger gamete (ovum) whereas males
produce the smaller gamete (sperm). Whether an organism develops as a female or as a
male is determined either by genetic or environmental variables. These mechanisms are
known as genetic sex determination (GSD) and environmental sex determination
(ESD), respectively [8]. GSD relies on the presence of one or more loci that drive the
sex determination process and which, in most cases, are located on sex chromosomes.
Heteromorphic sex chromosomes represent an amazing example of convergent
evolution that evolved independently in different lineages but share several common
features [9].
All mammals, birds and snakes rely on the presence of heteromorphic chromosomes to
determine male or female fate during development. Similarly, some reptilians,
amphibians and fish species also possess heteromorphic sex chromosomes, even though
ESD is equally common among these clades (Fig. 1). Mammals have an XY sex
chromosome system whereas birds and snakes have a ZW system. However, there is no
gene homology between the mammalian XY and the avian ZW [10]. Although both
birds and snakes have ZW systems, there is no gene homology between their sex
chromosomes [11]. The human X-chromosome is largely homologous to the chicken
chromosomes 1, 4 and 12 [12]; the chicken Z, to the human chromosomes 9 and parts of
the 5, 8 and 18 [10]; and the snake Z, to the human 3, 7, 10 and 17 and to the chicken 2
and 27 [11]. Another example of sex chromosome variation is the dragon lizard Pogona
vitticeps. This animal has a micro-ZW system that is not homologous to birds nor to
snakes sex chromosomes [13]. Interestingly, the gecko Gekko hokouensis also has a
8
ZZ/ZW female heterogametic system, which shares homology to the bird ZW [14].
However, it has been suggested that sex chromosomes in this gecko species evolved
only recently and independently from the bird sex chromosomes [15]. These different
examples illustrate how sex chromosomes have evolved several times in different
clades and even in different species.
Sex chromosomes evolved from a pair of autosomes
It has been proposed that these different sex chromosome systems evolved from
different pairs of autosomes from the common ancestor independently in each lineage
[16]. It has been suggested that the differentiation process between the proto-sex
chromosomes started when one of them acquired a sex-determining allele (Fig. 2). This
was subsequently followed by the accumulation of male-advantageous alleles
surrounding
the
novel
sex-determining
locus.
Furthermore,
a
chromosomal
rearrangement (for example an inversion), which might have occurred before or during
this process prevented recombination between the proto-X and proto-Y [17]. This
process was likely accelerated and exacerbated by the addition of one or more sexually
antagonistic genes to this area [9, 18]. A sexually antagonistic gene has contrasting
effects on the fitness of the two sexes [19], therefore, the gain of a gene that is
advantageous for males but disadvantageous for females in the rearranged fragment
could have favored selection against recombination [9]. Lack of recombination
consequently led to the degradation of the proto-Y, as it allowed mutations, deletions
and insertions to accumulate in the non-recombinant region. Therefore, once a mutation
arises in this area there is no possibility of reverting it due to the inability to repair a
non-recombinant region [17, 20]. This resulted in the differentiation of the ancestral
autosomal pair into heteromorphic sex chromosomes where the X largely maintained its
gene content while the Y underwent massive gene loss and heterochromatization.
9
Figure 2. Sex chromosomes evolved from a pair of autosomes in the common ancestor.
The differentiation process between the proto sex chromosomes (X and Y represented
in this figure) involved the acquisition of a sex determining allele as well as
accumulation of a number of male advantageous alleles on one of the chromosomes.
Additionally, suppression of recombination triggered the process of Y-chromosome
degradation. Figure adapted from [21].
10
Presumably, an analogous process led to the evolution of the ZW system in birds and
snakes. However, different pairs of autosomes became sex chromosomes independently
in each lineage (Fig. 3), which explains the lack of homology amongst the sex
chromosomes of different species.
There is extensive evidence that supports this theory of sex-chromosome evolution. Sex
chromosomes at different stages of differentiation have been identified. In birds, for
example, the Z chromosome is similar in size across most species while the Wchromosome varies significantly in size and heterochromatinization level [22-27].
Flightless ratites have a larger W chromosome that is similar in size to the Zchromosome, representing an early stage of sex chromosome evolution. In contrast,
among carnites the Z and W chromosomes are highly differentiated, with the W being
small and heterochromatic indicating a more advanced stage in evolution [28, 29]. A
similar scenario is also observed in snakes, where again the Z chromosome is similar in
size across species and the W varies greatly representing different stages of Wchromosome degradation [16].
Another piece of evidence supporting this theory is the existence of gametologous
genes. It is known that the X and Y gene content overlaps, thus it is possible to find a Xborne homologue for most of the genes located on the Y. The candidate
spermatogenesis gene Rbmy located on the Y chromosome of both humans and mice
evolved from the X-borne gene Rbmx [30-32]. Similarly Tspy represents a Y-borne gene
and Tspx is its X-borne counterpart [33]. Another example is the sex-determining region
Y gene (Sry), which is on the Y chromosome of therian mammals and triggers male sex
determination in these organisms. Sry evolved from its X-borne parent gene Sox3 [34].
These pairs of genes are called gametologous; they were homologous in the ancestral
autosomal pair and evolved independently due to suppression of recombination [35].
11
12
Figure 3. Sex chromosomes evolved from an ancestral pair of autosomes. This figure
represents the autosomes in the common ancestor and how the ZW from snakes, the
ZW from birds and the XY from therian mammals evolved from different autosomes,
whereas in birds and monotremes sex chromosomes evolved from the same ancestral
autosome.
13
Gametologous genes exist both in XY and in ZW systems. However, different pairs of
gametologous exist in different species. This is because each species might have lost a
different set of genes from the Y-chromosome, therefore, the content of sex
chromosomes that share ancestry might overlap but is not necessarily the same [21]. An
example of this is the X-borne gene Atrx, which is widely conserved among therian
mammals but only marsupials have a Y-borne Atrx gametolog, Atry [36]. However, it is
important to add that the degraded sex chromosomes, i.e. Y and W, carry other types of
genes too. Not all the genes on the Y chromosome are necessarily gametologous to Xborne gene. Y-chromosomes also carry amplicons of other genes present on the Y as
well as elements that arrived to this chromosome by retrotransposition from autosomes
[37]. The Y-borne gene Daz is an example of this. Daz is more closely related to an
autosomal gene and studies indicate that it was retrotransposed onto the Ychromosomes in the lineage of great apes [38, 39].
The therian XY sex chromosome system
Therian mammals (marsupials and placental mammals) have an XY/XX male
heterogametic system. The X chromosome is highly conserved in gene content and
order amongst placental mammals [40-42], except in the case of rodents in which the
gene order seems to be scrambled [43]. When comparing the gene content of the
placental X and the marsupial X it was discovered that the placental X is composed of
ancient and added regions [40]. These were called X-conserved region (XCR) and Xadded region (XAR), respectively (Fig. 4). The XCR is present both in marsupials and
placentals and it comprises the long arm of the therian X chromosome as well as the
region above the centromere [44]. The XAR includes the short arm of the placental X
and the pseudoautosomal regions. The XAR was found to be autosomal in marsupials
14
Figure 4. Human X and Y chromosomes. The human X chromosome consists of the
XCR region shared by all therian mammals and the XAR, which is absent in
marsupials. The X and Y chromosome retain homology in the PAR1 and PAR2, of 2.6
Mb and 320 kb respectively. In these regions the two chromosomes are able to pair and
recombine. Sry is located on the male specific Y region (MSY) and its X-borne parent
gene, Sox3, forms part of the XCR. The gray region on the Y chromosome represents
heterochromatin. Figure adapted from [40].
15
and completely represented in the chicken chromosome 1 [45]. Therefore, the XAR was
added to the placental X after their divergence from marsupials, around 100-180 million
years ago [46].
Similar to the X chromosome, the Y chromosome also possess a layered structure [47]
that reflects its evolutionary history. The Y chromosome bears five layers that go from a
high degree of divergence with respect to their X-borne gametologs (if any) to a lower
degree of divergence. The first layer, with the higher degree of divergence includes the
gene Sry, while the fifth layer is formed by the pseudoautosomal regions [47]. The
pseudoautosomal regions (PAR) are located on the tips of the X and Y chromosome.
These are areas of conserved homology between the X and the Y and consequently are
still able to pair and recombine during meiosis behaving as autosomal regions [48].
PARs are also considered evidence of the autosomal origin of sex chromosomes.
However, in therians they are part of the XAR, which means that they are absent in
marsupials [44, 49]. In humans and other great apes, PAR1 is located on the tip of the
short arm of the X and it covers 2.6 MB [50] whereas PAR2 is on the tip of the long
arm and is only 320 kb [51]. Interestingly, mice and rats have lost PAR1. Comparisons
of the gene content of the PARs amongst different mammalian species [52] have led to
the conclusion that in each species the homologous regions between the X and Y
chromosomes have reduced to a different extent due to independent Y-specific
mutations and rearrangement events in each lineage.
Given the process of degradation Y chromosomes have gone through, they are generally
much smaller than their X counterpart, which is very gene rich. For example, the human
X chromosome comprises 5% of the total genome and it contains around 1,000 genes
[53] with functions biased toward reproduction [54]. On the other hand, the human Y
chromosome represents only 2% of the genome and contains a few active genes [37].
16
The Y-chromosome is rich in pseudogenes and amplified copies, mainly due to the
suppressed recombination between X and Y [37]. From the 54 genes found on the Y
chromosome, 24 map to PAR1, 5 to PAR2 and 25 to the non-recombinant region [55],
called male specific Y region, i.e. MSY. This highly degenerated nature of Y
chromosomes in mammals has made sequencing this chromosome a technically
challenging task excluding them from most sequencing projects [56].
The prototherian multiple sex chromosome system
Similar to therian mammals, monotremes have a male heterogametic XY sex
chromosome system. However, this system is unique among mammals because it is
composed of multiple sex chromosomes that have no homology to the therian XY but
share extensive homology to the avian ZW system [6]. Echidna and platypus females
have ten X chromosomes that form five homologous pairs, whereas males have 9 and
10 sex chromosomes in echidna and platypus respectively [4, 5, 57]. During male
meiosis, the sex chromosomes form an alternating chain: X1Y1X2Y2X3Y3X4Y4X5Y5 in
platypus and X1Y1X2Y2X3Y3X4Y4X5 in echidna (Fig. 5d) [57], which segregates into Xbearing and Y-bearing sperm [4]. Sex chromosomes in the platypus chain pair through
the pseudoautosmal regions in an ordered fashion starting from the X5Y5 to the X1Y1
end [58]. PARs have been identified for all the chromosome pairing regions except for
the platypus X5Y5 [6], for the echidna X3Y3 and Y3X4 [57], possibly because they are
small chromosomes and the PARs would be very small as well.
Interestingly, echidna and platypus sex chromosomes are not completely homologous to
each other. Platypus chromosomes X1Y1X2Y2X3 are homologous to the echidna
X1Y1X2Y2X3, respectively. However, the platypus X5 is homologous to echidna X4, the
platypus Y5 is represented in the echidna Y3 and surprisingly the platypus X4 and Y3 are
17
represented in the echidna autosome 27 whereas the echidna Y4 and X5 are also
represented by autosomes in platypus (Fig. 5a,b) [57]. In addition to the changes on sex
chromosomes various autosomal rearrangements have occurred after the divergence of
platypus and echidna [57].
Monotremes share homology to the bird sex chromosomes and not to the therian XY [6,
57]. The XAR maps to the platypus chromosome 15 and 18 [6] and the XCR is
homologous to the platypus chromosome 6 [6, 59-61] (Fig. 5c) and echidna
chromosome 16. These chromosomes are interesting because they contain the gene
from which Sry evolved, Sox3 [60]. Interestingly, recent research (Casey and Daish
personal commnication) shows that chromosome 6 in the platypus stays in close
proximity to the sex chromosomes during prophase I in male meiosis. Additionally,
chromosome 6 is a remarkably heteromorphic autosome [57, 62] (Fig. 5c) and it has
been reported that length heteromorphism leads to segregation distortion [63].
Moreover, it has been suggested that non-random segregation during meiosis could
facilitate the transmission of sexually antagonistic alleles [64]. This warrants further
investigation of this proto-sex chromosome.
18
Figure 5. Monotreme sex chromosomes. (A) Echidna male sex chromosomes. The
numbers on the right of each chromosome indicate their homologs in the platypus. (B)
Platypus male sex chromosomes. The numbers on the right of each chromosome
indicate their homologs in echidna. (C) Platypus chromosome 6 is remarkably
heteromorphic as shown by G-banding. (D) Echidna male meiotic chain. The red signal
indicates the positions of the telomeres. Figure adapted from [57, 62].
19
Molecular aspects of mammalian sex determination
At the histological level, sex determination is highly conserved: testes are distinguished
by the presence of testis cords and ovaries are characterized by the presence of follicles
surrounding the developing eggs. The genes involved in the male and female sex
determination pathways are also highly conserved among vertebrates independently of
differences in the sex chromosome system or even in the type of sex determination, i.e.
GSD or ESD. These genes, however, show some variation in sequence, expression
patterns and timing. Particularly, the factor that triggers the sex determination cascade
varies greatly between species and it can be environmental or genetic. This section
describes some of the most relevant genes involved in the therian sex determination
pathway.
Sex determination genes in therian mammals
It has been shown that Sry (sex-determining region Y) acts as the testis determining
factor (TDF) in therian mammals [65]. This gene is the founder of the SOX family,
which is characterized by a high mobility group (HMG) domain. The HMG box is
responsible for DNA-binding, DNA-bending, protein interactions and nuclear import
and export among SOX genes [66]. Sry was discovered in mice and humans [65, 67]
over two decades ago. It was shown that replacement of the HMG box in the mouse Sry
for the HMG boxes of the SOX genes Sox3 or Sox9 was still able to induce female-tomale sex reversal when introduced into XX mice [68]. These findings show that the
HMG box from these different SOX genes are functionally interchangeable.
20
Sry is an intronless gene (except in marsupials) that encodes 204 amino acids from
which only the 80 that correspond to the HMG box are highly conserved among
therians, whereas the rest are highly divergent. As previously mentioned, Sry originated
from the intronless gene Sox3 [69-71]. Mutations in the human Sox3 cause X-linked
hypopituitarism, recessive hypoparathyroidism, mental retardation and significant
gonadal defects, including small testes [72, 73]. It has been shown that Sox3 expression
occurs predominately in type A undifferentiated spermatogonia and that loss of this
gene is associated with failure of the germ cell maturation. Thus Sox3 is directly related
to spermatogenesis [74]. Based on amino acid homology it has been proposed that a
portion of the Di-George syndrome critical region 8 (DGCR8) corresponding to its first
exon, was inserted upstream of the ancestral Sox3 on the proto-Y chromosome [75].
Therefore, Sry is a hybrid of a portion of DGCR8 and Sox3.
Sry spatial expression is specific to the bipotential gonad, where the process of sex
determination begins once the cells of the bipotential gonad commit to either male or
female development [76]. The bipotential gonad primordium originates from the
mesonephros, a primitive organ that functions as the embryonic kidney and forms part
of the developing urogenital system. In mouse, the urogenital system forms at 9-9.5
days post coitum (d.p.c.) and the bipotential gonad primordium at 10.5 d.p.c. Sry
expression starts in male-fated gonads and finishes at 12.5 d.p.c. [77] due to a negative
feedback loop by its target gene Sox9 [78, 79]. At this stage it is possible to distinguish
between a female and a male gonad because the male gonad is twice the size of the
female gonad and it has highly structured testis cords. These are composed of the male
supporting cells (Sertoli cells) and the primordial germ cells (PGC), and are surrounded
by the interstitium. By this time, the gonad in female is smaller and less organized than
in males as no structural tissue is observed [80]. Once the fate as male or female is
chosen, sex differentiation starts and the set of genes that will act in each specific sex
21
will repress the opposite sex pathway in order to reinforce the chosen one [76].
Hormones secreted from the supporting cells, such as the anti-Müllerian hormone
(AMH) in males, actively repress the opposite pathway.
Supporting cell precursors have their origin in the coelomic epithelium (CE) from cells
that express the steroidogenic factor 1 (Sf1) and migrate to the bipotential gonad. Sf1
expression starts in the CE during 9.5 d.p.c in both sexes [81]. It has been demonstrated
that Sf1 expression is essential for mice gonadal development in the two sexes [82],
however, humans heterozygous for Sf1 missense mutations show XY female sex
reversal highlighting its relevance for male sex determination [83]. The CE cells that
arrive to the bipotential gonad undergo an asymmetrical division in which one of the
daughter cells will turn into an interstitial cell and the other will become a supporting
cell precursor and will retain Sf1 expression [78, 84]. Once this decision has been taken
the supporting cell has to undergo a second decision, which is whether to become
Sertoli or granulosa cells. Sry expression at 10.5 d.p.c. leads to the supporting cell
precursors to become Sertoli cells.
Sry acts synergistically with Sf1 to activate Sox9 through the testis-specific enhancer of
Sox9 (TES) (Fig. 6) [85]. TES is a 3.2 kb regulatory motif located 13 kb upstream of the
Sox9 initiation site. TES contains a 1.9 kb region called TES core (TESCO) which is
sufficient for Sox9 gonadal specific expression. From these 1.9 kb, 1.4 kb are highly
conserved in human, dog and rat [85] and 180 bp are highly conserved in marsupials,
monotremes, birds, reptiles and amphibians [86]. This region is called the evolutionarily
conserved region (ECR) and it contains modules that predict regulatory roles for SOX,
TCF/LEF, Forkhead, DMRT and GATA proteins [86]. Therefore, it may have an
important role in the regulation of Sox9 across most vertebrates. After Sry expression is
repressed, Sf1 and Sox9 expression are highly maintained in Sertoli cells [81], which
22
means that Sox9 is regulated by other factors, such as Fgf9/Fgfr2 [76, 87] (Fig. 6).
SOX9 itself is very efficient binding TESCO in presence of SF1, both in vivo and in
vitro [85]. This suggests that SOX9 and SF1 might interact physically to regulate
TESCO, establishing a positive feedback loop. This idea has led to the proposition that
Sry evolved by mimicking Sox9 autoregulation [86]. Thus Sox9 expression consists of
three phases: initiation, promoted by Sf1 expression in the bipotential gonad;
upregulation, dependent on the synergistic action of SF1 and SRY; and maintenance
through autoregulation and positive feedback loops, such as the FGF9/FGFR2. 1,000
genes have been identified to act downstream of Sox9, from which many are
specifically expressed in Sertoli cells [88]. Those genes that are activated by SOX9
must be related to repression of bipotential gonad expressed-genes and female-pathway
genes [89].
It has been shown that Sox9 loss of function in male mice triggers male to female sex
reversal whereas gain of function in female mice triggers sex reversal in the opposite
direction [90, 91]. This shows that Sox9 is necessary for testis differentiation and that it
is the only gene downstream of Sry that is relevant for male sex determination [89]. Sry
might activate other genes beside Sox9, e.g. cerebelin4 (Cbln4), but their biological
function in testis differentiation is still unclear [92]. Another function preformed by Sry
is the inhibition of the !-catenin pathway, perhaps directly or through a downstream
target such as Sox9 [93]. Sry expression seems to be triggered by the genes Gata4 and
Wt1 as it has been demonstrated that they can act synergistically and activate human,
mouse and pig Sry promoters [94].
23
Figure 6. Sex determination in therian mammals. The Sox9 pathway, promoted by Sry
and Sf1, determines the formation of the male gonad and actively represses the female
pathway. Ovary development involves the activity of genes that actively repress the
male pathway, such as !-catenin. !-catenin expression is stabilized by Wnt4 and Rspo1
activity, and it represses Sox9 expression. Foxl2 is expressed after birth and it also
represses Sox9 activity. Figure adapted from [89].
24
In mice Sox3 is also expressed in the somatic cells of the embryonic gonad (overlapping
with Sry expression) as well as in the Sertoli cells of adult testes [34, 95]. Interestingly,
it was found that overexpression of Sox3 in the bipotential gonad of XX mice induces
female-to-male sex reversal [96]. It has been shown that this Sox3-mediated sex reversal
requires Sf1 expression and occurs only in a normal Sox9 context, as is the case of Sry
induced sex determination [96]. This implies that Sry and Sox3 act through a similar
mechanism and therefore are functionally interchangeable, suggesting that the
modifications that gave rise to Sry should have conferred it a specific early gonad
expression.
The time at which Sry is activated is crucial for its activity as testis determining factor.
It has been proven that induced Sry expression in XX mice after 11.5 d.p.c. fails to
induce complete female to male sex reversal [97, 98]. In the female pathway, formation
of the Müllerian duct starts between 11.5 and 12.5 d.p.c. through an invagination of the
surface epithelium of the mesonephros. The Müllerian duct gives rise to the oviduct,
uterus and the upper part of the vagina. In males, SOX9 and SF1 interact physically to
activate the anti-Müllerian hormone gene (Amh) [99]. This hormone is secreted by fetal
Sertoli cells and it represses the formation of the Müllerian duct. Therefore, if Sry does
not trigger Sox9 expression on time the Müllerian duct formation commences and it
results in female development [98, 100]. Interestingly, some exceptions exist in other
animals. For example, in chickens and alligators Amh expression precedes Sox9; hence
it is not possible that Sox9 activates Amh in these organisms [101, 102]. Moreover, it
has been shown that a male-specific duplicated copy of Amh is involved in male
development in the teleost fish Patagonian pejerrey (Odontesthes hatchery) [103],
which suggests some interesting alternative roles for Amh among vertebrates.
25
Despite the fact that most of the information presented in this section comes from
studies done in mice, the results can be extrapolated to other therian mammals with
some variations. For example, Sry in humans and pigs is expressed both during the
postnatal stage as well as during embryogenesis as in mice [104, 105]. Human Sry
expression is not restricted to Sertoli cells but it is also detected in germ cells [105].
However, the function of these genes is still conserved. For instance, human Sry and
goat Sry can promote male sex determination in mice under mouse regulatory sequences
[106, 107]. Another example is the previously discussed TES enhancer of Sox9 because
it contains the ECR region that is present in most vertebrates [86], suggesting a highly
conserved role.
Sex determination in monotremes
Monotremes have a unique reproductive biology that represents an amalgam of
mammalian and reptilian characteristics. Their female tracts are similar to those of
reptiles and marsupials; in the platypus the right ovary is inactive and the left is
functional similar to birds, whereas in echidna both ovaries are functional [108]. Males
have seminiferous tubules that resemble the structure of other amniote species, in which
at least four generations of germ cells are supported by a group of Sertoli cells.
Therefore, several stages of spermatogenesis can be visualized in a cross section of the
monotreme seminiferous tubules [108, 109]. Given their position as basal mammals and
their amazing mixture of features, monotremes are very important for comparative
genomic analysis to understand the evolution of sex determination and differentiation in
mammals.
There is nothing known about genes underlying sexual development in monotremes.
These prototherian mammals have an XY sex chromosome system in which males are
26
heterogametic. However, their multiple sex chromosomes are homologous to the bird
sex chromosomes. Several efforts have been made in order to identify orthologs to the
therian mammal and bird sex determination triggers in the platypus, such as Sox9,
Gata4, Wt1, Sf1, Atrx, Fgf9 and Wnt4; but their autosomal or pseudo-autosomal
locations [60, 110-112] excluded them from functioning as primary testis determining
factors. Therefore, the molecular bases underlying the initiation of sex determination in
monotremes are still unknown.
Dmrt1, the best candidate to be the bird sex determination trigger, has been mapped to
the platypus chromosome X5 and to the echidna X4 [4, 5, 57]. The location of this gene
in a X chromosome in monotremes might be the relic of a Dmrt1-controlled system
[113]. In birds, Dmrt1 is expressed in Sertoli and male germ cells [114] whereas in
mouse it is expressed in the supporting cells of both sexes [115, 116]. Dmrt1 expression
in
platypus
was
first
detected
only
in
adult
testis
through
RT-PCR.
Immunohistochemistry experiments showed that the DMRT1 protein is expressed in the
Sertoli cells as well as in granulosa cells, however lower in the later; hence, Dmrt1
expression is higher in males than in females but it is not sex-specific [111, 113]. These
findings imply that the function of Dmrt1 is highly conserved in the gonads of both
sexes in mammals, however, the presence of two Dmrt1 copies in male chicken versus
one copy in male platypus do not agree with a role as sex determining switch in
monotremes. However, we cannot exclude the possibility that a Dmrt1 Y-gametolog
might have diverged to an extent at which is no longer recognizable.
Several attempts have been made in order to identify Y-borne sex determining
candidate genes in monotremes. However, orthologs of genes involved in the therian or
bird sex determination pathways such as Sox9, Gata4, Wt1, Sf1, Atrx, Fgf9 and Wnt4
have been mapped to autosomal or pseudo-autosomal regions [60, 110-112] excluding
27
them from functioning as the primary testis determining factors. Sox9, which seems to
act universally in the vertebrate male sex determination pathway has the TES enhancer
that is highly conserved among marsupials, monotremes, birds, reptiles and amphibians
and it contains modules that predict regulatory roles for DMRT and SOX proteins,
among others [86]. The fact that the therian X-chromosome is represented by an
autosome in these animals led to the conclusion that Sry is absent in monotremes. This
was later confirmed by mapping Sox3 to chromosome 6 in the platypus [60]. However,
a different SOX protein might be still responsible of Sox9 activation. Efforts to identify
this putative monotreme sex determining candidate gene have been fruitless due to the
lack of Y-specific sequence, since the sequenced platypus genome comes from a female
[2]. Around 700 kb of platypus Y-specific sequence have been obtained [117]. Ychromosomes are the obvious male-specific niches in which we could find male sex
determination factors. Therefore, in order to facilitate the search for the monotreme sexdetermining gene it is necessary to extract and analyze more Y-specific sequence in
these animals. Moreover, investigating candidate genes belonging to the SOX family
could shed light on potential sex-determining triggers.
Sex ratios in captive animal populations
Sex ratio bias in populations of captive-bred animals has been described in several
mammalian species [118-120]. Even though the molecular mechanisms underlying this
phenomenon are still unknown, different theories have been proposed in order to
explain why this might happen [121-125]. In general, they all predict that skewing the
sex ratio should lead to an increase in the fitness of the mother [126]. There are a
number of examples where the factors influencing the sex ratio of captive animals are
known. For instance, it has been shown that group size affects the offspring sex in
lemurs; mothers housed in groups tend to produce more males whereas in isolation they
28
produce more females [118]. Another example is given by the captive pygmy
hippopotamus; these animals also show a female bias that has been related to high
levels of manipulation and feeding [119]. Accurately determining the sex ratio is a vital
part of animal biology in captivity as well as in the wild and it plays an important role in
conservation [127]. Early identification of sex ratio bias in captive-bred populations is
important for zoos in order to make decisions about housing animals as well as to
design their population management plans [128].
So far no data have been available about sex ratio bias of captive bred monotremes. In
echidnas identification of the animals sex is challenging due to the lack of obvious
sexual dimorphisms [129]. Therefore, developing our understanding of the genetics of
sex determination in monotremes could help us to develop genetic sexing techniques to
accurately and non-invasively identify the sex of captively bred echidnas and to
investigate whether they are affected by sex ratio distortions.
29
Aims of the Project
Monotremes occupy a key position in the evolution of mammals. In contrast to birds
and therian mammals, there is no information about the gene or group of genes that
trigger sex determination in these organisms. The aim of this study was to gain more
information about Y chromosomes and to identify potential sex-determining genes. The
first approach followed was to look at candidate regions, such as the Y chromosomes
and the autosome that shares homology to the therian X, chromosome 6 in the platypus.
By studying this autosome the aim was also to characterize the therian proto-sex
chromosome to obtain insights into the evolution of the XY system in mammals. The
next aim was to study candidate genes, particularly, Sox3 given its relationship to the
therian male sex-determining factor Sry. Finally, this study aimed to apply our
knowledge in monotreme sex chromosomes to aid zoos in genetically determining the
sex of captive bred echidnas and to investigate any potential sex ratio bias.
30
Summary
The thesis is divided in six chapters; the first chapter provides background information
on sex chromosomes and sex determination. In chapter 2, I present the results of the
first approach that was followed which consisted of identifying Y-chromosome genes.
Chapter 3 talks about the outcomes of investigating platypus chromosome 6, which was
chosen for analysis as this chromosomes shares extensive homology to the eutherian
mammal X. Chapter 4 focuses in the study of the candidate gene Sox3 – the Sry parent
gene in therian mammals – while Chapter 5 provides a practical application of these sex
determination studies onto ecology and conservation of monotremes. Finally, Chapter 6
summarises and discusses the major outcomes of work presented.
In chapter 2 I present the general approach, which was to gain more insight into gene
content of monotreme Y-chromosomes. Y-chromosomes are the most obvious place to
look for male-specific sex-determining genes given that they are male specific,
however, the platypus genome was obtained from a female and, as it is the case with
many other sequenced mammals, this meant that we were lacking information about Ychromosome genes. In order to obtain more information about genes on the platypus Ychromosomes I joined a collaboration with a research group in Lausanne, Switzerland;
our collaborators predicted a number of male-specific genes based on male and female
transcriptome data. I then experimentally validated this approach through PCR and
fluorescent in situ hybridization (FISH) experiments. I found that Amhy, predicted to be
in one of the platypus Y chromosomes, maps to the Y5. Since Y5 is proposed to be the
oldest Y-chromosome in the platypus and a Y-copy of Amh acts as the primary sex
determination trigger in the Patagonian pejerrey fish this renders Amhy as a likely sex
determination candidate in the platypus.
31
Chapter 3 describes my results regarding the second candidate region I investigated, the
platypus chromosome 6. This chromosome was of interest because it is homologous to
the therian X-chromosome and it associates with the monotreme sex chromosomes
complex at meiosis. I started by investigating the NOR heteromorphism of chromosome
6, showing that it is heteromorphic in males and females but it is more prominent in
females. I found a number of other sex-specific differences. For example, chromosome
6 is hypermethylated in females but not in males and the male NOR has a different
silver-staining pattern compared to the pattern observed in females. I next investigated
chromosome 6 segregated in male meiosis, finding that this autosome is non-randomly
segregated in X-bearing sperm; the short version of chromosome 6 segregates
preferentially with the X chromosomes. Further experiments suggested that this bias is
related to negative selection of X-bearing sperm carrying the long version of
chromosome 6. This is of interest since it has been proposed that non-random
segregation of autosomes could facilitate the transmission of sexually antagonistic
alleles and even participate in the evolution of new complex sex determination systems,
which raises the possibility that chromosome 6 may have commenced differentiation
prior to monotreme therian divergence.
In chapter 4 I focus on the Sox3 gene. This gene is located on the platypus chromosome
6 and, in therian mammals, Sry evolved from Sox3. Interestingly, I found that there is a
male specific variation of Sox3. This gene has four polyalanine tracts in humans, which
have been related to transcriptional activity. Expansions of as little as one residue in
polyalanine tracts have been associated with disease in humans. Platypus has only two
poly-alanine tracts and I found that the second poly-alanine tract (which corresponds to
the fourth polyalanine tract in humans) has a reduction of two residues exclusively in
males. I speculate that this variation may lead to a male-specific Sox3 function. In order
32
to answer this, I progressed to modify the mouse Sox3 to resemble platypus Sox3 and to
test its potential to trigger male sex-determination in vitro.
Chapter 5 describes work towards using our information of monotreme sex
chromosomes to assist the conservation and breeding of monotremes, particularly
echidnas. Early identification of the sex of these animals is important to zoos in order to
make captive management plans. However, due to the lack of obvious sexual
dimorphisms the only way to determine the sex of these animals is by palpation, which
is quite invasive for the echidna puggles. I successfully established a genetic sexing
technique, which allowed us to identify a striking bias in the sex ratio of these captively
bred animals. Also, through this technique I identified a deletion of Sox3 in the only
sexed male. This gives us the unique opportunity to investigate the sexual development
of an animal in which this gene is naturally deleted helping us to overcome the
limitations of genetically manipulating monotremes.
33
References
[1] Cortez D, Marin R, Toledo-Flores D, Froidevaux L, Liechti A, Waters PD, et al.
Origins and functional evolution of Y chromosomes across mammals. Nature.
2014;508:488-93.
[2] Warren WC, Hillier LW, Marshall Graves JA, Birney E, Ponting CP, Grutzner F, et
al. Genome analysis of the platypus reveals unique signatures of evolution. Nature.
2008;453:175-83.
[3] Griffiths M. The Platypus. Scientific American. 1988;285:84-91.
[4] Grützner F, Rens W, Tsend-Ayush E, El-Mogharbel N, O'Brien PC, Jones RC, et al.
In the platypus a meiotic chain of ten sex chromosomes shares genes with the bird Z
and mammal X chromosomes. Nature. 2004;432:913-7.
[5] Rens W, Grutzner F, O'Brien P C, Fairclough H, Graves JA, Ferguson-Smith MA.
Resolution
and
evolution
of
the
duck-billed
platypus
karyotype
with
an
X1Y1X2Y2X3Y3X4Y4X5Y5 male sex chromosome constitution. Proc Natl Acad Sci
U S A. 2004;101:16257-61.
[6] Veyrunes F, Waters PD, Miethke P, Rens W, McMillan D, Alsop AE, et al. Birdlike sex chromosomes of platypus imply recent origin of mammal sex chromosomes.
Genome Res. 2008;18:965-73.
[7] Wallis MC, Waters PD, Graves JA. Sex determination in mammals--before and after
the evolution of SRY. Cell Mol Life Sci. 2008;65:3182-95.
[8] Charlesworth B. The evolution of chromosomal sex determination. Novartis Found
Symp. 2002;244:207-19; discussion 20-4, 53-7.
[9] Ellegren H. Sex-chromosome evolution: recent progress and the influence of male
and female heterogamety. Nat Rev Genet. 2011;12:157-66.
[10] Nanda I, Zend-Ajusch E, Shan Z, Grutzner F, Schartl M, Burt DW, et al.
Conserved synteny between the chicken Z sex chromosome and human chromosome 9
34
includes the male regulatory gene DMRT1: a comparative (re)view on avian sex
determination. Cytogenet Cell Genet. 2000;89:67-78.
[11] Matsubara K, Tarui H, Toriba M, Yamada K, Nishida-Umehara C, Agata K, et al.
Evidence for different origin of sex chromosomes in snakes, birds, and mammals and
step-wise differentiation of snake sex chromosomes. Proc Natl Acad Sci U S A.
2006;103:18190-5.
[12] Kohn M, Hogel J, Vogel W, Minich P, Kehrer-Sawatzki H, Graves JA, et al.
Reconstruction of a 450-My-old ancestral vertebrate protokaryotype. Trends Genet.
2006;22:203-10.
[13] Ezaz T, Moritz B, Waters P, Marshall Graves JA, Georges A, Sarre SD. The ZW
sex microchromosomes of an Australian dragon lizard share no homology with those of
other reptiles or birds. Chromosome Res. 2009;17:965-73.
[14] Kawai A, Ishijima J, Nishida C, Kosaka A, Ota H, Kohno S, et al. The ZW sex
chromosomes of Gekko hokouensis (Gekkonidae, Squamata) represent highly
conserved homology with those of avian species. Chromosoma. 2009;118:43-51.
[15] Pokorna M, Giovannotti M, Kratochvil L, Kasai F, Trifonov VA, O'Brien PC, et al.
Strong conservation of the bird Z chromosome in reptilian genomes is revealed by
comparative
painting
despite
275
million
years
divergence.
Chromosoma.
2011;120:455-68.
[16] Ohno S. Sex Chromosomes and Sex linked Genes. Springer-Verlag: New York.
1967.
[17] Charlesworth B, Charlesworth D. The degeneration of Y chromosomes. Philos
Trans R Soc Lond B Biol Sci. 2000;355:1563-72.
[18] Innocenti P, Morrow EH. The sexually antagonistic genes of Drosophila
melanogaster. PLoS Biol. 2010;8:e1000335.
[19] Rice WR. Sex chromosomes and the evolution of sexual dimporphism. Evolution.
1984;38:735-42.
35
[20] Muller HJ. THE RELATION OF RECOMBINATION TO MUTATIONAL
ADVANCE. Mutat Res. 1964;106:2-9.
[21] Graves JA. Sex chromosome specialization and degeneration in mammals. Cell.
2006;124:901-14.
[22] Shetty S, Kirby P, Zarkower D, Graves JA. DMRT1 in a ratite bird: evidence for a
role in sex determination and discovery of a putative regulatory element. Cytogenet
Genome Res. 2002;99:245-51.
[23] Kasai F, Garcia C, Arruga MV, Ferguson-Smith MA. Chromosome homology
between chicken (Gallus gallus domesticus) and the red-legged partridge (Alectoris
rufa); evidence of the occurrence of a neocentromere during evolution. Cytogenet
Genome Res. 2003;102:326-30.
[24] Shibusawa M, Nishida-Umehara C, Tsudzuki M, Masabanda J, Griffin DK,
Matsuda Y. A comparative karyological study of the blue-breasted quail (Coturnix
chinensis, Phasianidae) and California quail (Callipepla californica, Odontophoridae).
Cytogenet Genome Res. 2004;106:82-90.
[25] Derjusheva S, Kurganova A, Habermann F, Gaginskaya E. High chromosome
conservation detected by comparative chromosome painting in chicken, pigeon and
passerine birds. Chromosome Res. 2004;12:715-23.
[26] Itoh Y, Arnold AP. Chromosomal polymorphism and comparative painting
analysis in the zebra finch. Chromosome Res. 2005;13:47-56.
[27] Raudsepp T, Houck ML, O'Brien PC, Ferguson-Smith MA, Ryder OA, Chowdhary
BP.
Cytogenetic
analysis
of
California
condor
(Gymnogyps
californianus)
chromosomes: comparison with chicken (Gallus gallus) macrochromosomes. Cytogenet
Genome Res. 2002;98:54-60.
[28] Marshall Graves JA, Shetty S. Sex from W to Z: evolution of vertebrate sex
chromosomes and sex determining genes. J Exp Zool. 2001;290:449-62.
36
[29] Takagi N, Sasaki M. A phylogenetic study of bird karyotypes. Chromosoma.
1974;46:91-120.
[30] Delbridge ML, Lingenfelter PA, Disteche CM, Graves JA. The candidate
spermatogenesis gene RBMY has a homologue on the human X chromosome. Nat
Genet. 1999;22:223-4.
[31] Mazeyrat S, Saut N, Mattei MG, Mitchell MJ. RBMY evolved on the Y
chromosome from a ubiquitously transcribed X-Y identical gene. Nat Genet.
1999;22:224-6.
[32] Tsend-Ayush E, O'Sullivan LA, Grutzner FS, Onnebo SM, Lewis RS, Delbridge
ML, et al. RBMX gene is essential for brain development in zebrafish. Dev Dyn.
2005;234:682-8.
[33] Delbridge ML, Longepied G, Depetris D, Mattei MG, Disteche CM, Marshall
Graves JA, et al. TSPY, the candidate gonadoblastoma gene on the human Y
chromosome, has a widely expressed homologue on the X - implications for Y
chromosome evolution. Chromosome Res. 2004;12:345-56.
[34] Collignon J, Sockanathan S, Hacker A, Cohen-Tannoudji M, Norris D, Rastan S, et
al. A comparison of the properties of Sox-3 with Sry and two related genes, Sox-1 and
Sox-2. Development. 1996;122:509-20.
[35] Garcia-Moreno J, Mindell DP. Rooting a phylogeny with homologous genes on
opposite sex chromosomes (gametologs): a case study using avian CHD. Mol Biol Evol.
2000;17:1826-32.
[36] Pask A, Renfree MB, Marshall Graves JA. The human sex-reversing ATRX gene
has a homologue on the marsupial Y chromosome, ATRY: implications for the
evolution of mammalian sex determination. Proc Natl Acad Sci U S A. 2000;97:13198202.
37
[37] Skaletsky H, Kuroda-Kawaguchi T, Minx PJ, Cordum HS, Hillier L, Brown LG, et
al. The male-specific region of the human Y chromosome is a mosaic of discrete
sequence classes. Nature. 2003;423:825-37.
[38] Saxena R, Brown LG, Hawkins T, Alagappan RK, Skaletsky H, Reeve MP, et al.
The DAZ gene cluster on the human Y chromosome arose from an autosomal gene that
was transposed, repeatedly amplified and pruned. Nat Genet. 1996;14:292-9.
[39] Yu YH, Lin YW, Yu JF, Schempp W, Yen PH. Evolution of the DAZ gene and the
AZFc region on primate Y chromosomes. BMC Evol Biol. 2008;8:96.
[40] Graves JA. The origin and function of the mammalian Y chromosome and Y-borne
genes--an evolving understanding. Bioessays. 1995;17:311-20.
[41] Raudsepp T, Lee EJ, Kata SR, Brinkmeyer C, Mickelson JR, Skow LC, et al.
Exceptional conservation of horse-human gene order on X chromosome revealed by
high-resolution radiation hybrid mapping. Proc Natl Acad Sci U S A. 2004;101:238691.
[42] Delgado CL, Waters PD, Gilbert C, Robinson TJ, Graves JA. Physical mapping of
the elephant X chromosome: conservation of gene order over 105 million years.
Chromosome Res. 2009;17:917-26.
[43] Bourque G, Pevzner PA, Tesler G. Reconstructing the genomic architecture of
ancestral mammals: lessons from human, mouse, and rat genomes. Genome Res.
2004;14:507-16.
[44] Glas R, Marshall Graves JA, Toder R, Ferguson-Smith M, O'Brien PC. Crossspecies chromosome painting between human and marsupial directly demonstrates the
ancient region of the mammalian X. Mamm Genome. 1999;10:1115-6.
[45] Nanda I, Shan Z, Schartl M, Burt DW, Koehler M, Nothwang H, et al. 300 million
years of conserved synteny between chicken Z and human chromosome 9. Nat Genet.
1999;21:258-9.
[46] Graves JAM. Breaking laws and explaining rules. Nature Genetics. 1996;12:121-2.
38
[47] Lahn BT, Page DC. Functional coherence of the human Y chromosome. Science.
1997;278:675-80.
[48] Helena Mangs A, Morris BJ. The Human Pseudoautosomal Region (PAR): Origin,
Function and Future. Curr Genomics. 2007;8:129-36.
[49] Toder R, Graves JA. CSF2RA, ANT3, and STS are autosomal in marsupials:
implications for the origin of the pseudoautosomal region of mammalian sex
chromosomes. Mamm Genome. 1998;9:373-6.
[50] Rappold GA. The pseudoautosomal regions of the human sex chromosomes. Hum
Genet. 1993;92:315-24.
[51] Freije D, Helms C, Watson MS, Donis-Keller H. Identification of a second
pseudoautosomal region near the Xq and Yq telomeres. Science. 1992;258:1784-7.
[52] Toder R, Glaser B, Schiebel K, Wilcox SA, Rappold G, Graves JA, et al. Genes
located in and near the human pseudoautosomal region are located in the X-Y pairing
region in dog and sheep. Chromosome Res. 1997;5:301-6.
[53] Ross MT, Bentley DR, Tyler-Smith C. The sequences of the human sex
chromosomes. Curr Opin Genet Dev. 2006;16:213-8.
[54] Saifi GM, Chandra HS. An apparent excess of sex- and reproduction-related genes
on the human X chromosome. Proc Biol Sci. 1999;266:203-9.
[55] Ross MT, Grafham DV, Coffey AJ, Scherer S, McLay K, Muzny D, et al. The
DNA sequence of the human X chromosome. Nature. 2005;434:325-37.
[56] Bachtrog D. Y-chromosome evolution: emerging insights into processes of Ychromosome degeneration.
[57] Rens W, O'Brien PC, Grutzner F, Clarke O, Graphodatskaya D, Tsend-Ayush E, et
al. The multiple sex chromosomes of platypus and echidna are not completely identical
and several share homology with the avian Z. Genome Biol. 2007;8:R243.
39
[58] Daish T, Casey A, Grutzner F. Platypus chain reaction: directional and ordered
meiotic pairing of the multiple sex chromosome chain in Ornithorhynchus anatinus.
Reprod Fertil Dev. 2009;21:976-84.
[59] Waters PD, Delbridge ML, Deakin JE, El-Mogharbel N, Kirby PJ, Carvalho-Silva
DR, et al. Autosomal location of genes from the conserved mammalian X in the
platypus (Ornithorhynchus anatinus): implications for mammalian sex chromosome
evolution. Chromosome Res. 2005;13:401-10.
[60] Wallis MC, Waters PD, Delbridge ML, Kirby PJ, Pask AJ, Grutzner F, et al. Sex
determination in platypus and echidna: autosomal location of SOX3 confirms the
absence of SRY from monotremes. Chromosome Res. 2007;15:949-59.
[61] Ezaz T, Stiglec R, Veyrunes F, Marshall Graves JA. Relationships between
vertebrate ZW and XY sex chromosome systems. Curr Biol. 2006;16:R736-43.
[62] McMillan D, Miethke P, Alsop AE, Rens W, O'Brien P, Trifonov V, et al.
Characterizing the chromosomes of the platypus (Ornithorhynchus anatinus).
Chromosome Res. 2007;15:961-74.
[63] Wang J, Chen PJ, Wang GJ, Keller L. Chromosome size differences may affect
meiosis and genome size. Science. 2010;329:293.
[64] Schwander T, Beukeboom LW. Non-random autosome segregation: a stepping
stone for the evolution of sex chromosome complexes? Sex-biased transmission of
autosomes could facilitate the spread of antagonistic alleles, and generate sexchromosome systems with multiple X or Y chromosomes. Bioessays. 2011;33:111-4.
[65] Sinclair AH, Berta P, Palmer MS, Hawkins JR, Griffiths BL, Smith MJ, et al. A
gene from the human sex-determining region encodes a protein with homology to a
conserved DNA-binding motif. Nature. 1990;346:240-4.
[66] Lefebvre V, Dumitriu B, Penzo-Mendez A, Han Y, Pallavi B. Control of cell fate
and differentiation by Sry-related high-mobility-group box (Sox) transcription factors.
Int J Biochem Cell Biol. 2007;39:2195-214.
40
[67] Gubbay J, Collignon J, Koopman P, Capel B, Economou A, Munsterberg A, et al.
A gene mapping to the sex-determining region of the mouse Y chromosome is a
member of a novel family of embryonically expressed genes. Nature. 1990;346:245-50.
[68] Bergstrom DE, Young M, Albrecht KH, Eicher EM. Related function of mouse
SOX3, SOX9, and SRY HMG domains assayed by male sex determination. Genesis.
2000;28:111-24.
[69] Stevanovic M, Lovell-Badge R, Collignon J, Goodfellow PN. SOX3 is an X-linked
gene related to SRY. Hum Mol Genet. 1993;2:2013-8.
[70] Foster JW, Graves JA. An SRY-related sequence on the marsupial X chromosome:
implications for the evolution of the mammalian testis-determining gene. Proc Natl
Acad Sci U S A. 1994;91:1927-31.
[71] Katoh K, Miyata T. A heuristic approach of maximum likelihood method for
inferring phylogenetic tree and an application to the mammalian SOX-3 origin of the
testis-determining gene SRY. FEBS Lett. 1999;463:129-32.
[72] Laumonnier F, Ronce N, Hamel BC, Thomas P, Lespinasse J, Raynaud M, et al.
Transcription factor SOX3 is involved in X-linked mental retardation with growth
hormone deficiency. Am J Hum Genet. 2002;71:1450-5.
[73] Rizzoti K, Brunelli S, Carmignac D, Thomas PQ, Robinson IC, Lovell-Badge R.
SOX3 is required during the formation of the hypothalamo-pituitary axis. Nat Genet.
2004;36:247-55.
[74] Raverot G, Weiss J, Park SY, Hurley L, Jameson JL. Sox3 expression in
undifferentiated spermatogonia is required for the progression of spermatogenesis. Dev
Biol. 2005;283:215-25.
[75] Sato Y, Shinka T, Sakamoto K, Ewis AA, Nakahori Y. The male-determining gene
SRY is a hybrid of DGCR8 and SOX3, and is regulated by the transcription factor CP2.
Mol Cell Biochem. 2010;337:267-75.
41
[76] Kim Y, Capel B. Balancing the bipotential gonad between alternative organ fates: a
new perspective on an old problem. Dev Dyn. 2006;235:2292-300.
[77] Hacker A, Capel B, Goodfellow P, Lovell-Badge R. Expression of Sry, the mouse
sex determining gene. Development. 1995;121:1603-14.
[78] Sekido R, Bar I, Narvaez V, Penny G, Lovell-Badge R. SOX9 is up-regulated by
the transient expression of SRY specifically in Sertoli cell precursors. Dev Biol.
2004;274:271-9.
[79] Chaboissier MC, Kobayashi A, Vidal VI, Lutzkendorf S, van de Kant HJ, Wegner
M, et al. Functional analysis of Sox8 and Sox9 during sex determination in the mouse.
Development. 2004;131:1891-901.
[80] Morrish BC, Sinclair AH. Vertebrate sex determination: many means to an end.
Reproduction. 2002;124:447-57.
[81] Ikeda Y, Takeda Y, Shikayama T, Mukai T, Hisano S, Morohashi KI. Comparative
localization of Dax-1 and Ad4BP/SF-1 during development of the hypothalamicpituitary-gonadal axis suggests their closely related and distinct functions. Dev Dyn.
2001;220:363-76.
[82] Luo X, Ikeda Y, Parker KL. A cell-specific nuclear receptor is essential for adrenal
and gonadal development and sexual differentiation. Cell. 1994;77:481-90.
[83] Lin L, Philibert P, Ferraz-de-Souza B, Kelberman D, Homfray T, Albanese A, et
al. Heterozygous missense mutations in steroidogenic factor 1 (SF1/Ad4BP, NR5A1)
are associated with 46,XY disorders of sex development with normal adrenal function. J
Clin Endocrinol Metab. 2007;92:991-9.
[84] Karl J, Capel B. Sertoli cells of the mouse testis originate from the coelomic
epithelium. Dev Biol. 1998;203:323-33.
[85] Sekido R, Lovell-Badge R. Sex determination involves synergistic action of SRY
and SF1 on a specific Sox9 enhancer. Nature. 2008;453:930-4.
42
[86] Bagheri-Fam S, Sinclair AH, Koopman P, Harley VR. Conserved regulatory
modules in the Sox9 testis-specific enhancer predict roles for SOX, TCF/LEF,
Forkhead, DMRT, and GATA proteins in vertebrate sex determination. Int J Biochem
Cell Biol. 2010;42:472-7.
[87] Kim Y, Bingham N, Sekido R, Parker KL, Lovell-Badge R, Capel B. Fibroblast
growth factor receptor 2 regulates proliferation and Sertoli differentiation during male
sex determination. Proc Natl Acad Sci U S A. 2007;104:16558-63.
[88] Nef S, Schaad O, Stallings NR, Cederroth CR, Pitetti JL, Schaer G, et al. Gene
expression during sex determination reveals a robust female genetic program at the
onset of ovarian development. Dev Biol. 2005;287:361-77.
[89] Sekido R, Lovell-Badge R. Sex determination and SRY: down to a wink and a
nudge? Trends Genet. 2009;25:19-29.
[90] Polanco JC, Koopman P. Sry and the hesitant beginnings of male development.
Dev Biol. 2007;302:13-24.
[91] Wilhelm D, Palmer S, Koopman P. Sex determination and gonadal development in
mammals. Physiol Rev. 2007;87:1-28.
[92] Bradford ST, Hiramatsu R, Maddugoda MP, Bernard P, Chaboissier MC, Sinclair
A, et al. The cerebellin 4 precursor gene is a direct target of SRY and SOX9 in mice.
Biol Reprod. 2009;80:1178-88.
[93] Tamashiro DA, Alarcon VB, Marikawa Y. Ectopic expression of mouse Sry
interferes with Wnt/beta-catenin signaling in mouse embryonal carcinoma cell lines.
Biochim Biophys Acta. 2008;1780:1395-402.
[94] Miyamoto Y, Taniguchi H, Hamel F, Silversides DW, Viger RS. A GATA4/WT1
cooperation regulates transcription of genes required for mammalian sex determination
and differentiation. BMC Mol Biol. 2008;9:44.
[95] Shen JH, Ingraham HA. Regulation of the orphan nuclear receptor steroidogenic
factor 1 by Sox proteins. Mol Endocrinol. 2002;16:529-40.
43
[96] Sutton E, Hughes J, White S, Sekido R, Tan J, Arboleda V, et al. Identification of
SOX3 as an XX male sex reversal gene in mice and humans. J Clin Invest.
2011;121:328-41.
[97] Vainio S, Heikkila M, Kispert A, Chin N, McMahon AP. Female development in
mammals is regulated by Wnt-4 signalling. Nature. 1999;397:405-9.
[98] Maatouk DM, DiNapoli L, Alvers A, Parker KL, Taketo MM, Capel B.
Stabilization of beta-catenin in XY gonads causes male-to-female sex-reversal. Hum
Mol Genet. 2008;17:2949-55.
[99] De Santa Barbara P, Bonneaud N, Boizet B, Desclozeaux M, Moniot B, Sudbeck
P, et al. Direct interaction of SRY-related protein SOX9 and steroidogenic factor 1
regulates transcription of the human anti-Mullerian hormone gene. Mol Cell Biol.
1998;18:6653-65.
[100] Chang H, Gao F, Guillou F, Taketo MM, Huff V, Behringer RR. Wt1 negatively
regulates
beta-catenin
signaling
during
testis
development.
Development.
2008;135:1875-85.
[101] Oreal E, Pieau C, Mattei MG, Josso N, Picard JY, Carre-Eusebe D, et al. Early
expression of AMH in chicken embryonic gonads precedes testicular SOX9 expression.
Dev Dyn. 1998;212:522-32.
[102] Western PS, Harry JL, Graves JA, Sinclair AH. Temperature-dependent sex
determination in the American alligator: AMH precedes SOX9 expression. Dev Dyn.
1999;216:411-9.
[103] Hattori RS, Murai Y, Oura M, Masuda S, Majhi SK, Sakamoto T, et al. A Ylinked anti-Mullerian hormone duplication takes over a critical role in sex
determination. Proc Natl Acad Sci U S A. 2012;109:2955-9.
[104] Parma P, Pailhoux E, Cotinot C. Reverse transcription-polymerase chain reaction
analysis of genes involved in gonadal differentiation in pigs. Biol Reprod. 1999;61:7418.
44
[105] Salas-Cortes L, Jaubert F, Barbaux S, Nessmann C, Bono MR, Fellous M, et al.
The human SRY protein is present in fetal and adult Sertoli cells and germ cells. Int J
Dev Biol. 1999;43:135-40.
[106] Lovell-Badge R, Canning C, Sekido R. Sex-determining genes in mice: building
pathways. Novartis Found Symp. 2002;244:4-18; discussion -22, 35-42, 253-7.
[107] Pannetier M, Tilly G, Kocer A, Hudrisier M, Renault L, Chesnais N, et al. Goat
SRY induces testis development in XX transgenic mice. FEBS Lett. 2006;580:3715-20.
[108] Grutzner F, Nixon B, Jones RC. Reproductive biology in egg-laying mammals.
Sex Dev. 2008;2:115-27.
[109] Lin M, Jones RC. Spermiogenesis and spermiation in a monotreme mammal, the
platypus, Ornithorhynchus anatinus. J Anat. 2000;196 ( Pt 2):217-32.
[110] Grafodatskaya D, Rens W, Wallis MC, Trifonov V, O'Brien PC, Clarke O, et al.
Search for the sex-determining switch in monotremes: mapping WT1, SF1, LHX1,
LHX2, FGF9, WNT4, RSPO1 and GATA4 in platypus. Chromosome Res.
2007;15:777-85.
[111] Tsend-Ayush E, Lim SL, Pask AJ, Hamdan DD, Renfree MB, Grutzner F.
Characterisation
of
ATRX,
DMRT1,
DMRT7
and
WT1
in
the
platypus
(Ornithorhynchus anatinus). Reprod Fertil Dev. 2009;21:985-91.
[112] Wallis MC, Delbridge ML, Pask AJ, Alsop AE, Grutzner F, O'Brien PC, et al.
Mapping platypus SOX genes; autosomal location of SOX9 excludes it from sex
determining role. Cytogenet Genome Res. 2007;116:232-4.
[113] El-Mogharbel N, Wakefield M, Deakin JE, Tsend-Ayush E, Grutzner F, Alsop A,
et al. DMRT gene cluster analysis in the platypus: new insights into genomic
organization and regulatory regions. Genomics. 2007;89:10-21.
[114] 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;461:267-71.
45
[115] Lei N, Karpova T, Hornbaker KI, Rice DA, Heckert LL. Distinct transcriptional
mechanisms direct expression of the rat Dmrt1 promoter in sertoli cells and germ cells
of transgenic mice. Biol Reprod. 2009;81:118-25.
[116] Pask AJ, Behringer RR, Renfree MB. Expression of DMRT1 in the mammalian
ovary and testis--from marsupials to mice. Cytogenet Genome Res. 2003;101:229-36.
[117] Kortschak RD, Tsend-Ayush E, Grutzner F. Analysis of SINE and LINE repeat
content of Y chromosomes in the platypus, Ornithorhynchus anatinus. Reprod Fertil
Dev. 2009;21:964-75.
[118] Perret M. Influence of Social Factors on Sex Ratio at Birth, Maternal Investment
and Young Survival in a Prosimian Primate. Behavioral Ecology and Sociobiology.
1990;27:447-54.
[119] Zschokke S. Distorted Sex Ratio at Birth in the Captive Pygmy Hippopotamus,
Hexaprotodon liberiensis. Journal of Mammalogy. 2002;83:674-81.
[120] Glatston AR. Sex ratio research in zoos and its implications for captive
management. Applied Animal Behaviour Science. 1997;51:209-16.
[121] Fisher RA. The genetical theory of natural selection. New York: Dover
Publications; 1930.
[122] Trivers RL, Willard DE. Natural selection of parental ability to vary the sex ratio
of offspring. Science. 1973;179:90-2.
[123] Myers JH. Sex Ratio Adjustment Under Food Stress: Maximization of Quality or
Numbers of Offspring? The American Naturalist. 1978;112:381-8.
[124] Gomendio M, Clutton-Brock TH, Albon SD, Guinness FE, Simpson MJ.
Mammalian sex ratios and variation in costs of rearing sons and daughters. Nature.
1990;343:261-3.
[125] Wiebe KL, Bortolotti GR. Facultative sex ratio manipulation in American
kestrels. Behavioral Ecology and Sociobiology. 1992;30:379-86.
46
[126] Thogerson CM, Brady CM, Howard RD, Mason GJ, Pajor EA, Vicino GA, et al.
Winning the genetic lottery: biasing birth sex ratio results in more grandchildren. PLoS
One. 2013;8:e67867.
[127] Lyttle TW. Segregation distorters. Annu Rev Genet. 1991;25:511-57.
[128] Faust LJ, Thompson SD. Birth sex ratio in captive mammals: Patterns, biases, and
the implications for management and conservation. Zoo Biology. 2000;19.
[129] Rismiller PD, McKelvey MW. Frequency of Breeding and Recruitment in the
Short-Beaked Echidna, Tachyglossus aculeatus. Journal of Mammalogy. 2000;81:1-17.
47
CHAPTER 2: Prediction of Y-linked genes in the platypus reveals Amhy as the
most likely sex-determining candidate gene
This chapter consists of one published paper.
Chapter overview
This paper describes work where a bioinformatics and molecular cytogenetic approach
was combined to identify and confirm novel Y-chromosome genes. This successfully
identified a large number of novel Y-borne genes across different mammalian species.
In the platypus, 25 Y-linked protein-coding genes were identified and I confirmed their
male-specific nature after establishing a novel FISH technique specifically to map short
PCR products. The establishment of this technique proved to be challenging and it
required several rounds of optimization. However, it provided a crucial result in this
study that was the physical mapping of Amhy onto platypus chromosome Y5. AMH is
known to repress the female developmental pathway in vertebrates and in some
organisms it acts as the primary sex-determining gene, rendering Amhy as the most
likely sex-determining gene in the platypus so far.
48
Publication:
Cortez, D., Marin, R., Toledo-Flores, D., Froidevaux, L., Liechti, A., Waters, P.D.,
Grutzner, F., and Kaessmann, H. (2014). Origins and functional evolution of Y
chromosomes across mammals. Nature 508, 488-493.
50
Cortez, D., Marin, R., Toledo-Flores, D., Froidevaux, L., Liechti, A., Waters, P.D.,
Grutzner, F., and Kaessmann, H. (2014). Origins and functional evolution of Y
chromosomes across mammals.
Nature, v. 508 (7497), pp. 488-493.
NOTE:
This publication is included between pages 50-51 in the print
copy of the thesis held in the University of Adelaide Library.
It is also available online to authorised users at:
http://dx.doi.org/10.1038/nature13151
CHAPTER 3: Non-random meiotic segregation of the therian proto sexchromosome in platypus may provide insights into differentiation of sex
chromosomes in mammals
This chapter consists of one manuscript in preparation.
Chapter overview
Platypus chromosome 6 shares extensive homology to the therian X chromosome and
hence represents the therian proto-sex chromosome. During meiosis the NOR,
chromosome 6 and the sex chromosome chain are closely associated. Chapter 3 and 4
describes work to investigate if chromosome 6 is segregated non randomly or shows
signs of sex chromosome differentiation.
Utilizing a number of molecular and cytogenetic techniques, I have identified sexspecific characteristics between chromosome 6 homologs. Moreover, I found that
chromosome 6 is non-randomly segregated during male meiosis. To our knowledge,
this is the first autosome shown to be non-randomly segregated during male meiosis in a
mammal. This may suggests that non-random segregation of chromosome 6 may have
facilitated its evolution as the therian sex chromosome.
51
Manuscript in preparation:
Deborah Toledo-Flores, R. Daniel Kortschak, Frank Grützner. Non-random
segregation of the therian proto-sex chromosome at platypus male meiosis.
53
Non-random segregation of the therian proto-sex chromosome at platypus male
meiosis
Deborah Toledo-Flores1, R. Daniel Kortschak1, Frank Grützner1
1
The Robinson Research Institute, School of Molecular and Biomedical Science, The University of
Adelaide, Adelaide, Australia
Abstract
Random segregation of chromosomes during meiosis is a widely accepted key aspect of
sexual reproduction. However there is increasing evidence that chromosomal
heteromorphisms can lead to non-random segregation at meiosis [1]. The platypus
features a large NOR heteromorphism on its singular NOR on chromosome 6. This
autosome shares extensive homology to the X chromosome of other mammals. Here we
investigated whether this NOR heteromorphism is associated with non-random meiotic
segregation of this proto-sex chromosome. First we determined that the heteromorphism
is found in both sexes although it is more prominent in females than in males. Our
results suggest that this is in part due to hypermethylation of the NOR region in females
but not in males. Importantly we found first evidence of non-random segregation of this
autosome in platypus male meiosis. Our data suggests that negative selection might act
against X-bearing sperm carrying the chromosome 6 with the large NOR. This work
provides a possible mechanism that may have influenced the evolution of chromosome
6 as a sex chromosome in therian mammals.
54
Introduction
The platypus is one of three extant monotremes. Its karyotype is arguably the most
unusual amongst mammals; with 21 pairs of autosomes [2] and an unusually large sex
chromosome complement that consists of X1X2X3X4X5Y1Y2Y3Y4Y5 in males and
X1X2X3X4X5X1X2X3X4X5 in females [3]. This sex chromosome complex shares
extensive homology with the birds Z chromosome [3-6] but not with the therian X [6,
7]. Instead, the therian X is largely homologous to platypus chromosome 6 [8-10].
Chromosome 6 is a very distinctive autosome carrying a large heteromorphic nucleolus
organizer region (NOR) located on its short arm [5, 10-12].
The NOR is the chromosomal locus around which nucleoli are formed [13, 14]. It
consists of clusters of tandemly repeated rRNA genes typically organized in a head-totail orientation [15-20]. Each rRNA gene codes for the 45S pre-rRNA precursor that is
subsequently cleaved forming the mature 18S, 5.8S and 25-28S (the exact size is
species-dependent), which will later assemble with the 5S into ribosome subunits.
rRNA gene expression is tightly regulated to ensure that the cell maintains a sufficient
amount of ribosome subunits to support protein synthesis levels during cell growth and
division [21-24].
In mammals, rRNA gene expression is controlled mainly through DNA methylation,
epigenetic modifications, number of rRNA genes and gene silencing due to positional
effects [25-30]; rRNA genes that have been hypermethylated and that are associated
with unacetylated histones are often inactive, whereas those that show DNA
hypomethylation coupled with histone acetylation are often active [28, 31]. Active
NORs form secondary constrictions in metaphase chromosomes. This is due to the
accumulation of RNA Pol I transcription factors on rRNA genes that were actively
55
transcribed during the previous interphase, which keeps them relatively decondensed
[13, 14, 32, 33]. These transcription factors contain acidic/argyophilic domains, hence
active NORs can be easily identified through silver staining [14, 34-39]. Differences in
NOR expression lead to both polymorphisms in the number of silver-stained NORs
(active NORs) [30, 40-46] as well as to NOR size heteromorphisms [47-51] between
individuals of the same species. Another factor influencing NOR size between
homologous chromosomes is unequal crossing over, since it can induce loss or tandem
duplications of rDNA regions [30, 52-54].
Interestingly, structural differences between homologous chromosomes, such as size
heteromorphism, have been shown to bias meiotic segregation [1]. In the nematode
Caenorhabditis elegans, for example, indels ranging from 1 kb to 7.3 Mb introduced in
any of its five autosomes produced a significant segregation bias; the shorter
chromosome in a homologous pair segregated preferentially with the X chromosome
whereas the longer segregated away from the X. They also showed a positive
correlation between indel size and transmission bias ratio [1].
In monotremes, both platypus and echidna, there are a number of autosomes that
present variation in size between homologous and between individuals [11, 12].
However, the platypus chromosome 6 bears the most remarkable heteromorphism given
its large NOR. In mammals NORs are commonly found in autosomes [55] with the
exception of marsupials [56] and some bats where NORs are on the sex chromosomes
[55]. Interestingly, in insects NORs are commonly found on sex chromosomes [57-61].
In this study we investigated whether this heteromorphism is sex-specific and whether it
is a result of differential DNA methylation or transcriptional activity. Importantly we
56
used this heteromorphism to trace chromosome 6 segregation during male meiosis and
found evidence of non-random segregation of chromosome 6 in sperm.
Results/Discussion
Chromosome 6 heteromorphism is observed in both sexes in platypus but it is more
prominent in females
Platypus chromosome 6 has been reported as a remarkably heteromorphic autosome
[11] however this observation has not previously been quantified or investigated
systematically in males or females. We first investigated whether this heteromorphism
was consistently found in both sexes. For accurate quantification we performed a twocolor FISH (fluorescence in situ hybridization) experiment utilizing chromosome 6specific BACs (bacterial artificial chromosomes) that flank the heteromorphic NOR
(Figure 1) to determine the length of this region on each homologous chromosome. The
measurements were normalized to different condensation levels by dividing each NOR
length by the average length of chromosome 1 in the corresponding metaphase.
This analysis was performed in fibroblast-derived metaphase spreads of three male and
female individuals each. 45-109 metaphases were scored per individual (Table 1), the
average length of each homologue was obtained and then a paired t-test was performed
to determine whether the population of long chromosome 6 had the same NOR mean
length as the population of short chromosome 6 in each individual. We found that the
lengths of the short and long chromosome 6 are significantly different in the 6 analyzed
individuals (Table 1). This shows that chromosome 6 is heteromorphic in both sexes.
57
We also analyzed the chromosome 6 length distribution. For this, we plotted the kernel
density distribution for the pooled short and long chromosome 6 of each individual. We
found that there is variation in the shape of the density distribution of different
individuals (Figure 2). Interestingly, in two out of three females we observed a bimodal
distribution (Figure 2A) whereas in males a shoulder-type distribution was observed in
two out of three cases (Figure 2B) but no bimodality was observed. The density graphs
thus suggest that the heteromorphism is more prominent in females than males.
Therefore, even though the difference in length between the short and long chromosome
6 is consistently significant in all the analyzed individuals (Table 1), the variation
between individuals and sexes results in different shapes of density distributions (Figure
2). This result raises the question of whether the variation can be attributed to a
difference in rDNA copy number or explained by a difference in condensation and
activity.
DNA hypermethylation contributes to the observed heteromorphism in females, but not
in males
In mammalian cells, extensive DNA methylation is a hallmark of silent rRNA gene
repeats [24, 26, 27] and it has been shown that inhibition of DNA methylation activates
rDNA transcription from previously silent cistrons [26]. Based on this, we decided to
investigate whether inhibition of DNA methylation had an impact on the observed
chromosome 6 heteromorphism. We treated male and female platypus fibroblasts with
5-aza-2’-deoxycytidine (5-aza-dC), a drug that inhibits cytosine methylation when
incorporated to DNA [26, 62]. Metaphase spreads were prepared from 5-aza-dC treated
cells and also from control untreated cells from the same individuals. The length of the
58
whole chromosome 6 was measured, as it was not possible to retrieve sharp and reliable
FISH signals in the metaphase spreads prepared from 5-aza-dC treated cells.
Consistent with our previous results, both male and female untreated cells showed a
statistically significant chromosome 6 heteromorphism (Figure 3). Interestingly, the
treated cells also showed a statistically significant heteromorphism in both sexes. We
then investigated whether there was any difference between the untreated short or long
chromosome 6 and their treated counter parts. Indeed the female long chromosome 6
remained unchanged whereas the short chromosome 6 was significantly elongated after
5-aza-dC treatment (Figure 3). In contrast, the 5-aza-dC treatment had no significant
effect in either chromosome 6 homologue in males (Figure 3). Therefore, only the
shorter female NOR is hypomethylated presumably resulting in reduction of its activity,
which may consequently increase condensation and contribute to the heteromorphism
observed in this sex.
It has been reported before that amongst homologous or non-homologous NORs, those
with the fewest ribosomal cistrons are preferentially inactivated [47, 50, 63, 64]. An
example of this is given by the preferential inactivation of the shorter NOR in interspecific hybrids [63]. Something similar may be occurring in the female NOR, where a
difference in the number of ribosomal cistrons on each homologous NOR might cause
the initial heteromorphism [52], and this could be exacerbated by preferential
inactivation of cistrons in the shorter NOR.
Silver staining shows different NOR activity between sexes
In order to investigate whether there are any sex-specific differences between the
activities of homologous NORs we performed silver staining of the proteins associated
59
with NOR activity on male and female platypus metaphase spreads. We found that in
females, both NORs are entirely covered by silver staining (Figure 4A). In males, we
observed that one of the homologous NORs bears a gap of unstained chromatin
surrounded by silver deposits over the rest of the NOR (Figure 4B). This is inconsistent
with the observation that demethylation affected the short female NOR and not the male
NOR. Therefore, the female short NOR must represent a mosaic of active and inactive
ribosomal cistrons [31, 63].
To investigate whether the gap in the male NOR is the result of localized rDNA
silencing or if the gap represents an area containing non-rDNA we performed a FISH
experiment using a 28S specific PCR probe (Figure S1A) on the same male. We
observed binding of the 28S probe to the entire NOR region with no evident gaps
(Figure S2 B,C). However, when we measured the intensity of the signal of the silver
staining and of the FISH we observed correspondence between the intensity plots for
both experiments (Figure S1 B,C). This is consistent with the observations of other
researchers that found that the size of the rDNA FISH signal is associated with the size
of the silver staining signal [65, 66]. Our FISH results suggest that the silver staining
gap observed in the male NOR might contain a lower number of rDNA genes as
indicated by the drop in FISH signal intensity. Alternatively, a mechanism other than
DNA methylation might be involved in the inactivation of the rDNA contained in this
region, as shown by silver staining. In horse, for example, it has been observed that
NOR hypomethylation can occur together with lack of silver staining deposits [30],
showing that DNA methylation does not always correlate with NOR activity.
60
Chromosome 6 heteromorphism is present in uncultured meiotic cells in platypus
In order to investigate segregation bias of the heteromorphism at meiosis we first had to
determine whether the different chromosome 6 homologs could be identified in sperm
nuclei. We measured the distance between NOR flanking BAC clone hybridization
signals in platypus sperm cells. The density distribution showed a clear bimodality
(Figure 5A). In addition, we analyzed this set with Bayesian Information Criterion
(BIC), which estimates the number of clusters that best represent the data. This analysis
indicated that the chromosome 6 measurements obtained from sperm are best
represented by two clusters (Figure 5B). This shows that the heteromorphism is not a
cell culture artifact and can be observed in uncultured cells. Importantly this allows us
to investigate segregation bias of chromosome 6 at male meiosis.
The short chromosome 6 segregates preferentially with the X chromosomes at platypus
male meiosis.
Heteromorphisms have been shown to bias meiotic segregation of autosomes with sex
chromosomes. In monotremes the large NOR associates closely with the sex
chromosomes complex (Casey and Daish, personal communication) raising the
possibility of non-random segregation. To identify X and Y bearing sperm we
performed a three-color FISH experiment; we used two chromosome 6 specific BACs
(Figure 6) and a third X- or Y-specific BAC. We then counted the number of X- or Ybearing sperm carrying the long and short versions of chromosome 6. We found that the
long and short chromosome 6 were randomly segregated in Y-bearing sperm (Table 2,
Figure7). Interestingly, we observed a significant bias in X-bearing sperm, which in
most cases carried the short version of chromosome 6. In the three individuals we
observed between 66 and 78% of the X-bearing sperm carrying the short version of
61
chromosome 6 and only between 21 and 33% carrying the longer version (Table 2,
Figure 7).
As previously discussed, a similar scenario was described before in C. elegans, where
length variations of autosomes affected their segregation. Consistent with our results in
the platypus, this study showed that the shorter version in a homologous pair segregated
preferentially with the X chromosome whereas the longer version segregated against it
[1]. Importantly, this study also showed that indels of as little as 1 kb were able to
induce the distortion, therefore, it is possible that even a subtle difference in the number
of ribosomal cistrons may result in a segregation bias in the platypus.
Depletion of X-bearing sperm with the long chromosome 6
The finding that there is a greater frequency of Y-bearing sperm compared to X-bearing
sperm (Table 3, Figure 8) might suggests that this is a case of gamete competition,
where a significant proportion of those male gametes carrying the X and the long
version of chromosome 6 must be somehow eliminated. If this were the case we would
expect to see a bias in X/Y sperm frequency. In order to investigate this, we counted the
frequency of X/Y sperm on the same three individuals in which we observed the
segregation bias. For this we performed sequential one-color FISH experiments. We
firstly performed FISH utilizing a red-labeled Y5 BAC (Figure 8A); we counted the
number of observed Y5 positive and negative sperm and recorded the location of the
scored cells. In the second FISH we used a X1 BAC also labeled in red (Figure 8B) and
we counted the number of X1 positive cells. As expected, most of the cells that were
negative for Y5 in the first FISH were positive for X1 in the second FISH. In the first
experiment we observed an overrepresentation of Y-bearing sperm (Figure 8C).
Similarly, we observed the same bias while comparing the percentage of X-bearing
62
sperm from the second experiment to the percentage of Y-bearing sperm from the first
FISH (Figure 8D). In the three studied males around 60% of the scored sperm were Y
positive, 30% were X positive and 10% of the sperm were uninformative with no
visible signal (Table 3). This supports the idea that there is a bias towards Y-bearing
sperm and this is consistent with our hypothesis that a proportion of X-bearing sperm
may be reduced by the removal of cells that bear the X and the long chromosome 6. We
evaluated our results under three different assumptions; (1) the total of X-sperm is
represented by the sum of X-positive and uninformative sperm, (2) Y sperm is
represented by the sum of Y-positive sperm and uninformative sperm and (3) we
assume that uninformative sperm represent cases in which either the X1 or Y5 were lost
in the scored sperm (Figure S2). In all these possible scenarios it is evident that the X/Y
bias exists in platypus sperm.
To explain the length distributions observed in males and females (Fig. 2) it is
necessary to consider the mode of inheritance of chromosome 6 from females (Fig. 9).
It is likely that females transmit both types of chromosome 6 (Fig. 9C). However, given
that no males were identified with a bimodal distribution of the chromosome 6 types,
there should be a mechanism eliminating or preventing presence of the two variants in
males. This might be through pre-zygotic mechanisms such as incompatibility between
oocytes carrying the short chromosome 6 and Y-bearing sperm carrying the long
chromosome 6 as well as between oocytes carrying the long chromosome 6 and Ybearing sperm carrying the short chromosome 6. Alternatively, XY zygotes carrying the
long and short chromosome 6 might be eliminated post-fertilization or these males may
have increased mortality (post-zygotic).
If this were the case, these mechanisms could also help to balance the ratio of males and
females. Given the fact that there is a bias towards Y-bearing sperm in the platypus
(Fig. 8), if no compensatory mechanisms exist there would be more males than females
63
in the population, which could put at risk this species and eventually lead to extinction
[67]. However, if there were increased mortality (or gamete incompatibility) of XY
males carrying the short and long chromosome 6, this could help to balance the ratio of
males and females. Future work may reveal more information about males and females
ratios in the wild to obtain and provide insights on whether the observed segregation
distortion and X/Y sperm bias has an impact on the platypus population.
Conclusion
Our findings show that the platypus chromosome 6, which is the proto-sex chromosome
in therian mammals, presents a number of sex-specific characteristics; its NOR is
heteromorphic predominantly in females, demethylation has an effect in females but not
males and there is a differential silver staining pattern in both sexes that suggests sex
specific NOR activity patterns. Importantly chromosome 6 is non-randomly segregated
in X-bearing sperm. Such sex-biased segregation of autosomes could facilitate the
transmission of sexually antagonistic alleles and even participate in the evolution of
new complex sex determination systems [68]. At this stage, we do not know whether
there is any relationship between our observations and the possible evolution of
chromosome 6 as a sex chromosome. However, to the best of our knowledge, this is the
first time non-random segregation of an autosome that has evolved as a sex
chromosome has been reported.
64
Materials and methods
BAC clones
BAC clones were obtained from the CUGI BAC/EST resource centre (Clemson, SC,
USA) from the library Oa_Ab or from the Children’s Hospital Oakland Research
Institute (CHORI, CA, USA) from the library CH236.
Preparation of chromosomes and sperm cells
Mitotic metaphase spreads were produced from established platypus fibroblast cell-lines
[3, 69-71]. Established sperm and meiotic cell suspensions were used [69]. For new
preparations, cells were obtained from extracted testes; the cells were then directly fixed
in methanol/acetic acid (3:1). Slides were prepared according to standard procedures.
Demethylation assay
Established cell lines were cultured in 50% Dulbecco’s Modified Eagle Medium High
Glucose 1x (Life technologies) supplemented with glutamine, heat-inactivated FCS and
Penicillin/Steptomycin; and 50% AmnioMAX-C100 (1X) (Life Technologies) at 32º.
Demethylation was performed for 72 hrs at a concentration of 15 µM 5-aza-dC. For
chromosome preparation information refer to Preparation of chromosomes and sperm
cells section.
Fluorescence in situ hybridization
BAC DNA (1 µg) was labeled with spectrum orange or green (Abbot Molecular) using
random primers and Klenow polymerase and hybridized by fluoresence in situ
hybridization (FISH) on female and male metaphase cells according to standard
procedures previously described in [69]. For FISH on sperm cells, one step was
introduced into the standard procedure; after pepsin treatment (10 min incubation in
65
0.01 % pepsin in 10 mM HCl at 37ºC), sperm slides were incubated in 8 mM DTT/PBS
at 37ºC for 8 min. Followed by the standard 10 min re-fixation in 1 x PBS, 50 mM
MgCl2, 1 % formaldehyde. Slides were then dehydrated in ethanol series followed by a
5 min denaturation at 75ºC in 70% formamide, 2x SSC and again dehydrated. Slides
were hybridized overnight in a moist chamber at 37ºC with a mix of DNA probe, 10-20
µg of sonicated genomic DNA and 50 µg of salmon sperm redissolved in 50%
formamide, 10% dextran sulfate, 2x SSC. Prior to hybridization the probe mix was
denatured for 10 min at 80ºC and preannealing of repetitive DNA sequences was
performed for 30 min at 37ºC. Post-hybridization, the slides were washed three times
for 5 min in 50% formamide, 2x SSC at room temperature followed by one wash for 5
min in 2x SSX at 42ºC, once for 5 min in 0.1x SSC at 60ºC and finally once for 5 min
in 2x SSC at 42ºC. Cells were counterstained with 1 µg/µl DAPI in 2x SSC for 1 min
and then mounted with one drop of Vectashield. Images were taken using a Zeiss
AxioImagerZ.1 epifluorescence microscope and the Zeiss Axiovision software.
Chromosome measurement and data analysis
In metaphase and sperm cells, FISH was performed with two chromosome 6 specific
BACs and the distance between the signals was measured. In 5-aza-dC treated
metaphase cells, the length of the whole chromosome was measured instead.
In both cases, measurements were taken with Olympus AnalySIS software by manually
drawing a line from one BAC signal to the other or from one end of the chromosome to
the other (5-aza-dC treated-cells). As described in [69], a macro was written to
incorporate information about the magnification to calibrate the measurements.
In metaphase spreads, an average length of the two chromatids was contained for each
chromosome followed by normalization for different levels of condensation.
Normalization was carried by dividing the chromosome length by the average length of
chromosome 1, which was used because it can be easily identified by eye.
66
In sperm cells, normalization was carried by dividing the distance between the
chromosome 6 signals by the total length of the sperm since it was previously shown
that chromatin extension depends on sperm elongation [69].
Measurements were processed using perl (v5.10.0) and statistical analyses were carried
in R (version 2.11.0). Clustering and BIC analyses were performed with the R package
mclust (version 3.4.8).
Silver staining
Silver staining on metaphase spreads was carried following standard procedures.
Briefly, two drops of gelatin solution (2% w/v gelatin / 1% v/v formic acid) plus two
drops of silver nitrate solution (50% w/v silver nitrate) were placed onto metaphasespread slides. Solutions were mixed by gently tilting the slide. Slides were then covered
with cover slips and placed on a 70ºC hotplate for approximately 2 min, or until golden
brown. Slides were mounted with a drop of Vectashield.
Acknowledgements
Thanks to Dr. Jonathan Tuke for advice in statistical analysis and to Aaron Casey for
very useful discussions and for providing a chromosome 6 BAC. Thanks to Prof.
Jeremy Timmis for very constructive feedback and for providing the used rDNA
primers. DT-F is funded by the National Council of Science and Technology from
Mexico (CONACYT). FG is an ARC research fellow.
Author contributions: DT-F performed sample processing, laboratory work and wrote
the manuscript. RDK helped in data interpretation and statistical analysis. FG
contributed to the design and supervision of the project as well as to the manuscript
preparation and data interpretation.
67
68
Figures
Figure 1. Two colour FISH with chromosome-6 specific BACs on platypus
metaphase spread.
(A) FISH on platypus metaphase cell with chromosome 6 specific BACs in red
(849O23) and green (171L02) hallmarking the heteromorphic NOR region. White
brackets indicate the length of the NOR. (B) Platypus chromosome 6 and chromosome
1 homologues. Brackets indicate how the measurements were performed.
69
Figure 2. Density distribution of chromosome 6 length measurements in metaphase
spreads of 3 males and 3 females.
The length of both chromosome 6 homologues was measured in a number of cells
(Table 1) and the density distribution for each individual was plotted. (A) Density plots
for three different females show variation but clear bimodality in individuals 1 and 3.
(B) Density plots for three different males show variation and a “shoulder” type of
distribution instead of the clear bimodality observed in females.
70
!"#$%&#'()*+,-./0#1/0)2345)(6#)/#)*(#/46(27(8#*()(2/'/29*36'#30
:('-,(6;#45)#0/)#30#'-,(6##
Effect of 5-aza-dC treatment on platypus chromosome 6
===#
!"*!#
Chromosome 6 – Normalized length
===#
===#
!"+!#
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,-.)/1#
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<3=52(#>"#?@-A-@8B#)2(-)'(0)#/:#9,-)+956#15,)52(8#C42/4,-6)6#(,/0=-)(6#)*(#:('-,(#6*/2)#1*2/'/6/'(#D"#
73:->65?@9# A5B# C9-CDC:3;# C9# E65:>EFB# GD.@D65B:B# C9# HF6:F.3"# (/5I5/;,# :.35:439:# BCJ9CGH59:6># 36@9J5:3B# :-3# BFigure 3. 5-aza-dC treatment of platypus cultured fibroblasts elongates the female
H-.@4@B@43#)#C9#K34563BL#A-3.35B#C:#-5;#9@#3M3H:#C9#:-3#K34563#6@9J#H-.@4@B@43#9@.#C9#4563#H366B"#<.35:439:#K5C63;
3.5B3# :-3# @DB3.N3;# -3:3.@[email protected]# BC9H3# :-3# 6@9J# 59;# B-@.:# H-.@4@B@43# )# 5.3# BCJ9CGH59:6># ;CM3.39:# C9# D@:-# B3O3
short chromosome 6.
:.35:3;#59;#F9:.35:3;#H366B"#P9#:-CB#J.5E-L#:-3#435BF.3439:B#A3.3#[email protected];#:@#:-3#B-@.:#F9:.35:3;#H-.@4@B@43#@9#3
B3O"#
Methylation was inhibited in platypus fibroblasts in culture. 5-aza-dC treatment
significantly elongates the short chromosome 6 in females (n = 127), whereas it had no
effect in the female long chromosome nor in male cells (n = 92). Treatment failed to
erase the observed heteromorphism since the long and short chromosome 6 are
significantly different in both sexes in treated and untreated cells.
71
Figure 4. Silver staining on female and male platypus metaphase spreads.
Panel (A) shows silver staining performed on a female metaphase spread, (B) shows the
same experiment on a male metaphase spreads. NORs are indicated by black arrows in
both cases. In the female we observed an uniform staining of the NOR as indicated both
in panel A and in C, where we show different examples of female chromosome 6 pairs
with a uniform silver staining on the NOR. In males, however, we can see both in panel
B and in the different examples depicted in panel D that while one of the NORs is
uniformly stained with silver, the second one presents a gap in the middle of the NOR.
72
Figure 5. Chromosome 6 heteromorphism is observed in platypus sperm.
(A) Density plot of chromosome 6 length measured in 127 platypus sperm cells of one
individual. The bimodality of this graph represents the existence of two populations of
chromosome 6 in platypus sperm. (B) Bayesian Information Criterion (BIC) estimates
the number of clusters that best represent the data. In this case, 2 clusters represent
chromosome 6 data better than 1 cluster. This is true assuming both equal (E) and
different (V) variance.
73
Figure 6. Three-color FISH was used to study chromosome 6 segregation in male
meiosis.
FISH was performed on platypus sperm using a green-labeled BAC (171L02) specific
to the long arm of chromosome 6 and a red-labeled BAC (849O23) specific to the short
arm. Either X (A) or Y (B) specific BACs (456I13 or 456I13, respectively) were labeled
in yellow (mix of green and red). The distance between the two chromosome 6 BACs
was measured, if it was above the individual’s chromosome 6 median value it was
classified as long and below, as short.
74
!"#$%&#'%()*#+%)(,('(,&#-#'&.)&./*&'#0)&1&)&23/445#67*%#*%&#8#+%)#
Percentage of short and long chromosome 6 on Y- and X- bearing sperm
+!"
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Percentage
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>8?@"A6B51"@89C@":81":8511"D?E151=:"?=D?4?DB/0@"?="C8?F8"F859G9@9G1"("C/@"G1/@B51DH"
I=":81"F/@1"9J"-KL1/5?=6"@M15GN"?="/00":81"F/@1@":8151"C/@"/="1OB/0"M59M95P9="9J"@895:"
/=D" 09=6" F859G9@9G1" (" 9L@1541DH" Q9C1415N" ?=" ," L1/5?=6" @M15G" C1" 9L@1541D" /" L?/@"
Figure
7. Percentage of short and long chromosome 6 on Y- and X-bearing
C?:8"/="941551M51@1=:/P9="9J"@895:","F859G9@9G1"("?="/00":81"F/@1@H"""
sperm.
This figure shows the three different individuals in which chromosome 6 was measured.
In the case of Y-bearing sperm, in all the cases there was an equal proportion of short
and long chromosome 6 observed. However, in X bearing sperm we observed a bias
with an overrepresentation of short X chromosome 6 in all the cases. Male #1 X-sperm
n = 66, Y-sperm n=105; male #2 X-sperm n = 71, Y-sperm n = 56; male #3 X-sperm n
= 59, Y-sperm n = 60.
75
Figure 8. XY sperm counting through two sequential 1-colour FISH experiments.
(A) A Y-specific BAC (285J12) was labeled in red and hybridized onto platypus sperm
spreads. The number of Y-positive and Y-negative sperm was counted. (B) The same
platypus sperm spreads were hybridized against a X-specific BAC (456I13) labeled in
red in a second FISH experiments. The same sperm cells recorded in experiment (A)
were observed and counted in this second FISH. (C) This graph reports the percentage
of Y positive sperm (Y) and Y negative sperm (no Y) observed in the first FISH
experiment (A), showing a clear bias towards Y positive sperm. (D) This graph reports
the percentage of X positive sperm (X) observed in the second 1-colour FISH
experiment (B), which was recorded as no Y in graph (C). The percentage no X
represents all the X-negative cells that were reported Y-positive in (C), and the reported
Non-informative percentage corresponds to sperm cells that did not yield a signal in
either experiment.
76
Figure 9. Potential modes of inheritance of platypus chromosome 6 variants in
females. (A) Assuming that females transmit only eggs that carry the short chromosome
6, the possible offspring will include males showing bimodal distribution that have not
been identified yet and only females showing bimodal distribution, however, we found
one case of non-bimodality in females. Therefore, this assumption does not fit our data.
(B) If females transmitted only the long version of chromosome 6, we would expect to
see bimodal males, but again no bimodality has been identified in males so far. Also,
this model does not allow the occurrence of non-bimodal females, therefore, it does not
explain our data either. (C) If females were to transmit both alleles we would be able to
observe all the distributions that we have observed in our studies. However, it would
also imply to have bimodal males. Therefore, a mechanism should exist to prevent the
development of bimodal males.
77
Figure S1. The signal intensity on the NOR after rDNA FISH resembles the signal
intensity obtained after silver staining. The chromosomes we present in this figure
correspond to the same male platypus. The silver staining chromosomes are
homologous as well as the FISH chromosomes, but they belong to different cells.
(A) A probe based on the human 28S rDNA was designed and used for DNA FISH. (B)
On the left we find the chromosome 6 homologue that presented a gap after silver
staining with its respective signal intensity graph, to the left we found the signal
intensity for a chromosome on which FISH rDNA was performed and we notice the
resemblance between the two intensity graphs. (C) On the left we observe the
chromosome in which FISH was performed with its respective signal intensity graph to
the left, showing similarity to the intensity graph of the silver staining chromosome on
the right.
78
Figure S2. Expected XY sperm counting differences assuming negative selection
against X-sperm carrying the long chromosome 6 versus observed XY sperm
counting differences under three different assumptions. Based on the data in Figure
3 and Table 1, the black column represents the difference Y-X sperm that we expect to
observe in each individual. The white, dark gray and light gray columns represent the
observed Y-X sperm differences based on the data presented in figure 4, under three
different assumptions. (1) We assume that X sperm is represented by the sum of X
positive sperm and non-informative sperm; (2) we assume X sperm is exclusively
represented by X positive sperm whereas Y sperm is Y positive sperm plus noninformative sperm; (3) we assume uninformative sperm represent cases in which either
the X1 or Y5 were lost in the scored sperm, therefore, we exclude them from the
calculation.
79
Tables
Table 1. Chromosome 6 NOR heteromorphism in platypus males and females.
N
P-value
N
P-value
Male #4
56
< 2.2e-16
Female #1
103
< 2.2e-16
Male #5
45
1.3e-12
Female #2
62
< 2.2e-16
Male #6
104
< 2.2e-16
Female #3
109
< 2.2e-16
Table 2. Percentage value of long and short chromosome 6 in X/Y sperm.
X-Sperm
Y-Sperm
N
Short %
Long %
N
Short %
Long %
Male #1
66
66.67
33.33
105
49.52
50.48
Male #2
71
78.87
21.13
56
50
50
Male #3
59
76.27
23.73
60
53.33
46.67
Table 3. XY sperm counting in two sequential 1-color FISH experiments showing a
bias towards Y sperm in all three analyzed individuals.
1st FISH
2nd FISH
Non-
N
Y%
no Y %
X%
Informative
Male #1
151
57
43
32.5
10.5
Male #2
109
59.6
40.4
30.3
10.1
Male #3
100
59
41
31
10
80
References
[1] Wang J, Chen PJ, Wang GJ, Keller L. Chromosome size differences may affect
meiosis and genome size. Science. 2010;329:293.
[2] Bick YAE, Sharman, G.B. The chromosomes of the platypus (Ornithorhynchus:
Monotremata). Cytobios. 1975;14:17-28.
[3] Grutzner F, Rens W, Tsend-Ayush E, El-Mogharbel N, O'Brien PC, Jones RC, et al.
In the platypus a meiotic chain of ten sex chromosomes shares genes with the bird Z
and mammal X chromosomes. Nature. 2004;432:913-7.
[4] El-Mogharbel N, Wakefield M, Deakin JE, Tsend-Ayush E, Grutzner F, Alsop A, et
al. DMRT gene cluster analysis in the platypus: new insights into genomic organization
and regulatory regions. Genomics. 2007;89:10-21.
[5] Rens W, O'Brien PC, Grutzner F, Clarke O, Graphodatskaya D, Tsend-Ayush E, et
al. The multiple sex chromosomes of platypus and echidna are not completely identical
and several share homology with the avian Z. Genome Biol. 2007;8:R243.
[6] Veyrunes F, Waters PD, Miethke P, Rens W, McMillan D, Alsop AE, et al. Birdlike sex chromosomes of platypus imply recent origin of mammal sex chromosomes.
Genome Res. 2008;18:965-73.
[7] Watson JM, Spencer JA, Riggs AD, Graves JA. The X chromosome of monotremes
shares a highly conserved region with the eutherian and marsupial X chromosomes
despite the absence of X chromosome inactivation. Proc Natl Acad Sci U S A.
1990;87:7125-9.
[8] Waters PD, Delbridge ML, Deakin JE, El-Mogharbel N, Kirby PJ, Carvalho-Silva
DR, et al. Autosomal location of genes from the conserved mammalian X in the
platypus (Ornithorhynchus anatinus): implications for mammalian sex chromosome
evolution. Chromosome Res. 2005;13:401-10.
81
[9] Wallis MC, Waters PD, Delbridge ML, Kirby PJ, Pask AJ, Grutzner F, et al. Sex
determination in platypus and echidna: autosomal location of SOX3 confirms the
absence of SRY from monotremes. Chromosome Res. 2007;15:949-59.
[10] Hore TA, Koina E, Wakefield MJ, Marshall Graves JA. The region homologous to
the X-chromosome inactivation centre has been disrupted in marsupial and monotreme
mammals. Chromosome Res. 2007;15:147-61.
[11] McMillan D, Miethke P, Alsop AE, Rens W, O'Brien P, Trifonov V, et al.
Characterizing the chromosomes of the platypus (Ornithorhynchus anatinus).
Chromosome Res. 2007;15:961-74.
[12] Murtagh C. A unique cytogenetics system in monotremes. Chromosoma.
1977;65:35-57.
[13] Heitz E. Die Ursache der gesetzmässigen Zahl, Lage, Form und Grösse
pflanzlicher Nukleolen Planta. 1931;12:775-884.
[14] McClintock B. The relation of a particular chromosomal element to the
development of the nucleoli in Zea mays. Z Zellforsch. 1934;21:294-328.
[15] Caburet S, Conti C, Schurra C, Lebofsky R, Edelstein SJ, Bensimon A. Human
ribosomal RNA gene arrays display a broad range of palindromic structures. Genome
Res. 2005;15:1079-85.
[16] Little RD, Braaten DC. Genomic organization of human 5 S rDNA and sequence
of one tandem repeat. Genomics. 1989;4:376-83.
[17] Schofer C, Weipoltshammer K, Almeder M, Wachtler F. Arrangement of
individual human ribosomal DNA fragments on stretched DNA fibers. Histochem Cell
Biol. 1998;110:201-5.
[18] Shiels C, Coutelle C, Huxley C. Analysis of ribosomal and alphoid repetitive DNA
by fiber-FISH. Cytogenet Cell Genet. 1997;76:20-2.
[19] Sorensen PD, Frederiksen S. Characterization of human 5S rRNA genes. Nucleic
Acids Res. 1991;19:4147-51.
82
[20] Wallace H, Birnstiel ML. Ribosomal cistrons and the nucleolar organizer. Biochim
Biophys Acta. 1966;114:296-310.
[21] Moss T, Langlois F, Gagnon-Kugler T, Stefanovsky V. A housekeeper with power
of attorney: the rRNA genes in ribosome biogenesis. Cell Mol Life Sci. 2007;64:29-49.
[22] Boisvert FM, van Koningsbruggen S, Navascues J, Lamond AI. The
multifunctional nucleolus. Nat Rev Mol Cell Biol. 2007;8:574-85.
[23] McKeown PC, Shaw PJ. Chromatin: linking structure and function in the
nucleolus. Chromosoma. 2009;118:11-23.
[24] Tucker S, Vitins A, Pikaard CS. Nucleolar dominance and ribosomal RNA gene
silencing. Curr Opin Cell Biol. 2010;22:351-6.
[25] Bird AP, Taggart MH, Gehring CA. Methylated and unmethylated ribosomal RNA
genes in the mouse. J Mol Biol. 1981;152:1-17.
[26] Santoro R, Grummt I. Molecular mechanisms mediating methylation-dependent
silencing of ribosomal gene transcription. Mol Cell. 2001;8:719-25.
[27] Stancheva I, Lucchini R, Koller T, Sogo JM. Chromatin structure and methylation
of rat rRNA genes studied by formaldehyde fixation and psoralen cross-linking. Nucleic
Acids Res. 1997;25:1727-35.
[28] Santoro R, Li J, Grummt I. The nucleolar remodeling complex NoRC mediates
heterochromatin formation and silencing of ribosomal gene transcription. Nat Genet.
2002;32:393-6.
[29] Guillen AK, Hirai Y, Tanoue T, Hirai H. Transcriptional repression mechanisms of
nucleolus organizer regions (NORs) in humans and chimpanzees. Chromosome Res.
2004;12:225-37.
[30] Slota E, Wnuk M, Bugno M, Pienkowska-Schelling A, Schelling C, Bratus A, et al.
The mechanisms determining the nucleolar-organizing regions inactivation of domestic
horse chromosomes. J Anim Breed Genet. 2007;124:163-71.
83
[31] McStay B. Nucleolar dominance: a model for rRNA gene silencing. Genes Dev.
2006;20:1207-14.
[32] Roussel P, Andre C, Comai L, Hernandez-Verdun D. The rDNA transcription
machinery is assembled during mitosis in active NORs and absent in inactive NORs. J
Cell Biol. 1996;133:235-46.
[33] Roussel P, Andre C, Masson C, Geraud G, Hernandez-Verdun D. Localization of
the RNA polymerase I transcription factor hUBF during the cell cycle. J Cell Sci.
1993;104 ( Pt 2):327-37.
[34] Matsui S, Sasaki M. Differential staining of nucleolus organizers in mammalian
chromosomes. Nature. 1973;246:148-50.
[35] Matsui S. Structural proteins associated with ribosomal cistrons in Xenopus laevis
chromosomes. Exp Cell Res. 1974;88:88-94.
[36] Funaki K, Matsui S, Sasaki M. Location of nucleolar organizers in animal and
plant chromosomes by means of an improved N-banding technique. Chromosoma.
1975;49:357-70.
[37] Howell WM, Denton TE, Diamond JR. Differential staining of the satellite regions
of human acrocentric chromosomes. Experientia. 1975;31:260-2.
[38] Goodpasture C, Bloom SE. Visualization of nucleolar organizer regions im
mammalian chromosomes using silver staining. Chromosoma. 1975;53:37-50.
[39] Hubbell HR. Silver staining as an indicator of active ribosomal genes. Stain
Technol. 1985;60:285-94.
[40] Kopp E, Mayr B, Czaker R, Schleger W. Nucleolus organizer regions in the
chromosomes of the domestic horse. J Hered. 1981;72:357-8.
[41] Switonski M, Marcolla P, Pienkowska A, Cholewinski G. Preliminary
investigation on interindividual variation of the nucleolar organizer regions (AgNORs)
in the horse karyotype Anim Sci Pap Rep. 1994;12:15-9.
84
[42] Forman M, Nguyen P, Hula V, Kral J. Sex chromosome pairing and extensive
NOR polymorphism in Wadicosa fidelis (Araneae: Lycosidae). Cytogenet Genome Res.
2013;141:43-9.
[43] Collares-Pereira MJ, Rab P. NOR polymorphism in the Iberian species
Chondrostoma lusitanicum (Pisces: Cyprinidae)--re-examination by FISH. Genetica.
1999;105:301-3.
[44] Castro J, Rodriguez S, Pardo BG, Sanchez L, Martinez P. Population analysis of an
unusual NOR-site polymorphism in brown trout (Salmo trutta L.). Heredity (Edinb).
2001;86:291-302.
[45] Zhuo L, Reed KM, Phillips RB. Hypervariability of ribosomal DNA at multiple
chromosomal sites in lake trout (Salvelinus namaycush). Genome. 1995;38:487-96.
[46] Schmid M, Feichtinger W, Weimer R, Mais C, Bolanos F, Leon P. Chromosome
banding in Amphibia. XXI. Inversion polymorphism and multiple nucleolus organizer
regions in Agalychnis callidryas (Anura, Hylidae). Cytogenet Cell Genet. 1995;69:1826.
[47] Caperta AD, Neves N, Morais-Cecilio L, Malho R, Viegas W. Genome
restructuring in rye affects the expression, organization and disposition of homologous
rDNA loci. J Cell Sci. 2002;115:2839-46.
[48] Araujo SM, Silva CC, Pompolo SG, Perfectti F, Camacho JP. Genetic load caused
by variation in the amount of rDNA in a wasp. Chromosome Res. 2002;10:607-13.
[49] Pinna-Senn E, Di Tada IE, Lisanti JA. Polymorphism of the Microchromosomes
and the Nucleolar Organizer Region in Pristidactylus Achalensis (Sauria:Iguanidae)
Herpetologica. 1987;43:120-7.
[50] Zurita F, Sanchez A, Burgos M, Jimenez R, Diaz de la Guardia R.
Interchromosomal, intercellular and interindividual variability of NORs studied with
silver staining and in situ hybridization. Heredity (Edinb). 1997;78 ( Pt 3):229-34.
85
[51] Mandrioli M, Manicardi GC, Bizzaro D, Bianchi U. NOR heteromorphism within a
parthenogenetic lineage of the aphid Megoura viciae. Chromosome Res. 1999;7:157-62.
[52] Wnuk M, Slota E, Kotylak Z, Bugno M. Factors responsible for modulation of
ribosomal RNA transcription. Folia Biol (Krakow). 2006;54:1-8.
[53] Schubert I, Wobus U. In situ hybridization confirms jumping nucleolus organizing
regions in Allium. Chromosoma. 1985;92:143-8.
[54] Gillings MR, Frankham R, Speirs J, Whalley M. X-Y Exchange and the
Coevolution of the X and Y Rdna Arrays in Drosophila melanogaster. Genetics.
1987;116:241-51.
[55] Hsu TC, Spirito SE, Pardue ML. Distribution of 18+28S ribosomal genes in
mammalian genomes. Chromosoma. 1975;53:25-36.
[56] Patel HR, Delbridge ML, Graves JAM. Organization and Evolution of the
Marsupial X Chromosome. In: Janine E. Deakin PDW, Jennifer A. Marshall Graves,
editor. Marsupial Genetics and Genomics: Springer; 2010.
[57] Gaspar VP, Shimauti EL, Fernandez MA. Chromosomal localization and partial
sequencing of the 18S and 28S ribosomal genes from Bradysia hygida (Diptera:
Sciaridae). Genet Mol Res. 2014;13:2177-85.
[58] Gerbi SA. Localization and characterization of the ribosomal RNA cistrons in
Sciara coprophila. J Mol Biol. 1971;58:499-511.
[59] Madalena CR, Amabis JM, Stocker AJ, Gorab E. The localization of ribosomal
DNA in Sciaridae (Diptera: Nematocera) reassessed. Chromosome Res. 2007;15:40916.
[60] Javier TF, Silva LAF, Rodarte RS, Leoncini O. Characterization of ribosomal
DNA (rDNA) in Drosophila arizonae. Genet Mol Biol. 2000;23.
[61] Marchi A, Pili E. Ribosomal RNA genes in mosquitoes: localization by
fluorescence in situ hybridization (FISH). Heredity (Edinb). 1994;72 ( Pt 6):599-605.
86
[62] Chen ZJ, Pikaard CS. Epigenetic silencing of RNA polymerase I transcription: a
role for DNA methylation and histone modification in nucleolar dominance. Genes
Dev. 1997;11:2124-36.
[63] Zurita F, Jimenez R, Burgos M, de la Guardia RD. Sequential silver staining and in
situ hybridization reveal a direct association between rDNA levels and the expression of
homologous nucleolar organizing regions: a hypothesis for NOR structure and function.
J Cell Sci. 1998;111 ( Pt 10):1433-9.
[64] Zurita F, Jimenez R, Diaz de la Guardia R, Burgos M. The relative rDNA content
of a NOR determines its level of expression and its probability of becoming active. A
sequential silver staining and in-situ hybridization study. Chromosome Res.
1999;7:563-70.
[65] Shubert I, Künzel G. Position dependent NOR activity in barley. Chromosoma.
1990;99:352-9.
[66] de Capoa A, Felli MP, Baldini A, Rocchi M, Archidiacono N, Aleixandre C, et al.
Relationship between the number and function of human ribosomal genes. Hum Genet.
1988;79:301-4.
[67] Lyttle TW. Segregation distorters. Annu Rev Genet. 1991;25:511-57.
[68] Schwander T, Beukeboom LW. Non-random autosome segregation: a stepping
stone for the evolution of sex chromosome complexes? Sex-biased transmission of
autosomes could facilitate the spread of antagonistic alleles, and generate sexchromosome systems with multiple X or Y chromosomes. Bioessays. 2011;33:111-4.
[69] Tsend-Ayush E, Dodge N, Mohr J, Casey A, Himmelbauer H, Kremitzki CL, et al.
Higher-order genome organization in platypus and chicken sperm and repositioning of
sex chromosomes during mammalian evolution. Chromosoma. 2009;118:53-69.
[70] Tsend-Ayush E, Kortschak RD, Bernard P, Lim SL, Ryan J, Rosenkranz R, et al.
Identification of mediator complex 26 (Crsp7) gametologs on platypus X1 and Y5 sex
87
chromosomes: a candidate testis-determining gene in monotremes? Chromosome Res.
2012;20:127-38.
[71] Tsend-Ayush E, Lim SL, Pask AJ, Hamdan DD, Renfree MB, Grutzner F.
Characterisation
of
ATRX,
DMRT1,
DMRT7
and
WT1
in
the
platypus
(Ornithorhynchus anatinus). Reprod Fertil Dev. 2009;21:985-91.
88
CHAPTER 4: Identification and characterization of a male-specific change in Sox3
in the platypus
This is a conventional thesis chapter.
Chapter overview
In this chapter I aimed to investigate the genes located on chromosome 6, focusing on
Sox3. The interest in this gene stems from the fact that Sry evolved from it and that it
has been shown to function in a similar way to Sry when expressed in the gonad.
Although circumstantial data suggested that Sox3 is on platypus chromosome 6, actual
information was lacking for this important gene. Furthermore, in light of the results
presented in the previous chapter Sox3 appears an interesting gene to look for any signs
of sex chromosome differentiation.
Platypus Sox3 location was confirmed to be on chromosome 6. Analysis of a fragment
of this gene in ten different individuals showed a male-specific variation on one of the
polyalanine tracts. Expansion and reduction of polyalanine tracts have been related to
disease in humans, therefore, a male-specific change in one of these regions could have
an impact on its transcriptional activity in a sex-specific fashion. Furthermore, given
that Sry evolved from Sox3 it is possible that this male-specific variant could be
involved in male sex determination or development. In the future, obtaining the fulllength Sox3 transcript in both sexes would be crucial to spot further sex-specific
variations. Moreover, testing the capability of the male-specific and female-occurring
Sox3 variants to activate Sox9 in vitro would be key to further confirm Sox9 possible
implication on male sex-determination.
89
Identification of a male-specific Sox3 allele in the platypus might suggest a role in
male sex determination
Abstract
Sox3 is the gene from which the therian male sex-determining gene (Sry) evolved. It is
normally expressed in the developing central nervous system and in the developing
urogenital ridge, as well as in adult gonads and neurogenic regions. Interestingly, it has
been shown that Sox3 can mimic Sry when ectopically expressed in the XX bipotential
gonad in mice. In platypus Sox3 is located on chromosome 6, which is the proto-sex
chromosome in therian mammals. The autosomal location of this gene confirmed the
lack of a Sry orthologue in the platypus. However, Sox3 has not been characterized in
this animal yet. Given the lack of information of Sox3 in the platypus, we decided to
characterize this gene. In this study we confirmed Sox3 location on chromosome 6 and
interestingly, we discovered a male-specific variant. The observed male-specific Sox3
has a reduced second polyalanine tract coding for seven residues instead of nine. We
analyzed five males and five females; none of the females carried the seven alanine
Sox3 whereas it was observed in four out of five males. Polyalanine tract reductions and
expansions have been related to disease in humans, with expansions of as little as one
residue producing a phenotype. This raises the interesting possibility that Sox3 might
have adopted a male-specific role in the platypus.
Introduction
Sox3 is the gene from which the therian male sex-determining gene (Sry) evolved [1, 2].
This is a very interesting gene that has been investigated in therian mammals and in
other vertebrates, but mostly overlooked in the platypus. Sox3 is a one-exon gene and it
contains a high mobility group (HMG) box, which is responsible for DNA-binding,
90
DNA-bending, protein interactions and nuclear import and export [3, 4]. This domain is
highly conserved across vertebrates, with around 86% identity among Sox3 orthologues
[5]. Interestingly, it has been shown that the HMG box from Sox3 and Sox9 can replace
Sry’s HMG box in sex determination [6]. SOX3 additionally contains a variable Nterminal region that bears homopolymeric runs of alanine (A), glycine (G) and proline
(P), and it has been suggested that this region is important for the modulation of SOX3
in different species [5].
Polyalanine tracts are of particular interest since expansions of these regions in SOX3
and several other transcription factors have been linked to disease in humans [7, 8].
Interestingly, expansions of as little as one amino acid have been associated to a
phenotype. For example, expansions from 16 to 17 residues in the first polyalanine tract
in ARX results in non-syndromic mental disability in humans [9]. Polyalanine tracts
have shown to be mammalian specific in SOX3 as well as in the HOX and GATA
protein families [5, 10]. SOX3 has four polyalanine tracts, the first tract is therianspecific and expansions of this tract have been associated with X-linked mental
retardation with hormone deficiency and hypopituitarism in humans [7, 11]. The second
and fourth tracts have been observed in all mammals including monotremes, which is
the only group missing the third tract [5]. Thereafter, monotremes have only two
polyalanine tracts that correspond to the second and fourth polyalanine tracts from the
eutherian SOX3 [12].
It is known that Sox3 is normally expressed in the developing central nervous system in
mouse [4, 13-15] and together with Sox1 and Sox2 participates in the initiation and
progression of neuronal differentiation [16]. Sox3 expression is also found within adult
neurogenic regions in the subventricular and subgranular zones as well as in a subset of
differentiated hypothalamic cells [17, 18]. On the other hand, Sox3 is also expressed in
91
the developing urogenital ridge [4] and later in the adult gonad in both males and
females [19, 20]. Unlike Sry, Sox3 is not required for sex determination but it is
important for normal oocyte development as well as for testis differentiation and
gametogenesis [20].
However, it has been shown that Sry and Sox3 are functionally interchangeable [21]. In
mice, SRY acts synergistically with SF1 to trigger male sex determination by upregulating Sox9 through its testis-specific enhancer (TES) [22]. SOX9 in turn activates
AMH, the anti-Müllerian hormone [23], which suppresses the formation of the
Müllerian duct repressing the female developmental pathway and allows the
progression of male development [24]. It has been shown that ectopical expression of
Sox3 in the bipotential gonad of XX mice leads to female-to-male sex reversal and that
this occurs only if there is also Sf1 expression as well as a normal Sox9 context [21].
This implies that Sry and Sox3 are able to induce male sex-determination through a
similar mechanism making them functionally interchangeable.
Sox3 has not been characterized yet in the platypus, however, mapping of this gene to
chromosome 6 [25] helped to exclude the possibility of a Sry orthologue in this animal.
It has been recently shown that chromosome 6 in the platypus shows non-random
segregation during male meiosis (Toledo Flores et al, in preparation) and this could
potentially facilitate the transmission of sexually antagonistic alleles [26]. Since
chromosome 6 is homologous to the therian X [25, 27, 28] we asked whether this
autosome could bear a male sex determination or sex development factor. Given that
Sry evolved from Sox3 we decided to investigate this gene in the platypus.
In this study we physically mapped and characterized platypus and echidna Sox3.
Importantly, we identified a variation in the platypus second polyalanine tract that
92
showed to be male-specific. This raises the interesting possibility that the Sox3 malespecific variant might have evolved a male-specific role in the platypus.
Materials and Methods
Phylogenetic analysis
Sequences of the following genes were included in the analysis: Sox1, Sox2, Sox3,
Sox14 and Sox21. The following species were used: Gallus gallus (GG),
Ornithorhynchus anatinus (OA), Monodelphis domestica (MD), Homo sapiens (HS)
and Caenorhabditis elegans (CE) as outgroup. The reference numbers of the used genes
are the following: Sox1 HS (NM_005986.2), Sox1 MD (XM_007501279), Sox2 GG
(D50603.1), Sox2 HS (NM_003106.3), Sox2 MD (XM_001368783.3), Sox2 OA
(XM_007669055.1), Sox3 GG (AB753847.1), Sox3 HS (NG_009387.1), Sox3 MD
(XM_007507371.1), Sox14 GG (NM_204761.1), Sox14 HS (NM_004189.3), Sox14
MD (XM_007493921.1), Sox14 OA (AY112710.1), Sox21 GG (AB026623.1), Sox21
HS (NM_007084.2), Sox21 MD (XM_007501393.1). The utilized Sox3 OA sequence
was obtained from Ultracontig 15 (release Oana-5.0 Ensembl gene build May 2006).
Sequences were aligned using MUSCLE [29] implemented in Geneious v5.3.4 using
default paramenters. MrBayes [30] was used to reconstruct the tree using 106 iterations
and GTR + gamma.
BAC clone isolation
We used the partial platypus Sox3 sequence identified in Ultracontig 15 (release Oana5.0 Ensembl gene build May 2006) to identify the Sox3 positive BAC clones KAAH752L02, KAAH-171I18 and KAAH-643C06 (http:// genome.wustl.edu/tools/blast).
93
RNA extraction
Adult male echidna and platypus tissues (testis, brain, liver) were used to isolate RNA
using standard protocols [31]. Briefly, total RNA was extracted from -80ºC frozen
tissues using TRIzol reagent (Invitrogen), according to the manufacturer’s instructions.
RNA was resuspended in nuclease free water and stored at -80ºC. RNA was later
treated with DNAse I (Invitrogen) to remove genomic DNA, precipitated and
resuspended in RNAse-free water.
cDNA synthesis
Samples of 1 µg of RNA were used to obtain cDNA from each tissue using a
SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen), as previously
described [31]. In summary, RNA was incubated for 5 min at 65ºC with 1 µl of 50 µM
oligo(dT) and 1 µl of 10 mM dNTP. This was followed by further incubation for 50
min at 50ºC with 2 µl of 10x RT buffer, 4 µl of 25 mM MgCl2, 2 µl of dithiothreitol
(DTT, 0.1 M), 1 µl RNaseOUTTM (40 U/µl), and 1 µl SuperScript III RT (200 U/µl).
Termination was performed at 85ºC for 5 min. Reverse transcription was followed by
RNase H treatment; 1 µl of RNase H was added to each reaction and incubated for 20
min at 37ºC. The cDNA samples were stored at -20ºC.
PCR and gene expression analysis
Each PCR had a final volume of 25 µl, which consisted of 0.5-1 µl (50 ng) of genomic
DNA, BAC DNA or cDNA, 5 µl of 5x PCR buffer with MgCl2 (Promega), 5 U/µl of
Taq DNA polymerase, 0.4 µM of each forward and reverse primers and 0.1mM dNTPs.
94
Initial denaturation was carried at 96ºC for 3 min followed by 35 cycles of: denaturation
at 96ºC for 30 s, annealing at 55-59ºC for 1 min and extension at 72ºC for 2 min. Final
extension was performed at 72ºC for 7 min. The primers used were SoxFGCACAACTCGGAGATCAGCAAG, SoxR-GGCAGGTACATGCTGATCATATC at
59ºC
(expected
size
~600bp)
or
the
gene-specific
Sox3F-
CTACTCGCTGATGCCCGACC and Sox3R-GAGCTGGGCTCCGACTTGAC at 59ºC
(expected size ~300bp), designed based on ENSOANG00000002448; !–actin-FGCCCATCTACGAAGGTTACGC and !–actin- AAGGTCGTTTCGTGGATACCAC
at 55ºC. A 1.5% agarose gel was used to visualize the products.
Preparation of mitotic chromosomes
Mitotic metaphase chromosomes and interphase preparations were made from
previously established platypus fibroblast cell lines [32]. Fibroblast cells were cultured
in DMEM Gibco BRL plus Hepes, 10% FCS, 2 mM L-glutamine (Invitrogen),
penicillin/streptomycin at 32ºC until they reached 75-85% confluence, changing media
two or three times per week. To harvest metaphase chromosomes, cells were treated
with 0.5 µg/ml colcemid for 2-3 hrs. Cells were then trypsinized and spun at 1,200 rpm
for 5 min to recover pellet. Supernatants were removed and the cell pellet was slowly
resuspended in 0.075 M KCl hypotonic and incubated at 37ºC for 10 min. Cells were
recovered after spinning at 1,200 rpm for 5 min and resuspended in fresh-made ice-cold
fixative, containing 75% methanol and 25% acetic acid. After washing the metaphase
preparation four to six times with fixative, cells were dropped in methanol-washed glass
slides and allowed to age overnight at -20ºC.
95
Fluorescence in situ hybridization
BAC DNA (1 µg) was labeled with spectrum orange or green (Abbott Molecular) using
random primers and Klenow polymerase and hybridized by fluoresence in situ
hybridization (FISH) on metaphase cells according to standard procedures previously
described in [33]. Briefly, slides were treated with 100 µg/ml RNase A/2x SSC for 30
min at 37ºC followed by pepsin treatment (10 min incubation in 0.01 % pepsin in 10
mM HCl at 37ºC). Next, slides were re-fixed for 10 min in 1 x PBS, 50 mM MgCl2, 1 %
formaldehyde and then dehydrated in ethanol series, denatured for 5 min at 75ºC in 70%
formamide, 2x SSC and again dehydrated. Slides were hybridized overnight in a moist
chamber at 37ºC with a mix of DNA probe, 10-20 µg of sonicated genomic DNA and
50 µg of salmon sperm redissolved in 50% formamide, 10% dextran sulfate, 2x SSC.
Prior to hybridization the probe mix was denatured for 10 min at 80ºC and preannealed
for 30 min at 37ºC. Post-hybridization, the slides were washed three times for 3 min in
50% formamide, 2x SSC at room temperature followed by one wash for 5 min in 2x
SSX at 42ºC, once for 5 min in 0.1x SSC at 60ºC and finally once for 5 min in 2x SSC
at 42ºC. Cells were counterstained with 1 µg/µl DAPI in 2x SSC for 1 min and then
mounted with one drop of Vectashield. Images were taken using a Zeiss AxioImagerZ.1
epifluorescence microscope and Zeiss Axiovision software.
Construct modification
Expression plasmid pcDNA3.1(+) containing mouse Sox3 was used as a template for
inverse PCR. Primers flanking the first polyalanine tract were used for standard PCR
reaction (see above) with the next modifications: TM of 65ºC, elongation for 8 min and
final
elongation
for
10
min;
the
used
primers
were
MM-1st_polyA_F96
AGCAGCCCGGTGGGCGTGG
and
MM-1st_polyA_R-
GCCCGGGGGCAGGAGGCCGC. PCR products were purified using QIAquick PCR
purification kit (QIAGEN) according to the manufacturer’s instructions. 20 ng of the
obtained inverse PCR product were self-ligated in a 20 µl ligation reaction with 2 µl
10x Buffer (NEB), 1 µl ligase T4 (NEB) and PEG4000 50% w/v. The ligation reaction
was performed overnight at 16ºC. 10 ng of ligated vector were used to transform 50 µl
of competent recA, endA E. coli cells by heatshock at 42ºC for 45-50 secs. Vector was
propagated and collected following standard procedures. Deletion was confirmed by
standard PCR and Sanger sequencing.
Results/Discussion
Sox3 is located on platypus chromosome 6
The first attempts to map Sox3 in the platypus using a Sox3 positive BACs did not
succeed due to unspecific signals in the FISH experiments [25]. To overcome this
problem, a BAC containing Sox3 contiguous sequence was used instead. This anchor
sequence was obtained from the Ultracontig 15 from the platypus genome (release
Oana-5.0 Ensembl gene build May 2006) and it comprised the region expanding
positions 1622 kb to 1799 kb. In this same ultracontig, a region matching Sox3 is
located around position 337.5 kb [25], however, it is surrounded by unresolved
sequence (Fig. 2). Besides this region, there are no direct matches to Sox3 in Ultracontig
15.
Chromosome 6 contains many other genes that have been mapped to the therian X
chromosome [34-36]; therefore, it is very likely that it contains Sox3 as well. However,
the existence of unresolved sequence surrounding Sox3 leads to the possibility that the
97
mapped BAC and Sox3 are not physically contiguous. Therefore, we first wanted to
confirm the physical location of Sox3. In order to do this, we first determined the
identity of the publicly available Sox3 sequence through a Bayesian phylogenetic
reconstruction, which show that the platypus Sox3 clusters together with Sox3 from
other species and not with other SOX genes, further confirming its identity (Fig. 3).
Next, we searched the platypus BAC sequence library of the Genome Institute at the
University of Washington (http:// genome.wustl.edu/tools/blast) with the confirmed
platypus Sox3 sequence. Through this, three different BACs containing fragments of
Sox3 were retrieved KAAH-752L02, KAAH-171I18 and KAAH-643C06 (Fig. 4A). We
physically mapped these three BACs, which colocalized on the long arm of platypus
chromosome 6 (Fig. 4B-D), in a region similar to that where the previous BAC
containing adjacent sequence has been localized, confirming previously reported work
[25].
Absence of detectable Sox3 mRNA expression in adult monotreme gonads
Next we wanted to investigate whether the expression is conserved in the monotreme
male gonad. As previously discussed, Sox3 is normally expressed in the developing
urogenital ridge [4] as well as in the adult gonad in both sexes [19, 20] and in specific
regions of the developing [4, 13-15] and adult brain [17, 18]. We prepared cDNA from
adult male platypus and echidna brain, testis and liver. Expression analysis was
performed using RT-PCR. As expected, no Sox3 expression was detected in liver of
either species. Surprisingly, we were able to observe mRNA expression in echidna brain
but no platypus brain expression and no mRNA was detected in gonads either (Fig. 5B).
This is surprising since Sox3 was previously detected in the adult gonad of both sexes in
mice and chickens [19, 20] and it has been proposed that it has an important function in
gametogenesis [20]. It is possible that Sox3 has a function earlier in development but to
98
investigate this we would require access to monotreme embryonic samples. The lack of
this type of material represents a limitation to this project.
The absence of Sox3 expression in the platypus brain is also surprising particularly after
observing expression in the echidna brain (Fig 5B). PCR amplification of the platypus
Sox3 gene has been a challenging task showing inconsistent PCR results. PCR
triplicates in the same individual showed that under the same conditions and using the
same reagents the reactions successfully amplified a product only sometimes, whereas
others it failed (Fig. 6). Interestingly, this has been observed only in the platypus but not
in the echidna. The lack of Sox3 expression in platypus brain, in contrast to the clear
Sox3 expression in echidna brain, could be the result of a failed RT-PCR or to the
inexistence of Sox3 expression in this tissue.
It is important to note, however, that these brain samples were taken more likely from
the brain cortex. It has been previously reported that in mice Sox3 expression in the
adult brain is limited to neuroprogenitor cells in the subventricular and subgranular
zones and in some differentiated hypothalamic cells [17]. Therefore, it is possible that
our platypus sample comes from a non-Sox3 expressing region.
A male-specific Sox3 variant shows a reduction of the second polyalanine tract
The published Sox3 sequence is incomplete; it comprises 660 bp that cover part of the
HMG box and the region coding for the polyalanine tracts. Several attempts using
genome-walking, TAIL-PCR and RACE-PCR were followed in order to amplify the
whole length of the platypus Sox3. However, these different attempts failed due to the
inconsistency of the Sox3 PCR, which is likely a result of the high GC content of this
gene (Fig. 6). Importantly, !–actin was used as a control to ensure that the genomic
99
DNA was suitable for PCR amplification. However a number of Sox3 fragments were
amplified from different individuals. Sequencing of the products revealed variation in
the polyanalanine tract of Sox3. Specifically this variant was defined by a reduced
second polyalanine tract (fourth tract in conserved terms). Previously in the literature it
was reported that the platypus second polyalanine tract contains nine residues [12],
however, we identify two common variants: a nine-alanine and a seven-alanine Sox3.
Next we identified and sequenced this region in ten animals. Interestingly, we found
that the seven-alanine Sox3 was found exclusively in males and not in females; five
females were sequenced and all of them carried a nine-alanine Sox3 variant whereas
four out of five males carried the seven-alanine Sox3 and one of them carried a fivealanine tract (Fig. 7). The Sox3 PCR product was cloned for four individuals, two males
and two females. Through cloning, we observed that the two males were heterozygous
and carried a seven- and nine-alanine Sox3. Whereas females carried both a nine- and
ten-alanine variants, confirming that females do not carry a seven-alanine Sox3.
This is very interesting because it has been shown that transcription factors are five
times more abundant in polyalanine tracts than other proteins [10], which suggests that
these tracts could be involved in transcriptional activity. Furthermore, the fact that
expansions of as little as one residue produce a phenotype [9] raises the possibility that
the male-specific Sox3 observed in platypus might function slightly differently in males
compared to females. Additionally, clinical and in vitro studies have shown that
deletions in the first polyalanine tract of the human Sox3 lead to increased
transcriptional activity [37].
Therefore, future work will aim to investigate the effects that the reduction in the
second polyalanine tract could have on the platypus Sox3 transcriptional activity in
vitro. We are particularly interested in investigating whether this variant could
100
participate in platypus sex determination by adopting a similar activity as Sry in therian
mammals in upreguating Sox9 via its TES enhancer. Therefore, we aim to test our
hypothesis that the Sox3 seven-alanine variant could efficiently activate Sox9 TES core
element (TESCO) using a reporter gene assay. This will consist in cloning the platypus
Sox3 variants into an expression vector that would be later transfected into cells
containing the TESCO enhancer linked to a luciferase reporter. Next, the relative
luciferase activity will be measured as an indication of the potential of Sox3 to activate
Sox9 through its TESCO enhancer. Other genes may be included in this assay, such as
Sf1 since it is known that it acts synergistically with Sry to activate Sox9 [22]. Similar
approaches have been followed in the past to test the potential of other male sexdetermining candidate genes to trigger Sox9 activity in the platypus [31].
Given the difficulty to amplify the entire Sox3 transcript, we have designed an
experiment that aims to mimic in a mouse Sox3 construct what we have observed in
platypus. Since platypuses have only two polyalanine tracts (Fig. 8A), which
correspond to the second and fourth polyalanine tracts in mouse (Fig. 8B), we have
decided to start by reducing the length of the mouse first polyalanine tract from 14 to
only six residues (Fig. 8C). Our aim is to later assess the impact this might have on
Sox3 transcriptional activity through the previously described reporter gene assay. The
next step would be to completely eliminate the first polyalanine tract and reduce the
length of the fourth polyalanine tract by two residues, from ten to eight (Fig. 8D) to
reproduce the change we observed in the platypus male-specific Sox3. We hypothesize
that there should be an increase in the transcriptional activity of the modified mouse
Sox3 construct (Fig. 8D). This will be tested using the described reporter gene assay.
Importantly, these samples were obtained from a platypus population in New South
Wales. Future work will aim to investigate whether the observed variant is specific to
101
this region or if it is widespread across different populations in Australia. Furthermore,
it would be important to find out what mechanisms are responsible of maintaining this
difference. Our previous work (chapter 3) has shown that chromosome 6 present
distinct characteristics between males and females as well as non-random segregation
during male meiosis. It is then possible that this chromosome commenced to
differentiate before monotremes and therians divergence. Suppression of recombination
between chromosome 6 homologues could then help to maintain the seven-alanine Sox3
variant in males, however, further work is necessary to investigate this.
Conclusion
Sox3 has shown to be able to trigger male sex determination through a similar
mechanism as Sry [21] and its location on the non-randomly segregated chromosome 6
makes it a worthy gene to study in terms of sex determination in the platypus. Our
finding of a male-specific variant of Sox3 raises interesting questions about whether it
might have potentially acquired a male-specific function that could be involved in male
sex determination or differentiation. Investigating the transcriptional activity of this
variant in vitro could provide us with insights about potential novel functions.
Additionally, it might help us to understand the function of the fourth polyalanine tract
in Sox3, which may have clinical relevance. Moreover, this could shed light on the
evolution of Sox3 as a sex-determining factor in therian mammals by looking at its
function in monotremes.
Acknowledgements
We would like to thank Arthur Ferguson and Belinda Turner from the Perth Zoo for
providing essential material for this study.
102
103
Figures
Figure 1. Sox3 HMG box and polyalanine tracts in mammals. This diagram shows the
structure of Sox3 in mouse, opossum, platypus and echidna. Mouse Sox3 has an HMG
box followed by 4 spaced polyalanine tracts. Opossum lacks the 1st polyalanine tract,
whereas platypus and echidna lack both the 1st and 3rd polyalanine tracts. In echidna, the
4th tract seems to be split in two by three serines.
104
Figure 2. Platypus Ultracontig 15 (release Oana-5.0 Ensembl gene build May 2006).
The identified Sox3 sequence is indicated at 337.5 kb and it is surrounded by unresolved
sequence indicated by “NNN”. The anchor sequence that was previously used to map
Sox3 [25] is between positions 1,622 kb and 1,799 kb.
105
Figure 3. Bayesian phylogenetic reconstruction of Sox genes. Species used for
Bayesian reconstruction: CE, Caenorhabditis elegans; MD, Monodelphis domestica;
GG, Gallus gallus; HS, Homo sapiens; OA, Ornithorhynchus anatinus. The OA Sox3
sequence used for this analysis was taken from the Ultracontig 15 in the platypus
genome (release Oana-5.0 Ensembl gene build May 2006). This phylogeny shows that
the platypus Sox3 clusters with the Sox3 of other species and not with any other Sox
gene, supporting that the sequence contained in the Ultracontig 15 is effectively Sox3.
106
Figure 4. Sox3 location on chromosome 6. (A) PCR on BACs 171I18, 752L02 and
643C06 using primers Sox_F and Sox_R that amplify platypus Sox3 confirmed these
BACs contain Sox3. (B) FISH onto female platypus metaphase spreads with 171I18
labeled in green and a known chromosome 6 BAC 849O23 labeled in red, confirms
location of 171I18 on chromosome 6. In panel (C) we observe the same FISH but with
BAC 752L02 labeled in green, and panel (D) shows colocalization of 752L02 and
643C06. This confirms that Sox3 is on platypus chromosome 6.
107
Figure 5. Sox3 expression in platypus and echidna adult tissues. (A) !–actin
amplification in echidna and platypus adult tissues was used as positive control. First
three samples belong to an adult male echidna, next three samples belong to an adult
male platypus. (B) RT-PCR with Sox3 specific primers (SoxF and SoxR) reveals
expression in echidna adult brain. Samples are presented in the same order as in (A).
Expected size: ~600 bp. Tes: testis; Li: liver; Br: brain; g: echidna genomic DNA; 1:
negative control. White arrow indicates Sox3 expected size.
108
Figure 6. Sox3 amplification in the platypus is inconsistent. (A) PCR amplification of
Sox3 from male and female platypus genomic DNA showed to be inconsistent.
Whenever the amplification was successful we retrieved a band of the expected size,
which after sequencing showed to be positive to Sox3. However, as indicated by the
triplicates for these four individual, PCR amplification was not successful in all the
cases. Used primers: Sox3F and Sox3R, expected size: ~300bp. (B) PCR amplification
of !–actin as a positive control.
109
Figure 7. A Sox3 male-specific allele was identified in the platypus. A fragment of
Sox3 was amplified in five different females and five different males. A Sox3 coding for
seven alanines on its 2nd polyalanine tract (4th in conserved terms) was detected only in
males and not in females.
110
Figure 8. Modified mouse Sox3 constructs to mimic platypus Sox3 structure. (A) The
platypus Sox3 polyalanine region consists of two polyalanine tracts whereas the mouse
Sox3 (B) features four polyalanine tracts. (C) To assess the impact that the lack of the
first polyalanine tract could have in the platypus, we reduced the length of the first
polyalanine tract in the mouse construct. (D) To fully mimic the structure of the
platypus Sox3 in mouse, we aim to delete the first polyalanine tract followed by
reduction of the fourth polyalanine tract.
111
Table 1. Cloning of PCR products reveals that four out of the ten analysed platypuses
are heterozygous for Sox3. Although three forms of Sox3 were identified, the 7-Ala
variant was observed exclusively in males.
7-Ala
9-Ala
10-Ala
!1
!
!
?
!2
!
!
?
"1
"
!
!
"2
"
!
!
112
References
[1] Foster JW, Graves JA. An SRY-related sequence on the marsupial X chromosome:
implications for the evolution of the mammalian testis-determining gene. Proc Natl
Acad Sci U S A. 1994;91:1927-31.
[2] Stevanovic M, Lovell-Badge R, Collignon J, Goodfellow PN. SOX3 is an X-linked
gene related to SRY. Hum Mol Genet. 1993;2:2013-8.
[3] Lefebvre V, Dumitriu B, Penzo-Mendez A, Han Y, Pallavi B. Control of cell fate
and differentiation by Sry-related high-mobility-group box (Sox) transcription factors.
Int J Biochem Cell Biol. 2007;39:2195-214.
[4] Collignon J, Sockanathan S, Hacker A, Cohen-Tannoudji M, Norris D, Rastan S, et
al. A comparison of the properties of Sox-3 with Sry and two related genes, Sox-1 and
Sox-2. Development. 1996;122:509-20.
[5] Mojsin M, Kovacevic-Grujicic N, Krstic A, Popovic J, Milivojevic M, Stevanovic
M. Comparative analysis of SOX3 protein orthologs: Expansion of homopolymeric
amino acid tracts during vertebrate evolution. Biochem Genet. 2010;48:612-23.
[6] Bergstrom DE, Young M, Albrecht KH, Eicher EM. Related function of mouse
SOX3, SOX9, and SRY HMG domains assayed by male sex determination. Genesis.
2000;28:111-24.
[7] Laumonnier F, Ronce N, Hamel BC, Thomas P, Lespinasse J, Raynaud M, et al.
Transcription factor SOX3 is involved in X-linked mental retardation with growth
hormone deficiency. Am J Hum Genet. 2002;71:1450-5.
[8] Brown LY, Brown SA. Alanine tracts: the expanding story of human illness and
trinucleotide repeats. Trends Genet. 2004;20:51-8.
[9] Shoubridge C, Gecz J. Polyalanine tract disorders and neurocognitive phenotypes.
Adv Exp Med Biol. 2012;769:185-203.
113
[10] Lavoie H, Debeane F, Trinh QD, Turcotte JF, Corbeil-Girard LP, Dicaire MJ, et al.
Polymorphism, shared functions and convergent evolution of genes with sequences
coding for polyalanine domains. Hum Mol Genet. 2003;12:2967-79.
[11] Woods KS, Cundall M, Turton J, Rizotti K, Mehta A, Palmer R, et al. Over- and
underdosage of SOX3 is associated with infundibular hypoplasia and hypopituitarism.
Am J Hum Genet. 2005;76:833-49.
[12] Lehoczky JA, Innis JW. Expanded HOXA13 polyalanine tracts in a monotreme.
Evol Dev. 2008;10:433-8.
[13] Wood HB, Episkopou V. Comparative expression of the mouse Sox1, Sox2 and
Sox3 genes from pre-gastrulation to early somite stages. Mech Dev. 1999;86:197-201.
[14] Uchikawa M, Kamachi Y, Kondoh H. Two distinct subgroups of Group B Sox
genes for transcriptional activators and repressors: their expression during embryonic
organogenesis of the chicken. Mech Dev. 1999;84:103-20.
[15] Brunelli S, Silva Casey E, Bell D, Harland R, Lovell-Badge R. Expression of Sox3
throughout the developing central nervous system is dependent on the combined action
of discrete, evolutionarily conserved regulatory elements. Genesis. 2003;36:12-24.
[16] Bylund M, Andersson E, Novitch BG, Muhr J. Vertebrate neurogenesis is
counteracted by Sox1-3 activity. Nat Neurosci. 2003;6:1162-8.
[17] Rogers N, Cheah PS, Szarek E, Banerjee K, Schwartz J, Thomas P. Expression of
the murine transcription factor SOX3 during embryonic and adult neurogenesis. Gene
Expr Patterns. 2013;13:240-8.
[18] Wang TW, Stromberg GP, Whitney JT, Brower NW, Klymkowsky MW, Parent
JM. Sox3 expression identifies neural progenitors in persistent neonatal and adult
mouse forebrain germinative zones. J Comp Neurol. 2006;497:88-100.
[19] Shen JH, Ingraham HA. Regulation of the orphan nuclear receptor steroidogenic
factor 1 by Sox proteins. Mol Endocrinol. 2002;16:529-40.
114
[20] Weiss J, Meeks JJ, Hurley L, Raverot G, Frassetto A, Jameson JL. Sox3 is required
for gonadal function, but not sex determination, in males and females. Mol Cell Biol.
2003;23:8084-91.
[21] Sutton E, Hughes J, White S, Sekido R, Tan J, Arboleda V, et al. Identification of
SOX3 as an XX male sex reversal gene in mice and humans. J Clin Invest.
2011;121:328-41.
[22] Sekido R, Lovell-Badge R. Sex determination involves synergistic action of SRY
and SF1 on a specific Sox9 enhancer. Nature. 2008;453:930-4.
[23] De Santa Barbara P, Bonneaud N, Boizet B, Desclozeaux M, Moniot B, Sudbeck
P, et al. Direct interaction of SRY-related protein SOX9 and steroidogenic factor 1
regulates transcription of the human anti-Mullerian hormone gene. Mol Cell Biol.
1998;18:6653-65.
[24] Mishina Y. The In Vivo Function of M"ºllerian-Inhibiting Substance During
Mammalian Sexual Development.
[25] Wallis MC, Waters PD, Delbridge ML, Kirby PJ, Pask AJ, Grutzner F, et al. Sex
determination in platypus and echidna: autosomal location of SOX3 confirms the
absence of SRY from monotremes. Chromosome Res. 2007;15:949-59.
[26] Schwander T, Beukeboom LW. Non-random autosome segregation: a stepping
stone for the evolution of sex chromosome complexes? Sex-biased transmission of
autosomes could facilitate the spread of antagonistic alleles, and generate sexchromosome systems with multiple X or Y chromosomes. Bioessays. 2011;33:111-4.
[27] Waters PD, Delbridge ML, Deakin JE, El-Mogharbel N, Kirby PJ, Carvalho-Silva
DR, et al. Autosomal location of genes from the conserved mammalian X in the
platypus (Ornithorhynchus anatinus): implications for mammalian sex chromosome
evolution. Chromosome Res. 2005;13:401-10.
115
[28] Hore TA, Koina E, Wakefield MJ, Marshall Graves JA. The region homologous to
the X-chromosome inactivation centre has been disrupted in marsupial and monotreme
mammals. Chromosome Res. 2007;15:147-61.
[29] Edgar RC. MUSCLE: multiple sequence alignment with high accuracy and high
throughput. Nucleic Acids Res. 2004;32:1792-7.
[30] Huelsenbeck JP, Ronquist F. MRBAYES: Bayesian inference of phylogenetic
trees. Bioinformatics. 2001;17:754-5.
[31] Tsend-Ayush E, Kortschak RD, Bernard P, Lim SL, Ryan J, Rosenkranz R, et al.
Identification of mediator complex 26 (Crsp7) gametologs on platypus X1 and Y5 sex
chromosomes: a candidate testis-determining gene in monotremes? Chromosome Res.
2012;20:127-38.
[32] Grutzner F, Rens W, Tsend-Ayush E, El-Mogharbel N, O'Brien PC, Jones RC, et
al. In the platypus a meiotic chain of ten sex chromosomes shares genes with the bird Z
and mammal X chromosomes. Nature. 2004;432:913-7.
[33] Tsend-Ayush E, Dodge N, Mohr J, Casey A, Himmelbauer H, Kremitzki CL, et al.
Higher-order genome organization in platypus and chicken sperm and repositioning of
sex chromosomes during mammalian evolution. Chromosoma. 2009;118:53-69.
[34] Ashley T. Chromosome chains and platypus sex: kinky connections. Bioessays.
2005;27:681-4.
[35] Ezaz T, Stiglec R, Veyrunes F, Marshall Graves JA. Relationships between
vertebrate ZW and XY sex chromosome systems. Curr Biol. 2006;16:R736-43.
[36] Veyrunes F, Waters PD, Miethke P, Rens W, McMillan D, Alsop AE, et al. Birdlike sex chromosomes of platypus imply recent origin of mammal sex chromosomes.
Genome Res. 2008;18:965-73.
[37] Alatzoglou KS, Kelberman D, Cowell CT, Palmer R, Arnhold IJ, Melo ME, et al.
Increased transactivation associated with SOX3 polyalanine tract deletion in a patient
with hypopituitarism. J Clin Endocrinol Metab. 2011;96:E685-90.
116
CHAPTER 5: Investigating sex bias in captive bred echidnas
This chapter consists of one manuscript and a brief letter, both in preparation.
Chapter overview
Determining the sex of captive born animals is of great relevance to zoos in order to
implement this information to their captive management plans. The echidna, is a
particularly difficult animal to sex given the lack of any obvious sexual dimorphisms
and due to the fact that sexual organs are internal. Adult echidnas can be sexed by
palpation, however, this is impossible on echidna puggles.
In this chapter, I describe how I utilized our knowledge about male specific genes to
develop a PCR based technique to genetically determine the sex of echidna puggles.
This revealed a strong female bias on captive born echidnas providing important hints
that suggest that factors during the captive breeding process might influence sex ratio.
Importantly, this noninvasive sexing technique can be potentially applied to wild
echidna and platypus populations, including the critically endangered long-beaked
echidna.
Moreover, I have identified a naturally occurring deletion of Sox3 in a male echidna.
This provides a unique opportunity to study the impact this deletion might have on
echidna sex development.
117
Manuscript in preparation:
Deborah Toledo-Flores, Wan Xian Kang, Arthur Ferguson, Belinda Turner,
Enkhjargal Tsend-Ayush, Shu Ly Lim, Frank Grützner. Genetic Sexing Reveals Female
Bias in Echidnas Born in Captivity
119
Genetic Sexing Reveals Female Bias in Echidnas Born in Captivity
Deborah Toledo-Floresa, Wan Xian Kanga, Arthur Fergusonb, Belinda Turnerb,
Enkhjargal Tsend-Ayusha, Shu Ly Lima, Frank Grütznera
a
The Environment Institute, School of Molecular and Biomedical Science, The
University of Adelaide, Adelaide, SA 5005, Australia
b
Perth Zoo, 20 Labouchere Road, South Perth, WA 6151, Australia
Abstract
Captive breeding of echidnas remains difficult and determining the sex of echidnas born
in captivity as soon as possible is important for husbandry and continued breeding
success. Accurately sexing juvenile echidnas is a challenging task due to the lack of
visible external genitalia. Here we have established a non-invasive PCR based
technique to determine the sex of echidnas using DNA from plugged hair. Genomic
DNA was extracted from as few as 10 echidna hair follicles and subsequently PCR
amplification of a male-specific gene was used for determining male or female sex. We
tested eight echidnas born in captivity at Perth Zoo and found a striking sex bias, with
seven females out of eight animals. In two individuals we confirmed the sex by
culturing peripheral blood lymphocytes for chromosomal analysis and Fluorescence in
situ hybridization (FISH) to identify an XX or XY sex chromosome constitution further
confirming the results obtained by PCR. In conclusion we have established a
straightforward and non-invasive molecular technique to reliably determine the sex of
young echidnas and discovered a striking bias towards female offspring in captive bred
echidnas.
120
Keywords: Echidna sexing, hair sample, blood sample, sex-specific PCR, FISH.
Introduction
Echidnas and platypuses are the only living monotremes and amongst Australia’s most
iconic animals. Echidnas are wide-spread in Australia and also found in New Guinea[1].
There are four species of echidnas; the short-beaked echidna (Tachyglossus aculeatus)
and three species of the critically endangered long-beaked echidna (Zaglossus bruijnii,
Z. bartoni and Z. attenboroughi) [2]. Echidnas are comparably easy to keep but difficult
to breed in captivity [3-5]. A considerable amount of planning and effort is required to
successfully breed Echidnas [8].
The London Zoo holds the first record of housing an echidna back in 1845 [3], after
which it became common practice to have echidnas as part of zoo collections. In 1908
the first captive-born echidna was reported in the Berlin Zoo, this came as a surprise
since the breeding was not planned. Unfortunately, the puggle died three months after
hatching [6]. A similar trend has been observed in a significant proportion of echidnas
born in captivity, by 2007 there were 20 echidnas born in international zoos and 7 born
in Australian zoos, with more than half dying before reaching 1 year of age [7]. Only
recently, the Perth Zoo reported that they were able to breed echidnas in captivity
during three consecutive years producing a total of five animals [8]. This breeding
success led to the question of how to accurately determine the sex of juvenile animals to
enable early population management succession planning.
In addition to the practical necessity of determining the sex of the animals bred in
captivity, there is the question of whether there is any bias in the sex ratio of the
offspring. Early identification of sex ratio bias is necessary so that population managers
121
at zoos can take compensatory measures into action [9], such as transferring known-sex
animals to other institutions for future breeding. Different cases of sex ratio bias in
captive animals have been described. For instance, ring-tailed and lesser mouse lemurs
show housing-dependent offspring sex ratio bias [10]. Also, captive populations of
pygmy hippopotamus present female-biased sex ratios associated with high feeding
intensity and “hands-on” husbandry, however, the mechanisms underlying the observed
segregation distortion remain unknown [11]. Similarly, this phenomenon has been
observed in a number of other primate, rodent, marsupial, cervid, bovid, carnivore,
chiropteran and cetacean species [12] but so far no data have been available about sex
ratio bias of captive bred echidnas.
Determination of the sex of echidnas, particularly juveniles, is not a straightforward task
since they do not have any externally obvious sexual dimorphism in body mass,
dimensions or color [1]. In addition, some of the known sex-specific traits such as the
female pouch or the male developed spur (without the sheath) are only established once
the echidna reaches sexual maturity, which means that the young cannot be sexed using
these traits. Even in adults, these characteristics cannot be unequivocally identified. The
female pouch, for example, is not well developed outside of the breeding season [3],
and it can be mistaken with the contraction of the longitudinal muscles of the abdomen
even in males [13]. In the case of the male spur, at least 25% of the adult male
population lose one of their spurs while 25% of mature females have one well
developed spur; this stems from the fact that both sexes have spurs while juveniles,
which are meant to regress in females during adulthood [13]. Furthermore, the
reproductive organs are internal [1]. During the breeding season it is possible to identify
reproductively active males because they show a degree of cloacal swelling, which is
thought to be due to enlarged testicles [5, 14]. However, this technique requires an
experienced eye and outside of the breeding season it is more difficult to accurately
122
determine the sex of individuals. Therefore, given the lack of obvious sexual
dimorphisms the only reliable way to determine the sex of an echidna is by palpation [1,
13, 15], which is quite invasive and impossible in echidna puggles.
Previously, a couple of fecal steroid analyses were established in echidna; however,
they had a maximum accuracy of 80% and one of them was reliable only during the
breeding season [16]. A number of non-invasive genetic sexing techniques available in
and outside breeding season have been established in other species. These techniques
rely on extraction of genomic DNA from unobtrusively collected samples such as hair
and feces [17-24]. Importantly, it has been shown that it is possible to obtain consistent
results from genomic DNA (gDNA) derived from hair, blood and other tissues [25-27].
To the best of our knowledge, no genetic techniques have been previously established to
non-invasively determine the sex of echidnas or platypuses.
Here we report the establishment of a non-invasive PCR-based method to genetically
determine the sex of echidna juveniles from genomic DNA extracted from small hair
samples. Also, we validated our results by performing the same method on genomic
DNA extracted from blood as well as through in situ hybridization with sex
chromosome-specific probes on metaphase spreads derived from peripheral blood
lymphocytes.
Materials and methods
Sample preparation and ethics
Hair samples with follicle attached were collected from the abdominal area of the
echidna under manual restraint using sterile tweezers and placed immediately in a
123
sterile 15mL Falcon® tube. Echidna hair samples were stored at room temperature postcollection and shipped in 70% ethanol.
Blood samples were collected from the venous beak sinus under general anesthetic, into
green-capped sodium heparin tubes. Blood samples were stored at room temperature
and shipped within one day of collection.
Since this was a non-invasive procedure which was done opportunistically during
routine examination no ethics permit had to be acquired.
Genomic DNA extraction from hair and blood samples
Hair follicles were cut under light microscope and each sample was placed in a tube
containing 0.5-1 ml of 70% ethanol. The tubes were briefly spun to collect follicles in
the bottom and ethanol was then removed. Follicles were allowed to dry at room
temperature. Hair follicles were incubated at 55ºC overnight while shaking at 700 rpm
in 300 µl of lysis buffer (50mM TrisChloride, pH 8; 100mM EDTA, pH 8; 100mM
NaCl; 1% SDS), 0.17µg/µl of proteinase K and 0.1M DTT. Once lysis was finalized,
0.6 µl of RnaseA (10 mg/ml) were added and the cell lysate was incubated for further
30 min at 37ºC. Samples were allowed to cool down at room temperature, after this
100µl of 7.5M ammonium acetate were added to the samples and they were vigorously
vortexed, followed by a 5 min incubation on ice. Samples were centrifuged for 5 min at
12,000 rpm, supernatants were then transferred to a new tube. DNA was then
precipitated at -20ºC overnight with 300 µl of 100% isopropanol in the presence of 0.5
µl of Glycogen solution (20 mg/ml). Centrifugation was carried at 12,000 rpm for 20
min, supernatants removed and the DNA pellet was resuspended in 10µl of 55ºC prewarmed Milli-Q water.
124
Genomic DNA from blood samples was extracted using the DNeasy Blood & Tissue
Kit (Qiagen) according to the manufacturer’s instructions. Concentrations were
determined using a NanoDrop 2000 spectophotometer (Thermo Scientific).
PCR and gel electrophoresis
Each PCR had a final volume of 25 µl, which consisted of 0.5-1µl of the extracted
genomic DNA (depending on the obtained concentration), 5µl of 5x PCR buffer with
MgCl2 (Promega), 5U/µl of Taq DNA polymerase, 0.4µM of each forward and reverse
primers and 0.1mM dNTPs. In the case of PCRs using hair-extracted gDNA, 2.5µl of
BSA were used per reaction. Initial denaturation was carried at 96ºC for 3 min followed
by 40 cycles of: denaturation at 96ºC for 30 s, annealing at 50-55ºC for 1 min and
extension at 72ºC for 2 min. Final extension was performed at 72ºC for 7 min. The
primers used were Crspy for-ACCAGTAAATGCTGTGAAACCTC, Crspy revTTCTTTTTATTGGCTGGTTCTGA
GCCCATCTACGAAGGTTACGC
[28]
at
50ºC
and
and
!–actin
!–actin
forrev-
AAGGTCGTTTCGTGGATACCAC at 55ºC. A 1.5% agarose gel was used to visualize
the products.
Culture of peripheral blood cells for chromosome analysis
500µl of echidna blood were cultured with 10ml of PB-MAX medium (Life
Technologies) for 72 hrs at 32ºC (due to monotremes lower body temperature),
according to the manufacturers suggested protocol. The 72 hr incubation was followed
by a further 30 min incubation with 0.5 µg/ml of colcemid. A cell pellet was then
obtained after a 5 min centrifugation at 1,200 rpm, supernatants were removed and cells
were slowly resuspended in 10 ml of 0.075M KCl hypotonic and incubated at 37ºC for
125
10 min. A cell pellet was recovered by spinning 5 min at 1,200 rpm, supernatants were
removed and the cells were slowly resuspended in 10ml of fresh, ice-cold fixative made
of 1 part of acetic acid and 3 parts of ethanol. Cells were then incubated for 10 min at
4ºC, washed 5 times with fixative and resuspended in a final volume of 0.5ml of
fixative. Cells were dropped in methanol-washed glass slides and allowed to age
overnight at -4ºC.
Fluorescence in situ hybridization of BAC clones
Y3-specific BAC EAmhy2 and Y3X4-PAR BAC 48g5 [29] were used to perform FISH
on echidna metaphase spreads under standard conditions, as previously described in
[30]. Images were taken with a Zeis AxioImager Z.1 epifluorescence microscope
equipped with a CCD camera and Zeiss Axiovision software.
Results
Echidna sexing through PCR on hair genomic DNA
We obtained small hair samples from eight echidnas (Fig. 1) born in captivity at the
Perth Zoo during a period of five years 2009 – 2014. Each of the samples consisted of
10 – 50 hairs from which only the follicles were used for the DNA extraction protocol
in order to minimize contaminants. The follicles have a characteristic conical shape
(Fig. 2) compared to the typical bulb shape found in humans. We observed some
variation in the efficiency of the DNA extraction protocol, however generally we
obtained 190.5 ng/µl from an extraction of only 10 hairs.
126
The sex of the individuals was then determined through PCR using primers that amplify
a platypus male-specific gene, Crspy. Crspy, the male specific copy of the Crsp70 gene
that is located on platypus chromosome Y5 [28], which has been shown to be
homologous to the echidna male specific chromosome Y3 [31]. These primers were
tested in echidnas whose sex was known, confirming that the platypus Crspy primers
amplify a male specific product and no products in females (Fig. 3). Running this PCR
on the DNA of eight echidna puggles showed that seven out of the eight captive-born
echidnas were females (Table 1). Importantly, !–actin primers were used as a positive
control.
Echidna sexing validation through PCR on blood genomic DNA and FISH
In order to validate our method we decided to back the PCR result with another method
to genetically determine the sex of the animals. Monotremes have a unique
heteromorphic sex chromosome system, which we have studied in detail [31-33]. We
accessed blood samples for two of the eight echidnas (ID 7 and 8, Table 1) to prepare
genomic DNA and metaphase chromosomes. PCR on the blood extracted DNA
confirmed the results obtained with the hair samples (Fig. 4).
To chromosomally sex the two animals we prepared metaphase spreads derived from
peripheral blood lymphocytes; we then performed DNA FISH using a Y3 specific BAC
together with a Y3X4-PAR BAC. In echidna number seven we observed two signals for
the red-labeled Y3X4-PAR BAC and no signals for the green-labeled Y3 BAC (Fig. 5A).
In the echidna number eight we observed two signals for the now green-labeled Y3X4PAR BAC and one signal for the red-labeled Y3 BAC, which colocalized with one of
the Y3X4-positive chromosomes (Fig. 5B). These results confirm the PCR based results.
127
Discussion
In this study we have established a reliable technique to genetically sex echidnas from
small hair samples using PCR-based amplification of the male-specific gene Crspy [28].
Our results, which have been confirmed by chromosomal sexing, are consistent with
previous studies that indicate that hair genomic DNA and DNA extracted from other
tissues perform reliably [18, 25-27]. Furthermore, we obtained biological confirmations
of the sex of three echidnas (number 1, 2 and 3) since they later had puggles or laid
eggs (Table 1). There is scope to use this technique also in other contexts including
study of sex ratios in wild populations where hair samples can be collected with hair
traps, as it was successfully established for the American pika (Ochotona princeps)
[25]. Adopting this approach may be particularly valuable in the study of the critically
endangered long-beaked echidnas.
Interestingly, this study also revealed a striking bias in the sex ratio of these echidnas
born in captivity. We observed that seven out of eight puggles were females (Table 1).
Distortions in the sex ratios of captive mammals are not unusual and several examples
have been described. Species such as the cotton-top tamarin (Saguinus Oedipus), the
Persian onager (Equus hemionus onager), the ring-tailed lemur (Lemur catta), the sloth
bear (Melursus ursinus) and the scimitar oryx (Oryx dammah) have shown male bias in
captivity [9] whereas others like the pygmy hippopotamus (Hexaprotodon liberiensis)
[11], the blue duiker (Cephalophus monticola) and the mangabey (Cercocebus
albigena) [9] have presented female bias. Several factors have shown to affect sex ratio,
for example, in the gray mouse lemur (Microcebus murinus) it was demonstrated that
group size affected the sex ratio, if the mothers were kept in isolation their offspring
was biased towards females, whereas if they were kept in groups they showed a male
128
bias instead [10]. Other studies have reported that in macaques with a matrilineal social
structure the social status of the mothers influences the sex ratio of their offspring; high
ranking females produce more daughters and lower ranks, produce more sons [34-36].
Also, in the pygmy hippopotamus, Hexaprotodon liberiensis, it was determined that
high feeding intensity coupled with “hands-on” husbandry favored production of
females [11].
A number of different hypotheses have been proposed in order to explain the observed
sex ratio biases. In general, they all predict that manipulation of the sex ratio by an
individual should lead to a greater offspring production (hence, increased fitness)
through the favored sex [37]. This was recently shown to be true, demonstrating that sex
ratio manipulation is quite common amongst species and that it represents a highly
adaptive evolutionary strategy in mammals [37]. One of the first hypothesis was the sex
allocation theory [38], which postulates that in species where the condition of the
mother influences the sex of the offspring there should be a higher production of the
evolutionarily less expensive sex (females) in poor quality mothers, which are females
with suboptimal nutrition and stressed. Other hypotheses such as the Trivers-Willards
hypothesis [39, 40] and the “cost of reproduction hypothesis” [41-44] also predict that
mothers in good conditions will produce more sons and poor mothers will produce more
females. The later suggests this is because mothers in poor conditions simply cannot
afford to produce offspring of the energetically costly sex (males).
In the case of the captive-born echidnas studied here we observed a bias towards
females that in the wild may be a sign of poor nutrition [12]. Stress is another factor that
has been associated with female bias [40]. However, stress often results in inhibition of
reproduction in several species [45-47] and the fact that these echidnas reproduced
successfully suggests that stress levels were low. Therefore, it could be possible that
129
females are costlier than males amongst echidnas, hence the favorable conditions in the
zoos may lead to a female bias [11]. More individuals over a longer period of time will
reveal if this trend is indeed a sex bias or a result of the relatively small sample size [9,
48]. It would be necessary to monitor the sex ratio in this population in the future, since
there have been cases in which populations showing strong sex ratio biases over a short
period of time have lost it or decreased it over time [49-51]. In addition, looking at
different variables such as diet composition (intake of fats versus carbohydrates) [52],
could also shed light on the potential factors influencing sex ratios in monotremes.
Conclusions and future directions
In this study we successfully established several methods to determine the sex of eight
echidnas born in captivity providing, to our knowledge, the first non-invasive genetic
sex determination technique applied to echidna juveniles, which has the potential of
being also applied to the study of wild echidnas. Future refinement will test other genes
on sex chromosomes to provide a positive result in both males and females. Importantly
we discovered a marked sex ratio bias in the studied group of echidnas born in captivity.
Future work on more animals for extended periods of time will test our hypothesis that
echidnas are vulnerable to a sex ratio bias during captive breeding.
Acknowledgements
Thanks to Prof. Steve Donnellan and Dr. Terry Bertozzi from the South Australian
Museum for advice on the hair genomic DNA extraction protocol and for providing hair
samples to test the protocol. DT-F is funded by the National Council of Science and
Technology from Mexico (CONACYT). FG is an ARC research fellow. Thanks to
Perth Zoo’s Veterinary staff for collection and shipment of samples.
130
Author contributions: DT-F performed sample processing, laboratory work and wrote
the manuscript. WXK performed sample processing and laboratory work. AF
contributed to the design of the project. BT collected samples and advised on echidna
biology. ET-A and SLL performed laboratory work. FG contributed to the design and
supervision of the project as well as to the manuscript preparation and data
interpretation.
131
Figures
Fig. 1. Echidna juvenile 7-months old, individual B20282 (Photo: Perth Zoo).
132
Fig. 2. Echidna hair follicle. Samples of 10-50 follicles were used from eight different
individuals.
133
Fig. 3. Sexing results for the eight analyzed echidnas. (A) Example of the PCR. On the
left, the gel for the amplification of the positive control !-actin. A band of 600bp was
obtained from male (!) and female (") control gDNA as well as from the hair extracted
gDNA of echidna number 6. On the right, the result of the sexing-PCR with Crspy
primers. A product is obtained from the male control (!) and no products are retrieved
from the female control (") or from echidna 6, showing that echidna 6 is female. (B)
PCR results for the sexing of the eight analyzed echidnas. On the left column, the
results of !-actin amplification and on the right, the sexing-PCR results. Amplification
of !–actin in all the individuals indicates that the lack of a Crspy band in females was
due to the inexistence of this gene in those individuals and not to a failure in our PCR
reactions. All the analyzed echidnas are females, except for echidna number 8. m: 100
bp marker; -: negative control.
134
Fig. 4. Sexing results through PCR using sex specific primers on genomic DNA from a
male control (!), a female control ("), individuals 7 and 8 (7,8) and a negative control
(-). B-actin primers were the positive control and Crspy primers were our sexing
primers. (A) The gDNA of individuals 7 and 8 were extracted from blood samples. (B)
The gDNA of individuals 7 and 8 were extracted from hair samples. m: 100 bp marker;
-: negative control.
135
Fig. 5. FISH on metaphase spreads prepared from short-term culture of echidna
peripheral blood lymphocytes. (A) FISH on metaphase spread from individual 7 with a
red-labeled Y3X4 BAC which yielded 2 signals and a green-labeled Y3 specific BAC
that yielded 0 signals. (B) FISH on metaphase spread from individual 8 with a green
labeled Y3X4 BAC that produced 2 signals and a red-labeled Y3 specific BAC which
produced 1 signal which co localized with one of the Y3X4 BAC positive
chromosomes.
136
Tables
Table 1 Information about the animals used in this study
ID
Individual
Sex
Age when
DOB
sampled
1
A80281
!
6.5 months
Sexing
Fertile
method
6/8/08
PCR
Yes (Mother of
B20282)
2
A80284
!
6 months
20/8/08
PCR
Yes (Mother of
B20300)
3
A70246
!
18 months
25/7/07
PCR
Yes
4
A90273
!
5 months
27/8/09
PCR
NA
5
A90272
!
5 months
24/8/09
PCR
NA
6
B10297
!
10 months
12/9/11
PCR
NA
7
B20282
!
19 months
17/8/12
PCR / FISH
NA
8
B20300
"
19 months
27/8/12
PCR / FISH
NA
Abbreviation: NA, not available.
137
References
[1] Rismiller PD, McKelvey MW. Frequency of Breeding and Recruitment in the ShortBeaked Echidna, Tachyglossus aculeatus. Journal of Mammalogy. 2000;81:1-17.
[2] IUCN. The IUCN Red List of Threatened Species. Version 2014.2.
<www.iucnredlist.org>. Downloaded on 21 October 2014. 2014.
[3] Jackson S. Australian Mammals: Biology and Captive Management: CSIRO
Publising: Melbourne; 2003.
[4] Temple-Smith P, Grant T. Uncertain breeding: a short history of reproduction in
monotremes. Reprod Fertil Dev. 2001;13:487-97.
[5] Johnston SD, Nicolson V, Madden C, Logie S, Pyne M, Roser A, et al. Assessment
of reproductive status in male echidnas. Anim Reprod Sci. 2007;97:114-27.
[6] Heck L. Echidna - Zuchtung im Berliner Zoologischen Garten. . Gesellschaft fur
Naturforshung Freunde Berlin. 1908:187-9.
[7] Perry L. Short-beaked echidna: births recorded in the ISIS studbook. Complied from
ISIS 4 database. 2007.
[8] Ferguson A, Turner B. Reproductive parameters and behaviour of captive shortbeaked echidna (Tachyglossus aculeatus acanthion) at Perth Zoo. Australian
Mammalogy. 2013;35:84-92.
[9] Faust LJ, Thompson SD. Birth sex ratio in captive mammals: Patterns, biases, and
the implications for management and conservation. Zoo Biology. 2000;19.
[10] Perret M. Influence of Social Factors on Sex Ratio at Birth, Maternal Investment
and Young Survival in a Prosimian Primate. Behavioral Ecology and Sociobiology.
1990;27:447-54.
[11] Zschokke S. Distorted Sex Ratio at Birth in the Captive Pygmy Hippopotamus,
Hexaprotodon liberiensis. Journal of Mammalogy. 2002;83:674-81.
138
[12] Glatston AR. Sex ratio research in zoos and its implications for captive
management. Applied Animal Behaviour Science. 1997;51:209-16.
[13] Rismiller PD. Overcoming a prickly problem. Australian Natural History.
1993;4:22-9.
[14] Beard RA, Grigg GC. Reproduction in the short-beaked echidna, Tachyglossus
aculeatus: field observations at an elevated site in south-east Queensland. Proceedings
of the Linnean Society of New South Wales. 2000;122:89-99.
[15] Rismiller PD. Field observations on Kangaroo Island echidnas (Tachyglossus
aculeatus multiacu- leatus) during the breeding season. In: Augee ML, editor. Platypus
and echidnas. Sydney, Australia: Royal Zoological Society of New South Wales; 1992.
p. 101-5.
[16] Oates JE, Bradshaw FJ, Bradshaw SD, Lonsdale LA. Sex identification and
evidence of gonadal activity in the short-beaked echidna (Tachyglossus aculeatus)
(Monotremata: Tachyglossidae): non-invasive analysis of faecal sex steroids. .
Australian Journal of Zoology. 2002;50:395-406.
[17] Dallas J, Carss D, Marshall F, Koepfli K-P, Kruuk H, Bacon P, et al. Sex
identification of the Eurasian otter Lutra lutra by PCR typing of spraints. Conservation
Genetics 2000;1:181-3.
[18] Pilgrim KL, McKelvey KS, Riddle AE, Schwartz MK. Felid sex identification
based on noninvasive genetic samples. Molecular Ecology Notes. 2005;5:60-1.
[19] Allen M, Engstrom AS, Meyers S, Handt O, Saldeen T, von Haeseler A, et al.
Mitochondrial DNA sequencing of shed hairs and saliva on robbery caps: sensitivity
and matching probabilities. J Forensic Sci. 1998;43:453-64.
[20] Vigilant L. An evaluation of techniques for the extraction and amplification of
DNA from naturally shed hairs. Biol Chem. 1999;380:1329-31.
139
[21] Ding B, Zhang Y-p, Ryder OA. Extraction, PCR amplification, and sequencing of
mitochondrial DNA from scent mark and feces in the giant panda. Zoo Biology.
1998;17:499-504.
[22] Machiels BM, Ruers T, Lindhout M, Hardy K, Hlavaty T, Bang DD, et al. New
protocol for DNA extraction of stool. Biotechniques. 2000;28:286-90.
[23] Reed JZ, Tollit DJ, Thompson PM, Amos W. Molecular scatology: the use of
molecular genetic analysis to assign species, sex and individual identity to seal faeces.
Mol Ecol. 1997;6:225-34.
[24] Faux CE, McInnes JC, Jarman SN. High-throughput real-time PCR and melt curve
analysis for sexing Southern Ocean seabirds using fecal samples. Theriogenology.
2014;81:870-4.
[25] Henry P, Henry A, Russello MA. A noninvasive hair sampling technique to obtain
high quality DNA from elusive small mammals. J Vis Exp. 2011.
[26] Li B, Zhang W, Shi X, Zhou X. Molecular sex identification of sable and stone
marten. Russian Journal of Ecology. 2013;44:137-41.
[27] Pilgrim KL, J. ZW, Schlexer FV, Schwartz MK. Development of a reliable method
for determining sex for a primitive rodent, the Point Arena mountain beaver
(Aplodontia rufa nigra). Conservation Genetics Resources. 2012.
[28] Tsend-Ayush E, Kortschak RD, Bernard P, Lim SL, Ryan J, Rosenkranz R, et al.
Identification of mediator complex 26 (Crsp7) gametologs on platypus X1 and Y5 sex
chromosomes: a candidate testis-determining gene in monotremes? Chromosome Res.
2012;20:127-38.
[29] Dohm JC, Tsend-Ayush E, Reinhardt R, Grutzner F, Himmelbauer H. Disruption
and pseudoautosomal localization of the major histocompatibility complex in
monotremes. Genome Biol. 2007;8:R175.
140
[30] Tsend-Ayush E, Dodge N, Mohr J, Casey A, Himmelbauer H, Kremitzki CL, et al.
Higher-order genome organization in platypus and chicken sperm and repositioning of
sex chromosomes during mammalian evolution. Chromosoma. 2009;118:53-69.
[31] Rens W, O'Brien PC, Grutzner F, Clarke O, Graphodatskaya D, Tsend-Ayush E, et
al. The multiple sex chromosomes of platypus and echidna are not completely identical
and several share homology with the avian Z. Genome Biol. 2007;8:R243.
[32] Grutzner F, Rens W, Tsend-Ayush E, El-Mogharbel N, O'Brien PC, Jones RC, et
al. In the platypus a meiotic chain of ten sex chromosomes shares genes with the bird Z
and mammal X chromosomes. Nature. 2004;432:913-7.
[33] Veyrunes F, Waters PD, Miethke P, Rens W, McMillan D, Alsop AE, et al. Birdlike sex chromosomes of platypus imply recent origin of mammal sex chromosomes.
Genome Res. 2008;18:965-73.
[34] Simpson MJ, Simpson AE. Birth sex ratios and social rank in rhesus monkey
mothers. Nature. 1982;300:440-1.
[35] Silk JB, Clark-Wheatley CB, Rodman PS, Samuels A. Differential reproductive
success and facultative adjustment of sex ratios among captive female bonnet macaques
(Macaca radiata). Animal Behaviour. 1981;29:1106-20.
[36] Meikle DB, Tilford BL, Vessey SH. Dominance Rank, Secondary Sex Ratio, and
Reproduction of Offspring in Polygynous Primates. The American Naturalist.
1984;124:173-88.
[37] Thogerson CM, Brady CM, Howard RD, Mason GJ, Pajor EA, Vicino GA, et al.
Winning the genetic lottery: biasing birth sex ratio results in more grandchildren. PLoS
One. 2013;8:e67867.
[38] Fisher RA. The genetical theory of natural selection. New York: Dover
Publications; 1930.
[39] Trivers RL, Willard DE. Natural selection of parental ability to vary the sex ratio
of offspring. Science. 1973;179:90-2.
141
[40] Love OP, Chin EH, Wynne-Edwards KE, Williams TD. Stress hormones: a link
between maternal condition and sex-biased reproductive investment. Am Nat.
2005;166:751-66.
[41] Myers JH. Sex Ratio Adjustment Under Food Stress: Maximization of Quality or
Numbers of Offspring? The American Naturalist. 1978;112:381-8.
[42] Gomendio M, Clutton-Brock TH, Albon SD, Guinness FE, Simpson MJ.
Mammalian sex ratios and variation in costs of rearing sons and daughters. Nature.
1990;343:261-3.
[43] Wiebe KL, Bortolotti GR. Facultative sex ratio manipulation in American kestrels.
Behavioral Ecology and Sociobiology. 1992;30:379-86.
[44] Martin JG, Festa-Bianchet M. Sex ratio bias and reproductive strategies: what sex
to produce when? Ecology. 2011;92:441-9.
[45] Tilbrook AJ, Turner AI, Clarke IJ. Effects of stress on reproduction in non-rodent
mammals: the role of glucocorticoids and sex differences. Rev Reprod. 2000;5:105-13.
[46] Rivier C, Rivier J, Vale W. Stress-induced inhibition of reproductive functions:
role of endogenous corticotropin-releasing factor. Science. 1986;231:607-9.
[47] Einarsson S, Brandt Y, Lundeheim N, Madej A. Stress and its influence on
reproduction in pigs: a review. Acta Vet Scand. 2008;50:48.
[48] Palmer AR. Quasireplicationandthecontract of error: lessons from sex ratios,
heritabilities and fluctuating asymmetry. Annual Review of Ecology and Systematics.
2000;31:441-80.
[49] Festa-Bianchet M, King WJ, Jorgenson JT, Smith KG, Wishart WD. The
development of sexual dimorphism: seasonal and lifetime mass changes in bighorn
sheep. Canadian Journal of Zoology. 1996;74:330-42.
[50] Kruuk LEB, Clutton-Brock TH, Albon SD, Pemberton JM, Guinness FE.
Population density affects sex ratio variation in red deer. Nature. 1999;399:459-61.
142
[51] Cockburn A, Legge S, Double M. Sex ratios in birds and mammals: can the
hypotheses be disentangled? In: Hardy I, editor. Sex ratios, concepts and research
methods. Cambridge, UK: Cambridge, University Press; 2002. p. 266-8.
[52] Rosenfeld CS, Roberts RM. Maternal diet and other factors affecting offspring sex
ratio: a review. Biol Reprod. 2004;71:1063-70.
143
Letter in preparation:
Deborah Toledo-Flores, Frank Grützner. Identification of a Sox3 deletion in a captivebred echidna.
145
Identification of a Sox3 deletion in a captive-bred echidna
Authors:
Deborah Fernanda Toledo-Flores and Frank Grützner
Affiliations:
The Robinson Research Institute, School of Molecular and Biomedical Science, The
University of Adelaide, Adelaide, Australia
Abstract:
In mice and humans, Sox3 deletions have been associated with X-linked
hypopituitarism. A deletion of the therian proto-Sry gene, Sox3, was identified in a
captive born echidna at the Perth Zoo following genetic sexing from hair genomic DNA
(Toledo-Flores et al, in preparation, chapter 5). Previous work indicates that in platypus,
a male-specific Sox3 allele could potentially be involved in male sex-determination
(Toledo-Flores and Grützner, in preparation, chapter 4). Work described in the first part
of this chapter revealed a marked female bias in captive bred echidnas. In this section
we describe the extraordinary finding that the only male identified in this population
lacked the Sox3 gene. The discovery of this deletion in a male echidna provides a
unique opportunity to investigate the role, if any, of Sox3 in male sexual development
and fertility.
Key words: Sex determination, echidna, Sox3, male deletion, captivity, female bias.
146
Over many decades, researchers have been interested in the mechanisms of sex
determination in monotremes. These unusual mammals have a rather peculiar sex
chromosome system composed of multiple sex chromosomes that have been found to be
homologous to sex chromosomes in birds and not in therian mammals [1-4]. So far the
best male sex-determining candidate in monotremes is Amhy located on one of the Y
chromosomes [5]. Sox3 is also a very interesting gene, since the therian male sex
determination gene Sry evolved from it [6-8]. Sox3 is on the platypus chromosome 6,
which represents the therian proto-sex chromosome [9-11]. The autosomal location of
this gene excluded the existence of a Sry orthologue in the platypus and therefore, the
relevance of Sox3 in platypus sex determination has been overlooked.
It has been recently described that there is a male specific allele of Sox3 in the platypus
(Toledo-Flores and Grützner, in preparation, chapter 4). Furthermore, it was found that
chromosome 6 in the platypus shows signs of divergence; it has a number of sex
specific characteristics and it is non-randomly segregated during male meiosis (Toledo
Flores et al, in prepartion, chapter 3). Therefore, it has been hypothesized that
chromosome 6 might have commenced differentiation prior to monotreme therian
divergence. This, together with the evidence of a male-specific Sox3 in the platypus,
might suggest that Sox3 is involved in monotreme sex determination. However, due to
the lack of the full Sox3 sequence in both platypus and echidna, no male-specific alleles
have yet been characterized in the later.
A recent study in captive born echidnas established a protocol to genetically sex
echidna juveniles from hair-extracted genomic DNA samples (Toledo-Flores et al, in
preparation, chapter 5). This study found a striking sex ratio bias in these captive born
animals, they observed that seven out of eight individuals were born genetically females
and only one was genetically male. A PCR utilizing Sox3 nested primers was performed
147
in two of the eight sexed echidna juveniles (echidna number seven and echidna number
eight). Fig. 1A shows firstly the control PCR using !-actin primers for a male and a
female echidna control genomic DNA as well as for echidna number seven and eight;
secondly, it shows the sexing PCR using the male specific primers amplifying Crspy
(both control and sexing PCRs were taken from Toledo Flores et al, in preparation,
chapter 5), which demonstrates that echidna seven is female and echidna eight is male;
thirdly, it shows two sets of results for amplification with primers Sox3 – 1 and Sox3 –
2. The later demonstrates that this Sox3 region is depleted in echidna number eight,
whereas it is present in the rest of the samples. Interestingly, this Sox3 region contains
part of the HMG domain as well as the only two poly-alanine tracts present in the
platypus Sox3 (Fig. 1B). Therefore, even though the rest of the Sox3 sequence was
present in this individual, deletion of these functional domains could dramatically affect
the activity of Sox3, if not completely abolish it.
The identification of this naturally occurring Sox3 deletion can thus provide a unique
opportunity to study the impact that such a modification/deletion could have in the sex
development and fertility of a male echidna. Identifying the parents of this animal and
their genotype, in addition to following the development of this individual as well as
measuring hormonal levels and investigating whether adjacent regions are also deleted
could make a significant contribution to our understanding of monotreme sex
determination and the role that Sox3 might play on it.
148
Figures
A
!
B
" 7
Sox3 - 1
CrspY
B-actin
8 -
!
" 7
8 -
HMG
!
" 7
1st PolyA
Sox3 - 2
8 -
!
"
7
8
-
2nd PolyA
Unknown genomic sequence
Sequence amplify by primers Sox3-1
Sequence amplify by primers Sox3-2
Figure 1. Sox3 deletion in a male echidna. (A) PCR results for an autosomal positive
control, !-actin; a male-specific gene, Crspy; and two sets of Sox3-specific primers,
Sox3 – 1 and Sox3 – 2. !: control male genomic DNA; ": control female genomic
DNA; 7: echidna number seven; 8: echidna number eight; - : negative control. (B)
Structure of echidna/platypus Sox3. The gray area represents the extend of unknown
sequence; the orange region, the sequence amplified by primers Sox3 – 1 and the blue
region, the sequence amplified by primers Sox3 – 2. The HMG domain and the two
polyalanine tracts are indicated by black lines labeled as HMG, 1st PolyA and 2nd PolyA,
respectively.
149
References
[1] Grutzner F, Rens W, Tsend-Ayush E, El-Mogharbel N, O'Brien PC, Jones RC, et al.
In the platypus a meiotic chain of ten sex chromosomes shares genes with the bird Z
and mammal X chromosomes. Nature. 2004;432:913-7.
[2] Rens W, Grutzner F, O'Brien P C, Fairclough H, Graves JA, Ferguson-Smith MA.
Resolution
and
evolution
of
the
duck-billed
platypus
karyotype
with
an
X1Y1X2Y2X3Y3X4Y4X5Y5 male sex chromosome constitution. Proc Natl Acad Sci
U S A. 2004;101:16257-61.
[3] Rens W, O'Brien PC, Grutzner F, Clarke O, Graphodatskaya D, Tsend-Ayush E, et
al. The multiple sex chromosomes of platypus and echidna are not completely identical
and several share homology with the avian Z. Genome Biol. 2007;8:R243.
[4] Veyrunes F, Waters PD, Miethke P, Rens W, McMillan D, Alsop AE, et al. Birdlike sex chromosomes of platypus imply recent origin of mammal sex chromosomes.
Genome Res. 2008;18:965-73.
[5] Cortez D, Marin R, Toledo-Flores D, Froidevaux L, Liechti A, Waters PD, et al.
Origins and functional evolution of Y chromosomes across mammals. Nature.
2014;508:488-93.
[6] Sinclair AH, Berta P, Palmer MS, Hawkins JR, Griffiths BL, Smith MJ, et al. A
gene from the human sex-determining region encodes a protein with homology to a
conserved DNA-binding motif. Nature. 1990;346:240-4.
[7] Foster JW, Graves JA. An SRY-related sequence on the marsupial X chromosome:
implications for the evolution of the mammalian testis-determining gene. Proc Natl
Acad Sci U S A. 1994;91:1927-31.
[8] Stevanovic M, Lovell-Badge R, Collignon J, Goodfellow PN. SOX3 is an X-linked
gene related to SRY. Hum Mol Genet. 1993;2:2013-8.
150
[9] Wallis MC, Waters PD, Delbridge ML, Kirby PJ, Pask AJ, Grutzner F, et al. Sex
determination in platypus and echidna: autosomal location of SOX3 confirms the
absence of SRY from monotremes. Chromosome Res. 2007;15:949-59.
[10] Waters PD, Delbridge ML, Deakin JE, El-Mogharbel N, Kirby PJ, Carvalho-Silva
DR, et al. Autosomal location of genes from the conserved mammalian X in the
platypus (Ornithorhynchus anatinus): implications for mammalian sex chromosome
evolution. Chromosome Res. 2005;13:401-10.
[11] Hore TA, Koina E, Wakefield MJ, Marshall Graves JA. The region homologous to
the X-chromosome inactivation centre has been disrupted in marsupial and monotreme
mammals. Chromosome Res. 2007;15:147-61.
151
CHAPTER 6: Significance and future directions
This chapter consists of a conventional significance and future directions section.
Chapter overview
In this chapter, I briefly summarize the main results of my projects and the significance
these results have on the field.
152
Significance and future directions
Monotremes represent one of the three major mammalian lineages. With genomic data
sets increasingly being accessible they have also become a valued species in
comparative genomics. Work on their sex chromosome system has revolutionized our
understanding of mammalian sex chromosome evolution. In addition they are the only
mammalian species in which the therian sex chromosome is autosomal. Sex
determination in monotremes has been a mystery and the work presented here provides
novel insights into possible genes involved in sex determination as well as a new
perspective into sex chromosome evolution in mammals. The mapping of Amhy onto a
Y chromosome provides a key result that makes this gene a worthy candidate as a gene
important for male sex determination in monotremes. This gives a new direction to the
field of monotreme sex determination making of Amhy the most likely sex determining
candidate gene in monotremes, potentially mimicking the function of male-specific
copy of amh in the teleost fish Patagonian pejerrey. Future work should focus on
finding functional evidence to support the involvement of Amhy in male sex
determination. The biggest challenge in these studies is the lack of platypus embryonic
material as well as the fact that genetic and hormonal manipulations in the platypus are
not possible. Therefore, it would be necessary to find a suitable organism or in vitro
system to model the effects of the platypus anti-Müllerian hormone.
Furthermore, by studying the therian proto-sex chromosome, chromosome 6, I
discovered several lines of evidence that this is not an ordinary autosome. Firstly I
found a unique case of non-random segregation of this autosome in platypus male
meiosis. Secondly the extent in NOR heteromorphism is different in males and females
and thirdly DNA demethylation has an effect on the heteromorphism only females. This
might suggest that chromosome 6 is undergoing a process of differentiation and also
153
might show differential epigenetic changes in males and females. Future work may
investigate the mechanisms underlying this segregation bias and identify further
evidence of differentiation. Particularly interesting would be to see if recombination
occurs along the whole length of chromosome 6.
One would predict that lack of recombination is responsible for my finding of a malespecific allele of Sox3 in the platypus. Since the observed variation consists of a two
amino acid reduction in one of the two platypus Sox3 polyalanine tracts, this raises the
possibility that such a change might confer a male-specific function that in turn could be
involved in male sex determination or differentiation. Given the GC-rich nature of this
gene and the fact that successful Sox3 amplification from platypus gDNA or cDNA is
highly inconsistent, manipulating this gene and further investigating its potential to
upregulate Sox9 activity in vitro has been technically challenging and the planned
cloning and testing of the Sox3 deletions in a TESCO reporter gene assay where beyond
the timeframe imposed by my candidature. In the future, it is necessary to obtain the full
length of Sox3 in the platypus to test through reporter gene assay whether it can up
regulate Sox9 in comparison to the female-occurring alleles. Alternatively, to mimic this
change in the human or mouse Sox3 and to assess how this alters its transcriptional
activity would also shed-light on the impact of this variation in the platypus.
Finally, I applied my research to assist zoos in determining the sex of young echidna
puggles. Importantly, this technique has the potential to be used to study echidnas in the
wild, including the critically endangered long-beaked echidna. This work revealed a
strong female sex bias of echidnas born in captivity. This highlights the need to monitor
sex rations in captively bred species as a possible indicator of the effects of husbandry
on the breeding population. I found the first reported naturally occurring Sox3 deletion
154
in monotremes. This provides a unique opportunity to study the effects of such a
deletion in a male echidna in vivo.
In conclusion, this work provided novel insights into sex chromosomes and sex
determination in monotremes. It also highlights the need to further research into
chromosome 6 as therian proto-sex chromosome. In addition to this basic research a
useful approach to sex echidnas has been developed and applied revealing unmarked
sex bias in a captive bred echidna population. This research provides more evidence for
the extraordinary genetics found in monotremes, which nonetheless provides vital
information about mammalian evolution. It also highlights that much is to be done to
understand the biology of these fascinating creatures at the molecular level.
155
Amendments
Page 1, paragraph 2
Read:
In addition, this autosome features a large heteromorphic NOR and associates with the
sex chromosomes during male meiosis (Casey and Daish personal communication).
Changed to:
In addition, this autosome features a large heteromorphic nucleolus organizer region
(NOR) and associates with the sex chromosomes during male meiosis (Casey and Daish
personal communication).
Page 71, figure 3
Modification:
Error bars and sample sizes (n) were added to figure 3.
Page 60, paragraph 1
Read:
We found that in females, both NORs are entirely covered by silver staining (Figure
3A). In males, we observed that one of the homologous NORs bears a gap of unstained
chromatin surrounded by silver deposits over the rest of the NOR (Figure 3B).
Changed to:
156
We found that in females, both NORs are entirely covered by silver staining (Figure
4A). In males, we observed that one of the homologous NORs bears a gap of unstained
chromatin surrounded by silver deposits over the rest of the NOR (Figure 4B).
Page 75, figure 7
Modification:
Average columns, error bars and sample sizes (n) were added to figure 7.
Page 91, third paragraph
Read:
It is known that Sox3 is normally expressed in the developing central nervous system [4,
13-15] and together with Sox1 and Sox2 participates in the initiation and progression
of neuronal differentiation [16].
Changed to:
It is known that Sox3 is normally expressed in the developing central nervous system in
mouse [4, 13-15] and together with Sox1 and Sox2 participates in the initiation and
progression of neuronal differentiation [16].
Page 95, first paragraph
Read:
The
primers
used
were
SoxF-GCACAACTCGGAGATCAGCAAG,
SoxR-
GGCAGGTACATGCTGATCATATC at 59ºC (expected size ~600bp) or Sox3FCTACTCGCTGATGCCCGACC and Sox3R-GAGCTGGGCTCCGACTTGAC at 59ºC
157
(expected size ~300bp);
!–actin-F-GCCCATCTACGAAGGTTACGC and !–actin-
AAGGTCGTTTCGTGGATACCAC at 55ºC. A 1.5% agarose gel was used to visualize
the products.
Changed to:
The
primers
used
were
SoxF-GCACAACTCGGAGATCAGCAAG,
SoxR-
GGCAGGTACATGCTGATCATATC at 59ºC (expected size ~600bp) or the genespecific
Sox3F-CTACTCGCTGATGCCCGACC
and
Sox3R-
GAGCTGGGCTCCGACTTGAC at 59ºC (expected size ~300bp), designed based on
ENSOANG00000002448; !–actin-F-GCCCATCTACGAAGGTTACGC and !–actinAAGGTCGTTTCGTGGATACCAC at 55ºC. A 1.5% agarose gel was used to visualize
the products.
Page 99, last paragraph
Read:
However, these different attempts failed due to the inconsistency of the Sox3 PCR,
which is likely a result of the high GC content of this gene (Fig. 6). However a number
of Sox3 fragments were amplified from different individuals.
Changed to:
However, these different attempts failed due to the inconsistency of the Sox3 PCR,
which is likely a result of the high GC content of this gene (Fig. 6). Importantly, !–actin
was used as a control to ensure that the genomic DNA was suitable for PCR
amplification. However a number of Sox3 fragments were amplified from different
individuals.
158
Page 100, paragraph 1
Read:
Specifically this variant was defined by a reduced second polyalanine tract.
Changed to:
Specifically this variant was defined by a reduced second polyalanine tract (fourth tract
in conserved terms).
Read:
Interestingly, we found that the seven-alanine Sox3 was found exclusively in males and
not in females; five females were sequenced and all of them carried a nine-alanine Sox3
variant whereas four out of five males carried the seven-alanine Sox3 (Fig. 7).
Changed to:
Interestingly, we found that the seven-alanine Sox3 was found exclusively in males and
not in females; five females were sequenced and all of them carried a nine-alanine Sox3
variant whereas four out of five males carried the seven-alanine Sox3 and one of them
carried a five-alanine tract (Fig. 7).
Page 108, figure 5
Modification:
The following sentence was added to the end of Fig. 5 legend: “White arrow indicates
Sox3 expected size.”
Page 110, figure 7
159
Legend read:
Figure 7. A Sox3 male-specific allele was identified in the platypus. A fragment of Sox3
was amplified in five different females and five different males. A Sox3 coding for seven
alanines on its 2nd polyalanine tract was detected only in males and not in females.
Legend changed to:
Figure 7. A Sox3 male-specific allele was identified in the platypus. A fragment of
Sox3 was amplified in five different females and five different males. A Sox3 coding for
seven alanines on its 2nd polyalanine tract (4th in conserved terms) was detected only in
males and not in females.
Page 127, first paragraph
Read:
The sex of the individuals was then determined through PCR using primers that amplify
a platypus male-specific gene, Crspy. Crspy, the male specific copy of the Crsp70 gene
that is located on platypus chromosome Y5 [28], which has been shown to be
homologous to the echidna male specific chromosome Y3 [31]. These primers were
tested in echidnas whose sex was known, confirming that the platypus Crspy primers
amplify a male specific product and no products in females (Fig. 3). Running this PCR
on the DNA of eight echidna puggles showed that seven out of the eight captive-born
echidnas were females (Table 1).
Changed to:
The sex of the individuals was then determined through PCR using primers that amplify
a platypus male-specific gene, Crspy. Crspy, the male specific copy of the Crsp70 gene
160
that is located on platypus chromosome Y5 [28], which has been shown to be
homologous to the echidna male specific chromosome Y3 [31]. These primers were
tested in echidnas whose sex was known, confirming that the platypus Crspy primers
amplify a male specific product and no products in females (Fig. 3). Running this PCR
on the DNA of eight echidna puggles showed that seven out of the eight captive-born
echidnas were females (Table 1). Importantly, !–actin primers were used as a positive
control.
Page 129, second paragraph
Read:
In general, they all predict that manipulation of the sex ratio by an individual should
lead to a greater offspring production through the favored sex, increasing in this way
its fitness [37].
Changed to:
In general, they all predict that manipulation of the sex ratio by an individual should
lead to a greater offspring production (hence, increased fitness) through the favored sex
[37].
Page 153, first paragraph
Read:
This gives a new direction to the field of monotreme sex determination. Future work
should focus on finding functional evidence to support the involvement of Amhy in male
sex determination.
161
Changed to:
This gives a new direction to the field of monotreme sex determination making of Amhy
the most likely sex determining candidate gene in monotremes, potentially mimicking
the function of male-specific copy of amh in the teleost fish Patagonian pejerrey. Future
work should focus on finding functional evidence to support the involvement of Amhy
in male sex determination.
Page 154, second paragraph
Read:
Given the GC-rich nature of this gene and the fact that successful Sox3 amplification
from platypus gDNA or cDNA is highly inconsistent, manipulating this gene and further
investigating its potential to upregulate Sox9 activity in vitro has been technically
challenging and the planned cloning and testing of the Sox3 deletions in a TECO
reporter gene assay where beyond the timeframe imposed by my candidature.
Changed to:
Given the GC-rich nature of this gene and the fact that successful Sox3 amplification
from platypus gDNA or cDNA is highly inconsistent, manipulating this gene and
further investigating its potential to upregulate Sox9 activity in vitro has been
technically challenging and the planned cloning and testing of the Sox3 deletions in a
TESCO reporter gene assay where beyond the timeframe imposed by my candidature.
162