The influence of aging on oocyte quality and the importance of mitochondria

by Krystal Trull

B.S. in Physiology and Neurobiology, University of Connecticut

A thesis submitted to

The Faculty of the College of Science of Northeastern University

in partial fulfillment of the requirements for the degree of Master of Science

July 24, 2014

Thesis directed by

Jonathan Tilly Professor and Department Chair of Biology

Abstract of Thesis

Female fertility peaks at 25 years of age, yet many women put off having children until

much later in life (Nybo Andersen et al. 2000; Hook 1981; Hamilton et al. 2013). Age is a major

determining factor in pregnancy outcomes and increased maternal age is correlated with

increased chances of aneuploidy and miscarriage (Robinson et al. 2001; Benadiva et al. 1996;

Battaglia et al. 1996). When older women encounter difficulties with having a successful

pregnancy, it is often due to poor oocyte quality as a result of mitochondrial dysfunction

(Eichenlaub-Ritter et al. 2011; Bentov et al. 2010; Jansen & Burton 2004). Mitochondria are the

energy-producing organelles of the cell and are important in producing adenosine triphosphate

(ATP) to meet the high energy demands of the oocyte. Mitochondria have their own DNA

(mtDNA) inherited maternally that encodes proteins necessary for oxidative phosphorylation and

subsequent ATP production. Along with ATP, mitochondria also produce harmful reactive

oxygen species (ROS) at a low, but significant rate (Takeo et al. 2013; Parsons et al. 1997).

With natural aging, mtDNA damages due to ROS can accumulate and disrupt normal cell

function and ATP production. Without adequate ATP supply, oocytes fail to fertilize, implant,

or develop properly during pregnancy. This eventual decline of oocyte quality occurs with

natural aging, but decreased fertility is also observed in diabetes and other metabolic disorders

(Moley et al. 1991; Sadler et al. 1988). As the number of women and couples seeking fertility

treatments increases, it is important to understand why pregnancy fails. In this thesis, oocyte

quality and its decline with maternal age are discussed, with a focus on the effects of

mitochondrial dysfunction. Factors and disorders affecting proper mitochondrial function are

introduced, and methods of both improving and preserving oocyte quality are reviewed.

Table of Contents

Abstract ii

Table of Contents iii

List of Abbreviations iv

Introduction 1

Chapter 1 Oocyte development and maturation 10

Chapter 2 Mitochondrial function and structure 15

Chapter 3 Mitochondrial dysfunction 21

Chapter 4 Oocyte quality decreases with age 31

Chapter 5 Strategies for improving oocyte quality 47

Figures 60

Literature Cited 68

List of Abbreviations

aCGH array comparative genomic hybridization

ADP adenosine diphosphate

AGE advanced glycation end-product

AL ad-libitum

APC/C anaphase promoting factor/cyclosome

ART assisted reproductive technologies

ATP adenosine triphosphate

AUGMENT autologous germline mitochondrial energy transfer

BMI body mass index

CGH comparative genomic hybridization

CoQ10 coenzyme Q10

CR caloric restriction

FADH2 flavin adenine dinucleotide

FISH fluorescence in situ hybridization

FSH follicle stimulating hormone

H2O2 hydrogen peroxide

ICSI intracytoplasmic sperm injection

IMM inner mitochondrial membrane

IVF in vitro fertilization

LH luteinizing hormone

m mitochondrial membrane potential

MAPK mitogen-activated protein kinase

MCC mitotic checkpoint complex

MG methylglyoxal

mtDNA mitochondrial DNA

MTOC microtubule-organizing center

NADH nicotinamide adenine dinucleotide

OMM outer mitochondrial membrane

OSC oogonial stem cell

PCOS polycystic ovarian syndrome

PI3K phosphoinositide 3-kinase

POL DNA polymerase gamma

PSSC premature separation of sister chromatids

RCS reactive carbonyl species

RNA ribonucleic acid

ROS reactive oxygen species

rRNA ribosomal RNA

SAC spindle assembly checkpoint

SIRT1 sirtuin 1

SOD superoxide dismutase

TCA tricarboxylic acid

TFAM mitochondrial transcription factor A

tRNA transfer RNA

Introduction

In today’s society, women are deciding to have children later than ever before. A record

number of women in their 20’s are putting off having families until later in life, and many are

choosing to start careers before having children (Hamilton et al. 2013). While the birth rates for

women in their 20’s were reported at a record low of 83.1 births per 1,000 women in 2012, birth

rates for women in their 30’s and 40’s have increased (Hamilton et al. 2013). Couples of all ages

seek fertility assistance for a number of reasons, but younger women are more likely to have

successful pregnancies on their own (Van Voorhis 2006). Women in their 20’s have

approximately a 9% chance of suffering from miscarriage at some point during pregnancy, but

this number jumps to 75% for women 40 years old and above (Nybo Andersen et al. 2000). This

has led to an increase in the number of couples and individuals seeking fertility assistance, and

sparks the conversation about the negative relationship between oocyte quality and maternal age.

It is well known that as a woman matures, her chances of successful pregnancy decrease.

Although women in their 40’s and 50’s have been known to have healthy pregnancies, female

fertility typically peaks at age 25 years and drops dramatically after age 35 (Nybo Andersen et al.

2000; Hook 1981).

Oocyte quality plays an important role in the determination of pregnancy success.

Though assisted reproductive technology (ART) methods often fail for older women using their

own oocytes, success can be achieved using donor oocytes from younger women, and post-

menopausal women have also served as successful surrogates (Sauer et al. 1995). This

demonstrates that decreased fertility appears to lie majorly within the oocyte itself rather than the

uterine environment. An increased understanding of the effects of aging and other metabolic

disorders on oocyte quality would give researchers and clinicians the ability to improve fertility

and pregnancy outcomes for many women. While many technologies have been developed to

increase the chances of pregnancy for patients, the efficiency rates of these methods are still low

and there is much room for improvement (CDC 2013). To increase egg quality in both ART and

natural conception, it is important to understand what exactly defines a “good” oocyte and the

mechanisms behind its ultimate failure.

Many mechanisms of fertilization and embryogenesis fail when oocyte quality is poor.

Depending on the degree of impairment, failure can occur during fertilization, implantation into

the uterine wall, or later in development. Decreased oocyte quality often results in aneuploidy,

the incomplete separation of chromosomes during meiosis. When this occurs, cells contain the

incorrect number of chromosomes needed for proper development. Aneuploidy can occur in any

of the 23 pairs of chromosomes, but most aneuploidy embryos do not survive. In fact, euploid

embryos are much more likely to develop to the blastocyst stage than aneuploid embryos (93%

vs. 21%) (Sher et al. 2007). Trisomies, a result of aneuploidy, are more often seen in

pregnancies of older women than younger women. The chances of a woman giving birth to a

child with trisomy 21, also known as Down syndrome, increase to 1 in 11 at age 49 (Hook 1981).

Aneuploidy and other problems that arise with advanced maternal age are associated with

an increase in the degree of mitochondrial dysfunction present in oocytes (Volarcik et al. 1998).

As a woman ages, oocyte quality declines until eventual ovarian failure, also known as

menopause. When oocytes begin to “go bad”, it is often due to accumulated mitochondrial

dysfunction (Chistiakov et al. 2014; Jansen & Burton 2004; Bentov et al. 2011; Kushnir et al.

2012). As a result, oocytes do not have enough energy available to perform necessary cellular

functions. This is disadvantageous for oocytes, since spindle formation and chromosomal

segregation, as well as other important stages of cell division, are highly energy-intensive

processes (reviewed in Dumont & Desai, 2012).

Mitochondria are the organelles within the cell that supply most of the necessary energy

needed for proper function in the form of adenosine triphosphate (ATP), and play a crucial role

in metabolism. Mitochondria contain their own DNA separate from nuclear DNA, and have

approximately 1-10 copies of circular mitochondrial DNA (mtDNA) per organelle (Anderson et

al. 1981). The 13 proteins encoded by mtDNA make up many of the electron transport chain

complexes needed for oxidative phosphorylation, and additional proteins necessary for

mitochondrial function are translocated from the nucleus (Schatz 1979). ATP is created through

the process of oxidative phosphorylation carried out across the electron transport chain within

the inner membrane of the mitochondria. Oxidative phosphorylation typically results in the

formation of ATP and water, but this process is not 100% efficient (Porter & Brand 1995).

When oxygen, the final electron acceptor, is incompletely reduced, superoxide radicals are

formed. Superoxide radicals can form reactive oxygen species (ROS) that disrupt normal

cellular activities and damage DNA molecules when not balanced with antioxidant defenses

(Halliwell 2007; Raha et al. 2000; Haigis & Yankner 2010). To compound the problem,

dysfunctional or damaged mitochondria produce increased levels of ROS, thereby damaging

more mitochondria in the process (Crompton 1999). MtDNA molecules are in such close

proximity to the production of ROS within the mitochondria that they are unable to avoid

damage. The nucleus employs certain protective mechanisms to preserve the integrity of nuclear

DNA from ROS-induced oxidative damage, but these are lacking in mtDNA. Unlike nuclear

DNA, mtDNA is not wound around protective histone proteins, contains no non-coding introns,

and lacks the repair mechanisms necessary to repair damage caused by ROS (reviewed in

Palikaras & Tavernarakis, 2014). This leaves mtDNA highly susceptible to mutations.

Whereas nuclear DNA is inherited from both parental lineages, and therefore able to

overcome genetic mutations to a certain degree, mtDNA is uniparental and only inherited from

the mother (Giles et al. 1980). Sperm mitochondria are destroyed shortly after fertilization by

autophagy (Kaneda et al. 1995; Thompson et al. 2003). For this reason, it is remarkable that a

woman of 25 years, whose oocytes have presumably experienced 25 years of ROS exposure, can

give birth to healthy children at a high rate of success (Hamilton et al. 2013). Because mtDNA

inheritance is uniparental, it is important to consider the degree of fertility of children born to

older mothers. The age of the maternal grandmother at the time of the mother’s birth is an

influential factor associated with an increased risk of Down syndrome (Malini & Ramachandra

2006; Aagesen et al. 1984). For every year after age 30, the risk of the grandchild having Down

syndrome increases by 30% (Malini & Ramachandra 2006; Aagesen et al. 1984).

Mitochondrial dysfunction can be measured in a number of ways. MtDNA copy number

can be used to predict which oocytes are more likely to complete successful fertilization and

embryogenesis through development (Wai et al. 2010). A threshold (about 50,000 copies in both

humans and mice) exists for the amount of mtDNA necessary for development, and oocytes with

an mtDNA copy number below the threshold are less likely to implant into the uterine wall

during blastocyst formation (Wai et al. 2010). From fertilization until implantation, mtDNA

replication is halted in mammalian zygotes. However, the early embryo is still growing and

developing at this time, and rapid cell division causes a dilution effect of mtDNA within

individual cells until replication can be restarted. If the original oocyte has mtDNA copy

numbers below the threshold, it is not able to recover from this rapid cell division in order to

continue embryogenesis (Wai et al. 2010).

The integrity of mtDNA is also important for normal mitochondrial function. When

mtDNA becomes damaged, the mitochondrion tries to rescue itself by fusing with another

healthy mitochondrion. This dilutes the defective mtDNA with healthier mtDNA and can

ameliorate its effects on the cell. If the damage is beyond repair or accumulates to an irreparable

degree, the mitochondrion is phagocytized through a process called mitophagy that clears the

damaged mitochondrion from the cell (Lemasters 2005). With age, this clearance mechanism

becomes impaired and increased numbers of dysfunctional mitochondria begin to aggregate

within cells (Wilding et al. 2001). As dysfunctional mitochondria aggregate, increased levels of

ROS are produced and damage is often compounded.

DNA polymerase- (POL) is an enzyme translocated from the nucleus that is

responsible for replicating mtDNA within mitochondria. Because POL is the only DNA

polymerase within mitochondria (Bebenek & Kunkel 2004), any mutations in the catalytic

subunits may lead to dysfunction of replication. Mitochondrial dysfunction can occur from a

number of mutations in Pol, causing decreased oocyte quality and mitochondrial diseases in

mutant mice (Chan & Copeland 2009).

Mitochondria contain both an inner and outer membrane, both of which are important for

functional oxidative phosphorylation. Maintenance of mitochondrial membrane potential is

correlated with the ability of the organelle to effectively produce ATP (Wilding et al. 2003; Van

Blerkom et al. 1995). Embryos with lower membrane potential are more likely to have random

segregation of chromosomes (Wilding et al. 2003). If the mitochondrial membrane potential is

decreased or disrupted, mitochondria lose the ability to create the electron gradients necessary to

drive ATP production. Therefore, mitochondrial membrane potential is typically used as an

indicator of mitochondrial dysfunction (Komatsu et al. 2014; Wilding et al. 2003).

It is of great importance to understand why oocytes fail and how it may be possible to

reverse their phenotypes to resemble younger, healthier oocytes. Diabetic or obese women often

have oocytes with similar phenotypes as older women, regardless of their age, and therefore have

an increased risk of aneuploidy and miscarriage (Sadler et al. 1988; Colton et al. 2002). Many

women also suffer from premature ovarian failure (menopause onset before age 40) for a number

of reasons, including genetic disorders like Turner syndrome, or chemotherapy and other cancer

treatments (Kalantaridou et al. 1998). Women with these conditions, as well as those with

fertility lost by natural aging, would certainly benefit from treatments aimed at increasing oocyte

quality.

While many improvements have been made in the last few decades in the field of assisted

reproduction, no method has been able to achieve 100% success. The successful use of young

donor eggs for aged women, or older surrogates for younger women with in vitro fertilization

(IVF) or intra-cytoplasmic sperm injection (ICSI) demonstrate that oocyte quality is more of a

determinant of pregnancy success than the receptivity of the host uterus; older patients using

oocytes donated from younger patients are able to carry successful pregnancies when using their

own aged oocytes fails (Sauer et al. 1995). With this knowledge, clinicians took a small amount

of cytoplasm from a young oocyte and transferred it into an oocyte of patient with previously

failed pregnancy attempts in the hopes of improving oocyte quality. Successful pregnancies

were achieved with this method (Cohen et al. 1998), but it was soon realized that these children

had inherited mtDNA from both the mother and the donor, resulting in mitochondrial

heteroplasmy (Brenner et al. 2000; Barritt et al. 2001). Because the long-term effects of this

condition are unknown, the US Food and Drug Administration put a moratorium on this

procedure in 2001. For this reason, cytoplasmic transfers are not a feasible method for

improving oocyte quality. However, it is generally accepted that the mitochondria within the

young oocyte are the rejuvenating factor within the cytoplasm. Therefore, improving

mitochondrial function to increase oocyte quality has become a recent focus of research.

One such improvement has been the use of autologous, healthy mitochondria from a

woman’s oogonial stem cells to increase the quality of her oocytes during ART procedures

(Woods et al. 2013). Oogonial stem cells are adult stem cells that produce new immature

oocytes postnatally in mammalian females (Johnson et al. 2004). Mitochondria within these

oogonial stem cells, which are from the same cell lineage as oocytes, would therefore provide the

best autologous source of mitochondria for improving oocyte quality in patients. These oogonial

stem cell mitochondria are less differentiated than oocytes residing in the ovary, and therefore

are less likely to have accumulated genomic damage that would cause dysfunction. The process,

coined “AUGMENT” (autologous germline mitochondrial energy transfer), is promising

(reviewed in Tilly & Sinclair, 2013), but extracting oogonial stem cells from female patients and

further isolating mitochondria is challenging and invasive. Therefore, emphasis has shifted from

transferring functional mitochondria into poor-quality oocytes to how to improve a woman’s

endogenous oocyte mitochondria.

Because mitochondrial function is impaired mostly by ROS, scientists have begun to

focus on how to counteract this damage. Antioxidant treatments have been promising as a way

to preserve oocyte quality in aged females, but can cause additional damage to the function of

reproductive somatic tissues in the process, thereby decreasing overall fertility (J J Tarín et al.

2002). Levels of the necessary electron transport chain component Coenzyme Q10 (CoQ10)

naturally decrease with age (Pignatti et al. 1980), which once again links aging with increased

mitochondrial dysfunction. CoQ10 supplements have been shown to increase oocyte quality and

menopausal cognitive effects in mice by affecting mitochondrial dysfunction and ROS

production (Gendelman & Roth 2012; Sandhir et al. 2014). Resveratrol, a dietary-restriction

mimetic, has also been suggested to improve oocyte quality by acting as a potent antioxidant.

This polyphenol has been shown to preserve oocyte quality in aging mice (Liu et al. 2013) as

well as improve anti-oxidation levels in bovine oocytes and embryos (Takeo et al. 2013).

Resveratrol acts through sirtuin 1 (SIRT1) to reduce oxidative stress by increasing mitochondrial

number and function, and SIRT1 inhibitors cause decreased fertilization (Takeo et al. 2013;

Takeo et al. 2014).

Another approach to preserving oocyte quality with age in mice is caloric restriction.

When diets are restricted, fertility is reduced, but it is able to rebound once normal feeding is

resumed (Ball et al. 1947; Nelson et al. 1985; Visscher et al. 1952; Lintern-Moore & Everitt

1978). When applied to aged mice with poor oocyte quality, this treatment resulted in marked

increases in the fertility of females compared to age-matched controls once the dietary

restrictions were removed (Selesniemi et al. 2008). Because obesity and diabetes are linked with

subfertility in women of any age (Purcell & Moley 2011; Wang & Moley 2010; Grindler &

Moley 2013; Jungheim & Moley 2010), it is not surprising that oocyte quality benefits from a

low-calorie diet. Because similar studies have yet to be conducted in humans, it is unclear

whether the treatment durations could be feasibly translated from mice to women.

Summary

Regardless of the underlying mechanisms, it is clear that female fertility eventually

reaches its limit by the fifth or sixth decade of life (Richardson et al. 1987). Aged oocytes have

different characteristics from younger oocytes, and understanding what makes them so different

may provide clues for improving quality. Oocytes from young, healthy females have a stable

mitochondrial membrane potential, mtDNA copy numbers well above threshold, and mtDNA

with minimal damage. Due to efficiently functioning mitochondria, adequate levels of ATP are

provided to the cell, allowing for normal spindle formation and equal chromosomal segregation

during meiosis. In contrast, poor quality oocytes show a reduced mitochondrial membrane

potential, mtDNA copy number, and ATP levels, often due to (or resulting in) increased mtDNA

damage. Because ATP production is disrupted, chances of aneuploidy and developmental failure

are increased. Methods to improve oocyte quality have been developed, but a single method that

achieves 100% success has not yet been discovered. A better understanding of the mechanisms

responsible for oocyte failure is clearly required to move forward in the field of ART and fertility

treatments.

This thesis will review the present literature on oocyte quality and how it is negatively

affected by age, with a focus on mitochondrial dysfunction. Factors that determine oocyte

quality, the consequences of poor quality, and current methods used to improve quality will also

be discussed.

CHAPTER 1: Oocyte development and maturation

It has long been believed that women are born with all of the oocytes they will ever have

throughout their lifetime (Zuckerman 1951). Because menopause, caused by reduced ovarian

function and eventual ovarian failure, occurs naturally with age, it was previously assumed that

menopause is the result of a depletion of healthy ovarian follicles (Richardson et al. 1987).

Many suggested that once a woman’s follicular reserve she was endowed with at birth was

exhausted, the ovaries were depleted and menopause was initiated (Richardson et al. 1987). For

decades this dogma remained unchallenged, yet females of many other vertebrate and

invertebrate species studied exhibit some form of germline stem cell and produce new oocytes

throughout adulthood (Kimble 1981; Hirsh et al. 1976; Lin & Spradling 1997; Nakamura et al.

2010), making mammals the exception to the rule.

In 2004, it was discovered that mitotic germ line stem cells are present within the ovaries

of adult female mice (Johnson et al. 2004). These oogonial stem cells have been isolated from

mouse ovaries and used to produce live offspring (Zou et al. 2009). Additionally, oogonial stem

cells have been found in the ovaries of adult women (White et al. 2012), suggesting that ongoing

germ cell renewal can occur in humans of both sexes. It is currently unclear the extent of which

these cells can be used to improve female fertility, but many studies have suggested using these

undifferentiated cells as reproductive therapies and possibly replacement oocytes. It has been

suggested that older women whose mature oocytes are damaged from aging may benefit from the

use of autologous oogonial stem cell replacements (reviewed in Tilly & Sinclair 2013). While

many therapies are currently practiced for assisted reproduction, this recent discovery reinforces

the notion that there is still much to be learned about female reproduction.

Development of primordial follicles to fertilization of the oocyte

The generation of oocytes first begins in the fetal ovary. Mitotic divisions increase the

number of germ cells within the fetal ovary, and incomplete cytokinesis creates the formation of

joined cells known as germ cell cysts (Pepling & Spradling 2001; Kezele et al. 2002). At

approximately the 11th or 12th week of gestation, follicles begin to form and meiosis begins

(Gondos et al. 1986; Martins da Silva et al. 2004). It has been suggested that retinoic acid

triggers the switch from mitosis to meiosis that defines the transition from oogonia to oocytes

(Bowles & Koopman 2007). This switch causes the germ cell cysts to dissociate and individual

oocytes become surrounded by a single layer of pre-granulosa somatic cells to form primordial

follicles (Kezele et al. 2002).

Before arresting in prophase I, oocytes must carry out synapsis and recombination of

homologous chromosomes to ensure successful future development and survival (reviewed in

Eichenlaub-Ritter, 1998). The location and number of crossovers during meiosis I are important

for preventing errors in the first meiotic division (reviewed in Hunt & Hassold, 2008).

Crossovers too close to either the telomeres or centromeres of the chromosomes can have

detrimental effects on future embryonic development and survival (Hassold et al. 2007). During

follicular formation, gap junctions are created between follicular somatic cells and the oocyte in

order to coordinate their functions. Atresia, a form of regulated cell death, occurs continuously

throughout follicular development and the majority of follicles die before ovulation (reviewed in

Hunt & Hassold, 2008; Martins da Silva et al., 2004). Those that do make it move on to the later

stages of follicular development. Oocytes within primordial follicles continue through meiosis

to prophase I, where cell division is arrested in the dictyate stage. Although it is unknown what

causes the recruitment of follicles from the primordial pool, many primordial follicles remain

dormant in humans for decades before further development and ovulation occur.

Once follicles are recruited from the primordial follicle pool, follicular development

resumes while the oocyte remains in prophase I. As the primordial follicles develop into primary

follicles, the granulosa cells become more cuboidal and a theca layer develops (Kezele et al.

2002). Granulosa cells are in direct contact with the oocyte, whereas theca cells form in the

surrounding stromal tissue. Gap junctions between somatic cells and also between somatic cells

and the oocyte are important during early follicular growth, as gonadotropin levels that support

development are only adequate after reaching sexual maturation (i.e. puberty) (Hutt & Albertini

2007; Downs 1995; Donahue & Stern 1968; Grazul-Bilska et al. 1997). Both the surrounding

somatic cells and oocytes rely on each other for bi-directional signaling to promote maturation

and development of the follicle (Knight & Glister 2006; Hutt & Albertini 2007; Woodruff &

Mayo 2005; Kezele et al. 2002). As the follicle grows, the number of cuboidal granulosa cell

layers increases and the oocyte grows larger. Once the follicle has developed to the pre-antral

stage, follicular fluid begins to form between the granulosa and theca cell layers (Hirshfield,

1991; reviewed in Peters, 1969). When a woman enters puberty, the oocyte is now exposed to

gonadotropins such as follicle stimulating hormone (FSH) that support the growth and ovulation

of large, Graafian follicles (Halpin et al. 1986; Cattanach et al. 1977; Kumar et al. 1997). This

transition is significant, as development from a pre-antral to antral follicle provides the oocyte

with the ability to resume meiosis (Sorensen & Wassarman 1976).

Mid-way through the menstrual cycle, the surge of luteinizing hormone (LH) induces the

completion of meiosis I and the ovulation of the mature oocyte from the ovary. Whereas oocytes

arrested in prophase I are characterized by an intact nuclear membrane known as the germinal

vesicle, oocytes that have completed meiosis I show germinal vesicle breakdown, condensation

of chromosomes, and the assembly of the bipolar spindle (Leung & Adahshi 2004). The

completion of meiosis I segregates homologous chromosomes into the oocyte and the first polar

body. After meiosis I, the oocyte becomes arrested again, this time in metaphase II with sister

chromatids aligned on the metaphase II plate. Meiosis II is completed and the second polar body

is formed once fertilization occurs. If fertilization is successful, pronuclei form around the

genetic material of the oocyte and the sperm. The first mitotic division is initiated once the

pronuclear envelopes dissolve and genetic material combine to form the zygote.

Spindle assembly in oocytes

In order for cell division to occur, oocytes must have proper spindle formation. Mitotic

somatic cells contain centrosomes from which the centrioles are formed on either side of the cell.

These centrioles assemble alpha and beta tubulin to make microtubules of the spindle (Vogt et al.

2008; Dumont & Desai 2012). These microtubules then extend towards the center of the cell,

continuously lengthening and shortening in attempts to attach to the kinetochores of

chromosomes. Once chromosomes are properly aligned and the correct connections are made

with microtubules, the cell proceeds into anaphase where the chromosomes are pulled apart from

their kinetochores and are equally separated into two daughter cells (Vogt et al. 2008; Dumont &

Desai 2012).

Cell division proceeds in a similar fashion in sperm, but oocytes have a slightly different

process. Oocytes lack centrosomes and therefore meiosis in oocytes is considered an

“acentrosomal” process. Rather than originating on the periphery of the cell, microtubule-

organizing centers (MTOCs) are adjacent to chromosomes throughout the cell. Once meiosis is

initiated, the MTOCs form a ball in the center of the cell and the chromosomes line up along its

center during metaphase. Microtubules must make proper connections with chromosomes to

enter into anaphase, although the checkpoint here is not as strict as it is in somatic cells and

sperm cells (LeMaire-Adkins et al. 1997). As microtubules are pulled towards opposing sides of

the cell, the chromosomes are pulled along with them, segregating the chromosome pairs into

two cells.

CHAPTER 2: Mitochondrial function and structure

Mitochondria are the major energy producers of eukaryotic cells. These organelles are

the site of many metabolic functions including the tricarboxylic acid (TCA) cycle (also known as

the Krebs cycle or the citric acid cycle) and oxidative phosphorylation, which lead to ATP

production. ATP is used by countless energy-dependent processes in the cell including cell

division, proteasome activity, and microtubule function (Goldberg & St John 1976; Bershadsky

& Gelfand 1981; Gottesman & Maurizi 1992; Robinson & Spudich 2000). Mitochondria are

made up of two membranes, the inner mitochondrial membrane (IMM) and the outer

mitochondrial membrane (OMM). The IMM is made up of many folded cristae, which increase

the surface area of the IMM to allow for additional production of ATP. Within the IMM lies the

mitochondrial matrix, the site of the TCA cycle. The electron transport chain is made up of

various protein complexes located across the IMM that produce a proton gradient as a result of

the transfer of electrons between complexes. Mitochondria then utilize this proton gradient

created within the intermembrane space to drive ATP production (reviewed in Lodish, Berk, &

Zipursky, 2000).

Mitochondrial DNA

As a result of the endosymbiosis of two prokaryotic cells that formed the first eukaryotic

cell, mitochondria contain approximately 1-10 copies of their own DNA distinct from the cell’s

nuclear DNA (Anderson et al. 1981). The importance of mitochondria in oocyte quality stems

from the fact that mitochondrial DNA (mtDNA) is maternally inherited, rather than nuclear

DNA, which is inherited as a combination of maternal and paternal DNA (Giles et al. 1980).

Although it is unclear exactly how the mechanism functions, it is known that upon fertilization,

paternal mtDNA is degraded by ubiquitin-proteasome machinery within the oocyte that

recognizes the sperm cell’s ubiquitin substrates, targeting the mitochondria for destruction

(Kaneda et al. 1995; Thompson et al. 2003). Mitochondrial DNA molecules from the mother

then go on to act as a template for all of the mtDNA found in the offspring. This 16.5kb circular

DNA strand contains 37 genes that encode 13 proteins, 22 transfer RNAs (tRNAs), and 2

ribosomal RNAs (rRNAs) (Anderson et al. 1981). Proteins encoded by mtDNA include subunits

of the electron transport chain that are essential for ATP production. Although mtDNA only

codes for 13 proteins, there are over 1500 found in mitochondria, indicating that the remaining

proteins are imported from the nucleus (Schatz 1979). Proteins found in the mitochondria are

specific to the cell type, which demonstrates cross-talk between mitochondria and nuclei to carry

out specific cell functions (Johnson et al. 2007).

mtDNA replication

In order for mtDNA to be successfully transmitted from mother to offspring, it must

undergo DNA replication at the appropriate time during development. mtDNA molecules

contain a heavy and a light strand that must both be replicated (reviewed in Shoubridge & Wai

2007). There are multiple theories on how exactly the circular mtDNA is replicated, two of

which being the coupled leading/lagging strand synthesis model and the asymmetrical synthesis

model (Yasukawa et al. 2005; Shadel & Clayton 1997). The coupled leading/lagging strand

model predicts that both the heavy and light strands are replicated simultaneously in opposite

directions from the same origin (Yasukawa et al. 2005). On the other hand, the asymmetric

synthesis model states that the heavy strand is synthesized first, and the light strand initiates

replication when the heavy strand replication has reached two thirds of completion (Shadel &

Clayton 1997). Unfortunately conclusive evidence is lacking to determine which model is

accurate, and many scientists disagree (Bogenhagen & Clayton 2003; Holt & Jacobs 2003).

Although oocytes contain more mitochondria per cell than any other cell type, the

immature mitochondria of oocytes and early embryos are rounded with few cristae and little

capacity for oxidative phosphorylation (Sathananthan et al. 2002; P May-Panloup et al. 2005;

Pikó & Matsumoto 1976; Jansen & de Boer 1998). It is not until embryonic cells begin to

develop that mitochondria become elongated and gain the ability to carry out typical metabolic

function (Trimarchi et al. 2000; Wilding et al. 2001; Houghton 2006). Until blastocyst

implantation, mtDNA replication is arrested and the expression of replication factors is

extremely low (Thundathil et al. 2005; Pascale May-Panloup et al. 2005; Spikings et al. 2007;

Pikó & Taylor 1987). MtDNA copy number decreases due to increased cellular division during

these stages of embryogenesis, further reducing the mitochondrial capacity of each cell to carry

out oxidative phosphorylation (Thundathil et al. 2005; Pikó & Taylor 1987). Replication of

mtDNA begins again when the blastocyst implants into the uterine wall and replication factors

are once again upregulated (Spikings et al. 2007; Thundathil et al. 2005; Pascale May-Panloup et

al. 2005; Pikó & Taylor 1987). This process of cell division without mtDNA replication

followed by the replication of the remaining mtDNA molecules is thought to create genetic drift

among mtDNA variants in attempts to prevent mutant mtDNA from populating the offspring

(Fan et al. 2008; Stewart et al. 2008). This creates a mitochondrial bottleneck through which the

random selection of mtDNA occurs (Hauswirth & Laipis 1982). Though the details of this

selection process are unclear, it may not be entirely random, as large mtDNA mutations are

typically not passed onto offspring as frequently as would be expected by chance (Keefe et al.

1995).

Metabolism – electron transport chain and oxidative phosphorylation

One of the most important roles of mitochondria is their function in cellular metabolism.

Many species use aerobic cellular respiration to convert glucose into ATP energy through

glycolysis, the TCA cycle, and oxidative phosphorylation (reviewed in Karp, 2013; Lodish,

Berk, & Zipursky, 2000). Glucose is first converted into pyruvate through glycolysis in the

cytoplasm of the cell and then to acetyl-CoA by pyruvate dehydrogenase (reviewed in Balaban et

al. 2005). Products of the TCA cycle such as NADH and FADH2 become oxidized and donate

electrons to the first steps of the electron transport chain to initiate oxidative phosphorylation.

Oxidative phosphorylation involves the transfer of electrons along a chain of proteins that

subsequently uses redox reactions to create a proton gradient across the IMM. This built up

gradient of protons then drives the final enzyme in the electron transport chain, ATP synthase.

When protons flow back down the generated concentration gradient and return to the

mitochondrial matrix, it drives the phosphorylation of adenosine diphosphate (ADP) by ATP

synthase to create ATP. Oxidative phosphorylation produces approximately 34 ATP per glucose

molecule, whereas aerobic respiration as a whole generates up to 38 ATP total (Rich 2003;

Boyer et al. 1977; Lodish et al. 2000).

Complex I, NADH dehydrogenase, is the first enzyme in the electron transport chain, and

is made up of proteins encoded by both nuclear and mitochondrial DNA. When NADH is

hydrolyzed into NAD+ and a proton, 2 electrons are donated to complex I, which drives 4

protons across the IMM into the intermembrane space. Complex II, succinate dehydrogenase,

acts as a second point of entry for electron donation and, like complex I, links oxidative

phosphorylation with the TCA cycle by oxidizing FADH2. This enzyme oxidizes succinate to

fumarate and reduces ubiquinone, which does not create as much energy as complex I, and

therefore does not contribute to the proton gradient across the IMM (Lodish et al. 2000; Karp

2013). Electrons from both complexes I and II are further transferred one at a time to coenzyme

Q10, also known as ubiquinone. This carrier then passes the electrons onto complex III,

coenzyme Q reductase, where 4 protons are transferred across the IMM into the intermembrane

space. From complex III, electrons are passed onto cytochrome c, which acts as a substrate for

complex IV. The final electron acceptor in the electron transport chain is oxygen, which

produces two water molecules as final end products of oxidative phosphorylation after receiving

4 electrons from complex IV. This reduction releases two final protons across the IMM to add to

the electrochemical gradient. Therefore, a total of 10 protons are released into the

intermembrane space for each pair of electrons transferred through oxidative phosphorylation to

create a concentration gradient across the IMM (Lodish et al. 2000). ADP is phosphorylated to

ATP when protons move down their electrochemical gradient to drive the function of ATP

synthase. ATP synthase is made up of two joined complexes, a rotating hexamer driven by a

transmembrane proton channel, which function together to drive ATP synthesis.

Reactive oxygen species

Although oxidative phosphorylation is important for cell survival, it is not 100% efficient

(Porter & Brand 1995). Electrons can leak out of the electron transport chain when they are not

sufficiently transferred between the complexes and can react with oxygen to create superoxide

radicals (O2-). Complexes I and III are known to be the main source of superoxide radicals in

mitochondria (Dröse & Brandt 2012; Lesnefsky et al. 2001; Lenaz 2012; Sugioka et al. 1988;

Wallace 2000). ROS are created when these superoxide radicals are not converted into hydrogen

peroxide (H2O2) or water via the antioxidant enzyme superoxide dismutase (SOD) (Halliwell

2007; Raha et al. 2000). While ROS are typically known as damaging molecules, they have also

been found to play a role in normal cell function at low levels by acting as second messengers in

MAPK and PI3K pathways, as well as the immune response (Devadas et al. 2002; Genestra

2007; Kwon et al. 2004; S.-R. Lee et al. 2002; Seo et al. 2005; Tobiume et al. 2001)

Although antioxidant defenses are in place to maintain homeostasis within the cell

(Chatterjee et al. 2007; Valko et al. 2006), increases in ROS production can over-power these

protection mechanisms, a process known as oxidative stress that causes cellular damage (Haigis

& Yankner 2010). Increased oxidative stress can damage proteins, lipids, and nucleic acids

within the vicinity of ROS production; therefore mitochondria are often the most affected

organelles (reviewed in Ray, Huang, & Tsuji, 2012). When too much oxidative stress occurs,

cells become unable to function (Genestra 2007). One facet of cellular aging has been attributed

to the accumulation of ROS in cells and the disruption of the oxidant/antioxidant balance

(Herrero & Barja 1997; Moghaddas et al. 2003). Therefore, it is important to understand how

the function of mitochondria influences oocyte quality with increased maternal age, and more

importantly, how oocyte quality can possibly be improved.

CHAPTER 3: Mitochondrial dysfunction

With advanced age, mitochondrial dysfunction increases in both somatic and germline

cells (Taylor et al. 2003; Michikawa 1999; Kang & Hamasaki 2005; Jansen & Burton 2004;

Tatone et al. 2010; Eichenlaub-Ritter et al. 2011). As cells age through natural processes, the

ability to produce ATP is gradually lost through several mechanisms, including decreased

mitochondrial membrane potential, decreased mtDNA copy number, and increased organelle

swelling (Wilding et al. 2003; De Boer et al. 1999; Müller-Höcker et al. 1996; Van Blerkom

2004). These phenomena contribute to impaired ATP production and reduced energy to carry

out basic cellular functions. Oocytes are perhaps the cells most affected by mitochondrial

dysfunction, as they have more mitochondria than somatic cells and provide the template for all

mitochondria within offspring (May-Panloup et al. 2007; Ashley et al. 1989).

The loss of normal mitochondrial function in oocytes is often a major determinant of

pregnancy failure in older women (Lee et al. 2014; May-Panloup et al. 2007; P May-Panloup et

al. 2005; Wilding et al. 2009; Jansen & Burton 2004; Komatsu et al. 2014; Van Voorhis 2006).

Because oocyte mitochondria provide the mtDNA for the entire offspring, healthy inherited

mitochondria are important for the survival of the embryo as well as for the health of the future

child (Giles et al. 1980). Without the preservation of inherited mtDNA, electron transport chain

proteins important for ATP production may be transcribed incorrectly in the offspring (Preston et

al. 2008). Mitochondria are the main producers of cellular ATP and are therefore important for

highly energy-dependent processes such as oogenesis and embryonic development (Van

Blerkom et al. 1995; Van Blerkom 2004; Wilding et al. 2009; El Shourbagy et al. 2006; Spikings

et al. 2007). This chapter will examine the factors involved in mitochondrial dysfunction and

how these abnormalities increase with advanced maternal age in the oocyte. Overall, many of

these factors lead to ATP depletion, which decreases the ability of oocyte fertilization,

implantation, and further embryogenesis to occur successfully (Van Blerkom et al. 1995).

Mitochondrial biogenesis vs. mitophagy

In order to regulate mitochondrial health, there must be a balance between the creation of

new mitochondria and the disposal of dysfunctional mitochondria. Mitochondrial biogenesis and

mitophagy, the processes by which mitochondria are created and removed, respectively, achieve

this homeostasis (Ding & Yin 2012). Mitochondrial biogenesis involves the generation of new

mitochondria, and is regulated by factors such as the cellular environment (i.e. available nutrients

and temperature), hormone stimulation, and growth factors (reviewed in Palikaras &

Tavernarakis, 2014). When mitochondria become damaged or impaired, they fuse with other

healthy mitochondria to alleviate any mtDNA damages with intact mtDNA of the healthy

mitochondrion (Lemasters 2005). If mitochondria become damaged beyond the point of fusion

rescue, they must be removed from the cell to prevent further harm to the surrounding healthy

mitochondria, especially when this damage causes increased ROS production (Amicarelli et al.

2003; Wang et al. 2013; Takeo et al. 2013). During mitophagy, mitochondria are tagged for

disposal and phagocytized by lysosomes to digest dysfunctional mitochondria (Ding & Yin,

2012; reviewed in Palikaras & Tavernarakis, 2014). The PINK1/Parkin pathway mediates

mitophagy (Lazarou et al. 2012; Narendra et al. 2008). When mitochondria are dysfunctional,

the PINK1 enzyme becomes stabilized on the OMM and recruits Parkin (Narendra et al. 2010).

This ligase then ubiquitylates additional proteins on the OMM that allow the mitochondria to be

recognized by autophagosomic machinery within the cell. The mitochondria are then

sequestered into autophagosomes that deliver the damaged organelles to lysosomes that degrade

and dispose of the mitochondria (Yoshii et al. 2011; Chan et al. 2011).

This balance of mitochondrial biogenesis and mitophagy keeps cells functioning properly

(reviewed in Palikaras & Tavernarakis 2014). However, with age, these processes begin to

breakdown and homeostasis can be lost (FIGURE 1) (Fan et al. 2008; Bereiter-Hahn et al. 2008;

Preston et al. 2008). Increased activity of mitophagy mechanisms decreases mitochondrial

numbers, which lowers the overall amount of available ATP and can over-stress the remaining

mitochondrial population (Dagda et al. 2008; Yan et al. 2012; Zhu et al. 2007). If the cell is

unable to recover, lack of abundant ATP can ultimately lead to cellular death. However,

insufficient levels of mitophagy can also have detrimental effects. When mitophagy is impaired,

mitochondria begin to aggregate, which increases the amount of damaged mitochondria and also

increased ROS production, further damaging the surrounding mitochondria (Cavallini et al.

2007; Masiero & Sandri 2010; Wilding et al. 2001). Accumulation of mitochondria with age is

present in many organisms (H.-C. Lee et al. 2002; T.-M. Lee et al. 2002; Preston et al. 2008;

Bereiter-Hahn et al. 2008), yet it is unknown whether it is caused by decreased mitophagy or

increased mitochondrial biogenesis (Van Blerkom 2004).

Mitochondrial morphology, size, and structures

Studies from human oocytes show that both mitochondrial volume and number increase

with age (Müller-Höcker et al. 1996; Steuerwald et al. 2000). In metaphase II oocytes of women

aged 27 to 39 years, increased age is associated with increases in quantity, volume, and area of

oocyte mitochondria (Müller-Höcker et al. 1996). Differences in mitochondrial profile area with

age are most apparent when subjects are separated into two age groups: 27-30 years old, and 31-

39 years old. Aging has also been linked to morphological changes in mitochondrial cristae,

membrane, and vacuole structure (Simsek-Duran et al. 2013). Metaphase II oocytes from

hamsters and mice show that mitochondrial structure is significantly altered in aged oocytes from

each species (Simsek-Duran et al. 2013). In addition to collapsed cytoplasmic lamellae and

abnormal cristae, aged oocyte mitochondria display decreased ATP production and decreased

mtDNA copy numbers (Simsek-Duran et al. 2013), suggesting that abnormal mitochondrial

morphology leads to impairment of mitochondrial function. It has been proposed that increases

in the overall mitochondrial volume and number are associated with a compensatory mechanism

in order to “rescue” the cell from decreased mitochondrial function with age (Müller-Höcker et

al. 1996). When additional mitochondria become dysfunctional and are not disposed of by

mitophagy, the cell attempts to maintain ATP production by increasing the size and number of

the remaining mitochondria (Müller-Höcker et al. 1996). However, if these remaining

mitochondria are already dysfunctional, increasing the existing mitochondrial profile only

exacerbates present issues, leading to further mitochondrial dysfunction and oocyte failure

(Wilding et al. 2001; Genestra 2007; Gustafsson & Gottlieb 2009).

mtDNA replication

In order to maintain sufficient levels of ATP within the cell, mitochondria must have

proper transcription and translation of the proteins that make up the electron transport chain

(Moghaddas et al. 2003; Taylor et al. 2003). Because the majority of proteins encoded by

mtDNA are necessary for electron transport chain function, it is important for cells to contain

adequate copies of functional mtDNA. During embryonic development, ATP is crucial for the

segregation of nuclear material and cell division, and the level of ATP present in blastomeres is

associated with mtDNA abundance (Van Blerkom et al. 2000). Oocytes must contain certain

levels of mtDNA in order to successfully reach implantation and continue future development

(Wai et al. 2010). This is because although mtDNA replication is paused between metaphase II

and blastocyst implantation, the embryo is rapidly dividing during this period (Pikó & Taylor

1987; Thundathil et al. 2005). Cellular division and chromosomal segregation are energy-

intensive processes, so if the original oocyte lacks proper mtDNA copy numbers, the embryo

will not have enough energy to continue development as mitochondria become increasingly

segregated with each cell division (Wai et al. 2010).

Competent oocytes have greater numbers of mtDNA than incompetent oocytes, and

impaired mtDNA replication during oocyte maturation arrests development around the 6-cell

stage (Spikings et al. 2007). Additionally, the fertility outcomes of incompetent oocytes can be

recovered by supplementing with mitochondria from competent oocytes (El Shourbagy et al.

2006). In one study, competence of individual porcine oocytes was evaluated by the presence or

absence of glucose-6-phosphate dehydrogenase (G6PD) activity (Spikings et al. 2007).

Incompetent oocytes have ongoing G6PD activity whereas competent oocytes lose this enzyme;

therefore the activity of G6PD can be utilized through staining assays to determine overall

oocyte competence (Rodríguez-González et al. 2002).

The influence of mtDNA replication on oocyte competence can be examined to

determine the effects of decreased copy numbers on embryonic development (Spikings et al.

2007). In one study, oocytes were exposed 2’, 3’-dideoxycytidine (ddC) during in vitro

maturation to inhibit mtDNA replication. ddC disrupts replication by incorporating into newly

formed mtDNA to cause termination of the nucleoside chain, and cannot be removed by

polymerase gamma exonucleases (Spikings et al. 2007). Surprisingly, no difference in

fertilization ability was found between ddC-treated and untreated competent oocytes, but major

differences occurred during further embryogenesis. Oocytes with mtDNA replication disrupted

by ddC failed to develop beyond the 6-cell stage (Spikings et al. 2007). This shows that while

mtDNA copy number may have little effect on fertilization, mtDNA replication during oocyte

maturation is important to increase copy number to supply future embryonic development.

mtDNA copy number threshold

While it is understood that a certain number of mtDNA copies are important for proper

embryonic development, it is crucial to understand the threshold that defines competent and

incompetent oocytes. This may be especially important in older women seeking to become

pregnant, as low mtDNA copy numbers in oocytes have been associated with increased maternal

age (De Boer et al. 1999). Previous research has shown that low mtDNA copy numbers do not

appear to impair fertilization, but do have negative effects on post-implantation development

(Wai et al. 2010; Spikings et al. 2007). Although healthy, wild-type mouse oocytes have a range

of 11,000 to 428,000 copies of mtDNA, only 9 out of 219 embryos examined in one study

contained less than 50,000 (Wai et al. 2010). This shows that while the number of mtDNA

copies within healthy oocytes varies considerably, a certain threshold for competence is most

likely present.

To study the effects of low mtDNA copy number on oocyte fertilization and embryonic

development, heterozygous Tfam knockout mice can be used to create mice with low levels of

mtDNA (Wai et al. 2010). The TFAM (mitochondrial transcription factor A) protein is

important for mtDNA transcription and mitochondrial chromatin packaging, and therefore has

been suggested as a regulator of mtDNA copy number (Kaufman et al. 2007; Ekstrand et al.

2004). Genetically manipulated Tfam knockout mice were used to examine the effects of low

oocyte mtDNA copy number on fertilization and embryonic development. Low-copy oocytes

show normal fertilization and preimplantation development, but arrest shortly after the start of

embryogenesis (Wai et al. 2010). Similar outcomes are observed when low-copy embryos are

transplanted into wild-type pseudopregnant females, indicating that the impairment lies within

the oocyte itself rather than the maternal host environment (Wai et al. 2010).

These experiments revealed an estimated requirement of at least 50,000 mtDNA copies

for murine oocytes to successfully progress through development after blastocyst implantation

(Wai et al. 2010); human metaphase II oocytes show a similar threshold, with reported healthy

mtDNA copy numbers ranging from 50,000 to 1,500,000 (Reynier et al. 2001; P May-Panloup et

al. 2005). While healthy mammalian oocytes contain a wide range of mtDNA copy numbers, a

certain threshold exists, below which the mtDNA population cannot rebound from the successive

rounds of cleavage necessary for embryonic development (Wai et al. 2010). Because mtDNA

copy numbers have been shown to decrease with maternal age (De Boer et al. 1999), this may be

why older women struggle with maintaining pregnancies beyond fertilization.

mtDNA integrity

For proper mitochondrial function, mtDNA levels must be adequate in both quantity and

quality (Wai et al. 2010; Copeland & Longley 2014). Like nuclear DNA, mtDNA is vulnerable

to mutations and damage in both somatic and germline cells, but lacks many of the protective

mechanisms that help maintain nuclear DNA integrity, such as histones and non-coding regions

(Parsons et al. 1997). Several DNA polymerases maintain the integrity of nuclear DNA, but it is

thought that these mechanisms are inefficient in repairing prokaryote-like mtDNA (Bentov et al.

2011). Only DNA polymerase- (POL) is known to function in mitochondria to maintain

mtDNA integrity (Bebenek & Kunkel 2004), so it is not surprising that mutations in Pol can

cause significant mitochondrial disorders (reviewed in Chan & Copeland, 2009). Experiments

with mutations in Pol show that loss of this polymerase leads to increased ROS production and

overall mitochondrial dysfunction, often observed in aging (Lewis et al. 2007; Yen et al. 1989;

Taylor et al. 2003).

mtDNA mutations

Natural aging causes mtDNA mutations in many somatic tissues, such as the brain, liver,

skeletal and cardiac muscle (Cortopassi et al. 1992; Cortopassi & Arnheim 1990; Corral-

Debrinski et al. 1992; Yen et al. 1989), as well as oocytes (Keefe et al. 1995). Even in tissues

such as germline cells that may remain quiescent for decades, low levels of damaging ROS build

up over time to cause gradual mtDNA damage (Parsons et al. 1997). When natural protective

mechanisms against oxidative damage are overwhelmed by the increase in ROS production with

age, ROS begin to cause damage to proteins, lipids, and nucleic acids (Herrero & Barja 1997;

Moghaddas et al. 2003; Haigis & Yankner 2010). Although mtDNA mutations can cause severe

cellular dysfunction, it is suggested that mutations must accumulate within 30-80% of the

mtDNA population in order for a phenotype to be expressed (Lewis et al. 2007). Studies of

mutations such as the common 4977 deletion have shown that mutations are more prevalent in

oocytes from aged women compared to younger women (Keefe et al. 1995). Surprisingly, large-

scale deletions such as mtDNA4977 are rarely passed on to offspring; the mechanisms of this

transmission barrier are unknown, but a genetic bottleneck for mtDNA selection has been

proposed (Jansen & Burton 2004; Shoubridge & Wai 2007; Chinnery et al. 2004; Steffann et al.

2006). Although mtDNA selection is random, a genetic bottleneck may sufficiently limit the risk

of large mutations passing onto offspring (Chan & Copeland 2009; Fan et al. 2008; Corral-

Debrinski et al. 1992).

Mitochondrial membrane potential

While mtDNA integrity is necessary for the transcription and translation of important

mitochondrial proteins involved in oxidative phosphorylation, the mitochondrial membrane

potential must also be maintained in order to successfully produce ATP and carry out additional

mitochondrial functions (Mitchell & Moyle 1967; Shigenaga et al. 1994; Voisine et al. 1999). A

high mitochondrial membrane potential (m) across the inner mitochondrial membrane is

required to create the proton gradient involved in ATP generation during oxidative

phosphorylation (Wilding et al. 2003). When the m is disrupted or decreased with age,

adequate amounts of ATP are no longer produced by mitochondria within the oocyte and several

cellular functions begin to fail (Komatsu et al. 2014). A strong association exists in human

embryos between low m and abnormal meiotic spindle apparatuses (Wilding et al. 2003).

When data from these studies are separated by donor age, oocytes from women over the age of

37 years show dramatic increases in rates of spindle abnormalities and chaotic mosaicism,

characterized by the random segregation of chromosomes during meiosis (Wilding et al. 2003).

These data indicate that not only do the oocytes of older women show decreased m, but this

low m with age also increases the chance of abnormalities during embryonic development.

Conclusions

As the main energy producers of the cell, mitochondria play a large role in the

maintenance of cellular health and viability. Age-related studies regularly focus on

mitochondrial function as a benchmark of cellular health as many diseases and cancers are

associated with mitochondrial dysfunction and mtDNA damage (reviewed in López-Otín,

Blasco, Partridge, Serrano, & Kroemer, 2013). Because oocytes contain more mitochondria than

any other cell type (Jansen & de Boer 1998), it is important to understand how mitochondrial

function and dysfunction affect oocyte quality. While it is understood that oocyte quality

eventually decreases with age (Fragouli & Wells 2011; Hook 1981; Kim et al. 2013; Nybo et al.

2000; Yoon et al. 1996; Gaulden 1992; Schwarzer et al. 2014; Hassold & Chiu 1985), it is not

clear the exact mechanism by which this happens. Advanced age leads to increased

mitochondrial dysfunction, which, through multiple pathways, ultimately results in reduced

levels of ATP (Michikawa 1999; Taylor et al. 2003; Jansen & Burton 2004; Müller-Höcker et al.

1996; Eichenlaub-Ritter et al. 2011; Yen et al. 1989). Without the ability to produce an adequate

energy supply, oocytes are unable to carry out basic cell functions.

CHAPTER 4: Oocyte quality decreases with age

Because mitochondria are the primary energy source for the oocyte, it is not surprising

that loss of mitochondrial function can have detrimental effects on oocyte quality (Takeo et al.

2013; Bentov et al. 2011; Ford 2013; Van Blerkom et al. 1995). When mitochondrial

dysfunction occurs, sufficient levels of ATP are no longer provided to carry out the cellular

functions of the oocyte and problems can arise in oogenesis and embryogenesis once an oocyte

becomes fertilized (May-Panloup et al. 2007; Van Blerkom et al. 1995). These problems

typically manifest as pre or post implantation developmental arrest, implantation failure,

spontaneous abortion, or developmental defects in the offspring resulting from aneuploidy

(Robinson et al. 2001; Gaulden 1992; Allen et al. 2009; Hassold & Chiu 1985; Kushnir et al.

2012; Keefe et al. 1995).

Many studies have observed the increased risk of aneuploidy that is often associated with

increased maternal age (Fragouli & Wells 2011; Yoon et al. 1996; Hook 1981; Nybo Andersen et

al. 2000; Kim et al. 2013). Aneuploidy is the term used to describe any cell or group of cells that

contain an incorrect number of chromosomes, typically indicated by one too many (trisomic) or

one too few (monosomic) in a given chromosome pair. While approximately 20% of all human

oocytes contain an abnormal number of chromosomes, oocytes from older women show

aberrations in up to 50% (Hassold et al. 2007; Fragouli et al. 2011).

Oocyte quality has been observed in many studies to decrease with age, prompting the

acceptance of what is known as the “maternal age effect” (Schwarzer et al. 2014; Rowsey et al.

2013; Gaulden 1992; Hassold & Chiu 1985). Although it is widely accepted that maternal age is

associated with a decline in oocyte quality, it is undecided as to what exactly contributes to this

phenomenon in oocytes (Rowsey et al. 2013; Hultén et al. 2010). This chapter will discuss how

mitochondrial dysfunction manifests in the aging oocyte and how the resulting abnormalities

affect birth outcomes, drawing from results and statistics generated from various published

literature. It will also examine a few of the current theories on the cause (or causes) of the

maternal age effect, and discuss briefly the rapid decline in oocyte quality observed in

pathologies separate from maternal aging alone.

Spindle assembly

Perhaps the most important component of cell division is the spindle. Made up of

microtubules composed of and tubulin dimers, the spindle is highly ATP-dependent and is

responsible for the segregation of chromosomes during cell division. Unlike the spindle

apparatus of somatic cells, oocytes lack centrosomes (reviewed in Dumont & Desai, 2012).

Instead of microtubules nucleating from centrioles within centrosomes, oocytes contain

microtubule-organizing centers (MTOCs) adjacent to chromosomes within the cell and create an

“inside-out” formation of the spindle poles (reviewed in Chapter 1).

Studies have focused on the mechanics of the spindle apparatus itself and how its

dynamics can be altered by maternal aging (Battaglia et al. 1996; Vogt et al. 2008; Howe &

FitzHarris 2013). In one study, Battaglia et al. (1996) examined metaphase II oocytes from two

populations of women: 40 to 45 years old (aged) and 20 to 25 years old (young). Using high-

resolution confocal microscopy to visualize fluorescently labeled chromatin and -tubulin, it was

discovered that oocytes from aged women displayed increased spindle disruptions and abnormal

chromosome segregation compared to oocytes from younger women. Conditions consistent with

aneuploidy were observed in 79% of aged oocytes, but only 17% of young oocytes displayed the

same phenotypes (Battaglia et al. 1996). These results indicate that the increase of spindle

abnormalities with age likely contributes to observed increases in aneuploidy, and it is suggested

that mitochondrial dysfunction contributes these disruptions in spindle function (Battaglia et al.

1996; Howe & FitzHarris 2013; Vogt et al. 2008).

Spindle assembly checkpoint

In most dividing cells, certain checkpoints are in place to ensure that aneuploidy does not

occur. One such checkpoint is the spindle assembly checkpoint (SAC) that prevents cells from

entering anaphase without proper spindle-chromosomal attachment (Musacchio & Salmon 2007;

Musacchio 2011). This signal must be silenced in order for anaphase to begin. In mitosis,

proteins of the mitotic checkpoint complex (MCC) are produced to bind to and block activation

of the anaphase promoting factor/cyclosome (APC/C) (Musacchio & Salmon 2007; Howe &

FitzHarris 2013). When correct spindle attachments are detected, the cell no longer produces

MCC, leaving the APC/C available to activate anaphase (Musacchio & Salmon 2007).

Meiosis in male gametes uses a similar mechanism to prevent aneuploidy in developing

sperm. When the correct spindle attachments are not present, the cell cycle is arrested and cell

death occurs (Musacchio 2011). However, this checkpoint in female gametes appears to be

much less stringent, allowing for aneuploidy to occur without delay of the cell cycle (LeMaire-

Adkins et al. 1997). This may be the reason why the occurrence of aneuploidy is much greater in

oogenesis than spermatogenesis (Pacchierotti et al. 2007). It has been shown that anaphase can

still proceed in oocytes where one or more chromosomes is incorrectly attached to the spindle

(Kouznetsova et al. 2007; Nagaoka et al. 2011), leaving the SAC extremely prone to errors

during oogenesis and further development (Gui & Homer 2012; Kolano et al. 2012). Therefore,

it is no surprise that as mitochondrial function declines with advanced maternal age, oocytes

lacking proper spindle assembly and/or the energy to make proper microtubule-chromosome

attachments have a greater chance of bypassing cell cycle checkpoints than other cell types.

Premature separation of sister chromatids and recombination failure

Aneuploidy can occur as the result of a variety of events, such as recombination failure,

lack of segregation of homologous chromosomes or sister chromatids (nondisjunction), and

premature separation of sister chromatids (PSSC) (FIGURE 2) (reviewed in So I Nagaoka et al.,

2012). Based on data from incidences of aneuploidy in spontaneous abortions, stillbirths, and

live births, it has been estimated that approximately 5% of all fertilized oocytes are aneuploid

(Hassold & Hunt 2001), although not all survive until birth. Oocytes must go through two

rounds of meiosis from the primordial germ cell stage to fertilization, which involves two cell

divisions without DNA replication (as seen in mitosis). As a result, two polar bodies are formed

– one during the first meiotic division and the other during the second (FIGURE 3). The correct

number of chromosomes must be segregated into each so that the oocyte contains the appropriate

amount of genetic material when combined with a sperm cell.

When chromosomes or chromatid pairs do not separate at the appropriate time during cell

division, it can cause too many or too few chromosomes within the developing embryo, leading

to aneuploidy. The extent of aneuploidy depends on the time and location at which the error

takes place. Errors in meiosis I can be caused by recombination failure, premature separation of

sister chromatids, and nondisjunction, whereas nondisjunction and PSSC cause errors in meiosis

II (FIGURE 2); however, trisomies are more commonly caused by errors in meiosis I (Gaulden

1992).

Many aneuploidies are caused by the failure of homologous chromosomes to recombine

or create successful “crossovers” between chromosomes during the early stages of meiosis in the

fetal ovary (Hassold & Hunt 2001). In order for meiotic recombination to be successful,

chromosome pairs must contain crossovers that allow for the exchange of genetic information

between the two chromosomes (Smith & Nicolas 1998). Joined pairs of homologous

chromosomes, known as bivalents, that lack crossovers are expected to result in aneuploidy 50%

of the time (Cheng et al. 2009). Studies have shown that more than 10% of all oocytes possess

one of these “crossover-less” bivalents, yet almost all bivalents in males have at least one

crossover joining the two homologous chromosomes (Cheng et al. 2009; Lynn et al. 2002).

Spontaneous abortion and miscarriage

Developmental arrest, failure to implant into the uterine wall, and spontaneous abortions

can all be induced by aneuploidy (Fragouli & Wells 2011). Each of these lead to miscarriage,

the incidence of which typically increases with advanced maternal age (Nybo et al. 2000). Most

spontaneous abortions (characterized by fetal death occurring between 6 and 20 weeks gestation)

are caused by random segregation errors during meiosis, and the frequency of these errors

increases as a woman ages (Robinson et al. 2001). The chromosomal abnormality attributed to a

woman’s first miscarriage does not influence the chance of the woman having an additional

spontaneous abortion with the same abnormality (Robinson et al. 2001). Therefore, the

likelihood of a woman having a certain abnormality is not dependent on which abnormalities

caused her previous spontaneous abortions, indicating that the segregation errors observed are

indeed random. Based on these results, scientists have concluded that maternal age is the main

factor involved in predicting which cohorts of women will experience more of these random

segregation errors that lead to spontaneous abortion (Robinson et al. 2001).

Additional studies have discovered that a woman in her early 20’s has approximately a

9% risk for miscarriage, and this risk increases to over 75% for a woman over 45 years of age

(Nybo et al. 2000). It is not surprising, therefore, that 2-3% of embryos from women in their

20’s carry trisomies, while 35% of embryos from women in their 40’s are trisomic (Hassold &

Chiu 1985). Embryos with trisomies in some chromosomal pairs are able to survive, but overall,

trisomic embryos are 3 times less likely to develop to the blastocyst stage than euploid embryos

(Sher et al. 2007). A study of clinically diagnosed miscarriages of women older and younger

than 35 years of age found that chromosomal abnormalities were present in 50% of the

miscarriages of younger women and 75% of the miscarriages of older women (Nybo Andersen et

al. 2000). These data show that while increased chromosomal abnormalities play a role in the

risk for aneuploidy, additional factors may also contribute to miscarriage. It has been suggested

that chromosomal abnormalities may make certain oocytes more susceptible to future insults by

additional environmental factors that can cause mtDNA damage or loss of regulatory hormones

(Hassold & Sherman 2000).

Trisomy 21 risks

While aneuploidy can occur in any of the 23 pairs of chromosomes, many aneuploid

embryos are arrested during development and do not survive to birth (Hassold & Hunt 2001).

Live births have been reported in cases of certain trisomies, such as 21 and sex-chromosomal

trisomies, although the risk for each depends on the chromosome (Risch et al. 1986). For

example, different trisomies display different correlations with maternal age, although the overall

trend for trisomies shows an increased risk with advanced age (Kim et al. 2013). The origin of

the trisomy also depends on the individual chromosome, with some deriving from meiosis I, and

others from meiosis II (reviewed in Nagaoka et al. 2012). Trisomies that occur in meiosis I are

typically a result of nondisjunction of the homologous chromosomes, recombination failure, or

premature separation of sister chromatids, whereas trisomies in meiosis II are thought to be

caused by nondisjunction of the sister chromatids (Hassold & Hunt 2001).

Trisomy 21, typically known as Down syndrome, is one of the more commonly observed

trisomies since it can arise from 3 different recombination error scenarios (reviewed in Nagaoka

et al. 2012) (FIGURE 4) (versus one scenario in other trisomies), and more than 5,000 children

born each year in the United States are affected (Canfield et al. 2006). Like many other

trisomies, the risk of Down syndrome increases with advanced maternal age (Kim et al. 2013).

While the risk of a woman aged 20 to 24 years giving birth to a child with Down syndrome is

approximately 1 in 1400, this risk increases exponentially with age. At 35 years old risk

increases to 1 in 350, at 40 years old risk increases to 1 in 60, at 45 years old risk increases to 1

in 25, and at 49 years old, this risk becomes 1 in 11 (Hook 1981; Yoon et al. 1996).

As mtDNA is inherited maternally from mother to daughter, it is easily understood how

this risk for trisomy 21 also appears to increase with an advanced grand-maternal age at the time

of the mother’s birth (Aagesen et al. 1984; Malini & Ramachandra 2006). Studies of this

increased risk found that women had a 30% increased risk for giving birth to a child with Down

syndrome for every year their own mother’s age exceeded 30 years old at the time of their birth

(Aagesen et al. 1984; Malini & Ramachandra 2006). These studies suggest that while it is

possible for women of advanced maternal age to give birth to healthy children, it may increase

their daughters’ chances of having oocytes with increased levels of chromosomal abnormalities.

Limitations to diagnosis and research

Although there are a vast number of studies on aneuploidy and chromosomal

abnormalities, results must be approached with a certain degree of caution. As many studies on

oocyte quality in humans are performed on individuals seeking fertility treatments, it is unclear

whether these women are merely more likely to harbor oocyte abnormalities and other reasons

for reduced infertility, or if their results do in fact represent human females as a whole. Factors

causing chromosomal abnormalities or increased risk of aneuploidy with age may simply be

restricted to sub fertile or infertile advanced-age women, or this group may be more susceptible

to influence by hormonal or regulatory factors later in life. Interestingly enough, it has also been

suggested that predictions or risks associated with age may be more closely connected to

biological age rather than chronological age (Kline et al. 2000). In a study of post-menopausal

women who had previously experienced spontaneous abortions in their lifetimes, it was

discovered that women with karyotyped trisomic aborted embryos typically entered menopause 1

year earlier than those women with chromosomally normal aborted embryos (Kline et al. 2000).

Either way, women and couples seeking assisted reproductive technologies do not necessarily

represent the general population in many studies. One exception to this commonality is the study

of spindle abnormalities by Battaglia et al. (1996). This study used oocytes from naturally

cycling women with no history of infertility or poor health; therefore, researchers were able to

remove any bias that may be introduced from cohorts of women seeking infertility treatments

and still observed dramatic differences with age (Battaglia et al. 1996).

An additional limitation to trisomy research is that many aneuploidy studies are

performed retrospectively, meaning that the trisomic child has already been born. This limits

scientists’ access to only a certain type of available subjects, and not every case of trisomy that

may have caused implantation failure or spontaneous abortion. Rather than examine aneuploidy

retrospectively, more information about the chromosomal abnormality is often gained when

oocytes are examined before implantation. This is often accomplished either by taking a biopsy

of a single blastomere of the early embryo, or analyzing the polar bodies produced during

meiosis (reviewed in Nagaoka et al. 2012). Although this is typically only performed as a

technique in assisted reproduction therapies when choosing embryos for transfer, it is perhaps the

most reliable method to date for determining the genetic profile of an embryo.

Despite caveats involved in this type of research, it must be noted that there are in fact

observable differences between oocytes from young and aged women, even if these women are

from a specific subset of the entire population. Though analysis techniques and restricted

variability in sample size may exaggerate the presence of certain aneuploidies, it is clear that

intrinsic differences are present before scientific intervention for such disparities to occur

between experimentals and controls.

Long meiotic arrest results in mtDNA damage

These differences between young and aged oocytes have often been attributed to what is

known as the maternal age effect. Trisomies, miscarriage, implantation failure, and

developmental arrest have all been observed to increase with maternal age (Vialard et al. 2007;

Gaulden 1992; Allen et al. 2009; Hassold & Chiu 1985; Kim et al. 2013; Robinson et al. 2001).

Some studies point to a single cause, while others adopt a multi-hit mechanism (reviewed in

Rowsey et al. 2013). While it is unclear what exactly causes this maternal age effect, it is

generally speculated that it is not the result of a single factor, but may be the work of several in

combination (reviewed in Rowsey et al. 2013). Many theories point to possible mechanisms by

which the maternal age effect acts to decrease oocyte quality, each of which most likely

contribute to the overall effect. One of these theories focuses on the amount of time oocytes

spend arrested in prophase during adult life (Wang & Höög 2006). Oocytes enter meiosis I

during fetal development and arrest in late prophase until the LH surge during ovulation

stimulates the continuation and completion of meiosis I. Because some women delay childbirth

until later in reproductive life, oocytes generated during fetal development may be exposed to

ROS that cause mtDNA damage for decades before ovulation (Wang & Höög 2006). Although

oocyte mitochondria are relatively inactive compared to somatic mitochondria, low levels of

ATP and ROS are still produced, and mtDNA damage can accumulate over time (Amicarelli et

al. 2003; Wang et al. 2013; Dai et al. 2014). Mitochondrial dysfunction in oocytes is often

associated with an increased incidence of longer time to fertilization, implantation failure, and

developmental arrest (Van Blerkom et al. 1995; Van Blerkom et al. 2000; Keefe et al. 1995). It

may be possible that with age, oocytes that have been exposed to more mtDNA damage may be

more likely to become aneuploid when meiosis resumes.

Cohesion loss

Another promising theory for the maternal age effect is the loss of certain proteins over

time that contributes to proper chromosomal segregation during meiosis. For instance, the

cohesion complex is made up of specific proteins that hold the chromatids together during

prophase (FIGURE 5). Cohesion proteins are responsible for securing the homologous

chromosomes as well as each sister chromosome to its pair to keep them together until the cell is

ready to segregate each into the dividing cell (reviewed in Hunt & Hassold 2010). In

homologous chromosomes, the chromatin complex is present around the arms of each to keep

the pairs together; in sister chromatids it is found around the kinetochores (Nasmyth 2011).

When the cell is signaled to enter into anaphase and segregate the chromosomes, cohesion is

released from the appropriate region of the pair (either the arms or the kinetochores), allowing

the chromosomes to segregate to their designated cellular poles. If cohesion is prematurely lost

or degraded, the chromosome pairs fail to join tightly enough to keep them together during

various cell cycle stages. This can cause PSSC, which has been shown to play a major role in

the occurrence of aneuploidy (Angell 1991).

In both Drosophila melanogaster and mice, loss of cohesion proteins results in increased

rates of aneuploidy (Jeffreys et al. 2003; Hodges et al. 2005) and this phenomenon has also been

observed in humans (Tsutsumi et al. 2014). In a recent study by Tsutsumi et al, fluorescence

microscopy was used to identify levels of two cohesion complex subunits specific to meiosis in

human oocytes, REC8 and SMC1 (2014). The study examined oocytes arrested in prophase I

from samples of ovarian tissue of women aged 19 years to 49 years divided into “29 and below”

and “40 and above” cohorts. Comparisons showed an overall decline with increased age in the

average intensity levels compared to the internal control (Tsutsumi et al. 2014). Because certain

cohesion proteins are produced during the S-phase of the cell cycle in the fetal ovary, it is not

surprising that with increased maternal age, levels of these proteins within oocytes generated

during fetal development decrease (Tsutsumi et al. 2014). In a related study, mice with mutated

Smc1b demonstrated increased levels of aneuploidy (Hodges et al. 2005), suggesting that

aneuploidy is caused by the loss of these cohesion proteins. To further support these findings,

naturally-aged mice show an increased distance between sister chromatid kinetochores when

compared to younger mice at metaphase, indicating a natural loss of cohesion proteins with age

(Chiang et al. 2010). Together, these studies show that the loss of cohesion contributes to the

maternal age effect in oocytes, and that the resulting increase in aneuploid embryos may be a

result of the lost ability for the cell to select against trisomic oocytes.

Carbonyl stress

In addition to increases in reactive oxygen species, the maternal age effect has also been

attributed to an increase in reactive carbonyl species (RCS). These molecules are products of

metabolic glycolysis and have many of the same functions as ROS (Mano 2012). However,

these carbonyls are more stable than ROS, which allows them to travel greater distances in the

cell and to have a more damaging effect. Through glycation of proteins, RCSs cause post-

translational modifications and impair protein function in the cell (Mano 2012). To counteract

damage generated by natural metabolism, cells contain glyoxylases such as GLO1 to inactivate

RCSs and prevent the formation of advanced glycation end-products (AGEs) (Thornalley, 2003;

Dobler, Ahmed, Song, Eboigbodin, & Thornalley, 2006). The most destructive RCS is

methylglyoxal (MG) that is derived from carbohydrate metabolism and can have detrimental

effects on mitochondrial function (Amicarelli et al. 2003; Thornalley 2008). Reactive carbonyl

species can also create a positive feedback loop with ROS production to further cellular damage;

ROS promote the final step of protein glycation by RCS, and AGEs promote pathways that

induce ROS formation (Tatone & Amicarelli 2013). The first study to link RCS and ovarian

dysfunction discovered that women with polycystic ovarian syndrome (PCOS) had higher levels

of AGEs than control women of the same age (Diamanti-Kandarakis et al. 2007). It was also

discovered in mice that the ovaries of older females display increased levels of AGEs and

decreased activity of GLO1, indicating that cells are no longer able to protect against AGE-

induced damage (Tatone et al. 2010). While these studies focused on the ovarian environment

rather than oocytes specifically, the data show that the microenvironment of the oocyte within

the aging ovary may also play a large role in maintaining quality throughout adult life.

Oogonial stem cells

While many of the previously mentioned studies point to accumulation of damage to

arrested oocytes during adult life, there is still the question of how aging affects germline stem

cells and how these contribute to the aging phenotype. Although research in the last decade has

shown that oogonial stem cells (OSCs) exist in mammals (Johnson et al. 2004; Zou et al. 2009;

White et al. 2012), many wonder why menopause occurs if not for the exhaustion of a fixed

follicular pool. It has been suggested that women undergo menopause due to the lack of

necessity to have children past the fifth and sixth decade of life (Cohen 2004), and/or to limit the

time of reproduction so that cellular function can be directed towards maintaining the health of

the mother later in life rather than allocating resources for the production of offspring (Kirkwood

2005). Through transplantation studies (Niikura et al. 2009) it has been demonstrated that the

health of the host ovarian niche is extremely important in predicting the success of oocyte

development and competence. Though ovarian tissue from older mice can successfully produce

immature follicles when transplanted into younger ovaries, the reverse procedure often fails

(Niikura et al. 2009). This indicates that while oogonial stem cells may continually contribute to

the follicular pool during post-fetal life, the aging ovarian environment may reduce the ability of

OSCs to produce viable follicles.

Diabetes and obesity affect oocyte quality

Although many couples and individuals seek fertility assistance due to advanced maternal

age, there are also other pathologies that affect oocyte quality in women of all ages. While

maternal aging is a heavily researched topic in the field of female reproduction, the effects of

maternal obesity and diabetes on oocyte quality have also been the focus of many studies (Wang

& Moley 2010; Diamond et al. 1989; Wang et al. 2009; Moley et al. 1991; Grindler & Moley

2013; Purcell & Moley 2011). Oocytes from aged women and diabetic women often display

similar phenotypes, indicating that technologies and therapies to improve oocyte quality may be

able to benefit both demographics. Mitochondrial structural abnormalities, mitochondrial

aggregation, decreased mitochondrial function, increased incidence of miscarriage, spindle

defects, and implantation failure are all characteristics of both aged and diabetic oocytes

(Simsek-Duran et al. 2013; Komatsu et al. 2014; Wilding et al. 2003; Robinson et al. 2001;

Gaulden 1992; Wang et al. 2009; Colton et al. 2002; Tarín et al. 2001). Metaphase II oocytes

from diabetic mice also show increased levels of aneuploidy (Wang et al. 2009) similar to

increases in aneuploidy with maternal age (Hassold & Hunt 2001; Fragouli & Wells 2011).

In addition to mitochondrial dysfunction, maternal diabetes can disrupt important cell-to-

cell communication between granulosa cells and the oocyte (Ratchford et al. 2008). Diabetic

oocytes transferred into non-diabetic host uterine environments as early as the one-cell stage still

display cellular defects (Diamond et al. 1989; Moley et al. 1991; Wyman et al. 2008), suggesting

that the negative effects attributed to maternal diabetes are able to influence oocyte quality in the

early stages of oogenesis and embryogenesis. At the blastocyst stage, embryos from diabetic

mothers show reduced levels of glucose transport with down-regulated expression of glucose

transport proteins (Moley 1999) which is consistent with increased levels of apoptosis (Moley et

al. 1998).

Obesity is another condition that affects oocyte quality and can become a fertility issue

for women of all ages. Almost one third of all women of reproductive age in the United States

are currently considered to have a body mass index (BMI) in the “obese” range of 30 and above

(Hillemeier et al. 2011; Flegal et al. 2010). While the causes for reduced fertility in obese

women is currently debated (reviewed in Purcell & Moley, 2011), it is clear that oocyte quality is

affected. Obese women are approximately three times less likely to conceive than normal weight

women, even when cycling normally, and display increased risks of miscarriage and birth defects

when conception occurs (Jungheim & Moley 2010; Rich-Edwards et al. 1994). Obese women

are also more likely to develop PCOS, which has further detrimental effects on fertility (Norman

et al. 2007).

Conclusions

Based on studies of maternal aging, diabetes, and obesity, it is clear that decreases in

oocyte quality can arise from a number of factors (Wang & Moley 2010; Purcell & Moley 2011;

Korkmaz et al. 2014). Treatments to improve oocyte quality in women of advanced age may

also contribute to helping women with diabetes or obesity overcome the subfertility or infertility

associated with these conditions. While the maternal age effect on oocyte quality is quite

established, it is still debated as to which factors play the largest role in determining oocyte

competence in advanced maternal age. It is possible that abnormalities during early oocyte

development make certain women more susceptible to subfertility or infertility during adult life,

or perhaps the maternal age effect is due to a breakdown of certain quality control mechanisms

present in younger ovaries (reviewed in Rowsey et al. 2013; reviewed in Nagaoka et al. 2012).

The deterioration of the ovarian microenvironment may also contribute to poor oocyte quality in

older women. Until these details are better understood, assisted reproductive treatments cannot

be completely optimized, and though risk factors may vary between individuals, this area of

study could certainly benefit from additional clarification on female human reproduction as a

whole.

CHAPTER 5: Strategies for improving oocyte quality

As more and more women delay childbirth until later in life, the need for successful

assisted reproductive technologies becomes more apparent. Birth rates in women in their 30’s

and early 40’s in 2012 increased from rates in 2011, while rates for every other age group

dropped, showing an increase in the number of women waiting to have children (Hamilton et al.

2013). In fact, approximately 20% of women now give birth to their first child after the age of

35, although one-third of these women experience some form of subfertility problem (CDC

2013). According to the Center for Disease Control 2011 ART Clinical Report, approximately

11-12% of all reproductive-age women in the United States have difficulty becoming pregnant or

maintaining pregnancy naturally and have sought out some sort of fertility treatment or testing in

their lifetimes (CDC 2013). While the number of women receiving fertility assistance is

growing, pregnancy success still often depends on the age of the female. Women less than 35

years of age using fertility treatments with their own oocytes typically experience a success rate

of 40%, but this number quickly declines to 22% at age 40 and 1% at age 44 (CDC 2013).

Women of advanced age seeking infertility assistance face increased risks of miscarriage,

aneuploidy, and implantation failure compared to younger women (te Velde & Pearson 2002).

Although many current fertility treatments have been successful, none have been able to reach

100% rates for all women. Because reasons for and degree of infertility can vary between

women, researchers and clinicians must be able to provide individual resources depending on

what is needed to assist reproduction (te Velde & Pearson 2002). Until research can advance

enough to produce individual treatments, current therapies must be improved to increase oocyte

quality and embryonic development in both women of advanced age and women with premature

ovarian failure. This chapter will discuss the methods used in typical assisted reproductive

treatments and highlight recent progress in the field to suggest possible future directions.

How is infertility/subfertility treated?

Assisted reproductive technologies typically first begin with hormone treatments to

promote follicle growth and ovulation. It is debated over whether or not these exogenous

hormones have adverse effects on oocyte quality, as many studies have observed increased in

aneuploidy and epigenetic changes in oocytes exposed to high doses (Doherty et al. 2000;

Khosla et al. 2001; Rivera et al. 2008; Market-Velker et al. 2010; Munne et al. 1997). However,

more recent methods have begun to see improvements in embryo quality with lower doses of

hormones, suggesting that the regulation and maintenance of the oocyte environment is highly

sensitive to change (Baart et al. 2007; Rubio et al. 2010). Once oocytes are collected, they are

fertilized by either in vitro fertilization (IVF) or intracytoplasmic sperm injection (ICSI)

(FIGURE 6). The first procedure involves mixing oocytes with sperm cells in a dish, while the

latter injects sperm directly into an oocyte (reviewed in Nagaoka et al. 2012). After fertilization,

the zygotes are then cultured until further use.

Preimplantation genetic diagnosis

After early stage embryos are collected, they are often screened for aneuploidy (Munné et

al. 2010). This ensures that the best blastocysts with the greatest degree of potential success are

transplanted into the mother. One of the polar bodies or blastomeres from the early blastocyst is

used for genomic analysis before implantation. For many years, fluorescence in situ

hybridization (FISH) has been used as the primary means for karyotyping trisomies within a

given blastocyst (Hultén et al. 2010). However, this method has many limitations, the first of

which being that many FISH analyses can only monitor a few chromosomes at a given time

(reviewed in Rowsey et al. 2013). While this is helpful when determining rates of specific

trisomies, additional aneuploidies in other chromosomes within the same cell that can have

negative effects may go unnoticed. Therefore, the use of FISH for genetic analysis has been

highly debated; many believe using FISH for preimplantation genetic diagnosis provides no

benefit (Genetics 2009), while other scientists report improvements in pregnancy outcomes

(Munné et al. 2010). These opposing results may depend on whether the aneuploidy could be

detected from the limited number of probes chosen since all 23 chromosomes cannot be

monitored with this method.

A more reliable method is comparative genomic hybridization (CGH) (reviewed in

Nagaoka, Hassold, & Hunt, 2012), which can detect aneuploidy in all 23 chromosomes of a

single cell. This method involves the amplification of experimental DNA, which is then

compared to normal reference DNA using various fluorochromes to identify specific areas on

each chromosome. The ratio of each is calculated and compared to determine any genetic

defects present in the experimental DNA and examines all 23 pairs of chromosomes rather than a

small subset in FISH protocols. Pairing this method with a microchip array (aCGH) containing

each fluorochrome probe decreases the length of time it takes to receive results, which can be

critical in the timing of in vitro fertilizations. While these methods of preimplantation genetic

screening can be beneficial for some, they typically do not show any improvements in pregnancy

success in women younger than 40 years old. However, for women over 40, preimplantation

genetic screening can improve outcomes by twice as much (Milán et al. 2010). This information

reinforces the idea that fertility treatments must be tailored to individual women and not every

treatment is ideal for the entire population.

Once the embryos with the highest chance of success have been chosen, they are either

transferred into the mother or cryo-frozen for later use. While freezing embryos may appear to

be a tempting procedure for young women to take preventative measures against decreased

fertility later in life, it is actually more cost-efficient to forgo these procedures and deal with

subfertility as it arises. A study of healthy women found that the cost of freezing embryos or

ovarian tissue at age 25 was more expensive than using assisted reproductive treatments at age

40 (Hirshfeld-Cytron et al. 2012). In addition, some women did not experience difficulties with

fertility at age 40 at all. These data further reinforce the idea that chronological age may not

directly predict biological age, and that there is much variability between individuals of the same

population.

How can oocyte quality be improved?

Ooplasmic transfer

While improving methods used in assisted reproductive treatments is extremely

important, it cannot be forgotten that a major underlying factor is oocyte quality. If oocyte

quality can be improved, it would reduce the need for improved ART protocols and may also

decrease the number of women reliant on them. When scientists discovered that older patients

could carry successful pregnancies if the oocyte was from a younger donor, it was determined

that the oocyte is the major driving factor in pregnancy success (Sauer et al. 1995). This led

scientists to believe that some component within young oocytes was lacking in older oocytes. To

improve older oocytes, cytoplasm from young oocytes was injected during IVF procedures in

attempts to improve oocyte quality (FIGURE 7). Successful live births were derived from these

experiments (Cohen et al. 1998), but the health of the resulting offspring was questioned when it

was discovered that they had mitochondrial heteroplasmy (Barritt et al. 2001; Brenner et al.

2000). Because mitochondria are inherited maternally, these children contained mitochondria

from both their mother and the woman who donated the young cytoplasm. The US Food and

Drug Administration put a moratorium on “ooplasmic transfer” in 2001 due to the suspicion that

this procedure involved genetic manipulation that had unknown results (Frankel & Chapman

2001; Parens & Juengst 2001). Mouse studies performed after this procedure was performed in

humans found that mitochondrial heteroplasmy caused reduced mitochondrial function in adult

offspring (Acton et al. 2007). While these procedures are no longer performed, they made it

clear that the mitochondria of the young oocyte were the factor that improved quality of older

oocytes, and focus began to shift towards improving mitochondria to maintain fertility later in

life.

AUGMENT – autologous germline mitochondrial energy transfer

To improve mitochondrial quality in the aged oocyte while circumventing mitochondrial

heteroplasmy, an autologous source of healthy mitochondria was needed. This became possible

with the discovery and isolation of oogonial stem cells within female mammals, including

humans (Johnson et al. 2004; White et al. 2012; Zou et al. 2009). OSCs allowed for the

development of autologous germline mitochondrial energy transfer (AUGMENT), which

involves the transfer of oogonial stem cell mitochondria to the oocyte of the same patient to

rejuvenate the energy supply for the dividing embryo (FIGURE 8). Not only does AUGMENT

provide an autologous supply of mitochondria to older oocytes, but it also supplies mitochondria

from the same developmental cell line. Since mitochondria from different tissues differ in

function and protein expression (Johnson et al. 2007; Riva et al. 2005), it is important to use

mitochondria that will respond to the correct developmental cues present within the oocyte

microenvironment. This is one reason why, for instance, mitochondria from other somatic

tissues are inefficient for AUGMENT (Takeda et al. 2005). Another reason why autologous

somatic cell mitochondria should not be transferred into oocytes is because of accumulated

mtDNA damage (Barja 2002; Copeland & Longley 2014; Corral-Debrinski et al. 1992). Just as

mtDNA damage accumulates within oocyte mitochondria, it also affects somatic mtDNA with

age. Therefore, oogonial stem cell mitochondria would provide the “purest” and most

developmentally competent supply of mitochondria to poor quality oocytes with dysfunctional

mitochondria. One caveat to this method is that the oogonial stem cells must be harvested from

the patient and the mitochondria must be carefully isolated for the transfer of mitochondria to

occur. Because these cells are found in the ovarian tissue, the process may be invasive and

difficult to perform. Therefore, many treatments have turned to improving existing mitochondria

rather than transferring healthy organelles into oocytes.

Antioxidants

Because ROS are considered the main culprit of mtDNA damage with age, many studies

have focused on the use of antioxidants as preventative treatments for maternal age effects on

oocyte quality. Reactive oxygen species are a natural byproduct of cellular metabolism and are

most often produced at Complex I and Complex III in the electron transport chain of

mitochondria (Lenaz 2012; Dröse & Brandt 2012). They play a role in many cell-signaling

pathways such as the MAPK pathway and the PI3K pathway, and are also involved in

maintaining the immune response (Devadas et al. 2002; Genestra 2007; Tobiume et al. 2001; S.-

R. Lee et al. 2002; Kwon et al. 2004; Seo et al. 2005). In order to keep high levels of ROS at

bay, cells produce antioxidants that counter ROS production and damage to DNA and proteins

(Valko et al. 2006; Chatterjee et al. 2007). These antioxidants include enzymatic and non-

enzymatic varieties, the latter of which is divided into metabolic and nutrient types. The main

enzymatic antioxidant is superoxide dismutase (SOD) that converts superoxide radicals into

H2O2, which is then converted into H2O and O2 by additional enzymatic antioxidants (Halliwell

2007). Non-enzymatic metabolic antioxidants include endogenous CoQ10, while exogenous

antioxidants are derived from nutrients such as vitamin E, C, and omega-3 fatty acids (Willcox et

al. 2004).

Although antioxidants appear to be a clear choice for maternal aging treatments, many

studies focused on increasing endogenous and exogenous antioxidants conflict as to whether or

not antioxidant treatments improve cellular longevity (reviewed in Dai, Chiao, Marcinek, Szeto,

& Rabinovitch, 2014). One study demonstrated that the overexpression of SOD in transgenic

mice surprisingly did not increase lifespan compared to wild type controls (Pérez et al. 2009).

While some studies show benefits to antioxidants in the treatment of certain pathologies, others

do not, implying that antioxidants may play different roles in the prevention or repair of different

disease progressions (Myung et al. 2013; Li et al. 2012).

In regards to female reproduction, antioxidants do not appear to have practical

applications in improving oocyte quality with increased maternal age. Two experiments in

which the diets of female mice were supplemented with the same dosages of vitamins C and E

showed varying benefits to antioxidant treatments (Juan J Tarín et al. 2002; J J Tarín et al. 2002).

While one study discovered that the oocytes of antioxidant-treated aged mice showed normal

chromosomal distribution in comparison to younger mice (Juan J Tarín et al. 2002), the other

demonstrated an extreme reduction in litter size and the survival of offspring (J J Tarín et al.

2002). These results show that while antioxidants may improve oocyte quality in aged females

at first glance, they may have a detrimental effect on the uterine environment and/or embryonic

development in the womb.

Coenzyme Q10

Coenzyme Q10 (CoQ10) is considered a non-enzymatic metabolic antioxidant that is

naturally found in the electron transport chain of mitochondria. It functions to transport

electrons from Complexes I and II to Complex III, and also transfers protons across the inner

mitochondria membrane to increase ATP production (Rosenfeldt et al. 2003; Pignatti et al. 1980;

Sandhir et al. 2014). As humans age, levels of CoQ10 decrease (Pignatti et al. 1980), so many

treatments to counteract mitochondrial dysfunction with age have focused on CoQ10 dietary

supplements. CoQ10 has shown improvements in many studies related to various mitochondrial

disorders such as hypertension, Parkinson’s disease, and macular degeneration, and has also been

effective in increasing mitochondrial activity in aged oocytes from mice (Bentov et al. 2010;

Feher et al. 2005; Winkler-Stuck et al. 2004; Rosenfeldt et al. 2003). CoQ10 supplements have

also shown increases in oocyte quality and menopausal cognitive effects in mice by affecting

mitochondrial dysfunction and ROS production (Gendelman & Roth 2012; Sandhir et al. 2014).

While these studies have yet to be performed on humans, preliminary work in bovine and mouse

models has produced a promising outlook for CoQ10 treatments.

Resveratrol

The polyphenol resveratrol has also been a focus of increasing mitochondrial function

and oocyte quality by acting as a potent antioxidant. Resveratrol is an activator of SIRT1, which

has a key role in many mitochondrial functions including mitophagy and biogenesis to reduce

oxidative stress levels (Gustafsson & Gottlieb 2009; Kang & Hwang 2009; Cantó et al. 2012).

The inhibition of SIRT1 has been shown to cause decreased fertility in mice (Takeo et al. 2013).

Although grapes, soybeans, and peanuts are all excellent sources of resveratrol, it is most

commonly known as the antioxidant found in red wine (Burns et al. 2002). While high levels of

alcohol consumption may counteract the benefits of resveratrol in red wine (Katsiki et al. 2014),

data generated from studies using the isolated polyphenol in improving oocyte quality have

shown promising results. One study examined oocytes cultured in vitro with resveratrol and

monitored SIRT1 protein expression, mitochondrial function, and fertility rates during IVF

(Takeo et al. 2014). Compared to controls, resveratrol increased SIRT1 expression, ATP

content, mtDNA copy number, and mitochondrial membrane potential in in vitro matured

oocytes, and also increased fertilization rates during IVF treatments (Takeo et al. 2014). These

data not only show the antioxidant capacity of resveratrol to improve mitochondrial health, but

also the ability to improve fertilization and quality of oocytes.

Recent resveratrol research also points to the possibility of providing improvements in

mitochondrial function with age to increase the quality of oocytes. A study in mice compared

oocyte quality and quantity in mice supplemented with resveratrol for 6-12 months to age-

matched controls as well as young mice (Liu et al. 2013). Analysis after the 12-month period

showed an increase in the follicular pool and overall number of oocytes in mice treated with

resveratrol compared to aged mice without resveratrol. Oocyte quality, based on the alignment

of chromosomes on the metaphase plate and proper spindle formation, was also improved

compared to controls (Liu et al. 2013). This resulted in great differences in reproductive

potential, as mice treated with resveratrol were able to maintain successful reproduction while

age-matched controls produced no litters (Liu et al. 2013). These studies of resveratrol show that

this polyphenol improves and maintains oocyte quality both in vitro and in vivo, respectively,

and may be useful as a possible treatment for women experiencing sub-fertility issues. However,

it is still unclear whether resveratrol must be supplemented throughout a female’s lifetime or if

intervention during adulthood is sufficient to reverse aging phenotypes.

Caloric restriction

Restricting caloric intake to improve female fertility is not a new concept (Ball et al.

1947; Visscher et al. 1952; Nelson et al. 1985; Lintern-Moore & Everitt 1978; Masoro 2005).

While continued dietary restriction reduces fertility, it can return once caloric intake is increased

back to normal levels (Visscher et al. 1952; Selesniemi et al. 2008; Ball et al. 1947). Many

studies have found that reducing calories upon weaning and later restoring an ad-libitum (AL)

regiment can improve litter size and oocyte numbers in aged mice compared to control mice fed

an AL diet throughout the lifespan (Ball et al. 1947; Visscher et al. 1952; Nelson et al. 1985;

Lintern-Moore & Everitt 1978; Merry & Holehan 1979). Although these studies conclude that

caloric restriction (CR) postpones the maternal age effect on fertility, it is unclear whether or not

this is due to the stunted growth caused by CR initiated at weaning (Merry & Holehan 1979). To

test this idea, Selesniemi et al. designed an experiment that introduced CR after the onset of

sexual maturity in mice and reinstated an AL-fed diet at 15.5 months of age (2008). Once

dietary restrictions were removed, aged mice previously on the CR diet showed marked increases

in fertility rates, judged by the number of litters and survival rates of pups compared to age-

matched controls (Selesniemi et al. 2008). These results were similar to those obtained from

younger mice in their reproductive prime (Selesniemi et al. 2008). This shows that caloric

restriction still provides benefits to female fertility when induced later in life, and reproductive

preservation is not solely due to the delay in puberty exhibited by initiating CR at weaning.

In addition to this study, Selesniemi et al. later analyzed oocyte quality a step further

(2011). Previous research has shown that CR affects oocyte quality through metabolic pathways

and oxidative phosphorylation, closely linked to mitochondrial function (Sohal et al. 1994; Barja

2002). Because mitochondrial function is closely linked to decreased oocyte quality with age,

Selesniemi et al. examined chromosomal integrity, spindle formation, and mitochondrial

aggregation in metaphase II oocytes of adult-onset CR-AL mice (2011). Compared to 12 month

old AL-fed control mice, oocytes from 12 month old mice maintained on CR for 11 months and

then AL-feeding for 1 month displayed spindle assemblies and rates of aneuploidy similar to 3

month old controls (Selesniemi et al. 2011). Although mitochondrial aggregation was increased

in oocytes of 12-month CR-AL mice compared to 3-month controls, disruptions were not as

severe as those seen in 12 month AL controls. Chromosomal misalignment increased in 3 month

and 12-month control oocytes from 10% to 65%, respectively, but less than 25% of oocytes

obtained from 12 month CR-AL showed the same abnormalities (Selesniemi et al. 2011). These

data show that while the specific mechanisms by which CR improves fertility in aged mice are

still unclear, it is likely due to increases in oocyte quality. It also remains unclear whether or not

this method could be applied to humans, because this work suggests that CR would have to be

initiated early on in adult life.

Conclusions

While the topic of declining female fertility with age is still a complex problem to be

solved, recent research has made great strides in understanding how exactly female reproduction

is affected by time. Oocyte quality plays a large role in the determination of future pregnancy

success, yet the extreme variability observed in individual cases of infertility and subfertility

indicate that other factors are at play (te Velde & Pearson 2002). The maternal age effect on

reproduction clearly exists, but it is not yet completely understood which genetic and lifestyle

influences are the cause of these observations with increased maternal age. Mitochondrial

dysfunction has been closely linked with age and is a strong candidate for the cause of decreased

oocyte quality with increased maternal age. As the major sources of ATP, mitochondria function

is important for spindle formation and chromosomal segregation, two of the most important

cellular events during oogenesis and embryogenesis (Battaglia et al. 1996; Kolano et al. 2012;

Schwarzer et al. 2014; Fragouli & Wells 2011; Qi et al. 2014). Without a proper supply of

functioning mitochondria, oocyte quality quickly decreases. Therefore it is important to further

understand how mitochondrial function is affected by increased maternal age and how these

issues may be addressed in the increasing population of older first-time mothers.

Because mitochondria contain their own DNA molecules separate from nuclear DNA, it

would be beneficial to target gene expression and protein translation in aged individuals to

determine the molecular differences between the two phenotypes. Mitochondrial DNA encodes

only 13 proteins, while the remaining are imported from the nucleus (Schatz 1979). Differences

in mitochondrial proteins may provide a clue as to which genes are up-regulated or down-

regulated with age. Perhaps ameliorating missing or accumulated proteins with age with certain

factors or enzymes would increase oocyte quality in older mothers. Researchers may also focus

on how to genetically modify mitochondrial or nuclear DNA within the oocyte to maintain

longevity of quality with age.

The discovery of OSCs has created extensive opportunities for research into a new source

of autologous oocytes perhaps less affected by maternal age than those generated during fetal life

(Johnson et al. 2004; White et al. 2012). Although oogonial stem cells provide a continuous

supply of immature oocytes during adulthood in mammalian females, the follicular pool is still

exhausted with age to cause the onset of menopause. Increased understanding of the

mechanisms behind how aging affects oogonial stem cells will surely shed new light in the field

of female reproduction.

Clearly there is still much to be discovered about female fertility and its dynamics with

age. Women are delaying childbirth further every year for various reasons, yet the biological

clock remains unchanged. While more and more couples seek assisted reproductive treatments,

it is still clear that there is no single treatment that is successful for every patient. Until the

subtleties of female reproduction are uncovered, the ability to preserve fertility until later in life

remains elusive.

Figure 1

Figure 1: An imbalance of mitochondrial biogenesis and mitophagy causes detrimental effects to the cell. This can be caused by an increased activation of one function without an increase in the other, or a decreased activation of one without the decrease of the other. If biogenesis is dominant, increased numbers of mitochondria cause increased ROS production, increased aggregation of excessive mitochondria, and an increased number of dysfunctional mitochondria due to the lack of efficient mitophagy. If mitophagy is dominant, the number of mitochondria will decrease, causing increased stress on the remaining mitochondria and a decreased level of ATP production overall.

Figure 2

Figure 2: Different kinds of chromosomal errors can cause aneuploidy. When chromosomes fail to recombine and form crossovers during prophase I, they do not line up together on the metaphase plate and can be segregated into the same cell. While there is a chance they can successfully separate into different cells, the lack of recombination between chromosomes can lead to increased risks for future aneuploidy. Premature separation of sister chromatids (PSSC) is the most common cause of aneuploidy. This results when one of the sister chromatids separates too early, leaving one homologous chromosome attached to a single sister chromatid. These can then go on to separate into the two cells as a single chromatid and three chromosomes rather than two sister chromatids in each. Nondisjunction describes the phenomenon during which homologous chromosomes or sister chromatids fail to separate at the appropriate times. This leads to both homologous chromosomes segregating into a single cell during meiosis I, or two sister chromatids segregating into a single cell during meiosis II, leaving the second cell with no chromosomes.

Figure 3

Figure 3: Segregation of oocyte chromosomes during meiosis I and meiosis II creates a single oocyte and two polar bodies. Oocytes arrested in prophase of meiosis I contain two homologous chromosome pairs joined by recombination. After ovulation is induced by the LH surge, the oocyte resumes meiosis I and segregates each homologous pair into the oocyte and the first polar body. The oocyte proceeds through meiosis II and becomes arrested in metaphase II. Once, fertilized, the oocyte divides again to release the second polar body, and becomes haploid in preparation for fusion with the sperm cell.

Figure 4

Figure 4: Trisomy 21 is caused by three different types of segregation failure. Lack of crossovers prevents the exchange of genetic material between chromosomes and can lead to aneuploidy. Crossovers too close to the telomeres (distal recombination) or the centromeres (proximal recombination) can also lead to increases in aneuploidy that result in trisomy 21.

Figure 5

Figure 5: Cohesion proteins are necessary for the correct timing of chromosomal segregation. During meiosis I and meiosis II, homologous chromosomes and sister chromatids must be kept together until the exact moment of separation. If they segregate too early or too late, aneuploidy occurs due to PSSC or nondisjunction, respectively. Cohesion proteins form a ring around the ends of sister chromatids and the centromeres of homologous chromosomes. Cohesion surrounding the ends of the sister chromatids is first broken down during anaphase I, and cohesion around the centromeres is lost during anaphase II. This allows for proper segregation during cell division.

Figure 6

Figure 6: Methods of assisted reproductive fertilization. Two methods of fertilization in assisted reproduction are in vitro fertilization (IVF) and intracytoplasmic sperm injection (ICSI). IVF involves the co-culture of oocytes with multiple sperm cells to achieve fertilization. Some couples require the use of ICSI, which involves injecting a single sperm directly into the oocyte.

Figure 7

Figure 7: Ooplasmic transfer causes mitochondrial heteroplasmy. In ooplasmic transfer, a small volume of young donor oocyte cytoplasm is injected into aged or poor quality oocytes during IVF procedures. This method appears to rejuvenate aged oocytes, but live offspring result in mitochondrial heteroplasmy. Because this was considered genetic modification, the FDA suspended the procedure in 2001.

Figure 8

Figure 8: Autologous germline mitochondrial energy transfer (AUGMENT). This procedure was inspired by the discovery of OSCs that supply renewable autologous mitochondria for women with poor oocyte quality. Transfer of mitochondria from these OSCs would supply the oocyte with new mitochondria, free of mtDNA damage that arises with increased maternal age.

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