Updating the markers for oocyte quality evaluation: intracellular temperature as a new index.

BACKGROUND: The developmental competence of an embryo is principally dictated by the oocyte. Usually, oocyte selection is based on morphological properties; however, all morphological criteria that are currently used for the grading and screening of oocytes are not able to eliminate the subjectivity. Despite recent studies of the molecular factors related to oocyte quality, it is technically difficult to develop an index based on these factors, and new indices that reflect intracellular conditions are necessary.
METHODS: Morphological and molecular factors influencing developmental competence were comprehensively reviewed, and intracellular temperature was evaluated as a new marker of oocyte quality. MAIN
FINDINGS: The intracellular temperature of mature oocytes was high in fresh oocytes and decreased with time after polar body release. Under the same conditions, the intracellular temperature and its distribution differed among oocytes, suggesting that temperature represents the state of each oocyte.
CONCLUSION: Intracellular temperature is advantageous as an objective and quantitative indicator of oocyte quality. Further studies should evaluate the link between temperature and cellular phenomena to establish its use as an indicator of quality.

INTRODUCTION

The oocyte is known to be a unique and highly specialized cell responsible for creating, activating, and controlling the embryonic genome, as well as supporting basic processes, such as cellular homeostasis, metabolism, and cell cycle progression in the early embryo.1 An oocyte is formed in the ovarian follicle and is the largest single cell. Meiosis in the mammalian oocyte is initiated during fetal development and is arrested at the diplotene stage of the first meiotic prophase. After stimulation by endogenous luteinizing hormone surge, oocyte meiosis resumes and progresses to the second meiotic phase with a dynamic change. Ovulation leads to the release of an oocyte into the oviduct where meiosis stops at metaphase II (MII) until fertilization.2, 3, 4 Ovulated oocytes are presumed to have acquired fertilization and developmental competence, which is related to the ability to undergo meiotic maturation, fertilization, embryonic development, and successful pregnancy. Developmental competence is gradually acquired during oogenesis, and the final stage is important for optimal development prior to ovulation because the synchronization between nuclear and cytoplasmic maturation in the oocyte is completed at this stage.5

Usually, for in vitro fertilization (IVF) and intracytoplasmic sperm injection (ICSI), oocyte selection is based on morphological parameters related to the cumulus cells, polar body, and cytoplasm.6, 7 It has been speculated that some morphological irregularities that are easily assessed under light microscopy may reflect a compromised developmental ability and could therefore be useful for selecting competent oocytes prior to fertilization.8 The first polar body (PB1) is the easiest indicator for judging nuclear maturation. However, studies using polarized light microscopy have shown that oocytes displaying a polar body may still be immature.9 In addition, just after the extrusion of PB1, oocytes do not acquire sufficient developmental competence, despite exhibiting the morphologic features of the MII stage.10 Furthermore, if the structure of the mitotic spindle collapses due to overmaturation, developmental competence decreases.10 These previous findings indicate that developmental competence changes, even in the MII oocyte.

Accurate sorting of mature oocytes that are healthy and have a high developmental competence will improve the pregnancy rate. However, the morphological criteria that are currently used for the grading and screening of oocytes are subjective and controversial, and they may not be related to the intrinsic competence of the oocyte.11, 12 The identification of objective and noninvasive molecular markers that predict oocyte ability is a major research goal. Factors related to the quality of oocytes are being elucidated at the molecular level, but it is technically difficult to develop an index based on the visualization of these factors. Accordingly, new indices that reflect intracellular conditions are necessary. In this review, an overall summary of morphological factors related to oocyte quality is provided, recent studies of molecular markers are reviewed, and intracellular temperature is introduced as a potentially effective marker.

FACTORS INFLUENCING OOCYTE QUALITY

Fresh matured oocytes with intact PB1 are enclosed within the zona pellucida, made up of glycoprotein, and consist of meiotic spindle with aligned chromosomes, microtubule‐organizing centers (pericentriolar materials, PCMs) located at the spindle poles, mitochondria, microfilaments, and regularly aligned cortical granules underneath the oocyte cortex in the cytoplasm. The zona pellucida is covered with abundant cumulus cells. Aging or overmaturation of oocytes is associated with numerous morphological and cellular alterations, including changes in the structure of the plasma membrane, zona pellucida, cytoskeleton, and mitochondria. It is also associated with displacement of the spindle, misalignment of chromosomes, and displacement of PB1 and cortical granules.4 The presence of clear PB1 and the attachment level of cumulus cells are major indicators used to determine the quality of oocytes because these are easy to observe using a microscope.

Many studies have reported that changes in oocyte constituents are linked to oocyte quality. We provide an overview of factors contributing to oocyte quality, focusing on the PB1, meiotic spindle, cumulus cells, mitochondria, and oxygen consumption.

First polar body (PB1)

The removal of cumulus cells from oocytes allows a detailed observation of the morphological characteristics. Extrusion of PB1 is a cellular landmark of meiotic maturation. Recent studies have investigated the correlation between PB1 morphology and oocyte competence; although PB1 does not participate in the developmental process, studies of mouse oocytes have supported this connection.7 PB1 morphology is frequently used to evaluate oocyte quality. Oocytes with an intact PB1 have high fertilization rates and a high oocyte quality, whereas those displaying a PB1 characterized by large size, irregular shape, rough surface, or fragmentation are developmentally less competent after IVF, yielding low pregnancy rates after embryo transfer.13, 14 PB1 degeneration occurs within a few hours after extrusion, and it is associated with oocyte aging.15, 16, 17 Some PB1 shows displacement from the MII spindle at nuclear maturation, and the distance between PB1 and MII spindle increases over time during oocyte aging. Moreover, the perivitelline space increases over time and facilitates the lateral displacement of the degenerating PB1.17

Meiotic spindle

The meiotic spindle is a chromosome distribution cytoskeletal structure, critically important for the accurate distribution of chromosomes to the dividing blastomeres, thereby ensuring accurate embryonic development.4 In fresh oocytes, spindles display a vertical orientation with respect to the oolemma and spindle poles associate with PCMs to create a compact bipolar spindle. This morphology changes during aging at the MII stage; the spindle becomes elongated and/or loses tension in its microtubules and becomes weak.18, 19, 20, 21 The shape of the spindle is determined by the position of the spindle pole and changes over time after spindle formation. Oocytes exhibiting a reduction in the distance between the PCMs at the spindle pole have a higher developmental potential than those exhibiting an increase in the distance between the PCMs.10 When the distance between PCMs is short, a small rhomboid spindle body is formed, and as the distance increases, a large spindle is formed. The morphology of the mitotic spindle is an important indicator of oocyte condition. Conventionally, spindles were mainly visualized by confocal microscopy, which requires cell fixation and hence cannot be applied to live cells. Alternatively, meiotic spindles can be observed directly using a polarization microscope.22

The molecular mechanisms underlying meiotic spindle in fresh oocytes have shown the importance of meiotic spindles in fertilization and embryonic development.23, 24 The meiotic spindle is essential for the accurate separation of homologous chromosomes or two sets of chromatids during germ cell division.25 Oocyte aging results in significant increases in premature chromosome separation, which is strongly associated with aneuploidy.26, 27 Aneuploidy is involved in inheriting too many or too few of any of the chromosomes. Most aneuploid embryos that inherit only one copy of an autosome develop severe abnormalities and die before pregnancy. In contrast, inheriting an extra copy of an autosome is also associated with severe developmental abnormalities and miscarriages. Chromosome 21 trisomy, the cause of Down's syndrome, is by far the most frequent aneuploidy affecting live births.28, 29, 30 Chromosomes in 2‐day‐old oocytes are no longer aligned at the spindle equator but are scattered within the degenerating spindle. In oocytes aged 3‐4 days, chromosomes become more decondensed and display nuclear alterations. Chromosome loss, fragmentation, or the clumping of chromosomes and chromatid separation have been observed in aged oocytes.18, 31, 32

Cumulus cells

Cumulus cells are critical for oocyte maturation, ovulation, and fertilization,33 and are a determinant of oocyte quality.34 Cumulus cells support energy production in the cumulus‐oocyte complex.35, 36 Additionally, cumulus cells that surround oocytes may protect against the damaging effects of reactive oxygen species (ROS).37 Recent studies suggest that the mitochondrial function of cumulus cells can directly influence the ability to achieve a successful pregnancy.38, 39 The identification of surrogate markers of oocyte competence and favorable reproductive outcomes in assisted reproductive technology is a goal of many transcriptome, proteome, and metabolome studies. The analysis of granulosa and cumulus cells is considered one of the best noninvasive strategies available today.40

Mitochondria

The mitochondrion is directly involved in many essential cellular functions, including energy production, management of ROS levels, and regulation of apoptosis. Mitochondria play an extremely important role in supplying the energy that is consumed during the maturation process.41, 42 The primary function of mitochondria is to synthesize adenosine triphosphate (ATP), the preferred energy source of cells. Synthesis of ATP in adequate amounts is critical for cell survival, and severe ATP deficiency often leads to apoptosis.43 Although several metabolic pathways of ATP production have been identified, most of the ATP generated from glucose is produced via mitochondrial oxidative phosphorylation (OXPHOS).44 All the complex processes that occur in the oocyte prior to ovulation and fertilization require energy, which is derived mainly from ATP production via OXPHOS.45 Moreover, higher ATP content in oocytes and embryos has been correlated with better reproductive results among infertile patients.46 In contrast, mitochondrial dysfunction has been implicated in decreased oocyte quality, and clinical and experimental data have suggested decreased oocyte quality as the main factor in the age‐related deterioration of reproductive capacity. However, the molecular mechanisms underlying this mitochondrion‐related decrease in oocyte quality remain poorly understood.47, 48

The distribution and organization of mitochondria during oocyte maturation are dynamic, and these changes may be related to mitochondrial function. Oocytes with higher concentrations of ATP have significantly higher fertilization and blastocyst rates.42, 49 Lower ATP content in oocytes is at least partially responsible for positive spindle formation in in vitro maturation mammalian oocytes.50, 51 Decreasing the ATP content in mouse oocytes by treatment with carbonyl cyanide p‐trifluoromethoxyphenylhydrazone, an inhibitor of OXPHOS, leads to a reduction in the percentage of oocytes with nuclear maturation, normal spindle formation, and chromosome alignment, evenly distributed mitochondria, and the ability to form blastocysts.52 ATP is extremely important for nuclear and cytoplasmic maturation events. Spindle formation and chromosome movements depend on the expression and activity of motor proteins, which use ATP as their energy source. Due to the critical role of energy metabolism in oocyte maturation, ATP content has been proposed as an indicator of the developmental potential of oocytes.53, 54, 55

Oxidative stress (OS) results from an imbalance between the production of ROS and neutralizing antioxidant molecules.56 In mammalian mature oocytes, OS causes substantial mitochondrial dysfunction, impacting both mitochondrial ATP synthesis and the activation of mitochondrial‐mediated apoptotic mechanisms.57, 58 Enhanced and unbalanced ROS production may be a predominant cause of impaired mitochondrial OXPHOS.59 External factors contribute to the higher OS observed in vitro, including exposure to visible light, non‐ideal pH and temperature, centrifugation, cryopreservation, culture medium composition, oxygen concentrations, and oocyte and embryo manipulation processes.60 mtDNA is particularly susceptible to several elements causing OS, and mtDNA disruption leads to critical loss of function and, ultimately, diminished capacity to generate ATP.58, 59 Oocyte mtDNA content increases until the stage that immediately precedes fertilization. In healthy embryos, the accumulated mtDNA is divided equally among all cells during embryogenesis.61, 62, 63, 64 Recent studies have proposed quantification of mtDNA in cumulus, granulosa, and trophectoderm cells as a promising strategy for predicting embryo quality and viability.62, 65 Since mtDNA content in cumulus cells is correlated with that in oocytes for each cumulus‐oocyte complex, it is suggested that the mitochondrial characteristics of the cells may serve as a marker of the oocyte quality.40 Furthermore, mutations or deletions in mtDNA have been correlated with organelle dysfunction, low ATP levels, and embryonic developmental arrest.66 With aging, mtDNA deficiency of luteinizing granulosa cells and cumulus cells increases, which leads to a decrease in pregnancy rate.66, 67 These findings corroborate the understanding that mitochondrial function of granulosa and cumulus cells directly influences embryonic development, as well as the maturation and fertilization of oocytes.68

Various mitochondrial anomalies have been linked to the age‐related deterioration of oocyte quality, and at least some of these may be reflective of the changes in specific mitochondrial subpopulations.69 The most prominent of these defects are atypical mitochondrial localization and aggregation, reduced mtDNA content, reduced membrane potential (consequently, bioenergetic capacity), increased OS, and increased frequency of mtDNA mutations and deletions.47, 50, 70, 71, 72, 73, 74, 75, 76, 77, 78, 79

MOLECULAR MARKERS RELATED TO OOCYTE QUALITY

Meta‐analysis of microarray studies

Multiple microarray studies have been performed to identify markers associated with oocyte quality and developmental competence in oocytes or both oocytes and cumulus cells.80, 81, 82, 83, 84 Based on a meta‐analysis of previously published microarray data for various models of oocyte and embryo quality, 63 candidate genes associated with oocyte quality across several species were identified. Biological networks and transcription factor regulation associated with oocyte quality were also identified.85 If factors that control oocyte quality and molecular mechanisms are clarified, it might provide a basis for objectively evaluating oocyte quality.

microRNAs

Gamete maturation requires extensive signaling between germ cells and their surrounding somatic cells. In the ovary, from the theca cells, mural granulosa cells, cumulus cells, and oocyte secrete factors that are critical for ovulation of high‐quality oocyte, throughout follicle growth and oocyte maturation. Recent studies of a variety of species have uncovered the presence of cell‐secreted vesicles in follicular fluid.86 These cell‐secreted vesicles contain small non‐coding regulatory RNAs called microRNAs, which can be shuttled between maturing gametes and surrounding somatic cells.87 In humans, it is known that extracellular microRNAs of follicular fluid are associated with fertilization ability and early embryo quality,85 although little is known about the exact mechanism by which microRNAs are loaded into these cell‐secreted vesicles or are transferred and modulate gene expression and function. However, recent studies suggest that microRNAs in cell‐secreted vesicles are involved in oocyte maturation. These microRNAs involved in gamete maturation are potential therapeutic targets and diagnostic markers associated with fertility.86

VISUALIZATION OF OOCYTE QUALITY BY INTRACELLULAR TEMPERATURE IMAGING

The effects of temperature within the cell have drawn recent attention. Temperature affects various physiological functions and is important for maintaining homeostasis. Cellular functions are fundamentally regulated by intracellular temperature.88, 89, 90 The biological reactions responsible for cellular functions occur either exothermically or endothermically at particular locations within a cell, such as inside organelles. Thus, temperature distributions inside a living cell reflect the functions of cellular components.91 Okabe et al developed a novel fluorescent polymer thermometer (FPT) that can diffuse throughout whole cells; using this intracellular temperature imaging approach, they were able to evaluate the thermal profiles of living cells.92 The temperature resolutions of FPT were 0.18‐0.58°C; it can detect differences of approximately 0.2°C in the cell. There appears to be a temperature variation in 1‐2°C within somatic cells. Additionally, the temperatures of the nucleus and centrosome in somatic cells are significantly higher than that of the cytoplasm, and the temperature gap between the nucleus and the cytoplasm differed depending on the cell cycle. The FPT also detects heat production from mitochondria. The heterogeneous temperature distribution is inherently related to basic cellular processes, such as the cell cycle and mitochondrial function.92

In oocytes, during the transition from the germinal vesicle to the MII stage, virtually all major organelles undergo important changes in structure, function, and/or distribution.93, 94, 95, 96, 97, 98, 99 The oocyte during MII arrest is in a highly dynamic state, with spindle microtubules keeping all of the chromosomes perfectly aligned on the metaphase plate via proteins, such as maturation promoting factor (MPF).100 Furthermore, in the mature oocyte at MII, substantial changes, such as spindle formation, chromosome alignment, and mitochondrial activity to acquire developmental competence, occur, with the potential for an elevated temperature. We investigated the intracellular temperature and its relationship to oocyte quality using the FPT developed by Okabe et al. Intracellular temperature in mature oocytes was higher in fresh oocytes immediately after PB1 extrusion, and the temperature decreased with time after polar body release (submitted data). The differences in oocyte intracellular temperature can correlate with developmental competence. Fresh oocytes had high‐temperature regions localized around the cell membrane and around the spindle (Figure 1).

Figure 1

Schematic representation of intracellular temperature in matured oocytes. A, Fresh oocyte, and B, overmatured or aged oocyte. Fresh oocytes had high‐temperature regions localized around the cell membrane and around the spindle. Red and yellow spots indicate high temperature, and blue and green spots indicate low temperature

The dynamic changes in the cytoskeleton are considered to contribute to intracellular thermal variations. The temperature of a centrosome, the main microtubule‐organizing center, is higher than that of the surrounding area in COS7 and HeLa cells.92 However, the thermogenic mechanism at the centrosome remains unclear. It is presumed to be caused by the hydrolysis of tubulin‐GTP, ATP‐driven motion of motor proteins, and phosphorylation/dephosphorylation of centrosomal proteins by kinase/phosphatase.101 In addition, repeated shortening and elongation of microtubules are necessary for the formation and maintenance of the spindle, and this repetition might be responsible for heat generation.

Furthermore, heat production by mitochondria is considered a factor influencing intracellular temperature. Substantial heat generation in the mitochondria was observed when HeLa cells expressing tsGFP1‐mito, a genetically encoded thermosensor specifically targeting the mitochondria, were treated with carbonyl cyanide 3‐chloro‐phenylhydrazone.102 Simultaneous visualization of tsGFP1‐mito in HeLa cells using JC‐1, a dye that visualizes high mitochondrial membrane potential, and ATeam, a genetically encoded ATP sensor, revealed high temperature in mitochondria with high membrane potential and that there was a positive correlation between ATP levels and membrane potential.103, 104 This result demonstrates that constitutive thermogenesis occurs via the respiratory chain or OXPHOS in a subpopulation of mitochondria in HeLa cells.101 Interestingly, even in oocytes collected under the same conditions, the intracellular temperature and temperature distribution differed among oocytes, suggesting that the temperature represents the state of each oocyte well. Taken together, these reports indicate that the intracellular temperature may be affected by the function of organelles such as microtubules and mitochondria. Thus, the intracellular temperature of oocytes can be a strong predictor of oocyte quality and developmental competence.

CONCLUSION

Individuals with identical morphological features can differ with respect to developmental competence. It is well known that oocyte quality determines the developmental potential of embryos after fertilization.6 Oocytes arrested at the MII stage are normally fertilized within a few hours after ovulation or PB1 emission. If fertilization does not occur within the proper time, the unfertilized oocyte undergoes a time‐dependent deterioration in quality, resulting in oocyte aging, a cause of fertilization failure. Sakai et al reported that the optimal period of fertilization can be specified based on the distance between the PCMs of the meiotic spindle.10 It is important to accurately determine the generation capacity of individual oocytes, but it is also necessary to perform IVF/ICSI according to the fertilization period.

Factors related to oocyte quality are becoming evident at the molecular level. However, it is not easy to accurately evaluate the developmental competence of mature oocytes based on morphology or molecular activity without damage. Typically, oocyte selection is based on microscopically determined morphological properties; however, all morphological criteria that are currently used for the grading and screening of oocytes are sometimes subjective and may not reflect the intrinsic competence of the oocyte. Moreover, morphological evaluation depends greatly on the experience and subjectivity of the observer and lacks quantitativeness. A major advantage of using intracellular temperature as a predictor is that it can be evaluated objectively and quantitatively using a temperature imaging system. If we can definitively prove that the temperature accurately reflects phenomena in the cell, it could be an indicator of oocyte quality. We are working toward elucidating the mechanism by which temperature influences cellular processes. In this study, FPT has been injected into oocytes to measure intracellular temperature, but in the future, noninvasive methods for temperature measurement should be developed for visualization.

DISCLOSURES

Conflict of interest: The author declares no conflict of interest. Human rights statement and informed consent: This article does not contain any experiment performed with human subjects. Animal studies: All institutional and national guidelines for the care and use of laboratory animals were followed. All the experiments were approved and conducted in accordance with the guidelines of the Committee of Animal Experiments of Hiroshima University, Hiroshima, Japan.