Abstract
Mitochondria, originally of bacterial origin, are highly dynamic organelles that have evolved a symbiotic relationship within eukaryotic cells. Mitochondria undergo dynamic, stage-specific restructuring and redistribution during oocyte maturation and preimplantation embryo development, necessary to support key developmental events. Mitochondria also fulfil a wide range of functions beyond ATP synthesis, including the production of intracellular reactive oxygen species and calcium regulation, and are active participants in the regulation of signal transduction pathways. Communication between not only mitochondria and the nucleus, but also with other organelles, is emerging as a critical function which regulates preimplantation development. Significantly, perturbations and deficits in mitochondrial function manifest not only as reduced quality and/or poor oocyte and embryo development but contribute to post-implantation failure, long-term cell function and adult disease. A growing body of evidence indicates that altered availability of metabolic co-factors modulate the activity of epigenetic modifiers, such that oocyte and embryo mitochondrial activity and dynamics have the capacity to establish long-lasting alterations to the epigenetic landscape. It is proposed that preimplantation embryo development may represent a sensitive window during which epigenetic regulation by mitochondria is likely to have significant short- and long-term effects on embryo, and offspring, health. Hence, mitochondrial integrity, communication and metabolism are critical links between the environment, the epigenome and the regulation of embryo development.
Introduction
Mitochondria are double membrane-bound organelles that generate ATP through oxidative phosphorylation (OXPHOS), the components of which are partially encoded by their own genome. They represent the most abundant organelle in the oocyte and undergo significant structural and positional changes during preimplantation development (Motta et al. 2000, Sathananthan & Trounson 2000). This dynamic nature is essential for modulating key cellular events, coincident with the changing energy requirements of the embryo (Gardner 1998), yet the significance of mitochondria during preimplantation embryo development extends beyond the generation of ATP. This review integrates recent advances in our understanding of the dynamics of mitochondria, and the interactions between mitochondria and the cytoskeleton, and other organelles, that ensure mitochondrial, and oocyte and embryo, integrity and facilitate signalling. The implications of deficits in mitochondrial dynamics and signalling for embryo viability and offspring health are also explored, with an emphasis on the emerging understanding that modulation of metabolic flux contributes to the regulation of the nuclear epigenetic landscape (Donohoe & Bultman 2012, Harvey et al. 2016), proposing a novel role for mitochondria as mediators of metaboloepigenetic processes that programme the oocyte and preimplantation embryo. Consequently, the utility of mitochondria as a biomarker is limited by deficiencies in our understanding of the physiological and cellular consequences of in vivo and in vitro manipulation of mitochondrial function, and the link between mitochondrial movement and metabolite signalling during preimplantation embryo development that mediate alterations in the epigenome.
Mitochondrial function and cellular homeostasis
Mitochondria are primarily recognised as the powerhouse of cells, synthesising ATP through OXPHOS. The electron transport chain (ETC) comprises five complexes (I–V) that regulate the transfer of nicotinamide adenine dinucleotide (NADH) and succinate through the ETC, and the generation of an electrochemical gradient across the inner mitochondrial membrane to drive ATP synthase (complex V). Respiratory activity is determined by the assembly of respiratory chain complexes into functional supercomplexes within the inner mitochondrial membrane (Cogliati et al. 2013), although single complexes are also believed to exist to enable flexibility and adaptation to environmental changes (Porras & Bai 2015). The environment created within the inner mitochondrial membrane is distinct from the cytosol (Tzagoloff 1982, Herrmann & Riemer 2010), hence the mitochondrion possesses shuttles which regulate specific metabolite levels between the cytoplasm and the mitochondria and contribute to metabolic homeostasis (Herrmann & Riemer 2010, Wang et al. 2014). Mitochondrial metabolism also plays a role in the synthesis of macromolecules, such as fatty acids, amino acids and nucleotides, to support proliferation (reviewed by Ahn & Metallo 2015). These diverse functions underscore the significance of mitochondria as regulators of cellular metabolism, however mitochondrial function extends beyond the traditional confines of metabolic regulation.
Historically, mitochondria have been perceived as signalling effectors, that is, mere bystanders in the response to various stimuli. However, mitochondria are increasingly viewed as active participants in numerous pathways, acting as initiators and transducers of signals through the modulation of metabolite availability and changes in redox state (Chandel 2014). Beyond the production of ATP via OXPHOS, mitochondria participate in numerous cellular functions, including Ca2+ homeostasis (Duchen 2000, Rizzuto et al. 2012), stress responses (Zhao et al. 2002, Jovaisaite et al. 2014), apoptosis (reviewed by Tait & Green 2013) and have been proposed to be involved in the regulation of chromosome segregation (Schon et al. 2000, Qian et al. 2012). A consequence of OXPHOS is the production of reactive oxygen species (ROS), yet in stark contrast to the damaging effects of ROS at high concentrations, physiological levels are necessary to act as signalling molecules to directly modulate numerous processes through the modification of kinases, transcription factor activity, growth factors and metabolic enzymes (reviewed Harvey et al. 2002, by Hamanaka & Chandel 2010, Lees et al. 2017). Consequently, mitochondria are central to cellular and metabolic homeostasis and are key players in diverse physiological (and pathological) processes.
Importantly, mitochondrial function has also been connected to cell specification (Chung et al. 2007), where lineage commitment necessitates functional mitochondrial adaptation to support differentiation. Cellular energy demand varies significantly depending on cell function and activity, therefore modulation of energy production to accommodate changing physiological demands is essential and likely underpins cell identity. Indeed, once considered mere by-products of cellular metabolism, recent evidence supports a role for mitochondrial metabolites as signalling molecules that regulate gene expression, through the regulation of epigenetics, and consequently drive lineage decisions (Shyh-Chang et al. 2013a, Folmes & Terzic 2014, Harvey et al. 2016). Mitochondrial activity is an absolute requirement for stem cell differentiation, as inhibiting mitochondrial activity with antimycin A or CCCP (carbonyl cyanide m-chlorophenyl hydrazine) prevents embryonic stem cell (ESC) differentiation (Mandal et al. 2011, Pereira et al. 2013). Mitochondrial metabolism in particular controls the levels of the key co-factors including acetyl-CoA, α-ketoglutarate (αKG) and NADH/NAD+, as well as other tricarboxylic acid (TCA) intermediates including citrate and succinate, which act as key substrates for epigenetic modifiers (Wallace & Fan 2010). Studies in ESC have begun to highlight the significance of nutrient regulation in establishing the epigenome (reviewed by Harvey et al. 2016), however, our understanding of the consequences of modulating nutrient and metabolic intermediate levels on the epigenetic landscape in the oocyte, and during preimplantation embryo development, remains rudimentary.
Dynamic regulation of the preimplantation epigenome
Preimplantation embryo development is characterised by extensive reprogramming of the epigenetic landscape to support and regulate stage-specific events including embryonic genome activation and lineage specification. Following fertilisation, the paternal genome undergoes a wave of active DNA demethylation (Mayer et al. 2000, Oswald et al. 2000). Subsequently, passive demethylation proceeds with successive cleavage divisions, persisting through to the morula stage (Monk et al. 1987, Santos et al. 2002, Zaitseva et al. 2007), with the exception of imprinted genes and some repetitive elements. Formation of the blastocyst results in the establishment of the first cell lineages, in which the pluripotent inner cell mass (ICM) exhibits higher levels of de novo methylation compared with the trophectoderm (Reik et al. 2003). In addition to DNA methylation, chromatin remodelling and histone modifications also play a critical role in programming the embryonic epigenetic landscape. However, in contrast to DNA methylation, the dynamics of acetylation (ac) and histone methylation (me) during early development is largely limited to studies on the expression of histone (de)acetylases, methyltransferases and demethylases (reviewed Shi & Wu 2009, by Mason et al. 2012). Immunofluorescent analyses have however documented asymmetry in a number of pronuclear modifications (Adenot et al. 1997, Santos et al. 2005, van der Heijden et al. 2005, Boskovic et al. 2012), along with the presence of other marks in both pronuclei (Zhou et al. 2014, Rollo et al. 2017) and stage-specific changes (Ma & Schultz 2008, Boskovic et al. 2012, Liu et al. 2016b). Consequently, normal development relies on coordinated changes in DNA methylation, and histone acetylation and methylation, which underpin the activation and silencing of specific genes (reviewed by Santos & Dean 2004, Lucas 2013, Messerschmidt et al. 2014, Marcho et al. 2015).
Aberrant epigenetic patterns have been correlated with alterations in the microenvironment surrounding the developing embryo, as well as with subsequent developmental failure (Lucas 2013, Vickaryous & Whitelaw 2005, Ma & Schultz 2008)). In vitro embryo culture has been shown to accelerate DNA demethylation of rabbit embryos (Reis e Silva et al. 2012) and significantly lower histone 3 lysine 4 trimethylation (H3K4me3; Wu et al. 2012), while exposure of in vivo derived bovine embryos to in vitro culture conditions induces significant hypermethylation or hypomethylation depending on the timing of exposure, and include alterations in metabolic pathways (Salilew-Wondim et al. 2018). Further, increased acetylation of histone 3 lysine 9 (H3K9ac) has recently been proposed as a biomarker of culture-induced perturbations (Rollo et al. 2017). Importantly, inhibition of histone deacetylase activity with trichostatin A reduces (Adenot et al. 1997) or inhibits (Ito et al. 2000) blastocyst development, and perturbations in the programming of the preimplantation epigenome have been shown to reduce implantation (Ding et al. 2012) and disturb post-implantation decidualisation (Gao et al. 2012). While it remains to be determined whether epigenetic perturbations persist into foetal tissues, combined, these studies reveal that the preimplantation embryonic epigenome is sensitive to alterations in the surrounding nutrient environment, with alterations negatively impacting on subsequent development. Significantly, the activity of epigenetic modifiers that regulate the deposition and removal of DNA and histone marks depends on the availability of specific metabolites. Conceivably, the changing energy requirements, and therefore metabolic pathway activity, of the preimplantation embryo may drive epigenetic programming, such that perturbed metabolic activity transmits alterations in the cellular microenvironment which modify the epigenetic landscape (Gardner & Harvey 2015).
Metabolic regulators of acetylation and methylation
Levels of histone acetylation are modulated by histone acetyltransferases (HATs) and histone deacetylases (HDACs). Acetylation by histone acetyltransferases requires acetyl-CoA, a metabolic intermediate whose availability dictates histone acetylation levels, and consequently gene transcription. High glycolytic rates drive acetyl-CoA synthesis in mitochondria, although acetyl-CoA can also be produced in the cytosol from citrate, fatty acid and ketone body oxidation and the catabolism of amino acids, or within the nucleus (Wellen et al. 2009). Recently, glucose flux through glycolysis has been shown to regulate ESC acetyl-CoA levels, and therefore histone acetylation (Moussaieff et al. 2015). Supplementing human ESC with acetate, a precursor of acetyl-CoA, increases acetylation levels, blocks histone deacetylation and reduces differentiation, while inhibition of acetyl-CoA production from glucose results in the loss of pluripotency (Moussaieff et al. 2015). Alterations in glucose use can therefore significantly modify not only downstream metabolite levels, but also alter histone acetylation, through altered acetyl-CoA levels, and subsequently regulate differentiation. The significance of this is such that incorrect timing of substrate availability in the early embryo may accelerate or delay key events through gene activation and is particularly relevant given the changing energy requirements of the preimplantation embryo (reviewed by Gardner & Harvey 2015) and the significance of glucose use as a marker of human embryo viability (Gardner et al. 2011). As alterations in glucose uptake can significantly alter acetyl-CoA availability, it will be important to understand the impact of altering acetyl-CoA levels during preimplantation development. A role for appropriate modulation of acetyl-CoA levels to support embryo development, potentially via the epigenome, is however plausible given the identification of the embryotrophic factor promoting blastocyst development within bovine serum albumin as the acetyl-CoA precursor citrate (Gray et al. 1992).
In contrast, repression of gene transcription is in part mediated through histone deacetylation. In addition to Class I and II HDACs, which are dependent on zinc as a co-factor for their activity, nicotinamide adenine dinucleotide (NAD+) availability regulates a family of conserved NAD+-dependent histone deacetylases known as sirtuins (SIRTs), which deacetylate targets involved in the modulation of cellular metabolism and mitochondrial biogenesis, cell survival, DNA repair, differentiation and ROS production (Houtkooper et al. 2012). Of note, alterations in sirtuin activity have been shown to alter embryo development. Inhibition of sirtuin activity with nicotinamide or sirtinol significantly reduces porcine (Kwak et al. 2012) and mouse (Kawamura et al. 2010) blastocyst development, conceivably through alterations in acetylation. It will however be important to examine the regulation of sirtuins, and sirtuin regulated changes in the epigeneome, during preimplantation development in response to nutrient availability, particularly given the expanded use of monophasic culture systems in preference to biphasic media. Equally, investigation of the significance of NAD+ in regulating sirtuin activity during preimplantation development is warranted. NAD+ is synthesised de novo from the amino acid tryptophan (Bender 1983) or recycled via the NAD+ salvage pathway from nicotinamide and derivatives of vitamin B3 (Canto et al. 2015). In order to ensure the continuation of glycolysis, NAD+ is also recycled via the conversion of pyruvate to lactate, the production of which increases with blastocyst development (Gardner & Harvey 2015). NADH reducing equivalents are also shuttled across the impermeable inner mitochondrial membrane via the malate aspartate shuttle (MAS) to maintain NAD+:NADH (Lu et al. 2008). Significantly, the activity of the MAS is responsible for oxidative metabolism during embryo development (Lane & Gardner 2005) and inhibition of MAS activity with amino-oxyacetate impairs embryo metabolism and ATP production, accompanied by reduced blastocyst development and placental and foetal growth (Mitchell et al. 2009, Wakefield et al. 2011). Whether these changes are accompanied by alterations in the foetal epigenome remains to be determined. These studies however highlight a role for NAD+ regulation in programming development, and a requirement for coordinated compartmentalisation of metabolites between the cytoplasm and mitochondria to maintain mitochondrial function and ensure normal development.
In a similar manner, the activity of DNA methyltransferases (DNMTs), which catalyse the addition of methyl groups to the 5′ position of cytosine residues within cytosine-guanine dinucleotides (CpG) and result in the formation of 5-methylcytosine (5mC), as well as histone methyltransferases which catalyse the transfer of 1 (me), 2 (me2) or 3 (me3) methyl groups to lysine and arginine residues on histones, rely on metabolic intermediate availability. The amino acid methionine is the precursor of the universal methyl donor S-adenosylmethionine (SAM), catalysed by methionine adenosyltransferases (MATs), that acts as a co-factor in methylation reactions (Finkelstein 1990). Methionine has been found to be essential for mouse and bovine blastocyst formation, as omission of methionine from culture reduces mouse blastocyst formation, and morula and blastocyst H3K4me3 (Sun et al. 2018), while addition of the methionine antagonist ethionine following embryonic genome activation (EGA) impairs blastocyst development and cell number (Menezo et al. 1989, Ikeda et al. 2012, Kudo et al. 2015) accompanied by a reduction in H3K9me3 modification of Nanog and Tead4 (Kudo et al. 2015). Importantly, this loss in developmental potential coincides with the increase in SAM synthesis that occurs following compaction (Menezo et al. 1989), suggesting a requirement for appropriate metabolite availability at the time of EGA. A requirement for SAM is also evidenced by studies that demonstrate that inhibition of MAT2A, which catalyses the formation of SAM, significantly reduces bovine and mouse blastocyst development and cell number and increases apoptosis (Ikeda et al. 2017, Sun et al. 2018). Conversely, SAM supplementation during bovine embryo culture enhances hatching, accompanied by significant alterations in the global DNA methylation landscape (Shojaei Saadi et al. 2016). Likewise, methionine deprivation reduces SAM levels in human ESC, accompanied by a rapid decrease in H3K4me3 and reduction in global DNA methylation, which can be abrogated through SAM supplementation, with changes in methylation potentiating subsequent differentiation (Shiraki et al. 2014). These data reveal that methionine is important not only for maintaining the pluripotent epigenome but its availability has the potential to alter the timing of lineage specification.
Maternal methionine supplementation has likewise been associated with alterations in bovine blastocyst gene expression (Penagaricano et al. 2013), demonstrating that even subtle changes in methionine availability can impact development. Indeed, methionine utilisation has been proposed as a biomarker of human embryo developmental potential (Houghton et al. 2002). Considerably, deficiencies in metabolites that intersect the folate and methionine cycles, including folate (O’Neill 1998, Kwong et al. 2010) and vitamin B12 (Sinclair et al. 2007), are also known to impact embryo development, and more significantly, alter post-natal growth and health and DNA methylation (Sinclair et al. 2007). Threonine, metabolised via the folate cycle, likewise regulates SAM levels in mouse ESC (Wang et al. 2009a, Shyh-Chang et al. 2013b). The cytoplasmic folate cycle intersects with mitochondrial formate availability (Bao et al. 2016), while mitochondrial NADH oxidation, and therefore TCA activity, has recently been linked to methionine metabolism (Lozoya et al. 2018), highlighting a link between mitochondrial activity and methylation. Mitochondrial one-carbon metabolism therefore integrates nutrient, amino acid metabolism and mitochondrial activity, such that differences in pathway flux can profoundly alter the epigenome.
Conversely, α-KG, a product of the oxidation of glucose and glutamine via the mitochondrial TCA not only modulates the activity of Ten-Eleven translocation (TET) dioxygenases (Xiao et al. 2012) but also regulates the activity of Jumanji (JMJ) demethylases which remove methyl groups from lysine residues (Tsukada et al. 2006). The TET family of enzymes (TET1, 2 and 3) use oxygen to catalyse the oxidative decarboxylation of α-KG, generating a reactive enzyme-bound intermediate which converts 5mC to 5-hydroxymethylcytosine (5hmC). At the pronuclear stage, Tet3 mediates active demethylation of the paternal genome (Gu et al. 2011, Iqbal et al. 2011, Wossidlo et al. 2011) with 5hmC progressively lost through to the 8-cell stage (Inoue & Zhang 2011). In contrast, H3K9me2 of maternal chromatin enables the recruitment of Dppa3 (also known as Stella) to exclude Tet3 from binding and enabling its passive demethylation (Wossidlo et al. 2011, Nakamura et al. 2012). Tet3 expression in porcine embryos is regulated by EGA and modulates Nanog (Gu et al. 2011, Lee et al. 2014b), suggesting an involvement in lineage specification. Tet1 is detectable throughout preimplantation development in the mouse (Ito et al. 2010) and from the 4-cell stage in porcine embryos (Lee et al. 2014b) and becomes enriched in the ICM compared with the trophectoderm at the blastocyst stage, being absolutely required for specification toward the ICM compartment (Ito et al. 2010). The differential expression pattern plausibly reflects the differences in metabolism between these two lineages. Indeed, glucose- and glutamine-derived α-KG modulate mouse ESC histone and TET-dependent DNA demethylation (Carey et al. 2015), suggesting that glycolytic activity supports DNA demethylation and plausibly that appropriate uptake of glucose is likely necessary to establish correct (de)methylation dynamics. This may in part explain the significance of the relationship between glucose uptake and embryo viability (Gardner & Leese 1987, Gardner et al. 2011). Indeed, in vitro culture has been associated with increased DNA methylation relative to in vivo derived embryos in both the mouse and rat (Zaitseva et al. 2007), suggesting dysregulation of demethylating enzymes. It will therefore be important to determine whether modulation of α-KG impacts embryo development, accompanied by changes in methylation.
Appropriate regulation of histone demethylation is also required for normal preimplantation embryo development, mediated by JMJ demethylases which remove the methyl groups from lysine residues in histones in a Fe(2+) and α-KG-dependent manner. Knockdown of maternal JMJD3 (also known as KDM6B), required for the demethylation of H3K27me3 from the 2-cell stage, significantly reduces mouse (Canovas et al. 2012) and bovine blastocyst development accompanied by a reduction in both ICM and trophectoderm cell numbers (Chung et al. 2017), with JMJD3 activity correlated with α-KG availability in mouse ESC (Carey et al. 2015). Moreover, knockdown of JMJD2C (Wang et al. 2010) and the JmjC demethylase KDM5B (Huang et al. 2015) impairs mouse and porcine blastocyst development respectively, accompanied by perturbations in histone methylation, while inhibition of H3K4 demethylation impairs mouse embryo cleavage at the time of EGA (Shao et al. 2008), further indicating that appropriate, stage-specific regulation of the epigenetic landscape is required for development.
Combined, these studies identify that metabolism, and in particular mitochondrial activity, acts as a primary interface between the environment and the epigenome (Fig. 1). As a consequence, perturbations in metabolic pathway activity alter the production and availability of co-factors required for epigenetic modifier activity, resulting in an altered, and heritable, nuclear epigenetic landscape (reviewed by Donohoe & Bultman 2012, Brown et al. 2015, Harvey et al. 2016) at the time of lineage specification when the epigenetic landscape is being programmed. Therefore, perturbations in the nutrient environment surrounding the oocyte and preimplantation embryo may modulate long-term offspring health and viability through alterations in metabolism that impact the programming epigenome. However, it is also important to consider the dynamic nature of mitochondria within the oocyte and early embryo, as a means to deliver not only ATP but also facilitate and regulate the deposition and removal of epigenetic modifications in a stage-specific manner.
Mitochondrial regulation of the preimplantation epigenome. Diet and the extracellular microenvironment regulate the availability of nutrients that modulate metabolic pathway activity and mitochondrial metabolism, regulating the supply of intermediates that act as co-factors for epigenetic modifying enzymes. Glycolytic flux and the activity of the mitochondrial malate aspartate shuttle (MAS) determines the NAD+:NADH ratio. NAD+ regulates the activity of the NAD+-dependent histone deacetylases (sirtuins), repressing gene transcription. Conversely, acetyl-CoA, generated from glucose-derived pyruvate or β-oxidation of fatty acids, acts as a co-factor for histone acetyltransferases (HAT) responsible for transcriptional activation. Similarly, alpha-ketoglutarate (αKG) derived from the mitochondrial tricarboxylic acid (TCA) cycle and glutamine (Gln) metabolism mediates histone and DNA demethylation catalysed by Jumanji (JMJ) and Ten-Eleven Translocation (TET) demethylases that convert 5-methylcytosine (5mC) to 5-hydroxymethylcytosine (5hmC) respectively, activating transcription. Folate-mediated one-carbon metabolism relies on a dietary source of folate to supply tetrahydrofolate (THF), as well as serine (Ser) or mitochondrial-derived formate for the supply of 1C units. Methionine (Met) metabolism is required for S-adenosylmethioninine (SAM) generation via the folate and methionine cycles. SAM acts as a methyl donor for histone (HMT) and DNA (DNMT) methyltransferases to mediate transcriptional repression. Beyond their historical role in providing ATP (adenosine triphosphate), the dynamic nature of mitochondria during the development of the oocyte and preimplantation embryo likely establishes nutrient gradients that finely programme the preimplantation epigenetic landscape to mediate key developmental events, including lineage specification. Consequently, perturbations in nutrient availability, mitochondrial activity or mitochondrial movement have the potential to significantly alter long-term development through the regulation of heritable changes in the epigenome prior to differentiation. 5-mTHF, 5-methyl tetrahydrofolate; ac, acetylation; Asp, aspartate; Glu, glutamate; Gly, glycine; GSH, glutathione; HCY, homocysteine; me, methylation; NAD+, nicotinamide adenine dinucleotide; NADH, reduced form of NAD+; OAA, oxaloacetate; OXPHOS, oxidative phosphorylation; SAH, S-adenosylhomocysteine; THF, tetrahydrofolate.
Citation: Reproduction 157, 5; 10.1530/REP-18-0431
Mitochondrial dysfunction and fertility
In order to support the dynamic changes that occur during preimplantation embryo development, mitochondria within the oocyte and early embryo have primarily been regarded as necessary to generate adequate ATP to fuel the first few days of embryo development (Dumollard et al. 2004). In line with this, the mammalian preimplantation embryo undergoes significant changes in its metabolism during development, where metabolic pathway activity and ATP content vary significantly between human oocytes and embryos (Leese et al. 1986, Chi et al. 1988, Hardy et al. 1989, Gott et al. 1990, Slotte et al. 1990, Conaghan et al. 1993, Gardner & Harvey 2015). Growing oocytes preferentially metabolise pyruvate via OXPHOS, while their somatic compartment, the cumulus cells, are more glycolytic, metabolising glucose to supply the pyruvate required by the oocyte (reviewed by Dumesic et al. 2015). Similarly, the early embryo is characterised by low levels of oxidative metabolism and oxygen consumption, utilising pyruvate, lactate and amino acids to support development (Houghton et al. 1996, Gardner & Wale 2013). In contrast, with the activation of the embryonic genome and the increased energy demand required for blastocoel formation, the blastocyst stage embryo exhibits both high levels of glycolysis and oxygen consumption (reviewed Mills & Brinster 1967, Wales 1986, Houghton et al. 1996, Thompson et al. 1996, by Gardner & Harvey 2015). These changing energy requirements therefore necessitate dynamic and coordinated regulation of metabolism to support ongoing development.
In further support of a requirement for mitochondrial ATP production during development, perturbations in mitochondrial activity have been found to impact embryo development and implantation potential (Van Blerkom et al. 1995), as well as foetal and placental growth (Wakefield et al. 2011). Oocytes containing higher levels of ATP result in significantly higher fertilisation rates and increased blastocyst development. Conversely, those deficient in ATP result in compromised development (Stojkovic et al. 2001, Nagano et al. 2006), reflecting the need for sufficient ATP to support embryo development. A reduced capacity of oocytes to generate ATP has been observed in aged mouse and hamster oocytes (Simsek-Duran et al. 2013) and has been associated with abnormal nuclear spindle and chaotic chromosomal distribution in humans (Wilding et al. 2003) with oocyte mitochondrial activity in particular known to decrease linearly with increasing age (Wilding et al. 2001), and vary with stimulation regimen (Cree et al. 2015). Alterations in ATP content manifest not only as poor or abnormal human oocyte quality but also as reduced embryo development and pregnancy rates (Van Blerkom et al. 1995). However, this represents an oversimplistic view of the impact of mitochondrial activity on the maturing oocyte and developing embryo and fails to consider the importance of these organelles in modulating other aspects of cell homeostasis.
An alternative explanation for reduced fertility with ageing, or environmental perturbations, beyond ATP availability, is the accompanying alteration in the provision of metabolic intermediates necessary for modulation of the epigenetic landscape. Indeed, oocyte ageing is accompanied by reduced 5mC and DNMT expression levels (Yue et al. 2012), H4K12 and H4K16 acetylation (Manosalva & Gonzalez 2009) and histone methylation (Manosalva & Gonzalez 2010). Taken together, it is plausible that alterations in metabolic activity mediate the changes in the epigenetic landscape of oocytes and/or the early embryo, impacting subsequent developmental competence. Surprisingly, relatively few studies have investigated specific reproductive tract metabolite levels associated with ageing or dietary changes. Alterations in human follicular fluid lipid profiles have been associated with ageing (Cordeiro et al. 2018), as well as with embryos that fail to cleave (O’Gorman et al. 2013), however whether epigenetic change is elicited by these environments remains unclear. Ageing likewise alters cumulus cell function, with a significant impact on transcripts (Al-Edani et al. 2014) and proteins (McReynolds et al. 2012) involved in metabolism, possibly reflecting a reduced ability to support the metabolic and epigenetic requirements of the oocyte. As acetylation has recently been shown to be regulated by lipids and lipid-derived acetyl-CoA (McDonnell et al. 2016), and circulating maternal lipids have been shown to programme the developing foetal DNA methylation landscape (Marchlewicz et al. 2016), alterations in follicular fluid lipid content with ageing may perturb the oocyte epigenetic landscape. Indeed, lipid-derived acetyl-CoA is more readily incorporated into histones than that derived from glucose (McDonnell et al. 2016), emphasising that metabolic plasticity results in epigenetic remodelling in response to nutrient availability. Oocyte NADH/FAD levels have also been correlated with ageing (Babayev et al. 2016), and mitochondrial dysfunction has been associated with reduced NADH/NADPH in human zygotes (Wilding et al. 2001, 2002, 2003), yet whether such changes impact oocytes from young individuals, or the regulation of the sirtuin NAD+-dependent deacetylases, remains to be determined. The effects of ageing on the levels of other metabolites within follicular or reproductive tract fluid, including acetyl-CoA, have similarly not been investigated. It will therefore be essential to examine metabolite perturbations associated with poor developmental competence and outcomes, along with consequences for long-term offspring health, to establish whether specific changes in metabolite content associated with ageing and nutrient availability result in epigenetic priming that plays a role in the reduction in developmental competence.
Mitochondrial morphology in the oocyte and preimplantation embryo
During preimplantation embryo development, mitochondria undergo dynamic, functional and structural changes, morphing from spherical organelles containing dense matrices and few peripheral arched cristae to long filamentous organelles with numerous transverse cristae that maximise the surface area available for OXPHOS (Zick et al. 2009). This contrasts with the reticular, elongated, electron- and cristae-rich morphology of somatic cells. A spherical morphology is also characteristic of ESC (St John et al. 2005, Cho et al. 2006, Chung et al. 2007), suggesting that it underpins the pluripotency of these cells (reviewed by Harvey et al. 2013). These mitochondrial changes therefore reflect the developmental stage, and metabolic requirements, of the embryo (reviewed by Bavister & Squirrell 2000, Lees et al. 2017), but more significantly likely also facilitate mitochondrial signalling and crosstalk.
The primary human oocyte contains many spherical mitochondria with few peripheral arched cristae dispersed throughout the cytoplasm (Motta et al. 2000). Mitochondria remain primarily spherical during folliculogenesis, with pale matrices and small vesicular cristae, clustered around the nucleus (Stern et al. 1971, Sathananthan & Trounson 2000, Van Blerkom 2004). While primordial oocytes contain approximately 10,000 mitochondria, this increases to around 100,000 in mature oocytes (Lanzavecchia & Mangioni 1964, Sathananthan et al. 1986, Jansen & de Boer 1998). By ovulation, mitochondria are the most prominent organelle in the oocyte cytoplasm (Motta et al. 2000). During early cleavage, elongated mitochondria with well-developed transverse cristae begin to emerge (Piko & Matsumoto 1976, Motta et al. 2000). As development proceeds, the proportion of elongated, cristate-rich mitochondria increases, such that by the blastocyst stage in the human and mouse, blastomeres contain a mixed population of predominantly spherical and elongated mitochondria in the ICM and trophectoderm respectively (Nadijcka & Hillman 1974, Mohr & Trounson 1982, Motta et al. 2000, Sathananthan & Trounson 2000, Lees et al. 2017).
Despite the perceived ‘immature’ morphology, oocyte mitochondria are highly active, producing the majority of energy required to support development. The structural changes are however essential for preimplantation embryo development to proceed (Van Blerkom 1989) and are consistent with a progressive activation of mitochondrial respiratory activity, inferred from increased oxygen consumption and ATP production via OXPHOS (Houghton et al. 1996, Thompson et al. 1996, Trimarchi et al. 2000, Sturmey & Leese 2003) and changes in cristae shape (Mohr & Trounson 1982, Sathananthan & Trounson 2000, Cogliati et al. 2013). Interestingly, human apical trophectoderm cells are characterised by elongated mitochondria with transverse cristae, largely located around the periphery (Mohr & Trounson 1982). This morphological difference in both mitochondrial architecture and activity likely reflects differences in energy demand, but may also facilitate the process of implantation or may play a role in regulating cell specification, plausibly by differential metabolite availability regulating the epigenome. Metabolites are increasingly recognised as regulators of pluripotency and cell fate (Chung et al. 2007, Shyh-Chang et al. 2013b, Shiraki et al. 2014, Moussaieff et al. 2015), with mitochondria at the heart of sensing and regulating lineage decisions and establishing cellular memory (reviewed by Harvey et al. 2016, Lees et al. 2017). It is tempting to speculate that mitochondria may be involved in the regulation of the earliest embryonic specification event, the establishment of a polarised epithelium, in a manner similar to that observed when hepatocytes initiate polarisation, necessitating mitochondrial elongation (Fu et al. 2013). Notably, ATP generated via OXPHOS is required for this process (Fu et al. 2013). Further, the Hippo pathway, which mediates the differentiation of the trophectoderm from the ICM (reviewed by Manzanares & Rodriguez 2013), also regulates mitochondria fusion and size (Nagaraj et al. 2012). These data suggest that mitochondrial structure and activity, and cell polarisation, may be intricately linked, such that deficiencies in mitochondrial dynamics and activity during early development may have a significant impact on cell allocation and developmental capacity beyond the provision of ATP.
Regulation of mitochondrial morphology
Mitochondrial morphology adapts to fluctuating cellular requirements by modulating changes in shape through fission and fusion (Frederick & Shaw 2007), continually dividing and fusing to form a dynamic interconnecting network necessary to maintain mitochondrial and mtDNA integrity (Suen et al. 2008). Fission and fusion events are mediated by a number of nuclear encoded proteins (reviewed by Wai & Langer 2016). Mitochondrial fusion not only enables the creation of extensive mitochondrial networks to increase energy supply (Chen et al. 2005) but also the complementation, and the mixing of mitochondrial content, between healthy and defective mitochondria (Youle & van der Bliek 2012). The outer mitochondrial membrane GTPase required for mitochondrial fusion; fuzzy onions (fzo; Hales & Fuller 1997) was identified as an essential regulator of spermatogenesis. The human homologs, mitofusin 1 and 2 (MFN1, MFN2 respectively) likewise display GTPase-dependent activity and mediate mitochondrial fusion (Santel & Fuller 2001, Chen & Chan 2004). Cells deficient in either Mfn1 or Mfn2 display considerably reduced levels of mitochondrial fusion and distinct types of fragmented mitochondria (Chen et al. 2003, 2005) and increased ROS (Lee et al. 2014a). Beyond their role in spermatogenesis, they also contribute to ongoing development. Mice deficient in either Mfn1 or Mfn2 die around embryonic day (E) 11.5. Mfn1 mutants are significantly smaller with a pronounced developmental delay prior to resorption, while Mfn2 mutant embryos display a severe disruption of the placental trophoblast giant cell layer (Chen et al. 2003). Mfn2 knockdown in oocytes reduces oocyte maturation and fertilisation and alters mitochondrial distribution and spindle morphology (Liu et al. 2016a), further implicating mitochondria in the regulation of chromosome segregation in the oocyte. Moreover, reduced Mfn2 expression following siRNA attenuates mouse blastocyst formation and significantly reduces the number of embryos progressing through the third cell division (Zhao et al. 2015). As this coincides with the activation of the embryonic genome, active mitochondrial fusion is likely required to support this milestone. Embryos that survive Mfn2 knockdown, exhibit decreased ATP and mtDNA levels, reduced mitochondrial membrane potential and increased levels of apoptosis (Zhao et al. 2015), further suggesting that adequate control of mitochondrial dynamics is necessary for continued development, however whether these knockout phenotypes were accompanied by alterations in the epigenome, particularly at the time of EGA, were not examined.
A third protein, Optic Atrophy 1 (OPA1), is also essential for inner mitochondrial membrane fusion (Olichon et al. 2003). Opa1 knockout is similarly embryonically lethal by E13.5 in the mouse, further supporting a role for mitochondrial dynamics in regulating development (Davies et al. 2007). Loss of OPA1 results in cristae widening, thereby playing a role in the regulation of mitochondrial cristae shape (Patten et al. 2014). Significantly, low ADP concentrations, characteristic of the late cleavage and morula stage embryo (Leese et al. 1984), were reported to modify cristae shape to a more expanded, less dense matrix more than 50 years ago (Hackenbrock 1966), however only recently has cristae shape and volume been found to modulate the rates of several TCA cycle enzymes (Lizana et al. 2008), as well as the mitochondrial membrane, redox-sensitive protein ROS modulator 1 (ROMO1; Norton et al. 2014) and cardiolipin (Ikon & Ryan 2017, reviewed by Quintana-Cabrera et al. 2018). How these pathways are regulated by changes in nutrient availability, particularly during embryo development, remains to be determined. Given the redox-sensitive nature of OPA1 (Garcia et al. 2018) and ROMO1, ROS, NAD+:NADH and ATP:ADP ratios, which are impacted by in vitro and in vivo nutrient availability, are likely important signalling molecules that link mitochondrial metabolism, cristae morphology and cell signalling. Conceivably, any form of oxidative stress or metabolic dysfunction induced by assisted reproductive technologies, including culture, may therefore interfere with mitochondrial dynamics.
Conversely, the process of mitochondrial fission is mediated in part by dynamin-related protein 1 (Drp1), a large GTPase and fission 1 (Fis1). Drp1 is a large dynamin-related GTPase located primarily in the cytoplasm, with punctuate localisation marking future sites of fission (Smirnova et al. 2001). Downregulation of Drp1 results in loss of mitochondrial DNA, decreased mitochondrial respiration and increased ROS production in HeLa cells (Parone et al. 2003), as well as reduced cell proliferation and cellular ATP. Knockout of Drp1 in mice leads to embryonic lethality at E11.5 accompanied by a reduction in size, plausibly attributable to the loss of placental giant cells (Ishihara et al. 2009, Wakabayashi et al. 2009). Perturbations in placental structure would contribute to reduced nutrient transport, thereby implicating mitochondria in regulating placental cell differentiation and fate. In addition, oocyte specific Drp1 knockout results in infertility (Udagawa et al. 2014), and both decreased expression of Drp1 and increased Mfn2 mRNA levels, favouring mitochondrial fusion, have been associated with fragmented human day 3 embryos (Otasevic et al. 2016), although whether this is a cause or consequence is unclear. Further, inhibition of mitochondrial fission with the specific Drp1 inhibitor Mdivi-1 reduces porcine blastocyst formation accompanied by reduced mitochondrial membrane potential and increased ROS (Yeon et al. 2015), with increasing expression during early porcine cleavage stages indicative of higher quality embryos (Yang et al. 2018). Significantly, disruption of mitochondrial fission via Drp1 siRNA results in chromosomal instability and over-amplification of centromeres in a number of cell lines (Qian et al. 2012), suggesting that the incidence of aneuploidy may reflect compromised mitochondrial dynamics, again linking mitochondrial function to chromosomal segregation. Additionally, ROS are also known to directly alter the epigenetic landscape (Hitchler & Domann 2009), such that conditions that result in the overproduction of ROS, including through the disruption of mitochondrial fission/fusion, may result in ROS-mediated changes to the epigenetic landscape. A second key component of mitochondrial fission is Fis1 (Fission 1), a transmembrane protein localised to the outer mitochondrial membrane, recruits Drp1 to areas of mitochondrial constriction, resulting in membrane fission (Stojanovski et al. 2004). However, no studies have addressed the role of Fis1 during mammalian embryo development.
Collectively, these data reveal that a balance between mitochondrial fission and fusion is required to regulate developmental progression and embryo viability, likely to ensure segregation of mitochondria between cells and ensure appropriate regulation of chromosome segregation, but also support appropriate mitochondrial activity and ROS levels. This balance can be perturbed by changes in nutrient availability, leading to mitochondrial fragmentation or hyperfusion and is dependent on mitochondrial membrane potential (Ishihara et al. 2003). Whether active fission and fusion events are altered in in vitro cultured embryos, or by alterations in nutrient availability, have not been examined. While knockout studies highlight the significance of regulators of mitochondrial dynamics during in vivo development, few studies have addressed changes that may be induced by loss of these proteins prior to implantation that may correlate with reduced potential or implantation failure. Equally, studies examining modulation of mitochondrial dynamics have not assessed whether changes in methylation or acetylation occur.
The dynamic nature of mitochondria: relocalisation to facilitate stage-specific epigenetic programming?
Mitochondrial localisation also changes significantly throughout oocyte and early embryo development. Given the role of mitochondrially derived metabolic intermediates in regulating the epigenome, mitochondrial trafficking may serve to not only transport mitochondria to sites of energy demand, and facilitate communication and interaction with other organelles to support embryo development, but also to maintain metabolite/signalling gradients with the nucleus.
Although subtle species differences exist, significant mitochondrial (re)distribution occurs in the developing oocyte (Takahashi et al. 2016), with distinct patterns of localisation in pronuclear oocytes and early cleavage stage embryos correlated with developmental potential (Wang et al. 2009c, Igosheva et al. 2010, Yu et al. 2010). In mouse and human oocytes, highly polarised mitochondria cluster around the oocyte cortex (Van Blerkom et al. 2003, Van Blerkom & Davis 2006, 2007), proposed to maintain sufficient ATP for fertilisation, but also to support Ca2+ regulation upon fertilisation. In contrast, mitochondria with low membrane potential are homogeneously distributed throughout the cytoplasm. The cortical localisation of active mitochondria may conceivably be required to support oocyte metabolism through close proximity of mitochondria to the source of pyruvate (the surrounding cumulus cells) given that the oocyte is unable to metabolise glucose (Sutton-McDowall et al. 2010) or establish nutrient gradients to modulate pronuclear dynamics. Localised ATP bursts have also been observed during mouse oocyte maturation, correlating with the redistribution of mitochondria (Yu et al. 2010), but may equally be indicative of fluctuations in other mitochondrially derived metabolites in a pulsatile manner.
Following fertilisation, mitochondria translocate to cluster around the two pronuclei (Motta et al. 2000, Squirrell et al. 2003). Mitochondrial translocation to a perinuclear arrangement has been observed in hamster (Barnett et al. 1996), mouse (Van Blerkom & Runner 1984), pig (Sun et al. 2001), rhesus macaque (Squirrell et al. 2003) and human (Noto et al. 1993, Van Blerkom et al. 2000) zygotes, providing ATP and other intermediates to facilitate syngamy and ensure an even organelle distribution between resultant blastomeres. Intriguingly, the relocation of mitochondria coincides with differential modulation of the paternal and maternal epigenomes, such that clustering likely establishes crosstalk between the nuclear and mitochondrial genomes, and may establish local metabolite domains to regulate epigenetic modifiers. Interestingly, nuclear expression of mitochondrial TCA enzymes, necessary to produce metabolites required for epigenetic remodelling at the time of EGA, has recently been described in mouse embryos (Nagaraj et al. 2017), and relies on pyruvate availability, indicating a need for temporal nutrient requirements to regulate the preimplantation epigenetic landscape. The dynamic changes to the epigenetic landscape that occur at syngamy may therefore necessitate translocation of mitochondria to the pronuclei to provide essential metabolites, including α-KG and NAD+ to support demethylase activity. Dumollard et al. (2007) reported that NAD(P)H levels significantly decreased following fertilisation, an inverse indicator of an increase in NAD+ availability, supporting this hypothesis. However, temporal changes in NAD(P)H levels during syngamy remains relatively unexplored. Significantly, a lack of mitochondrial organisation at this stage has been correlated with reduced implantation (Wang et al. 2009b), with mitochondrial aggregation characteristic of oocytes with reduced developmental potential (Igosheva et al. 2010), suggesting that these dynamic properties of mitochondria are essential for embryo development. Further, ovarian stimulation regimen has a substantial impact on mitochondrial distribution in the mouse (Ge et al. 2012) and human oocyte (Dell’Aquila et al. 2009), indicating that stimulation can significantly alter mitochondrial dynamics, which may have consequences for implantation, the induction of aneuploidies, viability and long-term health through the modulation of epigenetics.
Perturbations in mitochondrial movement and recruitment are also likely to lead to subcellular ATP depletion and interfere with mitochondrial Ca2+ signalling. Mitochondria play a role in Ca2+ storage and homeostasis (Duchen 2000), with sustained Ca2+ oscillations following fertilisation necessary for oocyte activation (Kline & Kline 1992) and embryo development (Stricker 1999). Mitochondrial localisation, and activity, interplays with Ca2+ signalling at fertilisation, such that mitochondrial ATP production is required for maintaining Ca2+ levels in the oocyte (Wakai et al. 2013) and for sustaining Ca2+ oscillations post-fertilisation (Dumollard et al. 2004). Significantly, mitochondrial movement along microtubule tracks is regulated by Ca2+ sensitive microtubule associated proteins, including kinesins, dyneins and myosins that drive motor-based movement (Varadi et al. 2004, Lawrence et al. 2016), suggesting that mitochondrial distribution relies on their own activity, as well as the integrity of the cytoskeletal network, to sustain localised signalling. Indeed, inhibition of microtubule polymerisation in human oocytes reduces germinal vesicle breakdown (GVBD) and prevents the completion of maturation, accompanied by the loss of mitochondrial redistribution (Takahashi et al. 2016), and cytoskeletal disruption in mouse eggs alters mitochondrial ATP production (Yu et al. 2010). Consequently, altered mitochondrial activity may reduce their ability to modulate Ca2+ dependent interactions with the cytoskeleton.
Connectivity and communication between mitochondria, the nucleus and other organelles, as well as the cytoskeleton, is therefore essential to the maintenance of cellular homeostasis in the oocyte and developing embryo, integrating diverse environmental signals as in other cell types (reviewed by Anesti & Scorrano 2006, Rowland & Voeltz 2012, Marchi et al. 2014) and eliciting signalling cascades that support preimplantation development. Loss, or misregulation, of these close interactions may therefore negatively impact the embryo’s ability to respond to stress and regulate mitochondrial function, or lead to precocious or delayed modification of the epigenetic landscape, particularly in response to changes in nutrient availability, thereby impacting long-term viability and health.
Limitations to assessing mitochondrial DNA content in isolation
While mitochondrial health and functionality relies on metabolic activity, distribution, morphology and density, an often-overlooked characteristic that separates mitochondria from other organelles is that each mitochondrion contains numerous copies of its own double stranded genome (mitochondrial DNA; mtDNA). mtDNA encodes 13 proteins involved in the respiratory chain, along with 22 transfer RNAs and 2 ribosomal RNAs (Anderson et al. 1981, Taanman 1999, Clayton 2000); however a significant proportion of genes required for mitochondrial biogenesis, homeostasis and regulation of OXPHOS are encoded by the nuclear genome. Coordinated signalling between the nucleus and mitochondrion is therefore essential to regulate mitochondrial function (reviewed elsewhere by St John et al. 2010, Harvey et al. 2011), and metabolic intermediate levels, and therefore the epigenome. Indeed, a role for mtDNA, and therefore mitochondrial activity, in regulating the nuclear epigenetic landscape is evidenced by a study examining mtDNA depletion which results in aberrant CpG methylation that can be reversed following repletion of mtDNA (Smiraglia et al. 2008). Thus, deficiencies in mtDNA may be paralleled by changes to the epigenome that reduce developmental competence.
The mature human and mouse oocyte contains an estimated 20,000 to 800,000 mtDNA copies (Steuerwald et al. 2000, Reynier et al. 2001, Barritt et al. 2002) and between 10,000 to 650,000 copies (Piko & Taylor 1987, Steuerwald et al. 2000, Aiken et al. 2008, Cree et al. 2008) respectively, with mtDNA levels remaining low following fertilisation, only increasing around the time of implantation (Thundathil et al. 2005, May-Panloup et al. 2005b, Bowles et al. 2007, Spikings et al. 2007, Aiken et al. 2008, Mtango et al. 2008). A threshold of 40, 000 to 50, 000 mtDNA copies in a mature mouse oocyte has been proposed as critical to support development post-implantation (Wai et al. 2010). While blastocysts still develop with as few as 4000 mtDNA copies, extrapolated from 8-cell embryos, those with low mtDNA levels failed to develop post-implantation (Wai et al. 2010). Likewise, while unfertilised human oocytes and their associated somatic cells harbour higher levels of mtDNA mutations and deletions (Keefe et al. 1995, Seifer et al. 2002, Hsieh et al. 2004), associated with reduced expression of mitochondrial genes (Hsieh et al. 2002), and oocytes from older females being more likely to possess deletions (Keefe et al. 1995, Jacobs et al. 2007, Hammond et al. 2016), how this relates to oocytes within the cohort that fertilise is difficult to determine. Significantly, levels vary considerably within one individual, and can be altered by ovarian stimulation regimen (Cree et al. 2015). Moreover, mtDNA copy number is often assumed to be an indicator of mitochondrial number, yet each mitochondrion can contain a varying number of mtDNA nucleoids (Gilkerson et al. 2013). Combined with species’ differences, and the use of discarded and poor-quality human samples, the utility and specificity of these thresholds is potentially limited.
Despite these limitations, mtDNA defects have been associated with ageing and the failure of mature oocytes to fertilise. mtDNA levels decline in oocytes with advancing maternal age (de Boar et al. 1999, Murakoshi et al. 2013) and unfertilised oocytes contain significantly fewer copies of mtDNA, suggesting that mtDNA copy number may be indicative of fertilisation potential (Reynier et al. 2001, Santos et al. 2006, Wai et al. 2010). Ovarian insufficiency has also been characterised by a significantly lower mtDNA content in oocytes (May-Panloup et al. 2005a), as well as accompanying cumulus cells (Boucret et al. 2015). Further, blastocysts from high fat diet (HFD) fed mice contain fewer mtDNA copy numbers and give rise to heavier foetuses in which both liver and kidney mtDNA content is significantly reduced (Wu et al. 2015), indicating that altered nutrition prior to conception can disrupt mitochondrial transmission. In addition to a lowered capacity for ATP generation, reduced mtDNA levels likely result in a diminished pool of TCA metabolites that are preferentially shunted toward ATP production to support cell survival in lieu of histone modification, predisposing the developing embryo to compensatory changes in physiology and growth. Indeed, mitochondrial DNA depletion has been shown to result in alterations in one-carbon metabolism (Bao et al. 2016) and TCA activity, altering transcription of genes involved in acetyl-CoA metabolism, and modifies amino acid metabolism, particularly that of methionine, consequently impacting on methylation (Lozoya et al. 2018). Conversely, enhanced mitochondrial OXPHOS capacity with increased mtDNA content may result in an increased availability of mitochondrial metabolites that results in hypermethylation and/or acetylation, likewise impacting downstream growth and physiology.
Accumulating evidence suggests that epigenetic changes also occur within mtDNA. While low levels (<5%) of mtDNA methylation were reported in mouse and human cells in the 1980s (Shmookler Reis & Goldstein 1983, Pollack et al. 1984), mitochondria were believed to lack histones, the major component of chromatin for epigenetic modification, such that the existence of mtDNA methylation was, and remains, controversial. While speculation remains around potential sample contamination, histones have been detected in the mitochondrial membrane (Choi et al. 2011). Combined with the identification of mtDNA methyltransferase enzyme 1 (mtDNMT1), associated with mitochondrial cytosine methylation (Shock et al. 2011) and the isolation of DNMT3a from mitochondria (Wong et al. 2013), it is possible that methylation actively takes place within mitochondria. In support of this, mitochondrial targeted expression of DNA methyltransferases is sufficient to induce mtDNA methylation (van der Wijst et al. 2017) and (de)methylation of the D-loop has been associated with a reduction in mtDNA copy number (Tong et al. 2017), which would reduce mitochondrial gene expression. The mtDNA methylation may consequently alter the availability of the co-factors necessary for the activity of nuclear epigenetic modifiers, through the modulation of mtDNA encoded genes and therefore OXPHOS activity, although direct evidence for this, and how mitochondrial methyltransferases are regulated, is currently lacking. Similarly, given that mitochondria, and mtDNA, are more susceptible to damage, and rely on cellular antioxidant systems to prevent damage, the mtDNA epigenome may be even more vulnerable than the nuclear epigenetic landscape. Consequently, it will be important to investigate the effects of not only culture, but other aspects of assisted reproductive technologies, particularly mitochondrial replacement, on the oocyte and embryo nuclear and mtDNA epigenome, particularly given the potential for altering offspring health.
Controversially, studies have emerged suggesting that a higher mtDNA copy number (e.g. Mitoscore >97; Diez-Juan et al. 2015), in blastomere or trophectoderm biopsies, or in cumulus cells, is an indicator of poor blastocyst quality and/or implantation success (Diez-Juan et al. 2015, Fragouli et al. 2015, Ogino et al. 2016, Desquiret-Dumas et al. 2017, Ravichandran et al. 2017), with the increase suggestive of underlying stress and compensatory mitochondrial biogenesis activated in an attempt to rescue development. This is however at odds with the metabolic requirements of the blastocyst stage embryo (Gardner & Harvey 2015). Moreover, these scores rely on an artificial reference threshold, and do not account for differences in mtDNA levels that exist between individual cells or cell types, which has been documented within the embryo (Cree et al. 2008). Indeed, mtDNA segregation has recently been shown to be non-random, rather relating to the level of OXPHOS and mitochondrial membrane potential (Otten et al. 2018), the activity of which differs considerably between the trophectoderm and the ICM (reviewed by Gardner & Harvey 2015). However, a direct determination of neither whether a consistent relationship exists in both the trophectoderm and ICM nor whether defective mitochondria are preferentially segregated into the trophectoderm relative to the ICM, or whether trophoblast biopsies presenting with higher mtDNA levels reflect a significant reduction in ICM levels that contribute to their demise has been established.
Conversely, studies have shown mtDNA content to have no correlation with implanting and non-implanting embryos (Treff et al. 2017, Victor et al. 2017), although significant mathematical correction may preclude the identification of a relationship. Further, there appears to be little predictive value at moderate (e.g. 75th percentile) mtDNA levels (Klimczak et al. 2018), with considerable range overlap apparent across all studies to date. Methodological differences between species and clinics, including but not limited to stimulation regimens and maternal age (Cree et al. 2015, Fragouli et al. 2015), developmental stage relating to cell number and volume and developmental kinetics (Ho et al. 2018), but also culture conditions that alter nutrient availability, likely impact the determination of mtDNA copy number, although oxygen concentration does not appear to be a factor (de Los Santos et al. 2018). Consequently, the utility of blastocyst mtDNA copy number as a biomarker of embryo viability, in isolation, is questionable and will result in embryos that would produce a viable pregnancy being classified as non-viable; a travesty for those with limited embryo numbers and opportunities to become pregnant. While the use of mtDNA copy number is likely to be somewhat informative, it presents an oversimplistic view of the potential significance of mitochondrial function as a biomarker, particularly in light of their role in mediating the epigenome. Rather, mitochondrial, and mtDNA, integrity, taking into account activity, content (both mitochondrial and mtDNA heteroplasmy and mutational load) and mitochondrial dynamics (including morphology and distribution) will need to be integrated, together with existing markers and epigenetic analyses, in order to be of predictive value and to establish a correlation with post-transfer outcomes.
Mitochondrial replacement therapy: proceed with caution
Mitochondrial replacement therapy (MRT), approved for use in the United Kingdom in 2015, has been proposed as a viable option to prevent the transmission of debilitating mitochondrial diseases. The technique involves the removal of donor nuclear material, and replacement with the diseased individual’s DNA, either via pronuclear, spindle or polar body transfer. However, the procedure is only applicable to cases where the mitochondrial disease is encoded by mtDNA, excluding the vast majority of cases in which mitochondrial disease is nuclear encoded. MRT pregnancies are currently ongoing, although controversially, the first baby born using the technique was reported in 2016 (Zhang et al. 2017).
However, the technology brings with it the potential for long-term health effects, particularly given the plethora of roles mitochondria have in regulating not only metabolism, but also localised signalling cascades and intracellular interactions as noted herein. Of further concern is the potential for the retention of diseased maternal mtDNA, resulting in heteroplasmy, which can remain undetected (Tachibana et al. 2013), or be detected at levels of up to 10% in foetal tissues (Zhang et al. 2017). Sustained heteroplasmy has been reported following cytoplasmic transfer into human oocytes (Brenner et al. 2000) with varying proportions of mtDNA observed in different tissues (Barritt et al. 2001), although whether the technique and/or heteroplasmy has directly affected offspring health and disease risk is unclear. Heteroplasmy has also been reported in cloned animals, with proportions ranging from 0.5 to 40% (Hiendleder et al. 1999, Takeda et al. 1999, 2003, Steinborn et al. 2000, Hiendleder et al. 2003), although these figures are likely underestimates given the lack of sensitivity of the techniques used (Hammond et al. 2016). Ma et al. (2015) reported that mitochondrial deficits could be corrected with somatic cell nuclear transfer and genetic manipulation, however varying (0–100%) levels of heteroplasmy remained detectable in these lines. While heteroplasmy can occur in normal individuals, and varies across tissues, it has been associated with mitochondrial disease and metabolic perturbations in mice (Acton et al. 2007, Sharpley et al. 2012) with severity depending on the threshold level of diseased mitochondria. Significantly, a disease phenotype can occur even at very low (<5%) levels of heteroplasmy for certain deficits (Chinnery et al. 1997, Sacconi et al. 2008). While the birth of live macaque offspring (Tachibana et al. 2009) facilitated the development and adoption of this technique, longitudinal, whole systems studies of the health of resultant offspring is warranted, combined with care to ensure the absence of mitochondrial carry over from the patient.
Beyond the potential for heteroplasmy, the technique neglects the requirement for coordinated communication between mitochondria and the nucleus, along with the dynamic regulation of mitochondrial movement during oocyte maturation and fertilisation. As proposed, the recruitment of mitochondria to the pronuclei preceding syngamy (Barnett et al. 1996, Squirrell et al. 2003) may be required to modulate the local metabolite environment to facilitate syngamy, subsequent cytokinesis and differentially modify the parental genomes, a process that may well be initiated prior to pronuclear migration. Further, given the significance of microtubules to mitochondrial distribution and chromosome segregation as noted above, it is therefore likely that a loss in microtubule network integrity will impact on further development via impairment of mitochondrial trafficking. Mitochondrial distribution following cytoplasmic transfer has been shown to depend on oocyte stage (Fulka 2004), possibly reflecting stage-specific changes in microtubule dynamics. Further, asymmetrical mitochondrial distribution at the pronuclear stage results in blastomeres with reduced mitochondrial inheritance and diminished ATP generating capacity (Van Blerkom et al. 2000), and plausibly altered paternal/maternal methylation characteristics that contribute to the observed reduction in developmental competence. Therefore, studies will need to address whether MRT methodologies alter mitochondrial and cellular metabolic regulation, as well as the embryo transcriptome and epigenome, to ensure minimal risk to resultant offspring health and the absence of transgenerational effects.
Conclusions
The involvement of mitochondria in regulating oocyte and embryo development goes beyond their role in generating ATP to support energy demanding processes. It is increasingly recognised that mitochondria are dynamic signalling organelles, trafficking to specific regions within the cell and communicating not only with the nucleus but also other organelles; interactions that facilitate signalling cascades and impact chromosome segregation, cytokinesis and lineage specification. Loss of mitochondrial function, including activity, organelle interactions and trafficking, and the mechanisms that modulate mitochondrial and mtDNA integrity, manifests as reduced development, and can result in failure to develop post-implantation. Combined with our evolving understanding of how metabolite availability and metabolic and mitochondrial pathway flux can drastically affect the availability of co-factors required for the activity of epigenetic modifiers, mitochondria are emerging as sensors that integrate not only metabolic and cell signalling cues but may in part explain the origins of poor developmental outcomes through heritable changes that lead to disease susceptibility. Therefore, while the developing embryo exhibits a degree of plasticity to changes in its extracellular environment, enabling development to the blastocyst stage to continue, it is nonetheless uniquely sensitive to perturbations in nutrient availability at a time when the epigenetic landscape is being established. The complexity of their diverse roles is of particular significance given the adoption of assisted reproductive technologies that detect or manipulate mitochondria or mtDNA numbers and potentially disrupt mitochondrial-organelle and -cytoskeletal communication and trafficking. This complexity necessitates the determination of mitochondrial characteristics in combination with epigenetic change in order to provide a more accurate understanding of the impact of mitochondrial deficits and manipulation on oocyte, embryo and offspring health, as well as their utility as biomarkers.
Declaration of interest
The author declares that there is no conflict of interest that could be perceived as prejudicing the impartiality of this review.
Funding
This research did not receive any specific grant from any funding agency in the public, commercial or not-for-profit sector.
Acknowledgements
The author would like to thank Professor David K Gardner and Dr Mark P Green for their valuable comments on the manuscript.
References
Acton BM, Lai I, Shang X, Jurisicova A & Casper RF 2007 Neutral mitochondrial heteroplasmy alters physiological function in mice. Biology of Reproduction 77 569–576. (https://doi.org/10.1095/biolreprod.107.060806)
Adenot PG, Mercier Y, Renard JP & Thompson EM 1997 Differential H4 acetylation of paternal and maternal chromatin precedes DNA replication and differential transcriptional activity in pronuclei of 1-cell mouse embryos. Development 124 4615–4625.
Ahn CS & Metallo CM 2015 Mitochondria as biosynthetic factories for cancer proliferation. Cancer and Metabolism 3 1. (https://doi.org/10.1186/s40170-015-0128-2)
Aiken CE, Cindrova-Davies T & Johnson MH 2008 Variations in mouse mitochondrial DNA copy number from fertilization to birth are associated with oxidative stress. Reproductive Biomedicine Online 17 806–813. (https://doi.org/10.1016/S1472-6483(10)60409-9)
Al-Edani T, Assou S, Ferrieres A, Bringer Deutsch S, Gala A, Lecellier CH, Ait-Ahmed O & Hamamah S 2014 Female aging alters expression of human cumulus cells genes that are essential for oocyte quality. BioMed Research International 2014 964614. (https://doi.org/10.1155/2014/964614)
Anderson S, Bankier AT, Barrell BG, de Bruijn MH, Coulson AR, Drouin J, Eperon IC, Nierlich DP, Roe BA & Sanger F et al. 1981 Sequence and organization of the human mitochondrial genome. Nature 290 457–465. (https://doi.org/10.1038/290457a0)
Anesti V & Scorrano L 2006 The relationship between mitochondrial shape and function and the cytoskeleton. Biochimica and Biophysica Acta 1757 692–699. (https://doi.org/10.1016/j.bbabio.2006.04.013)
Babayev E, Wang T, Sanchez T, Lowther K, Taylor HS, Sakkas D, Needleman D & Seli E 2016 Oocyte ageing is associated with altered metabolic stress response and lower mitochondrial DNA copy number that correlate with intracellular NADH and FAD measured by fluorescence lifetime imaging microscopy (FLIM). Fertility and Sterility 106 e69. (https://doi.org/10.1016/j.fertnstert.2016.07.204)
Bao XR, Ong SE, Goldberger O, Peng J, Sharma R, Thompson DA, Vafai SB, Cox AG, Marutani E & Ichinose F et al. 2016 Mitochondrial dysfunction remodels one-carbon metabolism in human cells. ELife 5. (https://doi.org/10.7554/eLife.10575)
Barnett DK, Kimura J & Bavister BD 1996 Translocation of active mitochondria during hamster preimplantation embryo development studied by confocal laser scanning microscopy. Developmental Dynamics 205 64–72. (https://doi.org/10.1002/(SICI)1097-0177(199601)205:1<64::AID-AJA6>3.0.CO;2-3)
Barritt JA, Brenner CA, Malter HE & Cohen J 2001 Mitochondria in human offspring derived from ooplasmic transplantation. Human Reproduction 16 513–516. (https://doi.org/10.1093/humrep/16.3.513)
Barritt JA, Kokot M, Cohen J, Steuerwald N & Brenner CA 2002 Quantification of human ooplasmic mitochondria. Reproductive Biomedicine Online 4 243–247. (https://doi.org/10.1016/S1472-6483(10)61813-5)
Bavister BD & Squirrell JM 2000 Mitochondrial distribution and function in oocytes and early embryos. Human Reproduction 15 (Supplement 2) 189–198. (https://doi.org/10.1093/humrep/15.suppl_2.189)
Bender DA 1983 Biochemistry of tryptophan in health and disease. Molecular Aspects of Medicine 6 101–197. (https://doi.org/10.1016/0098-2997(83)90005-5)
Boskovic A, Bender A, Gall L, Ziegler-Birling C, Beaujean N & Torres-Padilla ME 2012 Analysis of active chromatin modifications in early mammalian embryos reveals uncoupling of H2A.Z acetylation and H3K36 trimethylation from embryonic genome activation. Epigenetics 7 747–757. (https://doi.org/10.4161/epi.20584)
Boucret L, de la Barca JMC, Moriniere C, Desquiret V, Ferre-L’Hotellier V, Descamps P, Marcaillou C, Reynier P, Procaccio V & May-Panloup P 2015 Relationship between diminished ovarian reserve and mitochondrial biogenesis in cumulus cells. Human Reproduction 30 1653–1664. (https://doi.org/10.1093/humrep/dev114)
Bowles EJ, Lee JH, Alberio R, Lloyd RE, Stekel D, Campbell KH & St John JC 2007 Contrasting effects of in vitro fertilization and nuclear transfer on the expression of mtDNA replication factors. Genetics 176 1511–1526. (https://doi.org/10.1534/genetics.106.070177)
Brenner CA, Barritt JA, Willadsen S & Cohen J 2000 Mitochondrial DNA heteroplasmy after human ooplasmic transplantation. Fertility and Sterility 74 573–578. (https://doi.org/10.1016/S0015-0282(00)00681-6)
Brown HM, Tan T & Thompson JG 2015 Metaboloepigenetics: providing alternate hypotheses for regulation of gene expression in the early embryo. Animal Reproduction 12 437–443.
Canovas S, Cibelli JB & Ross PJ 2012 Jumonji domain-containing protein 3 regulates histone 3 lysine 27 methylation during bovine preimplantation development. PNAS 109 2400–2405. (https://doi.org/10.1073/pnas.1119112109)
Canto C, Menzies KJ & Auwerx J 2015 NAD(+) metabolism and the control of energy homeostasis: a balancing act between mitochondria and the nucleus. Cell Metabolism 22 31–53. (https://doi.org/10.1016/j.cmet.2015.05.023)
Carey BW, Finley LW, Cross JR, Allis CD & Thompson CB 2015 Intracellular alpha-ketoglutarate maintains the pluripotency of embryonic stem cells. Nature 518 413–416. (https://doi.org/10.1038/nature13981)
Chandel NS 2014 Mitochondria as signaling organelles. BMC Biology 12 34. (https://doi.org/10.1186/1741-7007-12-34)
Chen H & Chan DC 2004 Mitochondrial dynamics in mammals. Current Topics in Developmental Biology 59 119–144. (https://doi.org/10.1016/S0070-2153(04)59005-1)
Chen H, Detmer SA, Ewald AJ, Griffin EE, Fraser SE & Chan DC 2003 Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. Journal of Cell Biology 160 189–200. (https://doi.org/10.1083/jcb.200211046)
Chen H, Chomyn A & Chan DC 2005 Disruption of fusion results in mitochondrial heterogeneity and dysfunction. Journal of Biological Chemistry 280 26185–26192. (https://doi.org/10.1074/jbc.M503062200)
Chi MM, Manchester JK, Yang VC, Curato AD, Strickler RC & Lowry OH 1988 Contrast in levels of metabolic enzymes in human and mouse ova. Biology of Reproduction 39 295–307. (https://doi.org/10.1095/biolreprod39.2.295)
Chinnery PF, Howell N, Lightowlers RN & Turnbull DM 1997 Molecular pathology of MELAS and MERRF. The relationship between mutation load and clinical phenotypes. Brain 120 (Pt 10) 1713–1721. (https://doi.org/10.1093/brain/120.10.1713)
Cho YM, Kwon S, Pak YK, Seol HW, Choi YM, Park DJ, Park KS & Lee HK 2006 Dynamic changes in mitochondrial biogenesis and antioxidant enzymes during the spontaneous differentiation of human embryonic stem cells. Biochemical and Biophysical Research Communications 348 1472–1478. (https://doi.org/10.1016/j.bbrc.2006.08.020)
Choi YS, Hoon Jeong J, Min HK, Jung HJ, Hwang D, Lee SW & Kim Pak Y 2011 Shot-gun proteomic analysis of mitochondrial D-loop DNA binding proteins: identification of mitochondrial histones. Molecular Biosystems 7 1523–1536. (https://doi.org/10.1039/c0mb00277a)
Chung S, Dzeja PP, Faustino RS, Perez-Terzic C, Behfar A & Terzic A 2007 Mitochondrial oxidative metabolism is required for the cardiac differentiation of stem cells. Nature Clinical Practice Cardiovascular Medicine 4 (Supplement 1) S60–S67. (https://doi.org/10.1038/ncpcardio0766)
Chung N, Bogliotti YS, Ding W, Vilarino M, Takahashi K, Chitwood JL, Schultz RM & Ross PJ 2017 Active H3K27me3 demethylation by KDM6B is required for normal development of bovine preimplantation embryos. Epigenetics 12 1048–1056. (https://doi.org/10.1080/15592294.2017.1403693)
Clayton DA 2000 Transcription and replication of mitochondrial DNA. Human Reproduction 15 (Supplement 2) 11–17. (https://doi.org/10.1093/humrep/15.suppl_2.11)
Cogliati S, Frezza C, Soriano ME, Varanita T, Quintana-Cabrera R, Corrado M, Cipolat S, Costa V, Casarin A & Gomes LC et al. 2013 Mitochondrial cristae shape determines respiratory chain supercomplexes assembly and respiratory efficiency. Cell 155 160–171. (https://doi.org/10.1016/j.cell.2013.08.032)
Conaghan J, Handyside AH, Winston RM & Leese HJ 1993 Effects of pyruvate and glucose on the development of human preimplantation embryos in vitro. Journal of Reproduction and Fertility 99 87–95. (https://doi.org/10.1530/jrf.0.0990087)
Cordeiro FB, Montani DA, Pilau EJ, Gozzo FC, Fraietta R & Turco EGL 2018 Ovarian environment aging: follicular fluid lipidomic and related metabolic pathways. Journal of Assisted Reproduction and Genetics 35 1385–1393. (https://doi.org/10.1007/s10815-018-1259-5)
Cree LM, Samuels DC, de Sousa Lopes SC, Rajasimha HK, Wonnapinij P, Mann JR, Dahl HH & Chinnery PF 2008 A reduction of mitochondrial DNA molecules during embryogenesis explains the rapid segregation of genotypes. Nature Genetics 40 249–254. (https://doi.org/10.1038/ng.2007.63)
Cree LM, Hammond ER, Shelling AN, Berg MC, Peek JC & Green MP 2015 Maternal age and ovarian stimulation independently affect oocyte mtDNA copy number and cumulus cell gene expression in bovine clones. Human Reproduction 30 1410–1420. (https://doi.org/10.1093/humrep/dev066)
Davies VJ, Hollins AJ, Piechota MJ, Yip W, Davies JR, White KE, Nicols PP, Boulton ME & Votruba M 2007 Opa1 deficiency in a mouse model of autosomal dominant optic atrophy impairs mitochondrial morphology, optic nerve structure and visual function. Human Molecular Genetics 16 1307–1318. (https://doi.org/10.1093/hmg/ddm079)
de Boar KA, Jansen RPS, Leigh DA & Mortimer D 1999 Quantification of mtDNA copy number in the human secondary oocyte. Human Reproduction 14 2419. (https://doi.org/10.1093/humrep/14.9.2419a)
de Los Santos MJ, Diez Juan A, Mifsud A, Mercader A, Meseguer M, Rubio C & Pellicer A 2018 Variables associated with mitochondrial copy number in human blastocysts: what can we learn from trophectoderm biopsies? Fertility and Sterility 109 110–117. (https://doi.org/10.1016/j.fertnstert.2017.09.022)
Dell’Aquila ME, Ambruosi B, De Santis T & Cho YS 2009 Mitochondrial distribution and activity in human mature oocytes: gonadotropin-releasing hormone agonist versus antagonist for pituitary down-regulation. Fertility and Sterility 91 249–255. (https://doi.org/10.1016/j.fertnstert.2007.10.042)
Desquiret-Dumas V, Clement A, Seegers V, Boucret L, Ferre-L’Hotellier V, Bouet PE, Descamps P, Procaccio V, Reynier P & May-Panloup P 2017 The mitochondrial DNA content of cumulus granulosa cells is linked to embryo quality. Human Reproduction 32 607–614. (https://doi.org/10.1093/humrep/dew341)
Diez-Juan A, Rubio C, Marin C, Martinez S, Al-Asmar N, Riboldi M, Diaz-Gimeno P, Valbuena D & Simon C 2015 Mitochondrial DNA content as a viability score in human euploid embryos: less is better. Fertility and Sterility 104 534.e531–541.e531. (https://doi.org/10.1016/j.fertnstert.2015.05.022)
Ding YB, Long CL, Liu XQ, Chen XM, Guo LR, Xia YY, He JL & Wang YX 2012 5-aza-2’-deoxycytidine leads to reduced embryo implantation and reduced expression of DNA methyltransferases and essential endometrial genes. PLoS One 7 e45364. (https://doi.org/10.1371/journal.pone.0045364)
Donohoe DR & Bultman SJ 2012 Metaboloepigenetics: interrelationships between energy metabolism and epigenetic control of gene expression. Journal of Cellular Physiology 227 3169–3177. (https://doi.org/10.1002/jcp.24054)
Duchen MR 2000 Mitochondria and calcium: from cell signalling to cell death. Journal of Physiology 529 (Pt 1) 57–68. (https://doi.org/10.1111/j.1469-7793.2000.00057.x)
Dumesic DA, Meldrum DR, Katz-Jaffe MG, Krisher RL & Schoolcraft WB 2015 Oocyte environment: follicular fluid and cumulus cells are critical for oocyte health. Fertility and Sterility 103 303–316. (https://doi.org/10.1016/j.fertnstert.2014.11.015)
Dumollard R, Marangos P, Fitzharris G, Swann K, Duchen M & Carroll J 2004 Sperm-triggered. (Ca2+) oscillations and Ca2+ homeostasis in the mouse egg have an absolute requirement for mitochondrial ATP production. Development 131 3057–3067. (https://doi.org/10.1242/dev.01181)
Dumollard R, Ward Z, Carroll J & Duchen MR 2007 Regulation of redox metabolism in the mouse oocyte and embryo. Development 134 455–465. (https://doi.org/10.1242/dev.02744)
Finkelstein JD 1990 Methionine metabolism in mammals. Journal of Nutritional Biochemistry 1 228–237. (https://doi.org/10.1016/0955-2863(90)90070-2)
Folmes CD & Terzic A 2014 Metabolic determinants of embryonic development and stem cell fate. Reproduction, Fertility, and Development 27 82–88. (https://doi.org/10.1071/RD14383)
Fragouli E, Spath K, Alfarawati S, Kaper F, Craig A, Michel CE, Kokocinski F, Cohen J, Munne S & Wells D 2015 Altered levels of mitochondrial DNA are associated with female age, aneuploidy, and provide an independent measure of embryonic implantation potential. PLOS Genetics 11 e1005241. (https://doi.org/10.1371/journal.pgen.1005241)
Frederick RL & Shaw JM 2007 Moving mitochondria: establishing distribution of an essential organelle. Traffic. Traffic 8 1668–1675. (https://doi.org/10.1111/j.1600-0854.2007.00644.x)
Fu D, Mitra K, Sengupta P, Jarnik M, Lippincott-Schwartz J & Arias IM 2013 Coordinated elevation of mitochondrial oxidative phosphorylation and autophagy help drive hepatocyte polarization. PNAS 110 7288–7293. (https://doi.org/10.1073/pnas.1304285110)
Fulka H 2004 Distribution of mitochondria in reconstructed mouse oocytes. Reproduction 127 195–200. (https://doi.org/10.1530/rep.1.00093)
Gao F, Ma X, Rusie A, Hemingway J, Ostmann AB, Chung D & Das SK 2012 Epigenetic changes through DNA methylation contribute to uterine stromal cell decidualization. Endocrinology 153 6078–6090. (https://doi.org/10.1210/en.2012-1457)
Garcia I, Innis-Whitehouse W, Lopez A, Keniry M & Gilkerson R 2018 Oxidative insults disrupt OPA1-mediated mitochondrial dynamics in cultured mammalian cells. Redox Report 23 160–167. (https://doi.org/10.1080/13510002.2018.1492766)
Gardner DK 1998 Changes in requirements and utilization of nutrients during mammalian preimplantation embryo development and their significance in embryo culture. Theriogenology 49 83–102. (https://doi.org/10.1016/S0093-691X(97)00404-4)
Gardner DK & Leese HJ 1987. Assessment of embryo viability prior to transfer by the noninvasive measurement of glucose uptake. Journal of Experimental Zoology 242 103–105.
Gardner DK & Wale PL 2013 Analysis of metabolism to select viable human embryos for transfer. Fertility and Sterility 99 1062–1072. (https://doi.org/10.1016/j.fertnstert.2012.12.004)
Gardner DK & Harvey AJ 2015 Blastocyst metabolism. Reproduction, Fertility, and Development 27 638–654. (https://doi.org/10.1071/RD14421)
Gardner DK, Wale PL, Collins R & Lane M 2011 Glucose consumption of single post-compaction human embryos is predictive of embryo sex and live birth outcome. Human Reproduction 26 1981–1986. (https://doi.org/10.1093/humrep/der143)
Ge H, Tollner TL, Hu Z, Da M, Li X, Guan H, Shan D, Lu J, Huang C & Dong Q 2012 Impaired mitochondrial function in murine oocytes is associated with controlled ovarian hyperstimulation and in vitro maturation. Reproduction, Fertility, and Development 24 945–952. (https://doi.org/10.1071/RD11212)
Gilkerson R, Bravo L, Garcia I, Gaytan N, Herrera A, Maldonado A & Quintanilla B 2013 The mitochondrial nucleoid: integrating mitochondrial DNA into cellular homeostasis. Cold Spring Harbor Perspectives in Biology 5 a011080. (https://doi.org/10.1101/cshperspect.a011080)
Gott AL, Hardy K, Winston RML & Leese HJ 1990 Non-invasive measurement of pyruvate and glucose uptake and lactate production by single human preimplantation embryos. Human Reproduction 5 104–108. (https://doi.org/10.1093/oxfordjournals.humrep.a137028)
Gray CW, Morgan PM & Kane MT 1992 Purification of an embryotrophic factor from commercial bovine serum albumin and its identification as citrate. Journal of Reproduction and Fertility 94 471–480. (https://doi.org/10.1530/jrf.0.0940471)
Gu TP, Guo F, Yang H, Wu HP, Xu GF, Liu W, Xie ZG, Shi L, He X & Jin SG et al. 2011 The role of Tet3 DNA dioxygenase in epigenetic reprogramming by oocytes. Nature 477 606–610. (https://doi.org/10.1038/nature10443)
Hackenbrock CR 1966 Ultrastructural bases for metabolically linked mechanical activity in mitochondria. I. Reversible ultrastructural changes with change in metabolic steady state in isolated liver mitochondria. Journal of Cell Biology 30 269–297. (https://doi.org/10.1083/jcb.30.2.269)
Hales KG & Fuller MT 1997 Developmentally regulated mitochondrial fusion mediated by a conserved, novel, predicted GTPase. Cell 90 121–129. (https://doi.org/10.1016/S0092-8674(00)80319-0)
Hamanaka RB & Chandel NS 2010 Mitochondrial reactive oxygen species regulate cellular signaling and dictate biological outcomes. Trends in Biochemical Sciences 35 505–513. (https://doi.org/10.1016/j.tibs.2010.04.002)
Hammond ER, Green MP, Shelling AN, Berg MC, Peek JC & Cree LM 2016 Oocyte mitochondrial deletions and heteroplasmy in a bovine model of ageing and ovarian stimulation. Molecular Human Reproduction 22 261–271. (https://doi.org/10.1093/molehr/gaw003)
Hardy K, Hooper MA, Handyside AH, Rutherford AJ, Winston RM & Leese HJ 1989 Non-invasive measurement of glucose and pyruvate uptake by individual human oocytes and preimplantation embryos. Human Reproduction 4 188–191. (https://doi.org/10.1093/oxfordjournals.humrep.a136869)
Harvey AJ, Kind KL & Thompson JG 2002 REDOX regulation of early embryo development. Reproduction 123 479–486. (https://doi.org/10.1530/rep.0.1230479)
Harvey A, Gibson T, Lonergan T & Brenner C 2011 Dynamic regulation of mitochondrial function in preimplantation embryos and embryonic stem cells. Mitochondrion 11 829–838. (https://doi.org/10.1016/j.mito.2010.12.013)
Harvey AJ, Rathjen J & Gardner DK 2013 The metabolic framework of pluripotent stem cells and potential mechanisms of regulation. In Stem Cells in Reproductive Medicine: Basic Science and Therapeutic Potential, pp. 164–179. Eds Simón C, Pellicer A & Pera RR. Cambridge: Cambridge University Press. (https://doi.org/10.1017/CBO9781139540742.016)
Harvey AJ, Rathjen J & Gardner DK 2016 Metaboloepigenetic regulation of pluripotent stem cells. Stem Cells International 2016 1816525. (https://doi.org/10.1155/2016/1816525)
Herrmann JM & Riemer J 2010 The intermembrane space of mitochondria. Antioxidants and Redox Signaling 13 1341–1358. (https://doi.org/10.1089/ars.2009.3063)
Hiendleder S, Schmutz SM, Erhardt G, Green RD & Plante Y 1999 Transmitochondrial differences and varying levels of heteroplasmy in nuclear transfer cloned cattle. Molecular Reproduction and Development 54 24–31. (https://doi.org/10.1002/(SICI)1098-2795(199909)54:1<24::AID-MRD4>3.0.CO;2-S)
Hiendleder S, Zakhartchenko V, Wenigerkind H, Reichenbach HD, Bruggerhoff K, Prelle K, Brem G, Stojkovic M & Wolf E 2003 Heteroplasmy in bovine fetuses produced by intra- and inter-subspecific somatic cell nuclear transfer: neutral segregation of nuclear donor mitochondrial DNA in various tissues and evidence for recipient cow mitochondria in fetal blood. Biology of Reproduction 68 159–166. (https://doi.org/10.1095/biolreprod.102.008201)
Hitchler MJ & Domann FE 2009 Metabolic defects provide a spark for the epigenetic switch in cancer. Free Radical Biology and Medicine 47 115–127. (https://doi.org/10.1016/j.freeradbiomed.2009.04.010)
Ho JR, Arrach N, Rhodes-Long K, Salem W, McGinnis LK, Chung K, Bendikson KA, Paulson RJ & Ahmady A 2018 Blastulation timing is associated with differential mitochondrial content in euploid embryos. Journal of Assisted Reproduction and Genetics 35 711–720. (https://doi.org/10.1007/s10815-018-1113-9)
Houghton FD, Thompson JG, Kennedy CJ & Leese HJ 1996 Oxygen consumption and energy metabolism of the early mouse embryo. Molecular Reproduction and Development 44 476–485. (https://doi.org/10.1002/(SICI)1098-2795(199608)44:4<476::AID-MRD7>3.0.CO;2-I)
Houghton FD, Hawkhead JA, Humpherson PG, Hogg JE, Balen AH, Rutherford AJ & Leese HJ 2002 Non-invasive amino acid turnover predicts human embryo developmental capacity. Human Reproduction 17 999–1005. (https://doi.org/10.1093/humrep/17.4.999)
Houtkooper RH, Pirinen E & Auwerx J 2012 Sirtuins as regulators of metabolism and healthspan. Nature Reviews Molecular Cell Biology 13 225–238. (https://doi.org/10.1038/nrm3293)
Hsieh RH, Tsai NM, Au HK, Chang SJ, Wei YH & Tzeng CR 2002 Multiple rearrangements of mitochondrial DNA in unfertilized human oocytes. Fertility and Sterility 77 1012–1017. (https://doi.org/10.1016/S0015-0282(02)02994-1)
Hsieh RH, Au HK, Yeh TS, Chang SJ, Cheng YF & Tzeng CR 2004 Decreased expression of mitochondrial genes in human unfertilized oocytes and arrested embryos. Fertility and Sterility 81 912–918. (https://doi.org/10.1016/j.fertnstert.2003.11.013)
Huang J, Zhang H, Wang X, Dobbs KB, Yao J, Qin G, Whitworth K, Walters EM, Prather RS & Zhao J 2015 Impairment of preimplantation porcine embryo development by histone demethylase KDM5B knockdown through disturbance of bivalent H3K4me3-H3K27me3 modifications. Biology of Reproduction 92 72. (https://doi.org/10.1095/biolreprod.114.122762)
Igosheva N, Abramov AY, Poston L, Eckert JJ, Fleming TP, Duchen MR & McConnell J 2010 Maternal diet-induced obesity alters mitochondrial activity and redox status in mouse oocytes and zygotes. PLoS One 5 e10074. (https://doi.org/10.1371/journal.pone.0010074)
Ikeda S, Sugimoto M & Kume S 2012 Importance of methionine metabolism in morula-to-blastocyst transition in bovine preimplantation embryos. Journal of Reproduction and Development 58 91–97. (https://doi.org/10.1262/jrd.11-096H)
Ikeda S, Kawahara-Miki R, Iwata H, Sugimoto M & Kume S 2017 Role of methionine adenosyltransferase 2A in bovine preimplantation development and its associated genomic regions. Scientific Reports 7 3800. (https://doi.org/10.1038/s41598-017-04003-1)
Ikon N & Ryan RO 2017 Cardiolipin and mitochondrial cristae organization. Biochimica and Biophysica Acta 1859 1156–1163. (https://doi.org/10.1016/j.bbamem.2017.03.013)
Inoue A & Zhang Y 2011 Replication-dependent loss of 5-hydroxymethylcytosine in mouse preimplantation embryos. Science 334 194. (https://doi.org/10.1126/science.1212483)
Iqbal K, Jin SG, Pfeifer GP & Szabo PE 2011 Reprogramming of the paternal genome upon fertilization involves genome-wide oxidation of 5-methylcytosine. PNAS 108 3642–3647. (https://doi.org/10.1073/pnas.1014033108)
Ishihara N, Jofuku A, Eura Y & Mihara K 2003 Regulation of mitochondrial morphology by membrane potential, and DRP1-dependent division and FZO1-dependent fusion reaction in mammalian cells. Biochemical and Biophysical Research Communications 301 891–898. (https://doi.org/10.1016/S0006-291X(03)00050-0)
Ishihara N, Nomura M, Jofuku A, Kato H, Suzuki SO, Masuda K, Otera H, Nakanishi Y, Nonaka I & Goto Y et al. 2009 Mitochondrial fission factor Drp1 is essential for embryonic development and synapse formation in mice. Nature Cell Biology 11 958–966. (https://doi.org/10.1038/ncb1907)
Ito M, Sakai S, Nagata M & Aoki F 2000 Effect of histone deacetylase inhibitors on early preimplantation development in mouse embryo. Journal of Mammalian Ova Research 17 90–95. (https://doi.org/10.1274/jmor.17.90)
Ito S, D’Alessio AC, Taranova OV, Hong K, Sowers LC & Zhang Y 2010 Role of Tet proteins in 5mC to 5hmC conversion, ES-cell self-renewal and inner cell mass specification. Nature 466 1129–1133. (https://doi.org/10.1038/nature09303)
Jacobs L, Gerards M, Chinnery P, Dumoulin J, de Coo I, Geraedts J & Smeets H 2007 mtDNA point mutations are present at various levels of heteroplasmy in human oocytes. Molecular Human Reproduction 13 149–154. (https://doi.org/10.1093/molehr/gal112)
Jansen RP & de Boer K 1998 The bottleneck: mitochondrial imperatives in oogenesis and ovarian follicular fate. Molecular and Cellular Endocrinology 145 81–88. (https://doi.org/10.1016/S0303-7207(98)00173-7)
Jovaisaite V, Mouchiroud L & Auwerx J 2014 The mitochondrial unfolded protein response, a conserved stress response pathway with implications in health and disease. Journal of Experimental Biology 217 137–143. (https://doi.org/10.1242/jeb.090738)
Kawamura Y, Uchijima Y, Horike N, Tonami K, Nishiyama K, Amano T, Asano T, Kurihara Y & Kurihara H 2010 Sirt3 protects in vitro-fertilized mouse preimplantation embryos against oxidative stress-induced p53-mediated developmental arrest. Journal of Clinical Investigation 120 2817–2828. (https://doi.org/10.1172/JCI42020)
Keefe DL, Niven-Fairchild T, Powell S & Buradagunta S 1995 Mitochondrial deoxyribonucleic acid deletions in oocytes and reproductive aging in women. Fertility and Sterility 64 577–583. (https://doi.org/10.1016/S0015-0282(16)57796-6)
Klimczak AM, Pacheco LE, Lewis KE, Massahi N, Richards JP, Kearns WG, Saad AF & Crochet JR 2018 Embryonal mitochondrial DNA: relationship to embryo quality and transfer outcomes. Journal of Assisted Reproduction and Genetics 35 871–877. (https://doi.org/10.1007/s10815-018-1147-z)
Kline D & Kline JT 1992 Repetitive calcium transients and the role of calcium in exocytosis and cell cycle activation in the mouse egg. Developmental Biology 149 80–89. (https://doi.org/10.1016/0012-1606(92)90265-I)
Kudo M, Ikeda S, Sugimoto M & Kume S 2015 Methionine-dependent histone methylation at developmentally important gene loci in mouse preimplantation embryos. Journal of Nutritional Biochemistry 26 1664–1669. (https://doi.org/10.1016/j.jnutbio.2015.08.009)
Kwak SS, Cheong SA, Yoon JD, Jeon Y & Hyun SH 2012 Expression patterns of sirtuin genes in porcine preimplantation embryos and effects of sirtuin inhibitors on in vitro embryonic development after parthenogenetic activation and in vitro fertilization. Theriogenology 78 1597–1610. (https://doi.org/10.1016/j.theriogenology.2012.07.006)
Kwong WY, Adamiak SJ, Gwynn A, Singh R & Sinclair KD 2010 Endogenous folates and single-carbon metabolism in the ovarian follicle, oocyte and pre-implantation embryo. Reproduction 139 705–715. (https://doi.org/10.1530/REP-09-0517)
Lane M & Gardner DK 2005 Mitochondrial malate-aspartate shuttle regulates mouse embryo nutrient consumption. Journal of Biological Chemistry 280 18361–18367. (https://doi.org/10.1074/jbc.M500174200)
Lanzavecchia G & Mangioni C 1964 Etude de la structure et des constituants du follicule humain dans l’ovaire foetal. Journal de Microscopie 3 447–464.
Lawrence EJ, Boucher E & Mandato CA 2016 Mitochondria-cytoskeleton associations in mammalian cytokinesis. Cell Division 11 3. (https://doi.org/10.1186/s13008-016-0015-4)
Lee JY, Kapur M, Li M, Choi MC, Choi S, Kim HJ, Kim I, Lee E, Taylor JP & Yao TP 2014a MFN1 deacetylation activates adaptive mitochondrial fusion and protects metabolically challenged mitochondria. Journal of Cell Science 127 4954–4963. (https://doi.org/10.1242/jcs.157321)
Lee K, Hamm J, Whitworth K, Spate L, Park KW, Murphy CN & Prather RS 2014b Dynamics of TET family expression in porcine preimplantation embryos is related to zygotic genome activation and required for the maintenance of NANOG. Developmental Biology 386 86–95. (https://doi.org/10.1016/j.ydbio.2013.11.024)
Lees JG, Gardner DK & Harvey AJ 2017 Pluripotent stem cell metabolism and mitochondria: beyond ATP. Stem Cells International 2017 2874283. (https://doi.org/10.1155/2017/2874283)
Leese HJ, Biggers JD, Mroz EA & Lechene C 1984 Nucleotides in a single mammalian ovum or preimplantation embryo. Analytical Biochemistry 140 443–448. (https://doi.org/10.1016/0003-2697(84)90191-X)
Leese HJ, Hooper MA, Edwards RG & Ashwood-Smith MJ 1986 Uptake of pyruvate by early human embryos determined by a non-invasive technique. Human Reproduction 1 181–182. (https://doi.org/10.1093/oxfordjournals.humrep.a136376)
Liu Q, Kang L, Wang L, Zhang L & Xiang W 2016a Mitofusin 2 regulates the oocytes development and quality by modulating meiosis and mitochondrial function. Scientific Reports 6 30561. (https://doi.org/10.1038/srep30561)
Liu X, Wang C, Liu W, Li J, Li C, Kou X, Chen J, Zhao Y, Gao H & Wang H et al. 2016b Distinct features of H3K4me3 and H3K27me3 chromatin domains in pre-implantation embryos. Nature 537 558–562. (https://doi.org/10.1038/nature19362)
Lizana L, Bauer B & Orwar O 2008 Controlling the rates of biochemical reactions and signaling networks by shape and volume changes. PNAS 105 4099–4104. (https://doi.org/10.1073/pnas.0709932105)
Lozoya OA, Martinez-Reyes I, Wang T, Grenet D, Bushel P, Li J, Chandel N, Woychik RP & Santos JH 2018 Mitochondrial nicotinamide adenine dinucleotide reduced (NADH) oxidation links the tricarboxylic acid (TCA) cycle with methionine metabolism and nuclear DNA methylation. PLOS Biology 16 e2005707. (https://doi.org/10.1371/journal.pbio.2005707)
Lu M, Zhou L, Stanley WC, Cabrera ME, Saidel GM & Yu X 2008 Role of the malate-aspartate shuttle on the metabolic response to myocardial ischemia. Journal of Theoretical Biology 254 466–475. (https://doi.org/10.1016/j.jtbi.2008.05.033)
Lucas E 2013 Epigenetic effects on the embryo as a result of periconceptional environment and assisted reproduction technology. Reproductive Biomedicine Online 27 477–485. (https://doi.org/10.1016/j.rbmo.2013.06.003)
Ma P & Schultz RM 2008 Histone deacetylase 1 (HDAC1) regulates histone acetylation, development, and gene expression in preimplantation mouse embryos. Developmental Biology 319 110–120. (https://doi.org/10.1016/j.ydbio.2008.04.011)
Ma H, Folmes CD, Wu J, Morey R, Mora-Castilla S, Ocampo A, Ma L, Poulton J, Wang X & Ahmed R et al. 2015 Metabolic rescue in pluripotent cells from patients with mtDNA disease. Nature 524 234–238. (https://doi.org/10.1038/nature14546)
Mandal S, Lindgren AG, Srivastava AS, Clark AT & Banerjee U 2011 Mitochondrial function controls proliferation and early differentiation potential of embryonic stem cells. Stem Cells 29 486–495. (https://doi.org/10.1002/stem.590)
Manosalva I & Gonzalez A 2009 Aging alters histone H4 acetylation and CDC2A in mouse germinal vesicle stage oocytes. Biology of Reproduction 81 1164–1171. (https://doi.org/10.1095/biolreprod.109.078386)
Manosalva I & Gonzalez A 2010 Aging changes the chromatin configuration and histone methylation of mouse oocytes at germinal vesicle stage. Theriogenology 74 1539–1547. (https://doi.org/10.1016/j.theriogenology.2010.06.024)
Manzanares M & Rodriguez TA 2013 Development: Hippo signalling turns the embryo inside out. Current Biology 23 R559–R561. (https://doi.org/10.1016/j.cub.2013.05.064)
Marchi S, Patergnani S & Pinton P 2014 The endoplasmic reticulum-mitochondria connection: one touch, multiple functions. Biochimica and Biophysica Acta 1837 461–469. (https://doi.org/10.1016/j.bbabio.2013.10.015)
Marchlewicz EH, Dolinoy DC, Tang L, Milewski S, Jones TR, Goodrich JM, Soni T, Domino SE, Song PX & Burant CF et al. 2016 Lipid metabolism is associated with developmental epigenetic programming. Scientific Reports 6 34857. (https://doi.org/10.1038/srep34857)
Marcho C, Cui W & Mager J 2015 Epigenetic dynamics during preimplantation development. Reproduction 150 R109–R120. (https://doi.org/10.1530/REP-15-0180)
Mason K, Liu Z, Aguirre-Lavin T & Beaujean N 2012 Chromatin and epigenetic modifications during early mammalian development. Animal Reproduction Science 134 45–55. (https://doi.org/10.1016/j.anireprosci.2012.08.010)
Mayer W, Niveleau A, Walter J, Fundele R & Haaf T 2000 Demethylation of the zygotic paternal genome. Nature 403 501–502. (https://doi.org/10.1038/35000654)
May-Panloup P, Chretien MF, Jacques C, Vasseur C, Malthiery Y & Reynier P 2005a Low oocyte mitochondrial DNA content in ovarian insufficiency. Human Reproduction 20 593–597. (https://doi.org/10.1093/humrep/deh667)
May-Panloup P, Vignon X, Chretien MF, Heyman Y, Tamassia M, Malthiery Y & Reynier P 2005b Increase of mitochondrial DNA content and transcripts in early bovine embryogenesis associated with upregulation of mtTFA and NRF1 transcription factors. Reproductive Biology and Endocrinology 3 65. (https://doi.org/10.1186/1477-7827-3-65)
McDonnell E, Crown SB, Fox DB, Kitir B, Ilkayeva OR, Olsen CA, Grimsrud PA & Hirschey MD 2016 Lipids reprogram metabolism to become a major carbon source for histone acetylation. Cell Reports 17 1463–1472. (https://doi.org/10.1016/j.celrep.2016.10.012)
McReynolds S, Dzieciatkowska M, McCallie BR, Mitchell SD, Stevens J, Hansen K, Schoolcraft WB & Katz-Jaffe MG 2012 Impact of maternal aging on the molecular signature of human cumulus cells. Fertility and Sterility 98 1574–80.e5. (https://doi.org/10.1016/j.fertnstert.2012.08.012)
Menezo Y, Khatchadourian C, Gharib A, Hamidi J, Greenland T & Sarda N 1989 Regulation of S-adenosyl methionine synthesis in the mouse embryo. Life Sciences 44 1601–1609. (https://doi.org/10.1016/0024-3205(89)90455-4)
Messerschmidt DM, Knowles BB & Solter D 2014 DNA methylation dynamics during epigenetic reprogramming in the germline and preimplantation embryos. Genes and Development 28 812–828. (https://doi.org/10.1101/gad.234294.113)
Mills RM & Brinster RL 1967 Oxygen consumption of pre-implantation mouse embryos. Experimental Cell Research 47 337–344. (https://doi.org/10.1016/0014-4827(67)90236-4)
Mitchell M, Cashman KS, Gardner DK, Thompson JG & Lane M 2009 Disruption of mitochondrial malate-aspartate shuttle activity in mouse blastocysts impairs viability and fetal growth. Biology of Reproduction 80 295–301. (https://doi.org/10.1095/biolreprod.108.069864)
Mohr LR & Trounson AO 1982 Comparative ultrastructure of hatched human, mouse and bovine blastocysts. Journal of Reproduction and Fertility 66 499–504. (https://doi.org/10.1530/jrf.0.0660499)
Monk M, Boubelik M & Lehnert S 1987 Temporal and regional changes in DNA methylation in the embryonic, extraembryonic and germ cell lineages during mouse embryo development. Development 99 371–382.
Motta PM, Nottola SA, Makabe S & Heyn R 2000 Mitochondrial morphology in human fetal and adult female germ cells. Human Reproduction 15 (Supplement 2) 129–147. (https://doi.org/10.1093/humrep/15.suppl_2.129)
Moussaieff A, Rouleau M, Kitsberg D, Cohen M, Levy G, Barasch D, Nemirovski A, Shen-Orr S, Laevsky I & Amit M et al. 2015 Glycolysis-mediated changes in acetyl-CoA and histone acetylation control the early differentiation of embryonic stem cells. Cell Metabolism 21 392–402. (https://doi.org/10.1016/j.cmet.2015.02.002)
Mtango NR, Harvey AJ, Latham KE & Brenner CA 2008 Molecular control of mitochondrial function in developing rhesus monkey oocytes and preimplantation-stage embryos. Reproduction, Fertility, and Development 20 846–859. (https://doi.org/10.1071/RD08078)
Murakoshi Y, Sueoka K, Takahashi K, Sato S, Sakurai T, Tajima H & Yoshimura Y 2013 Embryo developmental capability and pregnancy outcome are related to the mitochondrial DNA copy number and ooplasmic volume. Journal of Assisted Reproduction and Genetics 30 1367–1375. (https://doi.org/10.1007/s10815-013-0062-6)
Nadijcka M & Hillman N 1974 Ultrastructural studies of the mouse blastocyst substages. Journal of Embryology and Experimental Morphology 32 675–695.
Nagano M, Katagiri S & Takahashi Y 2006 ATP content and maturational/developmental ability of bovine oocytes with various cytoplasmic morphologies. Zygote 14 299–304. (https://doi.org/10.1017/S0967199406003807)
Nagaraj R, Gururaja-Rao S, Jones KT, Slattery M, Negre N, Braas D, Christofk H, White KP, Mann R & Banerjee U 2012 Control of mitochondrial structure and function by the Yorkie/YAP oncogenic pathway. Genes and Development 26 2027–2037. (https://doi.org/10.1101/gad.183061.111)
Nagaraj R, Sharpley MS, Chi F, Braas D, Zhou Y, Kim R, Clark AT & Banerjee U 2017 Nuclear localization of mitochondrial TCA cycle enzymes as a critical step in mammalian zygotic genome activation. Cell 168 210–223.e11. (https://doi.org/10.1016/j.cell.2016.12.026)
Nakamura T, Liu YJ, Nakashima H, Umehara H, Inoue K, Matoba S, Tachibana M, Ogura A, Shinkai Y & Nakano T 2012 PGC7 binds histone H3K9me2 to protect against conversion of 5mC to 5hmC in early embryos. Nature 486 415–419. (https://doi.org/10.1038/nature11093)
Norton M, Ng AC, Baird S, Dumoulin A, Shutt T, Mah N, Andrade-Navarro MA, McBride HM & Screaton RA 2014 ROMO1 is an essential redox-dependent regulator of mitochondrial dynamics. Science Signaling 7 ra10. (https://doi.org/10.1126/scisignal.2004374)
Noto V, Campo R, Roziers P, Swinnen K, Vercruyssen M & Gordts S 1993 Fertilization and early embryology: mitochondrial distribution after fast embryo freezing. Human Reproduction 8 2115–2118. (https://doi.org/10.1093/oxfordjournals.humrep.a137992)
Ogino M, Tsubamoto H, Sakata K, Oohama N, Hayakawa H, Kojima T, Shigeta M & Shibahara H 2016 Mitochondrial DNA copy number in cumulus cells is a strong predictor of obtaining good-quality embryos after IVF. Journal of Assisted Reproduction and Genetics 33 367–371. (https://doi.org/10.1007/s10815-015-0621-0)
O’Gorman A, Wallace M, Cottell E, Gibney MJ, McAuliffe FM, Wingfield M & Brennan L 2013 Metabolic profiling of human follicular fluid identifies potential biomarkers of oocyte developmental competence. Reproduction 146 389–395. (https://doi.org/10.1530/REP-13-0184)
Olichon A, Baricault L, Gas N, Guillou E, Valette A, Belenguer P & Lenaers G 2003 Loss of OPA1 perturbates the mitochondrial inner membrane structure and integrity, leading to cytochrome c release and apoptosis. Journal of Biological Chemistry 278 7743–7746. (https://doi.org/10.1074/jbc.C200677200)
O’Neill C 1998 Endogenous folic acid is essential for normal development of preimplantation embryos. Human Reproduction 13 1312–1316. (https://doi.org/10.1093/humrep/13.5.1312)
Oswald J, Engemann S, Lane N, Mayer W, Olek A, Fundele R, Dean W, Reik W & Walter J 2000 Active demethylation of the paternal genome in the mouse zygote. Current Biology 10 475–478. (https://doi.org/10.1016/S0960-9822(00)00448-6)
Otasevic V, Surlan L, Vucetic M, Tulic I, Buzadzic B, Stancic A, Jankovic A, Velickovic K, Golic I & Markelic M et al. 2016 Expression patterns of mitochondrial OXPHOS components, mitofusin 1 and dynamin-related protein 1 are associated with human embryo fragmentation. Reproduction, Fertility, and Development 28 319–327. (https://doi.org/10.1071/RD13415)
Otten ABC, Sallevelt SCEH, Carling PJ, Dreesen JCFM, Drüsedau M, Spierts S, Paulussen ADC, de Die-Smulders CEM, Herbert M & Chinnery PF et al. 2018 Mutation-specific effects in germline transmission of pathogenic mtDNA variants. Human Reproduction 33 1331–1341. (https://doi.org/10.1093/humrep/dey114)
Parone P, Priault M, James D, Nothwehr SF & Martinou JC 2003 Apoptosis: bombarding the mitochondria. Essays in Biochemistry 39 41–51. (https://doi.org/10.1042/bse0390041)
Patten DA, Wong J, Khacho M, Soubannier V, Mailloux RJ, Pilon-Larose K, MacLaurin JG, Park DS, McBride HM & Trinkle-Mulcahy L et al. 2014 OPA1-dependent cristae modulation is essential for cellular adaptation to metabolic demand. EMBO Journal 33 2676–2691. (https://doi.org/10.15252/embj.201488349)
Penagaricano F, Souza AH, Carvalho PD, Driver AM, Gambra R, Kropp J, Hackbart KS, Luchini D, Shaver RD & Wiltbank MC et al. 2013 Effect of maternal methionine supplementation on the transcriptome of bovine preimplantation embryos. PLoS One 8 e72302. (https://doi.org/10.1371/journal.pone.0072302)
Pereira SL, Graos M, Rodrigues AS, Anjo SI, Carvalho RA, Oliveira PJ, Arenas E & Ramalho-Santos J 2013 Inhibition of mitochondrial complex III blocks neuronal differentiation and maintains embryonic stem cell pluripotency. PLoS One 8 e82095. (https://doi.org/10.1371/journal.pone.0082095)
Piko L & Matsumoto L 1976 Number of mitochondria and some properties of mitochondrial DNA in the mouse egg. Developmental Biology 49 1–10. (https://doi.org/10.1016/0012-1606(76)90253-0)
Piko L & Taylor KD 1987 Amounts of mitochondrial DNA and abundance of some mitochondrial gene transcripts in early mouse embryos. Developmental Biology 123 364–374. (https://doi.org/10.1016/0012-1606(87)90395-2)
Pollack Y, Kasir J, Shemer R, Metzger S & Szyf M 1984 Methylation pattern of mouse mitochondrial DNA. Nucleic Acids Research 12 4811–4824. (https://doi.org/10.1093/nar/12.12.4811)
Porras CA & Bai Y 2015 Respiratory supercomplexes: plasticity and implications. Frontiers in Bioscience 20 621–634. (https://doi.org/10.2741/4327)
Qian W, Choi S, Gibson GA, Watkins SC, Bakkenist CJ & Van Houten B 2012 Mitochondrial hyperfusion induced by loss of the fission protein Drp1 causes ATM-dependent G2/M arrest and aneuploidy through DNA replication stress. Journal of Cell Science 125 5745–5757. (https://doi.org/10.1242/jcs.109769)
Quintana-Cabrera R, Mehrotra A, Rigoni G & Soriano ME 2018 Who and how in the regulation of mitochondrial cristae shape and function. Biochemical and Biophysical Research Communications 500 94–101. (https://doi.org/10.1016/j.bbrc.2017.04.088)
Ravichandran K, McCaffrey C, Grifo J, Morales A, Perloe M, Munne S, Wells D & Fragouli E 2017 Mitochondrial DNA quantification as a tool for embryo viability assessment: retrospective analysis of data from single euploid blastocyst transfers. Human Reproduction 32 1282–1292. (https://doi.org/10.1093/humrep/dex070)
Reik W, Santos F, Mitsuya K, Morgan H & Dean W 2003 Epigenetic asymmetry in the mammalian zygote and early embryo: relationship to lineage commitment? Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences 358 1403 –1409; discussion 1409. (https://doi.org/10.1098/rstb.2003.1326)
Reis e Silva AR, Bruno C, Fleurot R, Daniel N, Archilla C, Peynot N, Lucci CM, Beaujean N & Duranthon V 2012 Alteration of DNA demethylation dynamics by in vitro culture conditions in rabbit pre-implantation embryos. Epigenetics 7 440–446. (https://doi.org/10.4161/epi.19563)
Reynier P, May-Panloup P, Chretien MF, Morgan CJ, Jean M, Savagner F, Barriere P & Malthiery Y 2001 Mitochondrial DNA content affects the fertilizability of human oocytes. Molecular Human Reproduction 7 425–429. (https://doi.org/10.1093/molehr/7.5.425)
Rizzuto R, De Stefani D, Raffaello A & Mammucari C 2012 Mitochondria as sensors and regulators of calcium signalling. Nature Reviews Molecular Cell Biology 13 566–578. (https://doi.org/10.1038/nrm3412)
Rollo C, Li Y, Jin XL & O’Neill C 2017 Histone 3 lysine 9 acetylation is a biomarker of the effects of culture on zygotes. Reproduction 154 375–385. (https://doi.org/10.1530/REP-17-0112)
Rowland AA & Voeltz GK 2012 Endoplasmic reticulum-mitochondria contacts: function of the junction. Nature Reviews Molecular Cell Biology 13 607–625. (https://doi.org/10.1038/nrm3440)
Sacconi S, Salviati L, Nishigaki Y, Walker WF, Hernandez-Rosa E, Trevisson E, Delplace S, Desnuelle C, Shanske S & Hirano M et al. 2008 A functionally dominant mitochondrial DNA mutation. Human Molecular Genetics 17 1814–1820. (https://doi.org/10.1093/hmg/ddn073)
Salilew-Wondim D, Saeed-Zidane M, Hoelker M, Gebremedhn S, Poirier M, Pandey HO, Tholen E, Neuhoff C, Held E & Besenfelder U et al. 2018 Genome-wide DNA methylation patterns of bovine blastocysts derived from in vivo embryos subjected to in vitro culture before, during or after embryonic genome activation. BMC Genomics 19 424. (https://doi.org/10.1186/s12864-018-4826-3)
Santel A & Fuller MT 2001 Control of mitochondrial morphology by a human mitofusin. Journal of Cell Science 114 867–874.
Santos F & Dean W 2004 Epigenetic reprogramming during early development in mammals. Reproduction 127 643–651. (https://doi.org/10.1530/rep.1.00221)
Santos F, Hendrich B, Reik W & Dean W 2002 Dynamic reprogramming of DNA methylation in the early mouse embryo. Developmental Biology 241 172–182. (https://doi.org/10.1006/dbio.2001.0501)
Santos F, Peters AH, Otte AP, Reik W & Dean W 2005 Dynamic chromatin modifications characterise the first cell cycle in mouse embryos. Developmental Biology 280 225–236. (https://doi.org/10.1016/j.ydbio.2005.01.025)
Santos TA, El Shourbagy S & St John JC 2006 Mitochondrial content reflects oocyte variability and fertilization outcome. Fertility and Sterility 85 584–591. (https://doi.org/10.1016/j.fertnstert.2005.09.017)
Sathananthan AH & Trounson AO 2000 Mitochondrial morphology during preimplantational human embryogenesis. Human Reproduction 15 (Supplement 2) 148–159. (https://doi.org/10.1093/humrep/15.suppl_2.148)
Sathananthan AH, Trounson AO & Wood C 1986 Atlas of Fine Structure of Sperm Penetration, Eggs and Embryos Cultured in Vitro. Philadelphia: Praeger.
Schon EA, Kim SH, Ferreira JC, Magalhaes P, Grace M, Warburton D & Gross SJ 2000 Chromosomal non-disjunction in human oocytes: is there a mitochondrial connection? Human Reproduction 15 (Supplement 2) 160–172. (https://doi.org/10.1093/humrep/15.suppl_2.160)
Seifer DB, DeJesus V & Hubbard K 2002 Mitochondrial deletions in luteinized granulosa cells as a function of age in women undergoing in vitro fertilization. Fertility and Sterility 78 1046–1048. (https://doi.org/10.1016/S0015-0282(02)04214-0)
Shao GB, Ding HM & Gong AH 2008 Role of histone methylation in zygotic genome activation in the preimplantation mouse embryo. In Vitro Cellular and Developmental Biology – Animal 44 115–120. (https://doi.org/10.1007/s11626-008-9082-4)
Sharpley MS, Marciniak C, Eckel-Mahan K, McManus M, Crimi M, Waymire K, Lin CS, Masubuchi S, Friend N & Koike M et al. 2012 Heteroplasmy of mouse mtDNA is genetically unstable and results in altered behavior and cognition. Cell 151 333–343. (https://doi.org/10.1016/j.cell.2012.09.004)
Shi L & Wu J 2009 Epigenetic regulation in mammalian preimplantation embryo development. Reproductive Biology and Endocrinology 7 59. (https://doi.org/10.1186/1477-7827-7-59)
Shiraki N, Shiraki Y, Tsuyama T, Obata F, Miura M, Nagae G, Aburatani H, Kume K, Endo F & Kume S 2014 Methionine metabolism regulates maintenance and differentiation of human pluripotent stem cells. Cell Metabolism 19 780–794. (https://doi.org/10.1016/j.cmet.2014.03.017)
Shmookler Reis RJ & Goldstein S 1983 Mitochondrial DNA in mortal and immortal human cells. Genome number, integrity, and methylation. Journal of Biological Chemistry 258 9078–9085.
Shock LS, Thakkar PV, Peterson EJ, Moran RG & Taylor SM 2011 DNA methyltransferase 1, cytosine methylation, and cytosine hydroxymethylation in mammalian mitochondria. PNAS 108 3630–3635. (https://doi.org/10.1073/pnas.1012311108)
Shojaei Saadi HA, Gagne D, Fournier É, Baldoceda Baldeon LM, Sirard MA & Robert C 2016 Responses of bovine early embryos to S-adenosyl methionine supplementation in culture. Epigenomics 8 1039–1060. (https://doi.org/10.2217/epi-2016-0022)
Shyh-Chang N, Daley GQ & Cantley LC 2013a Stem cell metabolism in tissue development and aging. Development 140 2535–2547. (https://doi.org/10.1242/dev.091777)
Shyh-Chang N, Locasale JW, Lyssiotis CA, Zheng Y, Teo RY, Ratanasirintrawoot S, Zhang J, Onder T, Unternaehrer JJ & Zhu H et al. 2013b Influence of threonine metabolism on S-adenosylmethionine and histone methylation. Science 339 222–226. (https://doi.org/10.1126/science.1226603)
Simsek-Duran F, Li F, Ford W, Swanson RJ, Jones HW Jr & Castora FJ 2013 Age-associated metabolic and morphologic changes in mitochondria of individual mouse and hamster oocytes. PLoS One 8 e64955. (https://doi.org/10.1371/journal.pone.0064955)
Sinclair KD, Allegrucci C, Singh R, Gardner DS, Sebastian S, Bispham J, Thurston A, Huntley JF, Rees WD & Maloney CA et al. 2007 DNA methylation, insulin resistance, and blood pressure in offspring determined by maternal periconceptional B vitamin and methionine status. PNAS 104 19351–19356. (https://doi.org/10.1073/pnas.0707258104)
Slotte H, Gustafson O, Nylund L & Pousette A 1990 ATP and ADP in human pre-embryos. Human Reproduction 5 319–322. (https://doi.org/10.1093/oxfordjournals.humrep.a137097)
Smiraglia DJ, Kulawiec M, Bistulfi GL, Gupta SG & Singh KK 2008 A novel role for mitochondria in regulating epigenetic modification in the nucleus. Cancer Biology and Therapy 7 1182–1190. (https://doi.org/10.4161/cbt.7.8.6215)
Smirnova E, Griparic L, Shurland DL & van der Bliek AM 2001 Dynamin-related protein Drp1 is required for mitochondrial division in mammalian cells. Molecular Biology of the Cell 12 2245–2256. (https://doi.org/10.1091/mbc.12.8.2245)
Spikings EC, Alderson J & St John JC 2007 Regulated mitochondrial DNA replication during oocyte maturation is essential for successful porcine embryonic development. Biology of Reproduction 76 327–335. (https://doi.org/10.1095/biolreprod.106.054536)
Squirrell JM, Schramm RD, Paprocki AM, Wokosin DL & Bavister BD 2003 Imaging mitochondrial organization in living primate oocytes and embryos using multiphoton microscopy. Microscopy and Microanalysis 9 190–201. (https://doi.org/10.1017/S1431927603030174)
St John JC, Ramalho-Santos J, Gray HL, Petrosko P, Rawe VY, Navara CS, Simerly CR & Schatten GP 2005 The expression of mitochondrial DNA transcription factors during early cardiomyocyte in vitro differentiation from human embryonic stem cells. Cloning and Stem Cells 7 141–153. (https://doi.org/10.1089/clo.2005.7.141)
St John JC, Facucho-Oliveira J, Jiang Y, Kelly R & Salah R 2010 Mitochondrial DNA transmission, replication and inheritance: a journey from the gamete through the embryo and into offspring and embryonic stem cells. Human Reproduction Update 16 488–509. (https://doi.org/10.1093/humupd/dmq002)
Steinborn R, Schinogl P, Zakhartchenko V, Achmann R, Schernthaner W, Stojkovic M, Wolf E, Muller M & Brem G 2000 Mitochondrial DNA heteroplasmy in cloned cattle produced by fetal and adult cell cloning. Nature Genetics 25 255–257. (https://doi.org/10.1038/77000)
Stern S, Biggers JD & Anderson E 1971 Mitochondria and early development of the mouse. Journal of Experimental Zoology 176 179–191. (https://doi.org/10.1002/jez.1401760206)
Steuerwald N, Barritt JA, Adler R, Malter H, Schimmel T, Cohen J & Brenner CA 2000 Quantification of mtDNA in single oocytes, polar bodies and subcellular components by real-time rapid cycle fluorescence monitored PCR. Zygote 8 209–215. (https://doi.org/10.1017/S0967199400001003)
Stojanovski D, Koutsopoulos OS, Okamoto K & Ryan MT 2004 Levels of human Fis1 at the mitochondrial outer membrane regulate mitochondrial morphology. Journal of Cell Science 117 1201–1210. (https://doi.org/10.1242/jcs.01058)
Stojkovic M, Machado SA, Stojkovic P, Zakhartchenko V, Hutzler P, Goncalves PB & Wolf E 2001 Mitochondrial distribution and adenosine triphosphate content of bovine oocytes before and after in vitro maturation: correlation with morphological criteria and developmental capacity after in vitro fertilization and culture. Biology of Reproduction 64 904–909. (https://doi.org/10.1095/biolreprod64.3.904)
Stricker SA 1999 Comparative biology of calcium signaling during fertilization and egg activation in animals. Developmental Biology 211 157–176. (https://doi.org/10.1006/dbio.1999.9340)
Sturmey RG & Leese HJ 2003 Energy metabolism in pig oocytes and early embryos. Reproduction 126 197–204.
Suen DF, Norris KL & Youle RJ 2008 Mitochondrial dynamics and apoptosis. Genes and Development 22 1577–1590. (https://doi.org/10.1101/gad.1658508)
Sun QY, Wu GM, Lai L, Park KW, Cabot R, Cheong HT, Day BN, Prather RS & Schatten H 2001 Translocation of active mitochondria during pig oocyte maturation, fertilization and early embryo development in vitro. Reproduction 122 155–163.
Sun H, Kang J, Su J, Zhang J, Zhang L, Liu X, Zhang J, Wang F, Lu Z & Xing X et al. 2018 Methionine adenosyltransferase 2A regulates mouse zygotic genome activation and morula to blastocyst transition. Biology of Reproduction Epub. (https://doi.org/10.1093/biolre/ioy194)
Sutton-McDowall ML, Gilchrist RB & Thompson JG 2010 The pivotal role of glucose metabolism in determining oocyte developmental competence. Reproduction 139 685–695. (https://doi.org/10.1530/REP-09-0345)
Taanman JW 1999 The mitochondrial genome: structure, transcription, translation and replication. Biochimica and Biophysica Acta 1410 103–123. (https://doi.org/10.1016/S0005-2728(98)00161-3)
Tachibana M, Sparman M, Sritanaudomchai H, Ma H, Clepper L, Woodward J, Li Y, Ramsey C, Kolotushkina O & Mitalipov S 2009 Mitochondrial gene replacement in primate offspring and embryonic stem cells. Nature 461 367–372. (https://doi.org/10.1038/nature08368)
Tachibana M, Amato P, Sparman M, Woodward J, Sanchis DM, Ma H, Gutierrez NM, Tippner-Hedges R, Kang E & Lee HS et al. 2013 Towards germline gene therapy of inherited mitochondrial diseases. Nature 493 627–631. (https://doi.org/10.1038/nature11647)
Tait SW & Green DR 2013 Mitochondrial regulation of cell death. Cold Spring Harbor Perspectives in Biology 5 a008706. (https://doi.org/10.1101/cshperspect.a008706)
Takahashi Y, Hashimoto S, Yamochi T, Goto H, Yamanaka M, Amo A, Matsumoto H, Inoue M, Ito K & Nakaoka Y et al. 2016 Dynamic changes in mitochondrial distribution in human oocytes during meiotic maturation. Journal of Assisted Reproduction and Genetics 33 929–938. (https://doi.org/10.1007/s10815-016-0716-2)
Takeda K, Takahashi S, Onishi A, Goto Y, Miyazawa A & Imai H 1999 Dominant distribution of mitochondrial DNA from recipient oocytes in bovine embryos and offspring after nuclear transfer. Journal of Reproduction and Fertility 116 253–259.
Takeda K, Akagi S, Kaneyama K, Kojima T, Takahashi S, Imai H, Yamanaka M, Onishi A & Hanada H 2003 Proliferation of donor mitochondrial DNA in nuclear transfer calves (Bos taurus) derived from cumulus cells. Molecular Reproduction and Development 64 429–437. (https://doi.org/10.1002/mrd.10279)
Thompson JG, Partridge RJ, Houghton FD, Cox CI & Leese HJ 1996 Oxygen uptake and carbohydrate metabolism by in vitro derived bovine embryos. Journal of Reproduction and Fertility 106 299–306.
Thundathil J, Filion F & Smith LC 2005 Molecular control of mitochondrial function in preimplantation mouse embryos. Molecular Reproduction and Development 71 405–413. (https://doi.org/10.1002/mrd.20260)
Tong H, Zhang L, Gao J, Wen S, Zhou H & Feng S 2017 Methylation of mitochondrial DNA displacement loop region regulates mitochondrial copy number in colorectal cancer. Molecular Medicine Reports 16 5347–5353. (https://doi.org/10.3892/mmr.2017.7264)
Treff NR, Zhan Y, Tao X, Olcha M, Han M, Rajchel J, Morrison L, Morin SJ & Scott RT Jr 2017 Levels of trophectoderm mitochondrial DNA do not predict the reproductive potential of sibling embryos. Human Reproduction 32 954–962. (https://doi.org/10.1093/humrep/dex034)
Trimarchi JR, Liu L, Porterfield DM, Smith PJ & Keefe DL 2000 Oxidative phosphorylation-dependent and -independent oxygen consumption by individual preimplantation mouse embryos. Biology of Reproduction 62 1866–1874. (https://doi.org/10.1095/biolreprod62.6.1866)
Tsukada Y, Fang J, Erdjument-Bromage H, Warren ME, Borchers CH, Tempst P & Zhang Y 2006 Histone demethylation by a family of JmjC domain-containing proteins. Nature 439 811–816. (https://doi.org/10.1038/nature04433)
Tzagoloff A 1982 Mitochondrial Structure and Compartmentalization, Mitochondria. Cellular Organelles. Boston: Springer, pp. 15–38.
Udagawa O, Ishihara T, Maeda M, Matsunaga Y, Tsukamoto S, Kawano N, Miyado K, Shitara H, Yokota S & Nomura M et al. 2014 Mitochondrial fission factor Drp1 maintains oocyte quality via dynamic rearrangement of multiple organelles. Current Biology 24 2451–2458. (https://doi.org/10.1016/j.cub.2014.08.060)
Van Blerkom J 1989 Developmental failure in human reproduction associ- ated with preovulatory oogenesis and preimplantation embryogenesis. In Ultrastructure of Human Gametogenesis and and Embryogenesis, pp. 125–180. Eds Van Blerkom J & Motta PM. Boston: Kluwer Academic Publishers, pp. 125–180. (https://doi.org/10.1007/978-1-4613-1749-4_5)
Van Blerkom J 2004 Mitochondria in human oogenesis and preimplantation embryogenesis: engines of metabolism, ionic regulation and developmental competence. Reproduction 128 269–280. (https://doi.org/10.1530/rep.1.00240)
Van Blerkom J & Runner MN 1984 Mitochondrial reorganization during resumption of arrested meiosis in the mouse oocyte. American Journal of Anatomy 171 335–355. (https://doi.org/10.1002/aja.1001710309)
Van Blerkom J & Davis P 2006 High-polarized (Delta Psi m(HIGH)) mitochondria are spatially polarized in human oocytes and early embryos in stable subplasmalemmal domains: developmental significance and the concept of vanguard mitochondria. Reproductive Biomedicine Online 13 246–254. (https://doi.org/10.1016/S1472-6483(10)60622-0)
Van Blerkom J & Davis P 2007 Mitochondrial signaling and fertilization. Molecular Human Reproduction 13 759–770. (https://doi.org/10.1093/molehr/gam068)
Van Blerkom J, Davis PW & Lee J 1995 ATP content of human oocytes and developmental potential and outcome after in-vitro fertilization and embryo transfer. Human Reproduction 10 415–424. (https://doi.org/10.1093/oxfordjournals.humrep.a135954)
Van Blerkom J, Davis P & Alexander S 2000 Differential mitochondrial distribution in human pronuclear embryos leads to disproportionate inheritance between blastomeres: relationship to microtubular organization, ATP content and competence. Human Reproduction 15 2621–2633. (https://doi.org/10.1093/humrep/15.12.2621)
Van Blerkom J, Davis P & Alexander S 2003 Inner mitochondrial membrane potential (DeltaPsim), cytoplasmic ATP content and free Ca2+ levels in metaphase II mouse oocytes. Human Reproduction 18 2429–2440. (https://doi.org/10.1093/humrep/deg466)
van der Heijden GW, Dieker JW, Derijck AA, Muller S, Berden JH, Braat DD, van der Vlag J & de Boer P 2005 Asymmetry in histone H3 variants and lysine methylation between paternal and maternal chromatin of the early mouse zygote. Mechanisms of Development 122 1008–1022. (https://doi.org/10.1016/j.mod.2005.04.009)
van der Wijst MG, van Tilburg AY, Ruiters MH & Rots MG 2017 Experimental mitochondria-targeted DNA methylation identifies GpC methylation, not CpG methylation, as potential regulator of mitochondrial gene expression. Scientific Reports 7 177. (https://doi.org/10.1038/s41598-017-00263-z)
Varadi A, Johnson-Cadwell LI, Cirulli V, Yoon Y, Allan VJ & Rutter GA 2004 Cytoplasmic dynein regulates the subcellular distribution of mitochondria by controlling the recruitment of the fission factor dynamin-related protein-1. Journal of Cell Science 117 4389–4400. (https://doi.org/10.1242/jcs.01299)
Vickaryous N & Whitelaw E 2005 The role of early embryonic environment on epigenotype and phenotype. Reproduction, Fertility and Development 17 335–340. (https://doi.org/10.1071/RD04133)
Victor AR, Brake AJ, Tyndall JC, Griffin DK, Zouves CG, Barnes FL & Viotti M 2017 Accurate quantitation of mitochondrial DNA reveals uniform levels in human blastocysts irrespective of ploidy, age, or implantation potential. Fertility and Sterility 107 34.e3–42.e3. (https://doi.org/10.1016/j.fertnstert.2016.09.028)
Wai T & Langer T 2016 Mitochondrial dynamics and metabolic regulation. Trends in Endocrinology and Metabolism 27 105–117. (https://doi.org/10.1016/j.tem.2015.12.001)
Wai T, Ao A, Zhang X, Cyr D, Dufort D & Shoubridge EA 2010 The role of mitochondrial DNA copy number in mammalian fertility. Biology of Reproduction 83 52–62. (https://doi.org/10.1095/biolreprod.109.080887)
Wakabayashi J, Zhang Z, Wakabayashi N, Tamura Y, Fukaya M, Kensler TW, Iijima M & Sesaki H 2009 The dynamin-related GTPase Drp1 is required for embryonic and brain development in mice. Journal of Cell Biology 186 805–816. (https://doi.org/10.1083/jcb.200903065)
Wakai T, Zhang N, Vangheluwe P & Fissore RA 2013 Regulation of endoplasmic reticulum Ca(2+) oscillations in mammalian eggs. Journal of Cell Science 126 5714–5724. (https://doi.org/10.1242/jcs.136549)
Wakefield SL, Lane M & Mitchell M 2011 Impaired mitochondrial function in the preimplantation embryo perturbs fetal and placental development in the mouse. Biology of Reproduction 84 572–580. (https://doi.org/10.1095/biolreprod.110.087262)
Wales RG 1986 Measurement of metabolic turnover in single mouse embryos. Journal of Reproduction and Fertility 76 717–725. (https://doi.org/10.1530/jrf.0.0760717)
Wallace DC & Fan W 2010 Energetics, epigenetics, mitochondrial genetics. Mitochondrion 10 12–31. (https://doi.org/10.1016/j.mito.2009.09.006)
Wang J, Alexander P, Wu L, Hammer R, Cleaver O & McKnight SL 2009a Dependence of mouse embryonic stem cells on threonine catabolism. Science 325 435–439. (https://doi.org/10.1126/science.1173288)
Wang Q, Ratchford AM, Chi MM, Schoeller E, Frolova A, Schedl T & Moley KH 2009b Maternal diabetes causes mitochondrial dysfunction and meiotic defects in murine oocytes. Molecular Endocrinology 23 1603–1612. (https://doi.org/10.1210/me.2009-0033)
Wang S, Lin C, Shi H, Xie M, Zhang W & Lv J 2009c Correlation of the mitochondrial activity of two-cell embryos produced in vitro and the two-cell block in Kunming and B6C3F1 mice. Anatomical Record 292 661–669. (https://doi.org/10.1002/ar.20890)
Wang J, Zhang M, Zhang Y, Kou Z, Han Z, Chen DY, Sun QY & Gao S 2010 The histone demethylase JMJD2C is stage-specifically expressed in preimplantation mouse embryos and is required for embryonic development. Biology of Reproduction 82 105–111. (https://doi.org/10.1095/biolreprod.109.078055)
Wang C, Chen H, Zhang J, Hong Y, Ding X & Ying W 2014 Malate-aspartate shuttle mediates the intracellular ATP levels, antioxidation capacity and survival of differentiated PC12 cells. International Journal of Physiology, Pathophysiology and Pharmacology 6 109–114.
Wellen KE, Hatzivassiliou G, Sachdeva UM, Bui TV, Cross JR & Thompson CB 2009 ATP-citrate lyase links cellular metabolism to histone acetylation. Science 324 1076–1080. (https://doi.org/10.1126/science.1164097)
Wilding M, Dale B, Marino M, di Matteo L, Alviggi C, Pisaturo ML, Lombardi L & De Placido G 2001 Mitochondrial aggregation patterns and activity in human oocytes and preimplantation embryos. Human Reproduction 16 909–917. (https://doi.org/10.1093/humrep/16.5.909)
Wilding M, Fiorentino A, De Simone ML, Infante V, De Matteo L, Marino M & Dale B 2002 Energy substrates, mitochondrial membrane potential and human preimplantation embryo division. Reproductive Biomedicine Online 5 39–42. (https://doi.org/10.1016/S1472-6483(10)61595-7)
Wilding M, De Placido G, De Matteo L, Marino M, Alviggi C & Dale B 2003 Chaotic mosaicism in human preimplantation embryos is correlated with a low mitochondrial membrane potential. Fertility and Sterility 79 340–346. (https://doi.org/10.1016/S0015-0282(02)04678-2)
Wong M, Gertz B, Chestnut BA & Martin LJ 2013 Mitochondrial DNMT3A and DNA methylation in skeletal muscle and CNS of transgenic mouse models of ALS. Frontiers in Cellular Neuroscience 7 279. (https://doi.org/10.3389/fncel.2013.00279)
Wossidlo M, Nakamura T, Lepikhov K, Marques CJ, Zakhartchenko V, Boiani M, Arand J, Nakano T, Reik W & Walter J 2011 5-Hydroxymethylcytosine in the mammalian zygote is linked with epigenetic reprogramming. Nature Communications 2 241. (https://doi.org/10.1038/ncomms1240)
Wu FR, Liu Y, Shang MB, Yang XX, Ding B, Gao JG, Wang R & Li WY 2012 Differences in H3K4 trimethylation in in vivo and in vitro fertilization mouse preimplantation embryos. Genetics and Molecular Research 11 1099–1108. (https://doi.org/10.4238/2012.April.27.9)
Wu LL, Russell DL, Wong SL, Chen M, Tsai TS, St John JC, Norman RJ, Febbraio MA, Carroll J & Robker RL 2015 Mitochondrial dysfunction in oocytes of obese mothers: transmission to offspring and reversal by pharmacological endoplasmic reticulum stress inhibitors. Development 142 681–691. (https://doi.org/10.1242/dev.114850)
Xiao M, Yang H, Xu W, Ma S, Lin H, Zhu H, Liu L, Liu Y, Yang C & Xu Y et al. 2012 Inhibition of alpha-KG-dependent histone and DNA demethylases by fumarate and succinate that are accumulated in mutations of FH and SDH tumor suppressors. Genes and Development 26 1326–1338. (https://doi.org/10.1101/gad.191056.112)
Yang SG, Park HJ, Kim JW, Jung JM, Kim MJ, Jegal HG, Kim IS, Kang MJ, Wee G & Yang HY et al. 2018 Mito-TEMPO improves development competence by reducing superoxide in preimplantation porcine embryos. Scientific Reports 8 10130. (https://doi.org/10.1038/s41598-018-28497-5)
Yeon JY, Min SH, Park HJ, Kim JW, Lee YH, Park SY, Jeong PS, Park H, Lee DS & Kim SU et al. 2015 Mdivi-1, mitochondrial fission inhibitor, impairs developmental competence and mitochondrial function of embryos and cells in pigs. Journal of Reproduction and Development 61 81–89. (https://doi.org/10.1262/jrd.2014-070)
Youle RJ & van der Bliek AM 2012 Mitochondrial fission, fusion, and stress. Science 337 1062–1065. (https://doi.org/10.1126/science.1219855)
Yu Y, Dumollard R, Rossbach A, Lai FA & Swann K 2010 Redistribution of mitochondria leads to bursts of ATP production during spontaneous mouse oocyte maturation. Journal of Cellular Physiology 224 672–680. (https://doi.org/10.1002/jcp.22171)
Yue MX, Fu XW, Zhou GB, Hou YP, Du M, Wang L & Zhu SE 2012 Abnormal DNA methylation in oocytes could be associated with a decrease in reproductive potential in old mice. Journal of Assisted Reproduction and Genetics 29 643–650. (https://doi.org/10.1007/s10815-012-9780-4)
Zaitseva I, Zaitsev S, Alenina N, Bader M & Krivokharchenko A 2007 Dynamics of DNA-demethylation in early mouse and rat embryos developed in vivo and in vitro. Molecular Reproduction and Development 74 1255–1261. (https://doi.org/10.1002/mrd.20704)
Zhang J, Liu H, Luo S, Lu Z, Chavez-Badiola A, Liu Z, Yang M, Merhi Z, Silber SJ & Munne S et al. 2017 Live birth derived from oocyte spindle transfer to prevent mitochondrial disease. Reproductive Biomedicine Online 34 361–368. (https://doi.org/10.1016/j.rbmo.2017.01.013)
Zhao Q, Wang J, Levichkin IV, Stasinopoulos S, Ryan MT & Hoogenraad NJ 2002 A mitochondrial specific stress response in mammalian cells. EMBO Journal 21 4411–4419. (https://doi.org/10.1093/emboj/cdf445)
Zhao N, Zhang Y, Liu Q & Xiang W 2015 Mfn2 affects embryo development via mitochondrial dysfunction and apoptosis. PLoS One 10 e0125680. (https://doi.org/10.1371/journal.pone.0125680)
Zhou N, Cao Z, Wu R, Liu X, Tao J, Chen Z, Song D, Han F, Li Y & Fang F et al. 2014 Dynamic changes of histone H3 lysine 27 acetylation in pre-implantational pig embryos derived from somatic cell nuclear transfer. Animal Reproduction Science 148 153–163. (https://doi.org/10.1016/j.anireprosci.2014.06.002)
Zick M, Rabl R & Reichert AS 2009 Cristae formation-linking ultrastructure and function of mitochondria. Biochimica and Biophysica Acta 1793 5–19. (https://doi.org/10.1016/j.bbamcr.2008.06.013)
