Download Evolution of meiosis genes in sexual vs. asexual Potamopyrgus

University of Iowa
Iowa Research Online
Theses and Dissertations
Spring 2015
Evolution of meiosis genes in sexual vs. asexual
Potamopyrgus antipodarum
Christopher Steven Rice
University of Iowa
Copyright 2015 Christopher Steven Rice
This thesis is available at Iowa Research Online: http://ir.uiowa.edu/etd/1739
Recommended Citation
Rice, Christopher Steven. "Evolution of meiosis genes in sexual vs. asexual Potamopyrgus antipodarum." MS (Master of Science)
thesis, University of Iowa, 2015.
http://ir.uiowa.edu/etd/1739.
Follow this and additional works at: http://ir.uiowa.edu/etd
Part of the Biology Commons
EVOLUTION OF MEIOSIS GENES IN SEXUAL VS. ASEXUAL
POTAMOPYRGUS ANTIPODARUM
by
Christopher Steven Rice
A thesis submitted in partial fulfillment
of the requirements for the Master of
Science degree in Biology
in the Graduate College of
The University of Iowa
May 2015
Thesis Supervisors: Associate Professor John M. Logsdon, Jr.
Associate Professor Maurine Neiman
Copyright by
CHRISTOPHER STEVEN RICE
2015
All Rights Reserved
Graduate College
The University of Iowa
Iowa City, Iowa
CERTIFICATE OF APPROVAL
MASTER'S THESIS
This is to certify that the Master's thesis of
Christopher Steven Rice
has been approved by the Examining Committee for the
thesis requirement for the Master of Science degree in
Biology at the May 2015 graduation.
Thesis Committee:
John M. Logsdon Jr., Thesis Supervisor
Maurine Neiman, Thesis Supervisor
Bryant McAllister
ACKNOWLEDGMENTS
I would like to acknowledge Dr. Maurine Neiman and Dr. John Logsdon
for their outstanding mentorship and guidance in the completion of this project. I
would like to thank all members of the Neiman and Logsdon labs, for your
incredibly helpful contributions and advice. I would like to thank Dr. Jeffrey Boore
for his assistance in our bioinformatics endeavors, the University of Iowa Biology
Department, and the National Science Foundation for their support. Lastly I'd like
to thank my thesis committee for their insight and assistance in the completion of
my research.
ii
ABSTRACT
How asexual reproduction affects genome evolution, and how organisms
that are ancestrally sexual alter their reproductive machinery upon becoming
asexual are both central unanswered questions in evolutionary biology. While these
questions have been addressed to some extent in organisms such as asexual clams,
rotifers, ostracods, arthropods, and fungi, the most powerful and direct tests of how
sex and its absence influence evolution requires direct comparisons between closely
related and otherwise similar sexual and asexual taxa. Here, I quantify the rates and
patterns of molecular evolution in the meiosis-specific genes Msh4, Msh5, and
Spo11 in multiple sexual and asexual lineages of Potamopyrgus antipodarum, a
New Zealand freshwater snail. Because asexual P. antipodarum reproduce
apomictically (without recombination), genes used only for meiosis should be under
relaxed selection relative to meiosis-specific genes in sexual P. antipodarum,
allowing me to directly study how asexuality affects the evolution of meiosisspecific genes. Contrary to expectations under relaxed selection, I found no
evidence that these meiosis-specific genes are degrading in asexual P. antipodarum;
instead they display molecular patterns consistent with purifying selection. The
presence of intact meiosis-specific genes in asexual P. antipodarum hints that the
asexuals may maintain the ability to perform meiosis despite reproducing
apomictically. Asexual meiotic capability suggests that some meiotic components
may persist or acquire a new role in these asexuals.
iii
PUBLIC ABSTRACT
How asexual reproduction affects genome evolution, and how ancestrally sexual
organisms alter their reproductive machinery upon becoming asexual are central
unanswered questions in evolutionary biology. While these questions have been addressed
to some extent in different asexual species, the most powerful tests of how sex and its
absence influence evolution requires direct comparisons between closely related and
otherwise similar sexual and asexual taxa. I address these questions by studying the
evolution of genes critical to sexual reproduction in sexual and asexual lineages of
Potamopyrgus antipodarum, a New Zealand freshwater snail. Because genes used only for
sexual reproduction should have little or no functional relevance for asexuals, I can use this
approach to study the effects of asexuality on the evolution of genes required only for
sexual reproduction, which has implications for genome evolution in the absence of sex. I
discovered that genes necessary for sexual reproduction show no evidence for being lost in
asexual P. antipodarum. These results suggest asexuality may result in the loss of some,
but not all, components of sexual reproductive machinery or that not enough time has
passed for meiosis genes to be lost. Comparing evolutionary patterns between sexual and
asexual organisms will help the scientific community understand the benefits and
limitations of reproducing sexually. Understanding the consequences of sexual
reproduction will illuminate fundamental biological and human health questions, ranging
from the importance of biological diversity to the source and transmission of disease to
dynamics between humans (who reproduce sexually) and parasites (which are often
asexual).
iv
TABLE OF CONTENTS
List of Tables ............................................................................................................................... vi
List of Figures ............................................................................................................................ vii
Chapter 1: Introduction .................................................................................................................1
Determining Evolutionary Consequences of Asexuality for Meiosis-Specific Genes ..............1
Evolution of Genes under Relaxed Selection.............................................................................2
Meiosis in Sexual Reproduction ................................................................................................3
Asexual Reproduction ................................................................................................................5
Using the Meiosis Detection Toolkit to Detect Evidence of Relaxed Selection........................6
Study System: Potamopyrgus antipodarum ..............................................................................8
Summary ..................................................................................................................................10
Chapter 2: Materials and Methods ..............................................................................................11
Genome Sequencing and Assembly .........................................................................................11
Gene Assembly and Annotation ..............................................................................................11
PCR Amplification and Gel Electrophoresis of Meiotic Genes...............................................13
Cloning of PCR Products .........................................................................................................20
Screening of Clones .................................................................................................................20
Plasmid Extraction and Preparation .........................................................................................23
Processing and Analysis of Sanger Sequences ........................................................................23
Chapter 3: Results .......................................................................................................................26
Accumulated Mutations of Sexual and Asexual Potamopyrgus antipodarum ........................26
Evolutionary Rates of Sexual vs. Asexual Genes ....................................................................26
Chapter 4: Discussion..................................................................................................................35
Chapter 5: Summary....................................................................................................................38
References ...................................................................................................................................39
v
LIST OF TABLES
Table
1. Expected Sequence Patterns for Corresponding Modes of Selection. ..............................9
2. Msh4 PCR Primers. .........................................................................................................14
3. Msh5 PCR Primers. .........................................................................................................15
4. Spo11 PCR Primers.........................................................................................................16
5. Stock and Working Concentration of PCR Reagents. ....................................................17
6. Thermocycler Program used in PCR Reactions. .............................................................18
7. Stock and Working Concentrations of PCR Screening Reagents. ..................................21
8. Thermocycler Program used in PCR Screening Reactions. ............................................22
9. IUPAC Degenerate Codes at Heterozygous Sites for Maximum Likelihood Analysis. .25
10. Open Reading Frames in Sexual and Asexual P. antipodarum. .....................................27
vi
LIST OF FIGURES
Figure
1. Mitochondrial Phylogeny of Potamopyrgus antipodarum....................................... 12
2. Gene Models and PCR Strategy for Meiosis-Specific Genes. ................................. 19
3. Msh4 Polymorphism. ............................................................................................... 28
4. Msh5 Polymorphism. ............................................................................................... 29
5. Spo11 Polymorphism. .............................................................................................. 30
6. Maximum Likelihood Analysis of Msh4 ................................................................. 31
7. Maximum Likelihood Analysis of Msh5 ................................................................. 32
8. Maximum Likelihood Analysis of Spo11. ............................................................... 33
9. Evolutionary Rate of Sexual and Asexual Msh5 Coding Region. ........................... 34
vii
Chapter 1: Introduction
Determining Evolutionary Consequences of Asexuality for Meiosis-Specific Genes
My thesis will address the consequences of asexuality for genome evolution by studying how
meiosis genes critical to sexual reproduction evolve in asexual lineages. I have compared meiotic
genes of multiple diverse obligate sexual and obligate asexual lineages of the snail
Potamopyrgus antipodarum. My research aims to capture the early evolutionary genomic
consequences of asexuality, which are largely unstudied in eukaryotes outside of arthropods,
clams, rotifers, ostracods, and fungi (reviewed by Nomark 2003, Schurko et al. 2009, Schurko et
al. 2010). By comparing young and relatively old asexual lineages I will provide novel
observations of evolution within meiosis genes hypothesized to be under relaxed selection in
asexuals. Comparing meiosis gene evolution between reproductive modes in P. antipodarum is a
powerful strategy in this context because asexual organisms typically do not require meiosis to
reproduce (Naumova et al. 2001, Hojsgaard et al.2013, Wang et al. 2014), meaning that genes
only used for meiosis should experience relaxed selective constraint relative to their sexual
counterparts.
How relaxed selection affects molecular evolution in meiosis genes has yet to be addressed in
systems that allow direct comparisons between sexuals and otherwise similar asexuals. P.
antipodarum features closely related, coexisting, and ecologically similar sexual and asexual
individuals, meaning I can directly compare the evolution of orthologs in sexually vs. asexually
transmitted genomes. These comparisons will provide powerful tests of hypotheses for
pseudogenization and decay of asexual meiosis genes.
The expectations of meiosis gene decay in asexuals are complicated by the fact that some
asexual lineages use alternate forms of meiosis, likely mediated by modification to subsets of
proteins within otherwise maintained core meiotic machinery (Srinivasan et al. 2010, Hanson et
al. 2013). These asexuals engender important evolutionary questions regarding evolution of
meiosis-specific genes following asexuality, and whether the retention of at least some
components of meiosis can allow asexual lineages to clear harmful mutations and increase the
efficacy of selection. By comparing the evolution of meiosis genes between sexual vs asexual,
1
and young vs old asexual P. antipodarum I can determine: 1) whether meiotic genes in asexual
lineages are in the act of accumulating mutations, which will illuminate whether meiosis is being
lost, modified, or maintained, 2) how observed changes to meiotic machinery compensate for
transitions to asexuality, and 3) extrapolate how changes to meiotic genes could explain observed
patterns of polymorphism and substitution in asexuals.
Evolution of Genes under Relaxed Selection
I used comparative analysis of the rates and patterns of molecular evolution in meiosis-specific
genes in obligately sexual and obligately asexual lineages of the snail Potamopyrgus
antipodarum to determine whether genes involved in meiosis evolve under relaxed selection in
asexuals. Genes under relaxed selection are genes that accumulate mutations with little to no
fitness cost to an organism. Over time, these genes acquire mutations that are expected to result
in pseudogenization and subsequent degradation as mutations continue to accumulate (Lynch &
Conery 2000, Zhang 2003, Lahti et al. 2009). Pseudogenes then, are the no-longer-functional
remnants of functional genes that have experienced a shift from selection to maintain the gene to
relaxed selective constraints. The gene-to-pseudogene transition happens when the gene is
rendered nonfunctional by one or more mutations (e.g. a frameshift or nonsense mutation) that,
because selection is relaxed, can occur with little or no fitness cost to the organism (Li et al.
1981, Gojobori et al. 1982). The pseudogene will continue to accumulate silent mutations until
the original functional gene is no longer identifiable. Multiple studies have assessed the fate of
genes under relaxed selective constraints following gene duplications (Mungpakdee et al 2008,
Sur et al 2013, Yampolsky & Bouzinier 2014,) and after the loss of ancestral characters such as
opsin genes in cave fish (Yokoyama et al 1995) or tooth loss in birds (Meredith et al 2014) .
These studies indicate the common fate of a gene under relaxed selection is pseudogenization
followed by sequence degradation. Even so, degradation and eventual loss of pseudogenes is not
always a foregone conclusion. For example, recent studies have suggested that some
pseudogenes maintain expression in some organisms (Birney et al. 2007), have roles in negative
(Tam 2008) and positive (Guo & Zhang 2011) gene regulation, and may be resurrected to
perform a particular function (Chen et al 2011).
2
Genes used only for meiosis are expected to experience relaxed selective constraints in asexuals
that do not use meiosis relative to the same genes in sexual counterparts. Some of these meiosisspecific genes are part of a Meiosis Detection Toolkit (MDT) (Schurko & Logsdon 2008) that
can and has been used to assess whether there is a potential for meiosis, which is supported by
the presence of conserved genes with open reading frames known only to perform vital meiotic
functions. I identified genes from this toolkit in the genomes of sexual and asexual P.
antipodarum and then used comparative analyses of the rates and patterns of molecular evolution
in these genes to: 1) assess their meiotic capability and 2) characterize the type and strength of
selection experienced by these genes. This allowed me to determine whether selection was
relaxed in asexuals relative to sexuals, and if genes had experienced pseudogenization in the
asexuals. This study is the first to my knowledge to assess the genic consequences of relaxed
selection for meiosis-specific genes in a natural system comprised of closely related and
coexisting sexual and asexual lineages. This study is also novel in its potential to capture
meiosis-specific genes in the act of degeneration in recently derived asexual lineages.
Meiosis in Sexual Reproduction
Sexual reproduction is the process by which distinct genetic material is combined
(conventionally by two mating types each contributing respective genetic material) to create
genetically unique offspring. This process is accomplished by the production of specialized
gametes, with one gamete type produced by each mating type, which fuse to create the initial
zygote. A successful mature gamete requires altered amounts of DNA (typically a reduction to
about half of the parental genetic material) and is usually genetically unique relative to either
parent. Eukaryotes have evolved a highly conserved specialized process of cellular division,
meiosis (see Dacks & Roger 1999, Villeneuve & Hillers 2001, Ramesh et al 2005, Malik et al
2008), that creates reduced gametes via a reductional division with novel combinations of
genetic material via recombination within progenitor cells (meiocytes). The reduction of ploidy
and novel allele combinations within each gamete followed by their subsequent fusion facilitates
restoration of ploidy and production of a new genetically unique offspring.
Reductional division in meiosis is typically accomplished by one phase of DNA replication at S
3
phase, creating chromatids in meiocytes, followed by two phases of cellular division known as
Meiosis I and Meiosis II. Meiosis I results in reduced meiocytes with recombinant chromosomes
via the process of crossover. Crossover first requires double-stranded breaks and resection of
chromosomes. Single strands of DNA at break points, created by resection, then invade
homologous chromosomes to initiate homologous repair by DNA polymerase culminating in
Holliday Junctions. After repair, Holliday Junctions are resolved to create recombinant or nonrecombinant chromosomes. Homologous chromosomes then segregate into daughter cells that
undergo a second division during Meiosis II separating sister chromatids (reviewed in Zickler &
Kleckner 1999). In animals, these daughter cells undergo final developmental stages of
spermatogenesis in males (Hecht 1998), oogenesis in females (Sagata 1996), or both
spermatogenesis and oogenesis within a hermaphroditic species (Singaravelu & Singson 2011),
to create mature gametes. External or internal fertilization facilitates the fusion of mature
gametes to complete the process of sexually producing a new individual.
Sexual reproduction has been extremely successful, evident in the ubiquity of sex throughout
eukaryotes despite its costs (Williams 1975, Smith 1978, Bell 1982). Because males cannot
directly produce offspring, the number of individuals with direct reproductive capability in a
dioecious sexual population is halved relative to asexuals (typically entirely female). If all else is
equal, this so-called "two-fold" cost of sex will result in substantially lower growth rate of the
sexual population relative to asexual competitors (Smith 1978). Energetic costs and exposure to
predators and parasites during copulation represents additional risks inherent in sexual
reproduction (Ronkainen & Yloneh 1994). From a genetic standpoint, recombination confers
additional costs via the potential to break up beneficial combinations of alleles at different loci
(Muller 1950). Recombination can also result in crossover and chromosomal segregation errors
that lead to aneuploidy, which itself often leads to fetus loss, debilitating fetal phenotypes, or
serious congenital defects. The existence of these substantial costs and risks associated with sex
suggest there must be significant benefits to sexual reproduction that would explain its ubiquity
in nature.
The predominance of sexual reproduction is attributed to several potential advantages. First, the
ability of sexual reproduction to produce and maintain genetic variation through meiotic
4
recombination can increase the efficiency of selection with respect to removing harmful
mutations and producing novel combinations of beneficial alleles (Smith 1978, Williams 1975,
Barton & Charlesworth 1998, Burt 2000, Connallon & Knowles 2007, Hill & Robertson 2007,
Hadany & Comeron 2008). Multiple studies have also shown that the variation generated by
sexual reproduction might help organisms escape parasites and diseases (Jokela et al. 2009,
Kerstes et al 2012, Mostowy & Engelstaedter 2012, Hogsdon & Otto 2012).
Asexual Reproduction
Asexual reproduction (parthenogenesis) is a mode of reproduction whereby females produce
genetically identical offspring. Asexuals typically avoid meiotic reduction (Suomalainen et
al.1976, Bell 1982, Mogie 1988), with the implication that genes used only for meiosis should
experience relaxed selection relative to these same genes in sexual relatives. To recapitulate,
asexual organisms should outcompete sexual counterparts due to higher rates of population
growth, maintenance of linkage between and reliable inheritance of beneficial allele
combinations, avoidance of meiotic errors resulting in aneuploidy, and the ability to reallocate
resources that otherwise should be needed for copulation or mating behavior to other processes,
and avoidance of risks associated with mating behavior like predator attack and disease
exposure. Even so, asexual lineages are typically short lived relative to sexual counterparts
(Williams 1975, Smith 1978, White 1978, Bell 1982) and seem to be recently derived from
sexual ancestors (Neiman et al 2005, Beck et al 2011). Why asexual lineages do not persist as
long as sexual lineages remains unclear (Schwander & Crespi 2009), but is often ascribed to
various negative genomic consequences of the absence of sex. These consequences include
genome-wide linkage disequilibrium which, while efficient at maintaining alleles that are
beneficial when linked, requires beneficial allele combinations to independently occur within a
single lineage (Kondrashov 1993, De Visser 2007, Desai et al 2007). Lack of recombination as a
consequence of asexuality has also been shown to result in mutation accumulation due to an
inability to clear harmful mutations from genomes (Lynch et al. 1993, Neiman et al 2010, Henry
et al 2012). These findings stress the importance of meiosis, and in particular recombination, in
sexual reproduction.
5
Meiosis-specific genes both tend to be conserved in eukaryotes and play an important functional
role in sexual reproduction (Villeneuve & Hillers 2001). As such, genes that are important to
meiosis (and are part of the Meiosis Detection Toolkit) should be maintained by selection in
sexual lineages. Asexual lineages are derived from sexual ancestors and should thus have
meiosis genes. Because these genes should experience relaxed selection following the transition
to asexuality, I predict that comparisons of sequence evolution in these genes between sexual and
asexual lineages will reveal signatures of relaxed functional constraint (e.g. non-synonymous
substitutions, nonsense mutations, and frame shift mutations) on meiosis-specific genes in
asexuals relative to sexuals.
Using the Meiosis Detection Toolkit to Detect Evidence of Relaxed Selection
The Meiosis Detection Toolkit (MDT) is a set of core meiosis proteins whose only chronological
and spatial expression is within cells undergoing meiotic division, and whose functional roles
have been empirically determined to only be in meiotic events such as crossover and
recombination (Schurko & Logsdon 2008). Because these genes are only involved in and are
absolutely critical to perform meiosis they are deemed meiosis-specific. The MDT is used to
determine if an organism is capable of meiosis by virtue of being able to locate homologues of
most, if not all, meiosis-specific genes in the organisms genome with an open reading frame.
Conversely, the inability to detect either meiosis-specific homologues with an open reading
frame or the homologues themselves indicates an inability to perform meiosis as relaxed
selection has removed meiosis-specific genes through deletion and/or pseudogenization followed
by sequence degradation. The MDT has been successfully applied in a variety of taxa such as
Trichomonas vaginalis (Malik et al. 2008), Daphnia (Schurko et al. 2009), choanoflagellates
(Carr et al. 2010), arthropods (Schurko et al.2010), mycorrhizal fungi (Rosendahl 2012), and
monogonot rotifers (Hanson et al. 2013) indicating the MDT can be successfully applied to
search genomes for these meiosis-specific genes, and can be used to infer if organisms are
potentially capable of Meiosis. I have applied the MDT to determine the presence/absence of
meiosis-specific genes in sexual and asexual lineages of our model system Potamopyrgus
antipodarum, and determined the asexual state of sequence degradation relative to sexuals in
meiosis-specific genes: Msh4, Msh5, and Spo11. These three genes are crucial in the initiation
6
and resolution of crossover events and proper homologous segregation, making them exemplar
meiosis-specific candidates for genes pseudogenizing under relaxed selection in asexuals.
Gene pseudogenization is one of several fates meiotic genes may undergo following a change in
reproductive mode (Normark et al 2003, Lunt 2008). A signature of pseudogenization is a nearequal occurrence of non-synonymous polymorphism per non-synonymous site (πa) and
synonymous polymorphism per synonymous site (πs) across the entire span of the gene. This
pattern would suggest relaxed selection because non-synonymous mutations are not being
preferentially removed and are nearly equivalent in frequency to neutral synonymous mutations.
Frameshift and nonsense mutations are also expected to accumulate as an altered or truncated
amino acid sequence would not have a deleterious effect on fitness. A second possible (though
likely rare) fate following relaxed selection would be neofunctionalization, or gain of function,
of a previously meiotic gene. Elevated levels of πa relative to πs are consistent with this
phenomenon, reflecting positive or diversifying selection for amino acid variants. It is also
possible that meiotic genes may be maintained by purifying selection in asexual lineages
(Charlesworth et al 1993, Som & Reyer 2007, D’Souza et al. 2010). Under purifying selection
most non-synonymous mutations will be eliminated from a population, meaning that πa will be
relatively low compared to πs. Selection favoring the maintenance of meiotic genes may be
generated by the presence of a meiotic step during the asexual production of gametes, e.g.
meiotic divisions that occur but afterwards restore ploidy (Marescalchi et al 1993, Liu et al 2007,
Sekin & Tojo 2010), or the existence of a secondary function of meiotic genes that is not
meiosis-specific (Lydall et al 1996, Matthews et al 1998, Peters et al 2010).
I have used the MDT to assess the type and magnitude of selection on meiosis-specific genes in
sexual and asexual P. antipodarum and have obtained evidence concerning the degree of relaxed
selection in asexual P.antipodarum. I accomplished these goals by quantifying the frequency of
πa relative to πs, quantifying the frequency of frameshift or nonsense mutations, and comparing
rates of evolution between sexual and asexual lineages using maximum likelihood analysis. I
then inferred the selective pressure on candidate meiosis genes by identifying expectations of
selective pressures, outlined in the previous paragraph, that were consistent with observed
sequence patterns (see Table 1 for summary of modes of selection and their associated patterns).
7
Study System: Potamopyrgus antipodarum
Potamopyrgus antipodarum is a freshwater snail native to New Zealand that is well suited for
addressing how relaxed selection influences molecular evolution in meiosis genes because
obligately sexual and asexual individuals often coexist and compete (Winterbourn 1970, Lively
1987, Lively 1992), allowing for direct and powerful comparisons between sexuals and asexuals
(Dybdahl & Lively 1995). While the reliable production of sexual offspring by sexual female P.
antipodarum and asexual offspring by asexual female P. antipodarum indicate that reproductive
mode is genetically determined (Wallace 1985, Phillips & Lambert 1989), how new asexual
lineages are produced from sexual ancestors and the specific mechanism by which asexual P.
antiopodarum produce offspring remain unclear. Evidence gathered from allozyme genotyping
of different P. antipodarum populations with high or relatively low male frequencies has
suggested that populations of high male frequency are sexual, while populations with very low
frequencies of males are populations where females reproduce apomictically (Phillips & Lambert
1989).
Despite these uncertainties, it is evident that new asexual P. antipodarum lineages have been
separately derived from sexual ancestors on multiple occasions (Neiman and Lively 2004,
Neiman et al 2011) allowing me to perform multiple experiments comparing sexual and asexual
meiotic genes. Because multiple transitions to asexuality have occurred at different times each
asexual lineage represents a unique experiment in studying the evolution of meiosis genes under
relaxed selection which may show different patterns in different lineages. I have used this study
system to sequence a set of core meiotic genes across several sexual and asexual lineages to
investigate evolutionary consequences of relaxed selection.
8
Table 1: Expected Sequence Patterns for Corresponding Modes of Selection.
Selection
Frameshift and Nonsense
Mutations
π𝑎
π𝑠
Rates of Evolution (relative to
sexual lineages)
Relaxed
Present
~1
Elevated
Purifying
Absent
<1
Proportional
Positive
Absent
>1
Elevated
9
Summary
Evolutionary consequences of asexuality on genome evolution and how asexuals utilize their
ancestral reproductive machinery is not well understood, despite the light this can shed on the
ubiquity, benefits, and limitations of sexual reproduction through comparisons of sexual and
asexual genome evolution. I have aimed to illuminate the genomic consequences of asexuality by
performing comparative evolutionary analysis of meiosis-specific genes between sexual and
asexual lineages of the snail P. antipodarum. By characterizing the selective pressures and
evolutionary patterns of meiosis genes in asexual lineages I address the consequences of
asexuality on a host of meiosis genes absolutely essential for sexual reproduction, but whose
function is not particularly necessary for asexual reproduction. The general lack of necessity for
meiosis in asexuality makes meiosis-specific genes powerful candidates for studying genome
evolution associated with asexuality because these genes are likely to be either placed under
relaxed selection or undergo some modification, non-exclusively resulting in meiosis-specific
gene loss or accumulation of non-synonymous mutations. I compared asexual lineages of
varying ages to capture the evolution of meiosis-specific genes ‘in the act’, providing novel data
regarding the process of mutation accumulation over time in asexually transmitted genomes. In
particular, I use molecular and bioinformatics techniques to infer: 1) the type and magnitude of
selective pressure on meiosis-specific genes in asexuals, 2) how observed changes to meiotic
machinery accommodate the transition to asexuality, and 3) extrapolate how changes to meiosisspecific genes explain observed patterns of polymorphism and substitution in asexual lineages.
Based on these results, I will then draw connections between observed evolutionary patterns in
meiosis-specific genes and the evolution of meiosis in asexual P. antipodarum.
10
Chapter 2: Materials and Methods
Genome Sequencing and Assembly
In collaboration with the rest of the P. antipodarum genome consortium, I used high-throughput
Illumina sequencing to sequence genomic libraries produced from three asexual P. antipodarum
lineages (Kaniere, Waikaremoana, and Poerua) and two sexual lineages (Ianthe and
Alexandrina). A lineage is defined as a line of snails descended from a single female. We used
the P. antipodarum mitochondrial phylogeny (Figure 1) to attempt to obtain sequence data from
lineages representing the range of genetic diversity in the species. This design also allowed us to
use comparisons between sexual vs. asexual, and old vs. new asexual lineages to capture patterns
of genome evolution across different times following transitions to asexuality.
We extracted DNA by dissecting away the head tissues of three snails from each of the five
lineages, pooling the heads together within each lineage, and then using a modified phenolchloroform protocol for DNA extraction. Next, we used the Nextera library preparation protocol
to create five libraries (one for each lineage), with approximately 200-600 base pair (bp) insert
sizes for each lineage. Each of the five libraries was barcoded and run in one lane of Illumina HiSeq. The five barcoded libraries were also pooled and run together in a sixth lane. All samples
were pair-end sequenced on the Illumina Hi-Seq, yielding sequence lengths of 101bp in each
direction. We used the CLC bio genomics workbench to filer and assemble sequence reads
within each lineage to create five genome assemblies (one for each lineage) ranging from ~15x
to ~17x coverage.
Gene Assembly and Annotation
I first created meiotic gene models by using meiotic protein queries from NCBI to locate genome
contigs with a matching DNA sequence within each of the five genome assemblies. Proteins of
genes with both meiotic and mitotic functions were also used as queries to locate matching
genome contigs as a control, since genes involved in meiosis and mitosis are expected to still be
11
Figure 1: Mitochondrial Phylogeny of Potamopyrgus antipodarum.
Maximum likelihood tree of concatenated mitochondrial genomes using a T93G model. Sexual
lineages are indicated in white, asexual lineages are black. Lineages sampled for genome
sequencing are indicated with the name of the lake from which they were sampled. Genomes not
sampled for genome sequencing are only indicated by a square. Tree provided by Joel
Sharbrough.
12
under purifying selection in asexuals. I computationally extracted matching contigs and then
used Sequencher (version 5.2) to assemble contigs from all five P. antipodarum genomes for
each specific gene to create one consensus sequence model for each gene. I then aligned each of
these models to sequences in the NCBI database to ensure that: 1) all exons in annotated NCBI
sequences aligned to meiosis gene models in the correct order, 2) meiosis gene models aligned to
their homologous genes in NCBI, indicating no incorporation into models of ectopic contigs
from other genes, and 3) meiotic gene models had the best alignments to sequences in NCBI
from other protostomes, indicating no foreign DNA contaminants. Next, I annotated each gene
model by computationally locating reads in a P. antipodarum transcriptome that aligned to
meiosis gene model queries. The transcriptome I used was based on RNA extracted from ovary
tissue of reproductively active female sexual and asexual Potamopyrgus antipodarum and
sequenced using Illumina technology. I computationally extracted reads from our transcriptome
and transferred these reads to Sequencher (version 5.2). I then aligned matching ovary
transcriptome reads back to each gene assembly to create annotated gene models with validated
intron-exon boundaries. Finally, I focused on annotated meiosis-specific gene models to design
PCR primers for each meiosis-specific gene.
PCR Amplification and Gel Electrophoresis of Meiotic Genes
I designed primers for the PCR amplification of Msh4, Msh5 and Spo11 (see Tables 2, 3, & 4)
with ~50% GC content and a melting temperature of ~56○C. Because these genes were to long
(~10 kilobases) to clone in one PCR reaction, I performed PCR (Tables 5 & 6) segmentally
across ~2,000 base pair overlapping increments along genes (See Figure 2). PCR amplification
was performed on DNA from six asexual and three sexual lineages for Msh4, six asexual and
three sexual lineages for Msh5, and 13 asexual and five sexual lineages for Spo11, these lineages
were ultimately used in comparative analyses. After PCR, I used gel electrophoresis with LE
agarose on 10µl of the 37µl of PCR product to determine whether amplified DNA was the
expected size, and assess whether ectopic amplification occurred in negative controls that
contained all reagents minus P. antipodarum DNA. If negative controls showed no DNA
amplification and PCR yielded amplified DNA of ~2,000bp I used low-melt gel electrophoresis
to purify the remaining PCR product. I then excised purified PCR product from the low melt gel
13
Table 2: Msh4 PCR Primers.
14
Table 3: Msh5 PCR Primers
15
Table 4: Spo11 PCR Primers
16
Table 5: Stock and Working Concentration of PCR Reagents.
Reagent
Stock Concentration
Working Concentration
MgCl2
25 mM
1.9mM
dNTPs
10 mM each
0.25mM each
5x
1x
5 U/µL
0.025 U/µL
2.5 U/µL
0.002 U/µL
5x Flexi buffer
Pro Taq
Pfu
Sdd H2O
1.314mM
Forward Primer
50 µM
0.8µM
Reverse Primer
50 µM
0.8µM
2 ng/µL
2ng
DNA
17
Table 6 : Thermocycler Program used in PCR Reactions.
Step
Temp. (○C )
Time (minutes)
1
95.0
2:00
2
94.0
1:00
3
53.5
1:30
4
72.0
3:30 (or 1 minute per kb)
5
Go to Step 2, Repeat 39x
6
72.0
10:00
7
4.0
Hold indefinitely
18
Figure 2: Gene Models and PCR Strategy for Meiosis-Specific Genes.
Annotated gene models created from tblastn searches of genome and transcriptome assemblies.
Introns: gray boxes, exons: black boxes, forward and reverse primers: white boxes. Forward
primers are distinguished with an ‘F’ followed by a specific number; reverse primers are
distinguished with an ‘R’ followed by a specific number. Step-wise amplification of genes
shown by white boxes below models, with a letter designation and size. Amplification of Msh4
(figure2a) was done in 7 PCR reactions. Msh5 (figure 2b) was done in 4 PCR reactions and
Spo11 (figure 2c) in 2 PCR reactions.
19
and stored gel slices containing amplified DNA of ~2,000bp at 4oC for future use.
Cloning of PCR Products
I cloned purified PCR products using the Strataclone cloning protocol that I modified by first
heating low melt gel slices at 65○C for 10 minutes, until completely melted. Next, I added 0.25µl
of vector, 0.75µl of cloning buffer, and 2.5µl of the melted agarose to each cloning reaction (for
details on cloning vector see StrataClone PCR Cloning Kit developed by Agilent Technologies).
I placed cloning reactions on the benchtop for ~15 minutes to allow integration of PCR into the
vector, creating a recombinant plasmid. I then added ~12µl of competent E. coli to each reaction,
which were then immediately placed onto ice for 10 minutes and then heat-shocked at 42○C for
45 seconds to transform plasmids into E. coli. Reactions were then immediately placed back onto
ice for two minutes to allow the porous membranes of competent E. coli cells to seal. I then
placed 125µl of LB medium into each reaction, which were subsequently incubated at 37○C for
two hours to allow for E. coli growth; culture plates containing LB medium and Kanamycin were
also incubated. After two hours of incubation I plated bacteria onto culture plates containing LB
and Kanamycin, and allowed E. coli to grow overnight.
Screening of Clones
The plasmids I used for cloning contained: 1) a Kanamycin-resistance gene, and 2) a DNA
insertion site within a LacZ reporter gene. The incorporation of PCR product into plasmids
disrupts the LacZ gene, producing white bacterial colonies that are Kanamycin-resistant. I
screened white colonies of E. coli that had grown on culture plates containing Kanamycin using
PCR with vector-specific primers to verify that PCR amplified DNA from low melt gels had
incorporated into plasmids (Tables 7 & 8). Using gel electrophoresis I determined if PCR screens
had amplified DNA with the length of low melt PCR product plus 200 vector base pairs,
indicating successful DNA incorporation into the plasmid. Colonies that I screened were
simultaneously grown on new culture plates containing LB and Kanamycin so I could extract
20
Table 7: Stock and Working Concentration of PCR Screening Reagents.
Reagent
Stock Concentration
Working Concentration
T3 vector primer
50µM
1.25 µM
T7 vector primer
50µM
1.25 µM
10 mM each
0.25 mM each
5x
1x
ProMEGA MgCl2
25mM
1.25 mM
Neb Taq
5 U/µL
0.0125 U/µL
dNTPs
ProMEGA 5x Flexi Buffer
dH2O
0.3736 mM
21
Table 8: Thermocycler Program used in PCR Screening Reactions.
Step
Temp. (○C )
Time (minutes)
1
95.0
2:00
2
94.0
1:00
3
53.5
1:30
4
72.0
3:30 (or 1 minute per kb)
5
Go to Step 2, Repeat 39x
6
72.0
10:00
7
4.0
Hold indefinitely
22
confirmed recombinant plasmids from white colonies. For each P. antipodarum lineage, I
selected ~three bacterial colonies for each PCR reaction to provide higher confidence in
sequencing results via independent replications of sequence in multiple clones.
Plasmid Extraction and Preparation
I transferred clones that tested positive for recombinant plasmids in PCR screens from culture
plates into test tubes containing liquid LB medium and Kanamycin that were then incubated at
37○C overnight. Following incubation, I used the Zyppy Plasmid Miniprep Kit and protocol (see
Zyppy Plasmid Miniprep Kits by Zymo Research) to extract plasmids from transformed
competent cells. Next, I used spectrophotometry to determine the DNA concentration of each
plasmid extraction and then submitted plasmids for Sanger sequencing.
Processing and Analysis of Sanger Sequences
I used Sequencher (version5.2) to process Sanger reads by trimming vector sequence and
manually call ambiguous bases. I then used Sequencher to assemble and create separate
consensus sequences from Sanger reads of six asexual and three sexual lineages for Msh4, six
asexual and three sexual lineages for Msh5, and 13 asexual and five sexual lineages for Spo11. I
annotated Sanger consensus sequences by aligning them to meiosis gene models that had been
annotated with transcriptome reads and made from genome contigs. I used these annotated
Sanger consensus sequences to identify nonsense or frameshift mutations by visually inspecting
gene assemblies.
Next, I calculated the frequency of nonsynonymous polymorphisms per nonsynonymous site (πa)
relative to the frequency of synonymous polymorphisms per synonymous site (πs) for Msh4,
Msh5 and Spo11 in sexual and asexual lineages. I began by concatenating the exon sequences
within each lineage to generate a lineage consensus for each gene and aligned these consensus
sequences with the Muscle algorithm in MEGA (version six) (Tamura et al. 2013). I then used
this alignment and DNAsp (version five) (Librado & Rozas 2009) to calculate πa/ πs with a 150bp
sliding window with three bp steps for the genes Msh4 and Msh5. Window sizes that were
23
smaller than this did not capture either πa or πs resulting in ratios of zero or undefined ratios,
respectively. Window sizes that were larger than this did not provide enough resolution to assess
whether selection was relaxed across the entire gene, or at particular sites. In other words, I
chose a window size of 150bp with three bp steps because this range gave the greatest number of
comparisons between πa and πs, providing a clearer picture as to selective pressures on meiosisspecific genes. Additionally while I obtained Sanger sequence data from six asexual lineages for
both Msh4 and Msh5, one asexual lineage was missing a substantial piece of coding sequence for
Msh4 and a different asexual lineage was missing coding sequence for Msh5 because of
technical difficulties in PCR amplification across the entire gene. I omitted the two asexual
lineages missing Sanger sequence data from this analysis so that the same lineages were being
compared in Msh4 and Msh5, avoiding any potential elevated πa/πs between these two genes
caused by confounding effects of phylogenetic dependence. For Spo11, I used a window size of
30bp and 3bp steps because I only had 318bp of coding Sanger sequence data, making a window
size of 150bp too large to obtain any resolution across available coding sequence.
I then used MEGA (Tamura et al. 2013) to quantify and compare the rates and patterns of
meiotic gene evolution between sexual and asexual lineages. I used IUPAC ambiguity codes to
represent heterozygous sites within consensus sequences of each lineage (Table 9). I then used
maximum likelihood analysis and a HYK+G model as implemented in MEGA to generate
phylogenetic trees of Msh4, Msh5 and Spo11 from consensus sequences that included coding
and noncoding sequence. I chose the HYK+G model because it gave the highest bootstrap
confidence values when creating phylogenetic trees. I omitted the asexual lineages missing
Sanger sequence data for Msh4 and Msh5 from the creation of their respective phylogenetic
trees. I did not remove them both from Maximum Likelihood analysis of Msh4 and Msh5 in
order to maximize the number of sexual vs. asexual comparisons. This does make sexual vs.
asexual rates of evolution between Msh4 and Msh5 less comparable because the lineages in these
analyses are not exactly the same, opening the possibility that differentially elevated asexual
branch lengths between Msh4 and Msh5 are caused by confounding effects of phylogenetic
dependence. However, if selection is indeed relaxed on all meiosis-specific genes in asexuals
then all branch lengths should be longer relative to each sexual lineage, regardless of the
phylogenetic distribution of lineages within this species.
24
Table 9: IUPAC Degenerate Codes at Heterozygous Sites for Maximum Likelihood
Analysis.
Only A and G are used here as examples. However the same basic rules applied for all IUPAC
degenerate symbols for any given combination of bases at a site.
1
Base1
(Frequency)
A(>25%)
Base2
(Frequency)
G(>25%)
Degenerative
Symbol
R
2
A(75%)
G(25%)
a
3
A(>75%)
a
4
A(>75%)
G(<25%)
*Independently replicated
in another clone
G(<25%)
*Only present in 1 clone
Site
25
A
Chapter 3: Results
Accumulated Mutations of Sexual and Asexual P. antipodarum
I did not detect any frameshift or nonsense mutations in any of the genes that I studied (Table
10). My sliding window comparisons of nonsynonymous relative to synonymous polymorphism
(πa/πs) revealed that most sites had a ratio well below one in all lineages, indicating a low rate of
retention of nonsynonymous relative synonymous polymorphism. When I compared the
cumulative πa/πs of each site across the coding region of meiosis-specific genes asexuals were
higher compared to sexuals, however neither estimate of πa/πs met neutral expectations (Figures
3-5). This result could indicate the presence of less efficient purifying selection in asexuals,
which has previously been reported in this system (Neiman et al. 2010) as well as other
freshwater asexual snails (Johnson & Howard 2007, Crummett et al. 2013). I did find that πa/πs
was considerably higher in Msh4 relative to the other genes (Figure 3). Altogether, the outcomes
of these analyses are consistent with purifying selection of meiosis-specific genes in asexual
lineages.
Evolutionary Rates of Sexual vs. Asexual Meiosis Genes
A phylogenetic comparison of the rate of evolution of each of the meiosis-specific genes in
sexual vs. asexual P. antipodarum revealed that the genes had similar branch lengths in sexuals
and asexuals, (Figures 6-8) suggesting that reproductive mode is not affecting the rate at which
these genes evolve. The only potential exception to this pattern came from Msh5. While there
were relatively long branch lengths for this gene in some asexual vs. sexual lineages (Figure 7),
additional comparisons between coding regions of sexual and asexual lineages indicates that the
rates of evolution of Msh5 are not different within exons in sexuals vs. asexuals (Figure 9). This
implies that polymorphisms driving the elevated Msh5 branch lengths in some asexuals are
predominantly in introns that are selectively neutral, as opposed to asexual polymorphisms in
Msh5 exons that show similar evolutionary rates to sexuals. Since there is an a priori prediction
that Msh5 exons in sexuals are under purifying selection my result indicates purifying selection
is also acting on the exons of Msh5 in asexuals.
26
Table 10: Open Reading Frames in Sexual and Asexual P. antipodarum.
This table shows all genes that have been bioinformatically assembled from genome assemblies,
representing work from myself, Cindy Toll, Matthew Wheat, and Keagan Kavanaugh. Genes that
have more than two sexual and more than 3 asexual lineages have assemblies from genome
contigs and Sanger sequence data. Genes in black background are meiosis-specific; genes in
white background have roles in meiosis and mitosis. Numbers show the number of lineages that
have open reading frames, with no evidence of either frameshift or nonsense mutations.
27
Figure 3: Msh4 Polymorphism.
Comparison of nonsynonymous (πa) polymorphism relative to synonymous (πs) polymorphism in
three sexual and four asexual lineages. Two asexual lineages are omitted from this analysis, due
to poor coverage for the gene Msh4 and Msh5 respectively. I omitted asexuals in order to make
this analysis comparable between the two genes Msh4 and Msh5. For this analysis I used a
window of 150bp and 3bp steps, the location of these windows relative to the exons of the gene
is indicated below the x-axis. The inset graph shows the total ratio of πa to πs across the entire
coding region in sexuals vs. asexuals. The difference between total πa/πs of sexual vs. asexual is
not significant (NS) (Fisher’s Exact Test, p >0.05).
28
Figure 4: Msh5 Polymorphism.
Comparison of πa relative to πs in sexual and asexual lineages. The same four asexual and three
sexual lineages from analysis of Msh4 were compared using a window of 150bp and 3bp steps,
the location of these windows relative to the exons of the gene is indicated below the x-axis. The
difference between total πa/πs of sexual vs. asexual is not significant (NS) (Fisher’s Exact Test, p
>0.05).
29
Figure 5: Spo11 Polymorphism.
Comparison of πa relative to πs in both sexual and asexual lineages. For this analysis 5 sexual and
13 asexual lineages were used to compare polymorphism across four exons. Reproductive modes
were compared using a window of 30bp and 3bp steps, the location of these windows relative to
the exons of the gene is indicated below the x-axis. The difference between total πa/πs of sexual
vs. asexual is not significant (NS) (Fisher’s Exact Test, p >0.05).
30
Figure 6: Maximum Likelihood Analysis of Msh4.
Branch length of sexual (white) and asexual (black) lineages was determined using an HKY+G
model with 100 bootstrap replicates. The numbers correspond to lineage/ploidy identities as
follows: 1: Lady (2x), 2: Grasmere (2x), 3: Ianthe Field Collected (2x), 4: Brunner (3x), 5:
Okoreka (3x), 6: Kaniere (3x) omitted due to stretches of zero Sanger sequence coverage, 7:
Grasmere (3x), 8: Poerua (4x), 9: Gunn (3x).
31
Figure 7: Maximum Likelihood Analysis of Msh5.
Branch length of sexual (white) and asexual (black) lineages was determined using an HKY+G
model with 100 bootstrap replicates. The numbers correspond to lineage/ploidy identities as
follows: 1: Lady (2x), 2: Grasmere (2x), 3: Ianthe Field Collected (2x), 4: Brunner (3x), 5:
Okoreka (3x), 6: Kaniere (3x), 7: Grasmere (3x), 8: Poerua (4x), 9: Gunn (3x) omitted due to
stretches of zero Sanger sequence coverage.
32
Figure 8: Maximum Likelihood Analysis of Spo11.
Branch length of sexual (white) and asexual (black) lineages was determined using an HKY+G
model with 100 bootstrap replicates. The numbers correspond to lineage/ploidy identities as
follows: 1: Lady (2x), 2: Grasmere (2x), 3: Ianthe (2x), 4: Brunner (3x), 5: Okareka (3x), 6:
Kaniere (3x), 7: Grasmere (3x), 8: Poerua (4x), 9a/b: Gunn (3x), 10: Taylor (3x), 11: Te Anau
(3x), 12: Brunner (4x), 13: Tarawera (3x), 14: Rotoroa (2x), 15: Ianthe Lab Lineage (2x) .
33
Figure 9: Evolutionary Rate of Sexual and Asexual Msh5 Coding Region.
Branch length of sexual (white) and asexual (black) lineages was determined using an HKY+G
model with 100 bootstrap replicates. The numbers correspond to lineage/ploidy identities as
follows: 1: Lady (2x), 2: Grasmere (2x), 3: Ianthe Field Collected (2x), 4: Brunner (3x), 5:
Okoreka (3x), 6: Kaniere (3x), 7: Grasmere (3x), 8: Poerua (4x), 9: Gunn (3x) omitted due to
stretches of zero Sanger sequence coverage.
34
Chapter 4: Discussion
The absence of observed frameshift or nonsense mutations in meiosis genes of asexual P.
antipodarum suggests that these mutations are deleterious and selectively removed from
populations before they can be detected. Additionally, the low frequency of πa relative to πs in
sexuals and asexuals indicates that selection is removing nonsynonmous polymorphisms. Finally,
maximum likelihood-based comparisons of evolutionary rates between sexual and asexual
lineages revealed that these meiosis-specific genes evolve at similar rates in sexual and asexual
P. antipodarum. Because meiosis genes in sexual organisms are expected to experience purifying
selection, the similar rates of evolution in sexual and asexual P. antipodarum suggests that
purifying selection is also acting on asexuals. My results can be interpreted in several ways: 1)
Asexual P. antipodarum are engaging in cryptic sex that has yet to be observed, 2) Asexuals P.
antipodarum are automictic, and 3) Asexual lineages of P. antipodarum are so recently derived
that relaxed selection has not yet translated into a discernable increase in the rate of mutation
accumulation in meiosis-specific genes relative to sexual counterparts. The remainder of my
discussion will aim to address these possibilities in light of what is known about P. antipodarum,
meiosis in automictic asexuals, and evolution of genes under relaxed selection.
Several studies have used molecular tools to detect evidence of cryptic sex in organisms
previously thought to be obligately asexual (Villate et al. 2010). Similar studies of microsatellite
(Weetman et al. 2002) and minisatellite (Hauser et al. 1992) inheritance in P. antipodarum,
however, have found no evidence for recombination or gene flow in asexual P. antipodarum.
Additional lack of evidence for recombination in this system comes from allozyme genotyping
of natural P. antipodarum populations that are completely or almost completely female with low
male frequencies <10%, to populations that have higher male frequencies of 20-50%
(Winterbourn 1970, Lively 1987, Wallace 1992, Lively & Jokela 2002). These low male
populations have been found to almost exclusively reproduce by apomictic parthenogenesis,
while the higher male populations usually reproduce sexually (Phillips and Lambert 1989). The
absence of evidence for recombination and gene flow in asexual P. antipodarum suggests that it
is unlikely that selection is maintaining meiosis-specific genes that enable asexual P.
antipodarum to at least occasionally engage in canonical sex. It is also possible that the apparent
maintenance of meiosis-specific genes in asexual P. antipodarum reflects automixis, whereby
35
gametogenesis in asexual organisms incorporates components of the meiotic process preceded by
premeiotic endoduplication coupled with endomitosis or followed by gamete fusion, to restore
ploidy (Marescalchi et al. 2003, Lampert et al. 2007, Mogie 2013). The possibility of automixis
is again confounded by the lack of evidence for recombination in asexual P. antipodarum.
Asexual P. antipodarum do on rare occasions produce males (Neiman et al. 2012); these males
produce morphologically normal but often aneuploid sperm via an apparently modified meiotic
process (Soper 2013). The fact that these males do seem to use some elements of meiosis
suggests that there may be retention of meiotic processes in their asexual female mothers. The
production of rare asexual males also raises the possibility of contagious asexuality (Sandrock et
al. 2011, Jaquiery et al. 2014).
Infectious asexuality has been heavily studied in parasitoid wasps (Sandrock & Vorburer 2011),
aphids (Jaquiery 2014), daphnia (Paland et al. 2005, Sandrock et al. 2011), and rotifers (Tucker
2013). Contagious asexuality involves a recessive meiosis-suppressing asexual allele that
segregates in the sexual population, until a homozygous female is produced with the asexual
phenotype, starting the first asexual lineage. An asexual lineage will continue to reproduce and
on rare occasions bear males. These asexually-produced males will be able to fertilize sexual
females and as such, transmit the asexuality-inducing allele to the offspring, initiating a new
asexual lineage. Contagious asexuality is supported in P. antipodarum because the variable
ploidy across asexual P. antipodarum sperm suggests some components of meiosis are
suppressed, possibly via a meiosis-suppressing element that induces the asexual phenotype.
Additionally, asexually-produced male P. antipodarum make sperm that appear morphologically
similar to sexual males and will copulate with sexual females (Soper et al. 2015 in press), which
supports rare asexually-produced males transmitting their meiosis-suppressing allele to a sexual
females egg. The elevated ploidy in asexual P. antipodarum supports the possibility of aneuploid
sperm fertilizing a sexual euploid egg, and the multiple independent transitions to asexuality in
P. antipodarum are consistent with rare asexually-produced males repeatedly fertilizing sexual
females to found new asexual lineages.
The final possibility for why meiosis-specific genes in asexual P. antipodarum do not appear to
be degrading is that selection is relaxed in asexual P. antipodarum that are too young relative to
other ancient asexual taxa who have experienced asexuality long enough to have accumulated
36
more nonsynonymous mutations in their meiosis-specific genes than their sexual counterparts
(Pellino et al.2013). Other studies that have evaluated meiosis gene evolution in asexual plants
(Pellino et al. 2013), degradation of eyes in cave fish (Yokoyama et al. 1995), and degradation of
teeth in birds (Meredith et al. 2014) have found elevated rates of nonsynonymous relative to
synonymous substitutions, as well as frameshift and nonsense mutations, in genes that are
exclusively associated with traits whose ancestral function are no longer relevant to that
organisms fitness. Selection on these ancestral traits has been relaxed in these systems for
millions of years. By contrast, the oldest asexual P. antipodarum lineage is only around 500,000
years old, suggesting that asexual P. antipodarum may simply be too recently derived from
sexual ancestors for the molecular signatures of relaxed selection to become apparent.
37
Chapter 5: Summary
Comparing evolutionary patterns between sexual and asexual P. antipodarum provides a
powerful means of evaluating the evolutionary consequences of asexuality. By sequencing and
performing comparative analysis of meiosis-specific genes between sexual and asexual lineages
of P. antipodarum, I have provided evidence that meiosis-specific genes in asexual P.
antipodarum are not in the process of degrading. Potential explanations for this surprising result
include the relatively recent derivation of most asexual P. antipodarum lineages or because
asexuals are in some way utilizing meiosis genes. For example, perhaps some components of
meiosis that are not necessarily associated with recombination are being used to create sperm in
asexually-produced male P. antipodarum that contagiously spread asexuality. Regardless, this
study has provided exciting evidence that meiosis gene evolution in asexual taxa is not an
inevitable process of gene degradation under relaxed selection. Elucidating why meiosis-specific
genes are not being lost in asexuals will be the next step in determining consequences for the
evolution of meiosis genes in asexual reproduction.
38
References
Barton, N.H., Charlesworth, B. (1998) Why sex and recombination? Science 281: 1986-1990.
Beck, J., Windham, M., Pryer, K. (2011) Do asexual polyploidy lineages lead to short
evolutionary lives? A case study from the fern genus Astrolepis. Evolution 65: 3217-3229.
Bell, G. (1982) The masterpiece of nature. The evolution of genetics and sexuality. University of
California press, Berkeley, California.
Birney, E. et al. (2007) Identification and analysis of functional elements in 1% of the human
genome by the ENCODE pilot project. Nature 447: 799-816.
Burt, A. (2000) Perspective: sex, recombination, and the efficacy of selection-was Weismann
right? Evolution 54: 337-351.
Carr, M., Leadbeater, B., Baldauf, S. (2010) Conserved meiotic genes point to sex in the
choanoflagellates. Journal of Eukaryotic Microbiology 57: 56-62.
Charlesworth, D., Morgan, M., Charlesworth, B. (1993) Mutation accumulation in finite
populations. Journal of Heredity 84: 321-325.
Charlesworth, D., Morgan, M., Charlesworth, B. (1993) Mutation accumulation in finite
outbreeding and inbreeding populations. Genetical Research 61: 39-56.
Chen, S., Ma, K., Zeng, J. (2011) Pseudogene: lessons from PCR bias, identification and
resurrection. Molecular Biology Reports 38: 3709-3715.
Connalon, T., Knowles, L. (2007) Recombination rate and protein evolution in yeast. Bmc
Evolutionary Biology 7.
Crummett, L., Sears, B., Lafon, D., Wayne M. (2013) Parthenogenetic populations of the
freshwater snail Campeloma limum occupy habitats with fewer environmental stressors than
their sexual counterparts. Freshwater Bioogy 58: 655-663.
D’Souza, T., Storhas, M., Schulenburg, L., Beukeboom, L., Michiels, N. (2010) The costs and
benefits of occasional sex: theoretical predictions and a case study. Journal of Heredity 1010:
S34-S41.
Dacks, J., Roger, A. (1999) The first sexual lineage and the relevance of facultative sex. Journal
of Molecular Evolution 48: 779-783.
De Visser, J. (2007) Bacterial solutions to the problem of sex. Plos Biology 5: 1844-1846.
Desai, M., Fisher, D., Murray, A. (2007) The speed of evolution and maintenance of variation in
asexual populations. Current Biology 17: 385-394.
39
Dybdahl, M., Lively, C. (1995). Diverse, endemic and polyphyletic clones in mixed populations
of a fresh-water snail (Potamopyrgus antipodarum). Journal of Evolutionary Biology 8: 385-398.
Gerstein, M., Zheng, D. (2006) The real life of pseudogenes. Scientific American 295: 48-55.
Gojobori, T., Li, W., Gruar, D. (1982) Patterns of nucleotide substitution in pseudogenes and
functional genes. Journal of Molecular Evolution 18: 360-369.
Guo, X., Zheng, D. (2011) Regulatory roles of novel small RNAs from pseudogenes. Noncoding
RNAs in Plants (RNA Technologies): 193-208.
Hadany, L., Comeron, J., Schlichting, C., Mousseau, T. (2008) Why are sex and recombination
so common? Year in Evolutionary Biology 1133: 26-43.
Hanson, S., Schurko, A., Hecox-Lea, B., Welch, D., Stelzer, CP., Logsdon, J. (2013) Inventory
and phylogenetic analysis of meiotic genes in monogonont rotifers. Journal of Heredity 104:
357-370.
Hauser, L., Carvalho, G., Hughes, R., Carter, R. (1992) Clonal structure of the introduced freshwater snail Potamopyrgus antipodarum (Prosobranchia, Hydrobiidae), as revealed by DNA
fingerprinting. Proceeding of the Royal Society B-Biological Sciences 249: 19-25.
Hecht, N. (1998) Molecular mechanisms of male germ cell differentiation. Bioessays 20: 555561.
Henry, L., Schwander, T., Crespi, B. (2012) Deleterious mutation accumulation in asexual
Timema insects. Molecular Biology and Evolution 29: 401-408.
Hill, W., Robertson, A. (2007) The effect of linkage on limits to artificial selection (reprinted).
Genetics Research 89: 311-336.
Hogsdon, E., Otto, S. (2012) The red queen coupled with directional selection favours the
evolution of sex. Journal of Evolutionary Biology 25: 797-802.
Hojsgaard, D., Martinez, E., Quarin, C. (2013) Competition between meiotic and apomictic
pathways during ovule and seed development results in clonality. New Phytologist 197: 336-347.
Jaquiery, J., et al. (2014) Genetic control of contagious asexuality in the pea aphid. PLoS
Genetics 10: e1004838.
Johnson, S., Howard, R. (2007) Contrasting patterns of synonymous and nonsynonymous
sequence evolution in asexual and sexual freshwater snail lineages. Evolution 61: 2728-2735.
Jokela, J., Dybdahl, M., Lively, C. (2009) The maintenance of sex, clonal dynamics, and hostparasite coevolution in a mixed population of sexual and asexual snails. American Naturalist
174:S43-S53.
40
Kerstes, M., Berenos, C., Schmid-Hempel, P., Wegner, K. (2012) Antagonistic experimental
coevolution with a parasite increases host recombination frequency. Bmc Evolutionary Biology
12.
Kondrashov, A. (1993) Classification of hypotheses on the advantage of amphimixis. Journal of
Heredity 84: 372-387.
Lahti, D., et al. (2009) Relaxed selection in the wild. Trends in Evology and Evolution 24: 487496.
Lampert, K., et al. (2007) Automictic reproduction in interspecific hybrids of Poeciliid fish.
Current Biology 17: 1948-1953.
Li, W., Gojobori, T., Nei, M. (1981) Pseudogenes as a paradigm of neutral evolution. Nature
292: 237-239.
Librado, P., Rozas, J. (2009) DnaSP v5: A software for comprehensive analysis of DNA
polymorphism data. Bioinformatics 25: 1451-1452.
Liu, Q., Thomas, V., Williamson, V. (2007) Meiotic parthenogenesis in a root-knot nematode
results in rapid genomic homozygosity. Genetics 176: 1483-1490.
Lively, C. (1987) Evidence from a New Zealand snail for the maintenance of sex by parasitism.
Nature 328: 519-521.
Lively, C. (1992) Parthenogenesis in a freshwater snail-reproductive assurance versus parasitic
release. Evoltuion 46: 907-913.
Lively, C., Jokela, J. (2002) Temporal and spatial distribution of parasites and sex in a freshwater
snail. Evolutionary Ecology Research 4: 219-226.
Lunt, D. (2008) Genetic tests of ancient asexuality in root knot nematodes reveal recent hybrid
origins. Bmc Evolutionary Biology 8: 194.
Lydall, D., Nikolsky, T., Bishop, D., Weinert, T. (1996) A meiotic recombination checkpoint
controlled by mitotic checkpoint genes. Nature 383: 840-843.
Lynch, M., Burger, R., Butcher, D., Gabriel, W. (1993) The mutational meltdown in asexual
populations. Journal of Heredity 84: 339-344.
Lynch, M., Conery, J. (2000) The evolutionary fate and consequences of duplicate genes.
Science 290: 1151-1155.
Malik, S., Pightling, A., Stefaniak, L., Schurko, A., Logsdon, J. (2008) An expanded inventory
of conserved meiotic genes provides evidence for sex in Trichomonas vaginalis. PLoS One 3:
e2879.
41
Marescalchi, O., Pijnacker, L., Scali, V. (1993) Autoictic parthenogenesis and its genetic
consequence in Bacillus atticus atticus (Insecta Phasmatodea). Invertebrate Reproduction and
Development 24: 7-12.
Marescalchi, O., Scali, V. (2003) Automictic parthenogenesis in the diploid-triploid stick insect
Bacillus atticusand its flexibility leading to heterospecific diploid hybrids. Invertebrate
Reproduction and Development 43: 163-172.
Matthews, L., Carter, P., Thierry-Mieg, D., Kemphues, K. (1998) ZYG-9, a Caenorhabditis
elegans protein required for microtubule organization and function, is a component of meiotic
and mitotic spindle poles. Journal of Cell Biology 141: 1159-1168.
Meredith, R., Zhang, G., Gilbert, M., Jarvis, E., Springer, M. (2014) Evidence for a single loss of
mineralized teeth in the common avian ancestor. Science 346: 1336.
Mogie, M. (1988) A model for the evolution and control of generative apomixis. Biological
Journal of the Linnean Society 35: 127-153.
Mogie, M. (2013) Premeiotic endomitosis and the costs and benefits of asexual reproduction.
Biological Journal of the Linnean Society 109: 487-495.
Mostowy, R., Engelstadter, J. (2012) Host-parasite coevolution induces selection for conditiondependent sex. Journal of Evolutionary Biology 25: 2033-2046.
Muller, H. (1950) Our load of mutations. American Journal of Human Genetics 2: 111-176.
Mungpakdee, S., et al. (2008) Differential evolution of the 13 Atlantic salmon hox clusters.
Molecular Biology and Evolution 25: 1333-1343.
Naumova, T., et al. (2001) Reproductive development in apomictic populations of Arabis
holboellii (Brassicaceae). Sexual Plant Reproduction 14: 195-200.
Neiman, M., Hehman, G., Miller, J., Jogsdon, J., Taylor, D. (2010) Accelerated mutation
accumulation in asexual lineages of a freshwater snail. Molecular Biology and Evolution 27:
954-963.
Neiman, M., Jokela, J., Lively, C. (2005) Variation in asexual lineage age in Potamopyrgus
antipodarum, a New Zealand snail. Evolution 59: 1945-1952.
Neiman, M., Larkin, K., Thopson, A., Wilton, P. (2012) Male offspring production by asexual
Potamopyrgus antipodarum, a New Zealand snail. Heredity 109: 57-62.
Neiman, M., Lively, C. (2004) Pleistocene glaciation is implicated in the phylogeographical
structure of Potamopyrgus antipodarum, a New Zealand snail. Molecular Ecoloty 13: 30853098.
42
Neiman, M., Paczesniak, D., Soper, D., Baldwin, A., Hehman, G. (2011) Wide variation in
ploidy level and genome size in a New Zealand freshwater snail with coexisting sexual and
asexual lineages. Evolution 65: 3202-3216.
Nomark, B., Judson, O., Moran, N. (2003) Genomic signatures of ancient asexual lineages.
Biological Journal of the Linnean Society 79: 69-84.
Paland, S., Colbourne, J., Lynch, M. (2005) Evolutionary history of contagious asexuality in
Daphnia pulex. Evolution 59: 800-813.
Pellino, M., et al. (2013) Asexual genome evolution in the apomictic Ranunculus auricomus
complex: examining the effects of hybridization and mutation accumulation. Molecular Ecology
22: 5908-5921.
Peters, N., et al. (2013) Control of mitotic and metioic centriole duplication by the Plk4-related
kinase ZYG-1. Journal of Cell Science 123: 795-805.
Phillips, N., Lambert, D. (1989) Genetics of Potamopyrgus antipodarum (Gastropoda,
Prosobranchia) – evidence for reproductive modes. New Zealand Journal of Zoology 16: 435445.
Ramesh, M., Malik, S., Logsdon, J. (2005) A phylogenomic inventory of meiotic genes:
evidence for sex in giardia and an early eukaryotic origin of meiosis. Current Biology 15: 185191.
Ronkainen, H., Ylonen, H. (1994) Behavior of cyclic bank voles under risk of mustelid
predation-do females avoid copulations. Oecologia 97: 377-381.
Rosendahl, S. (2012) The first glance into the Glomus genome: an ancient asexual scandal with
meiosis? New Phytologist 193: 546-548.
Sagata, N. (1996) Meiotic metaphase arrest in animal oocytes: its mechanisms and biological
significance. Trends in Cell Biology 6: 22-28.
Sandrock, C., Schirrmeister, B., Vorburger, C. (2011) Evolution of reproductive mode variation
and host associations in a sexual-asexual complex of aphid parasitoids. Bmc Evolutionary
Biology 11: 348.
Sandrock, C., Vorburger, C. (2011) Single-locus recessive inheritance of asexual reproduction in
a parasitoid wasp. Current Biology 21: 433-437.
Schurko, A., Logsdon, J. (2008) Using a meiosis detection toolkit to investigate ancient asexual
“scandals” and the evolution of sex. Bioessays 30: 579-589.
Schurko, A., Logsdon, J., Eads, B. (2009) Meiosis genes in Daphnia pulex and the role of
parthenogenesis in genome evolution. Bmc Evolutionary Biology 9: 78.
43
Schurko, A., Mazur, D., Logsdon, J. (2010) Inventory and phylogenomic distribution of meiotic
genes in Nasnoia vitripennis and among diverse arthropods. Insect Molecular Biology 19: 165180.
Schwander, T., Crespi, B. (2009) Twigs on the tree of life? Neutral and selective models for
integrating macroevolutionary patterns with microevolutionary processes in the analysis of
asexuality. Molecular Ecology 18: 28-42.
Sekine, K., Tojo, K. (2010) Automictic parthenogenesis of a geographically parthenogenetic
mayfly, Ephoron shigae (Insecta: Ephemeroptera, Polymitarcyidae). Biological Journal of the
Linnean Society 99: 335-343.
Singaravelu, G., Singson, A. (2011) New insights into the mechanism of fertilization in
nematodes. International Review of Cell and Molecular Biology 289: 211-238.
Smith, M. (1978) The evolution of sex. Cambridge University Press, Cambridge, UK.
Som, C., Reyer, H. (2007) Hemiclonal reproduction slows down the speed of Muller’s ratchet in
the hybridogenetic frog Rana esculenta. Journal of Evolutionary Biology 20: 650-660.
Soper, D., Neiman, M., Savytskyy, O., Zolan, M., Lively, C. (2013) Spermatozoa production by
triploid males in the New Zealand freshwater snail Potamopyrgus antipodarum. Biological
Journal of the Linnean Society 110: 227-234.
Srinivasan, D., Fenton, B., Jaubert-Possamai, S., Jaouannet, M. (2010) Analysis of meiosis and
cell cycle genes of the facultatively asexual pea aphid, Acyrthosiphon pisum (Hemiptera:
Aphididae). Insect Molecular Biology 19: 229-239.
Suomalainen, E., Saura, A., Lokki, J. (1976) Evolution of parthenogenetic insects. Evolutionary
Biology 9: 209-257.
Sur, S., Saha, S., Tisa, L., Borhra, A., Sen A. (2013) Characterization of pseudogenes in
members of the order Frankineae. Journal of Biosciences 38: 727-732.
Tam, O., et al. (2008) Pseudogene-derived small interfering RNAs regulate gene expression in
mouse oocytes. Nature 453: 534-538.
Tamura, K., Stecher, G., Peterson, D., Filipski, A., Kumar, S. (2013) MEGA6: molecular
evolutionary genetics analysis version 6.0. Molecular Biology and Evolution 30: 2725-2729.
Tucker, A., Ackerman, M., Eads, B., Xu, S., Lynch, M. (2013) Population-genomic insights into
the evolutionary origin and fate of obligately asexual Daphnia pulex. Proceedings of the National
Academy of Sciences of the United States of America 110: 15740-15745.
Vileneuve, A., Hillers, K. (2001) Whence meiosis? Cell 106: 647-650.
44
Villate, L., Esmenjaud, D., Van Helden, M., Stoeckel, S., Plantard, O. (2010) Genetic signature
of amphimixis allows for the detection and fine sale localization of sexual reproduction events in
a mainly parthenogenetic nematode. Molecular Ecology 19: 856-873.
Wallace, C. (1985). On the distribution of the sexes of Potamoprgus jenkinsi (Smith). Journal of
Molluscan Studies 51: 290-296.
Wallace, C. (1992). Parthenogenesis, sex and chromosomes in Potamopyrgus. Journal of
Molluscan Studies 58: 93-107.
Wang, C., Tseng, C. (2014) Recent advances in understanding of meiosis initiation and the
apomictic pathway in plants. Frontiers in Plant Science 5: 497.
Weetman, D., Hauser, L., Carvalho, G. (2002) Reconstruction of microsatellite mutation history
reveals a stron and consistent deletion bias in invasive clonal snails, Potamopyrgus antipodarum.
Genetics 162: 813-822.
White, M. (1978) Modes of speciation. W.H. Freem and Company, San Francisco, California.
Williams, G. (1975) Sex and evolution. Princeton University Press, Princeton, New Jersey.
Winterbourn, M. (1970) The New Zealand species of Potamopyrgus (Gartropoda: Hybrodiidae).
Malacologia 10: 283-321.
Yampolsky, L., Bouzinier, M. (2014) Faster evolving Drosophila paralogs lose expression rate
and ubiquity and accumulate more non-synonymous SNPs. Biology Direct 9: 2.
Yokoyama, S., Meany, A., Wilkens, H., Yokoyama, R. (1995) Initial mutational steps toward
loss of opsin gene-function in cavefish. Molecular Biology and Evolution 12: 527-532.
Zhang, J. (2003) Evolution by gene duplication: an update. Trends in Ecology and Evolution 18:
292-298.
Zhang, Z., Harrison, P., Liu, Y., Gerstein, M. (2003) Millions of years of evolution preserved: a
comprehensive catalog of the processed pseudogenes in the human genome. Genome Research
13: 2541-2558.
Zickler, D., Kleckner, N. (1999) Meiotic chromosomes: integrating structure and function.
Annual Review of Genetics 33: 603-754.
45