Research Progress on Mammalian Oocyte Vitrification: From Damage Mechanisms to Optimization Strategies

Simple Summary

Oocyte vitrification is an effective method for preserving female germplasm. However, current vitrification procedures for mammalian oocytes can still induce cellular oxidative stress, apoptosis, abnormal spindle distribution, and alterations in epigenetic modifications, thereby affecting subsequent developmental potential. In recent years, numerous studies have focused on incorporating antioxidants and novel cryoprotectants to improve oocyte vitrification protocols and enhance efficiency. Nevertheless, research on the mechanisms underlying cryodamage remains fragmented, and strategies for improving oocyte vitrification outcomes still require further synthesis. This review begins by outlining the fundamentals of oocyte cryo-sensitivity and further summarizes the causes and mechanisms of cryodamage during oocyte vitrification, the factors influencing oocyte survival and reproductive outcomes following vitrification, strategies for regulating vitrification-induced stress in oocytes, a comparison of vitrification processes and outcomes across different species, as well as the limitations of current research and future perspectives.

Abstract

With the continuous advancement in reproductive biology, oocyte vitrification has become a critical technology for preserving female germplasm and protecting it from environmental disruptions. This technique also eliminates temporal and spatial constraints in animal embryo engineering research. However, during the vitrification of animal oocytes, exposure to low temperatures and high concentrations of cryoprotectants can cause various forms of damage, including cytoskeletal disruption, spindle abnormalities, mitochondrial dysfunction, apoptosis, oxidative stress and epigenetic modifications. These issues are now understood to severely restrict the subsequent developmental competence of oocytes, resulting in lower cleavage and blastocyst formation rates than those of fresh oocytes. Currently, the mechanisms of cryodamage in vitrified oocytes remain poorly understood, and standardized strategies to enhance vitrification efficiency have yet to be firmly established. This review provides a formal overview of the physiological factors underlying oocyte sensitivity to vitrification, alongside the mechanisms of cryodamage and the variables influencing post-thaw survival and reproductive success. It evaluates strategies for mitigating vitrification-induced stress, compares interspecies differences, and addresses current research limitations. By identifying future directions, this review offers new insights for optimizing mammalian oocyte cryopreservation techniques.

1. Introduction

Advances in animal embryo engineering have established oocyte vitrification as a key strategy for preserving female germplasm and mitigating losses due to major diseases or infections [1]. This technique also overcomes temporal and spatial constraints associated with in vitro embryo culture, embryo transfer, and somatic cell nuclear transfer, thereby facilitating progress in embryonic genetic engineering [2,3]. In the field of animal husbandry, the vitrification of livestock oocytes enables the establishment of germplasm resource banks, thereby accelerating genetic improvement programs in large animals such as pigs, cattle, and sheep [4]. Moreover, it provides technical support for the international exchange of genetic resources while reducing transportation costs by minimizing the need for live-animal shipment [5]. Importantly, long-term preservation of oocytes through vitrification holds significant value for conserving genetic resources of endangered species and maintaining biodiversity [6,7].

Vitrification, first proposed by Rall and Fahy [8] in 1985, was successfully applied to mouse embryo cryopreservation. By exposing cells to high concentrations of cryoprotectants (CPAs) to induce gradual dehydration, then cooling them ultra-rapidly in liquid nitrogen, this technique avoids ice crystal formation by solidifying the solution into a glass-like state at sufficiently high cooling rates [9]. The first successful vitrification of human oocytes resulting in a live birth utilized a solution of 7.1 M ethylene glycol (EG) and 0.6 M sucrose [10]. Following ongoing optimization of vitrification solutions, a protocol combining permeable and non-permeable cryoprotectants (e.g., 15% EG, 15% Dimethyl Sulfoxide (DMSO), and 0.5 M sucrose) has been developed [11,12,13]. Studies have demonstrated that, compared with slow freezing, vitrification of human oocytes can significantly improve the oocyte survival rate (vitrification group 84.7% vs. slow-freezing group 58%) [14]. Owing to its advantage of minimizing ice crystal-induced mechanical damage, vitrification has been extensively applied in the cryopreservation of oocytes from various mammalian species, including economically important livestock such as cattle [15,16], sheep [17], pigs [18,19,20], horses [21], as well as model organisms like mice [22,23,24], and humans [25]. While vitrification is more effective than slow freezing for oocyte preservation [14], the process remains particularly challenging due to the unique structural and physiological characteristics. These challenges arise from the unique physiological characteristics of oocytes, including their large volume, high water content, low membrane permeability, extreme sensitivity of the meiotic spindle to low temperatures, and high intracellular lipid content [2,26,27,28]. These characteristics collectively render oocytes far more vulnerable to cryopreservation-induced damage than other cell types.

In recent years, significant efforts have been undertaken to optimize vitrification protocols for mammalian oocytes to enhance cryopreservation efficiency. However, several challenges in oocyte vitrification remain unresolved [29,30]. On the one hand, the mechanisms underlying cryoinjury have yet to be fully elucidated, particularly those involving oxidative stress, apoptosis, cytoskeletal and spindle disruption, zona pellucida hardening, and epigenetic alterations [30,31,32,33,34,35,36]. On the other hand, effective strategies for mitigating vitrification-induced stress also lack a comprehensive summary. Furthermore, a systematic comparison of vitrification processes and outcomes across species remains lacking. Notably, the cryopreservation efficiency of oocytes from large farm animals, such as pigs, cattle, and sheep, remains relatively low [37]. This review aims to elucidate the mechanisms underlying cryoinjury in mammalian oocytes, identify key factors that affect vitrification success, summarize effective strategies to mitigate vitrification-induced stress, and compare vitrification outcomes across species. The goal is to provide a reference for optimizing vitrification protocols for mammalian oocytes, improving the cryopreservation efficiency of oocytes from large animals, accelerating animal breeding programs, and facilitating the conservation of germplasm resources in endangered species.

2. Literature Search and Selection

A systematic literature search was conducted utilizing the PubMed and Web of Science databases for studies published between 1985 and 2026. To ensure a comprehensive retrieval, the following Boolean search string was employed: (vitrification) AND (oocyte OR oocytes) AND (mouse OR mice OR bovine OR cattle OR ovine OR sheep OR porcine OR pig OR human OR humans) AND (antioxidant OR “apoptosis inhibitor” OR “ice recrystallization inhibitor” OR delipidation OR “biosynthetic material” OR “automated device” OR cryoprotectant). The inclusion criteria focused on original research articles investigating mammalian oocyte vitrification, cryoinjury mechanisms, or optimization strategies (e.g., nanomaterials, biosynthetic materials, and antioxidants), provided they reported key outcomes such as survival, cleavage, or blastocyst rates. Conversely, non-English articles, conference abstracts, and duplicates were excluded. The selection process involved an initial independent screening of titles and abstracts by two authors, followed by a rigorous full-text assessment, with any discrepancies resolved through consultation with a third author. From an initial yield of 1058 articles, 178 met the eligibility criteria and were ultimately included. These studies were systematically categorized into themes, including oocyte physiology, cryoinjury mechanisms, efficiency factors, alleviation strategies, and cross-species comparisons.

3. Physiological Basis of Oocyte Sensitivity to Vitrification

The sensitivity of mammalian oocytes to vitrification is fundamentally determined by their unique physiological characteristics. First, oocytes possess a large volume relative to their limited plasma membrane surface area, resulting in high intracellular water content and reduced dehydration efficiency during vitrification [26]. Consequently, residual intracellular water may form ice crystals, leading to structural damage [2]. Second, the oocyte exhibits low permeability to cryoprotectants; high concentrations may induce cytotoxicity, whereas insufficient concentrations fail to achieve adequate dehydration, thereby increasing the risk of intracellular ice formation [37]. Third, the meiotic spindle of mature oocytes is acentrosomal and exquisitely sensitive to thermal fluctuations. Even minor cooling triggers microtubule depolymerization, which can result in chromosome misalignment and subsequent aneuploidy [27,38]. Fourth, the high lipid content of oocytes increases their susceptibility to chilling injury, and cryopreservation-induced alterations in lipid droplet morphology can further compromise subsequent developmental competence [28]. Fifth, oocyte vitrification can trigger premature cortical granule exocytosis, resulting in zona pellucida hardening prior to fertilization, thereby impairing sperm penetration even when the oocyte survives [37,39]. Besides, osmotic stress and cryoprotectant toxicity during freezing can disrupt membrane fluidity [26]. Such membrane damage is considered a major factor contributing to the reduced developmental potential of oocytes following cryopreservation [40,41]. Collectively, these physiological characteristics constitute the intrinsic basis for oocyte sensitivity to vitrification. These inherent biological constraints predispose oocytes to cryoinjury, warranting a comprehensive understanding of their sensitivities to systematically elucidate the mechanisms underlying cryoinjury.

4. Causes and Mechanisms of Oocyte Vitrification Injury

Oocytes represent the largest category of cells in mammals. Owing to their characteristics of large volume, high water content, and high intracellular lipid content, they are highly susceptible to structural damage during cryopreservation. Vitrification reportedly initiates oxidative stress in oocytes, disrupting intracellular metabolic homeostasis and triggering excessive reactive oxygen species (ROS) production [42]. Excessive ROS directly targets intracellular DNA, proteins, and lipids; among these, lipid peroxidation damages spindle microtubules and microfilaments, leading to spindle depolymerization [43]. Concurrently, oxidative stress impairs mitochondrial function by reducing ATP synthesis, further exacerbating metabolic dysfunction. Under sustained oxidative stress, abnormal structural changes occur in the glycoproteins of the oocyte zona pellucida, resulting in decreased elasticity and zona hardening [44]. Ultimately, the combined effects of oxidative stress, mitochondrial dysfunction, and metabolic disturbances induce abnormalities in epigenetic processes (e.g., DNA methylation and histone modifications) in oocytes, which ultimately lead to embryonic developmental arrest [45].

4.1. Oxidative Stress

Under physiological conditions, the organism eliminates excessive intracellular ROS via the endogenous antioxidant defense system, thereby maintaining the balance of the intracellular redox status [46]. There are three major types of ROS in cells: hydrogen peroxide (H₂O₂), superoxide anion (O₂⁻) and hydroxyl radical (•OH). ROS production during vitrification and warming of oocytes involves multiple pathways, including cryo-stress, osmotic changes, ice crystal formation, mitochondrial dysfunction, endoplasmic reticulum stress, and imbalances in enzymatic and non-enzymatic antioxidant defense systems [26,33,47] (Figure 1). These factors induce excessive ROS production, which damages mitochondria and reduces ATP synthesis. Numerous studies have demonstrated that vitrification treatment of oocytes increases intracellular H₂O₂ levels and decreases glutathione (GSH) levels. Vitrified porcine oocytes showed significantly decreased GSH levels and increased H₂O₂ after warming [47]. Vitrification also impaired glucose uptake and decreased GSH and ATP levels, while increasing ROS in mouse oocytes [31]. In a study using the open pulled straw (OPS) device (was a vitrification carrier made in the laboratory) to vitrify porcine metaphase II (MII) oocytes, significant mitochondrial dysfunction was observed, characterized by diminished mitochondrial membrane potential (MMP), decreased ATP production, elevated ROS levels, and dysregulated expression of key apoptosis-related markers, including caspase-3, caspase-8, and caspase-9 [48]. In mammalian oocytes, H₂O₂ is generated from superoxide produced by mitochondria and reduced nicotinamide adenine dinucleotide phosphate (NADPH) oxidase [49]. With a relatively long half-life, H₂O₂ is further reduced to the more destructive •OH via the Fenton reaction at high concentrations [49], thereby inducing intracellular oxidative stress damage. Overall, vitrification induces overproduction of ROS, leading to lipid peroxidation, DNA damage, and oxidative stress, thereby compromising oocyte survival and developmental competence. Supplementing culture media with antioxidants such as resveratrol, N-acetylcysteine, or ascorbic acid can mitigate oxidative damage. In the future, oxidative stress can be alleviated by supplementing antioxidants or by developing additives that combine antioxidants with other small molecules (e.g., apoptosis inhibitors and regulators of mitochondrial function), thereby achieving efficient oocyte vitrification.

Figure 1.

Schematic diagram of ROS generation sources during oocyte vitrification. EG, ethylene glycol; DMSO, dimethyl sulfoxide; ER, endoplasmic reticulum; NADPH, reduced nicotinamide adenine dinucleotide phosphate; O₂, Oxygen; H₂O₂, hydrogen peroxide; -O₂⁻, superoxide anion; H₂O, water.

4.2. Cytoskeletal and Spindle Disruption

Vitrification can adversely affect the cytoskeleton and spindle structure of oocytes. Guo et al. [50] utilized the OPS device for the vitrification of bovine germinal vesicle (GV) stage oocytes, demonstrating that this procedure can induce varying degrees of cytoskeletal damage. Wei et al. [32] vitrified bovine MII and GV stage oocytes and observed that the expression levels of the cytoskeleton-related genes gap junction protein alpha 1 (GJA1), cytokeratin 8 (CK8), and beta-actin (ACTB) were higher in MII stage oocytes than in GV stage oocytes. Utilizing the cryotop device (Kitazato Corp., Tokyo, Japan), vitrification has been shown to induce structural abnormalities in spindle microtubules of bovine MII oocytes and elevate ROS levels [51]. The oocyte spindle, composed primarily of α and β-tubulin microtubules, is highly sensitive to low temperatures, and its integrity is crucial for proper chromosome segregation during meiosis [52]. Vitrified mouse oocytes reduce microtubule density, disrupting pKIFF11 localization and inhibiting the formation of focused spindle poles via acentriolar microtubule-organizing centers (aMTOCs) [53]. Mammalian somatic cells complete chromosome segregation during mitosis through multiple pathways involving centrosomes [54]. However, oocytes lack canonical centriole structures; the two poles of the spindle are formed via microtubule-organizing centers [55]. Due to the absence of centrosomes, oocytes are highly vulnerable to structural damage and spindle depolymerization during vitrification, caused by extreme osmotic shifts, cryo-injury, and cryoprotectant toxicity [56]. Post-warming, the cell must reassemble the spindle. Without centrioles, oocytes rely on the relocalization of multiple, dispersed MTOCs and associated proteins (e.g., mDia2, pericentrin) to the chromosomal poles, a process that is error-prone and often incomplete. Accordingly, even if morphological recovery occurs, functional defects in the spindle persist, ultimately compromising embryonic developmental potential [57]. Porcine GV stage oocytes subjected to vitrification using the cryolock device (Importadora Mexicana de Materiales para Reproducción Asistida, Sociedad Anónima de Capital Variable, Mexico) exhibited decreased viability (97% in the control group vs. 78% in the vitrification group) and maturation rates (79% in the control group vs. 40% in the vitrification group), along with impaired actin filament organization and chromosomal integrity [30]. Furthermore, research using a mouse model revealed that spindle depolymerization during oocyte vitrification is associated with activation of cathepsin B within lysosomes [58]. These impairments induce abnormal chromosome alignment, along with decreased oocyte survival rate and developmental competence. To address gaps in existing research, future studies could leverage multi-omics approaches to elucidate the regulatory pathways underlying enhanced cytoskeletal stability in MII stage oocytes, thereby improving oocyte vitrification efficiency.

4.3. Mitochondrial Dysfunction

As crucial cellular organelles, mitochondria supply energy and maintain intracellular Ca²⁺ homeostasis [59]. Under physiological conditions, mitochondrial Ca²⁺ homeostasis, ATP synthesis, and ROS production are tightly regulated, maintaining a delicate balance [60]. Mitochondria are composed of inner and outer membranes, with continuous electron transfer across the inner membrane [61]. Electrons are transported through respiratory chain complexes I-IV and ultimately combine with oxygen [62]. This process establishes an electrochemical gradient across the membrane, known as the MMP, that drives oxidative phosphorylation, converting ADP to ATP [63]. However, compared to mammalian somatic cells, oocytes exhibit lower basal ATP levels. During vitrification and warming, extreme temperature fluctuations severely impair mitochondrial function. Vitrified mouse MII oocytes showed decreased mitochondrial copy numbers, altered distribution, and reduced ATP production [64,65]. Similar cytoskeletal damage has also been reported in other species, such as pigs and cattle [66,67]. Vitrification-induced oxidative stress impairs respiratory chain function, creating a vicious cycle of ROS accumulation that leads to critical molecular degradation, lipid peroxidation, and ultimately, cell death [33]. Furthermore, studies indicate that vitrification alters mitochondrial thermostasis and impairs mitochondrial quality in mouse oocytes [12]. These studies collectively demonstrate that vitrification impairs mitochondrial function through oxidative stress, altered MMP, and reduced ATP production. Future research may improve vitrification efficiency by deciphering temperature-mediated mitochondrial damage and stage-specific mitochondrial tolerance.

4.4. Damage to the Zona Pellucida

The zona pellucida (ZP) is now understood to play a critical role in early oocyte development [68]. Mouse oocytes with an intact ZP exhibit significantly higher post-vitrification survival rates compared to ZP-deficient oocytes, which is closely associated with their enhanced mechanical stability [69], highlighting the structural importance of the ZP during vitrification. Ultrastructural analysis of human oocytes following cryopreservation revealed a significant attenuation in cortical granule density [70]. Cryoprotectants and freezing–thawing processes can induce elevated intracellular Ca²⁺ levels, leading to premature cortical granule exocytosis and subsequent ZP hardening [34]. Premature release of cortical granules causes hardening of the ZP in oocytes, leading to a significant reduction in fertilization rate after cryopreservation. Studies in human oocytes have demonstrated abnormal reductions in cortical granule number and density using a cryotip device (Paillettes Crystal 133 mm; CryoBioSystem, Paris, France), potentially contributing to ZP hardening [39,71]. These findings collectively indicate that vitrification-induced ZP damage can impair sperm–oocyte interaction, reduce fertilization rates, and decrease early embryonic development. Therefore, in human reproductive medicine, intracytoplasmic sperm injection (ICSI) is commonly used to fertilize vitrified oocytes, thereby effectively circumventing the decline in fertilization rate caused by zona pellucida hardening. Similarly, this methodology is applicable to the vitrification of oocytes from endangered species, where it may mitigate the attenuation of developmental rates associated with zona pellucida hardening.

4.5. Induction of Apoptosis

Mammalian oocyte apoptosis occurs primarily through the mitochondria-mediated intrinsic pathway and the death receptor-mediated extrinsic pathway [72]. Current evidence suggests that granulosa cell apoptosis disrupts the supply of critical signaling molecules and small metabolites required for oocyte meiotic progression, thereby increasing the susceptibility of mammalian oocytes to apoptosis [73,74]. Meanwhile, oocytes at different developmental stages exhibit varying sensitivities to the activation of apoptotic pathways. Studies have demonstrated that, compared with mature rat oocytes, immature oocytes are more susceptible to H₂O₂-induced apoptosis, accompanied by phenotypic changes, including membrane degeneration and cell shrinkage [75]. In addition, elevated Ca²⁺ levels inside and outside the cell are major contributors to oocyte apoptosis. Sustained high intracellular Ca²⁺ concentrations induce oocyte apoptosis by increasing intracellular ROS levels or triggering FAS receptors [76,77]. Gao et al. [35] reported that supplementing vitrification/warming solutions with 3-methyladenine in mouse immature oocytes led to significantly higher mRNA and protein levels of beclin-1 (an autophagy marker) and caspase-3 compared to fresh oocytes. The surge in calcium ions within oocytes during vitrification elevates hydrogen peroxide levels. The persistent accumulation of ROS upregulates the Bax/Bcl-2 expression ratio, alters MMP, triggers cytochrome release, and ultimately activates the caspase cascade [78,79]. In summary, vitrification induces apoptosis via pathways including Fas/FasL, TNFR/TNF-α, and ROCK. Upregulation of apoptosis-related genes may serve as an early indicator of reduced developmental potential in vitrified oocytes. To mitigate apoptosis during oocyte vitrification, future research could focus on developing non-toxic cryoprotectants and targeted apoptosis inhibitors to suppress ROS accumulation and the subsequent activation of apoptotic signaling pathways.

4.6. Alterations in Epigenetic Modifications

Accumulating evidence indicates that oocyte vitrification is associated with widespread epigenetic perturbations; however, the extent to which these alterations are functionally relevant remains incompletely understood. Several studies have reported disrupted DNA methylation patterns following vitrification, accompanied by the downregulation of imprinted genes such as insulin-like growth factor 2 receptor (IGF2R), protein phosphatase 1 regulatory subunit 9A (PPP1R9A), and paternally expressed 3 (PEG3) [80]. In parallel, alterations in histone modifications have been consistently observed across species. Furthermore, Spinaci et al. [36] observed that vitrification of porcine oocytes induced alterations in site-specific histone acetylation, notably at histone H3 lysine 9 (H3K9ac), H4 lysine 5 (H4K5ac), and H4 lysine 21 (H4K21ac), as well as a reduction in H3 lysine 9 trimethylation (H3K9me3). Recent studies further suggest that vitrification may disrupt histone lactylation, as evidenced by the downregulated expression of lactate dehydrogenase A (LDHA), lactate dehydrogenase B (LDHB), and E1A binding protein p300 (EP300) in vitrified mouse oocytes [81]. Meanwhile, Vitrification reduced Sirtuin (SIRT1) expression, leading to aberrant H3K9 acetylation, DNA methylation, and expression of imprinted genes such as gene trap locus 2 (Gtl2) and Peg3 [82]. Although vitrification reportedly disrupts epigenetic regulation, including DNA methylation and histone modifications, direct comparisons remain challenging due to variations in experimental design, species, and analytical techniques. In addition, vitrification can cause vacuolization [83], increase mitochondria-smooth endoplasmic reticulum (M-SER) aggregates and mitochondria-vesicle complexes [39] and alter Ca²⁺ distribution, ultimately leading to morphological abnormalities and reduced developmental competence.

5. Key Factors Affecting Post-Vitrification Survival and Reproductive Success

The success of mammalian oocyte vitrification is determined by a complex interplay of multiple variables rather than any individual factor in isolation. These factors include the developmental stage of oocytes [32], the type and concentration of cryoprotective agents [84,85], vitrification devices and operational conditions (such as equilibration temperature and duration) [86], intracellular lipid content [87], cooling medium temperature [50,88], sample volume [89], physicochemical properties and osmolarity of vitrification solutions, as well as operator dependent technical variability. Therefore, accurate identification and optimization of the key limiting factors are essential for improving vitrification outcomes in mammalian oocytes.

5.1. Oocyte Developmental Stage

Among the factors limiting mammalian oocyte vitrification efficiency, the developmental stage of the oocyte is a key determinant. For farm animals, such as pigs, cattle, and sheep, understanding how oocytes respond to vitrification across developmental stages is crucial for optimizing protocol design and improving survival rates. From a morphological perspective of mammalian oocytes, the ooplasm diameter of immature oocytes is smaller than that of mature oocytes, and immature oocytes have lower developmental competence [90]. In vitrified mouse GV and MII oocytes using the cryolock device (Cat. No. CL-R-CT, Biotech, Inc., California, USA), the incidence of abnormal meiotic spindles was higher in MII oocytes [91]. MII oocytes contain cold-sensitive meiotic spindles, making them more susceptible to low-temperature damage during vitrification, whereas GV oocytes can avoid spindle damage [92,93]. Conversely, when bovine GV and MII-stage oocytes were vitrified, GV-stage oocytes exhibited a lower survival rate (25.90%) compared with MII-stage oocytes (35.60%) [32]. These findings suggest significant differences in cryopreservation efficiency between GV and MII oocytes across mammalian species. This may be attributed to the distinct morphological and structural features of mammalian oocytes at different developmental stages. For instance, compared with mature mammalian oocytes, immature oocytes exhibit characteristics such as reduced microtubule post-translational modifications, lower adenosine triphosphate (ATP) levels, decreased amounts of maturation-promoting factor, impaired glutathione synthesis capacity, and sluggish energy metabolism [94,95], which lead to marked differences in their post-vitrification performance. Therefore, appropriately supplementing exogenous substances according to the structural characteristics of different developmental stages of mammalian oocytes can significantly improve the efficiency of oocyte vitrification.

5.2. Types of Cryoprotectants

CPAs are commonly classified into permeating and non-permeating agents. Permeating CPAs, such as EG and DMSO, inhibit ice crystal formation by disrupting hydrogen bonding between water molecules [96]. Brewer et al. [97] demonstrated that a combination of EG and propylene glycol (PROH) for cryopreserving MII oocytes yielded higher MMP and lower ROS compared to an EG and DMSO regimen. However, the cytotoxic effects of permeating CPAs remain a significant concern in cell freezing. Studies have shown that even low concentrations of DMSO (around 0.1%), particularly in vitro, may induce significant alterations in transcriptomic, proteomic, and epigenetic profiles in human somatic cells [98]. Within the normal concentration range for oocyte vitrification (not exceeding 15%), the inclusion of DMSO exerts highly detrimental effects on oocytes [84]. These findings collectively indicate that the toxic effects of permeable CPAs (DMSO and EG) can impair subsequent oocyte development. Consequently, there is an urgent need to identify non-permeable cryoprotectants that can either fully replace or partially reduce the proportion of permeable CPAs in vitrification protocols.

Non-permeating CPAs, due to their high molecular weight, cannot enter cells easily. Instead, they increase solution viscosity, promote rapid cellular dehydration, and reduce the rate of ice crystal formation. Common non-permeating cryoprotectants include sucrose, ficoll, trehalose, polyvinylpyrrolidone, and antifreeze proteins (AFPs). Such cryoprotectants primarily promote cellular dehydration by increasing extracellular osmotic pressure. Among these, sucrose and trehalose are commonly used [99] and can maintain the balance between the rapid dehydration process prior to cell freezing and the rehydration process after thawing. Supplementing trehalose during vitrification could enhance the membrane stability of ovine oocytes during both cryopreservation and subsequent heat exposure [85]. This effect may be attributed to the ability of sucrose and trehalose to alter the osmotic pressure of the extracellular fluid of oocytes and reduce osmotic damage to oocytes [100]. In summary, during oocyte vitrification, selecting the optimal combination of CPAs based on species-specific characteristics is crucial for improving vitrification efficiency.

5.3. Vitrification Devices and Equilibration Temperature and Times

It is well-established that the selection of vitrification devices during the process determines the cooling and warming rates, which are among the main factors limiting vitrification efficiency. Common vitrification devices include the cryoloop [101], OPS, cryotop, hemi-straw [102], electron microscopy grids [103], solid surface vitrification (SSV) [104], Microfluidic vitrification device [105] and 3D photopolymerized device [106]. Representative images of each device are shown in Figure 2. These devices employ different materials and offer distinct advantages (Table 1). Notably, Yagoub et al. developed a 3D photopolymerization-based device in 2022 capable of achieving a 1000-fold reduction in CPA volume while streamlining the procedural workflow for mouse oocyte vitrification [106]. In addition, researchers have developed automated vitrification devices that enable the automated vitrification of mouse oocytes through linear CPA loading and precise regulation of cryoprotective solution volume, and results showed that oocytes vitrified with this automated device had higher survival (80.44% vs. 73.35%), cleavage (54.17% vs. 43.73%), and blastocyst rates (32.95% vs. 23.67%) compared with the manual cryotop device [107]. In research on mammalian oocyte vitrification, ongoing efforts have aimed to reduce the volume of cryoprotectants to improve vitrification efficiency. There are significant differences in the cooling rates of different vitrification carriers. Simulation experiments have demonstrated that cryoloop has the highest efficiency, reaching up to 100,000 °C/min [108], while the cooling rate of cryotop ranges from 37,500 °C/min [109] to 40,000 °C/min [86]. Although cryoloop offers superior cooling rates, practical applications have demonstrated that cryotop is more suitable for sheep oocytes, with a significantly higher survival rate (83.84%) compared to the traditional straw group (63.43%) [110]. Notably, a higher cooling rate during oocyte vitrification is associated with better vitrification outcomes. This finding accounts for the significant differences in the survival rates of cold-sensitive oocytes observed with different vitrification carriers.

Figure 2.

Representative images of different oocyte vitrification devices: (A) cryoloop, reproduced from [101]; (B) cryotop; (C) open pulled straw (OPS); (D) hemi-straw, adapted from [102]; (E) electron microscopy grids, adapted from [103]; (F) solid surface vitrification (SSV); (G) microfluidic vitrification device, reproduced from [105]: (a) Schematic representation of the vitrification system configuration; (b) The microfluidic chip and Cryotop are fixed on the operation stage; (c) Schematic diagram of the transfer of oocytes to the Cryotop; (H) 3D photopolymerized device, reproduced from [106]. All reproduced/adapted images are distributed under the Creative Commons Attribution 4.0 International License (CC BY 4.0).

Beyond the choice of vitrification carrier, exposure conditions to CPAs also significantly influence the functionality and subsequent development of vitrified oocytes. Among these conditions, exposure temperature and duration are critical, and interspecies differences are particularly pronounced. Prolonging equilibration to 10 min reduced the survival and blastocyst rates of mouse oocytes [111]. Exposing bovine oocytes to equilibration solution for 150 s at 38.5 °C improved oocyte quality and blastocyst rates after vitrification/warming [112]. Before vitrifying goat oocytes with the cryotop device, exposure to 10% DMSO or 10% EG for 1 or 3 min resulted in higher viability and developmental rates [113]. Similarly, vitrified/warmed sheep MII oocytes using a 0.25 mL straw (angled-cut straw tip used as a vitrification carrier) showed no significant difference in survival rates after 5, 7, or 10 min of equilibration [114]. This phenomenon may be attributed to increased oocyte permeability and CPA toxicity at higher temperatures [115,116]. The above studies overlap in their assertion that the efficiency of vitrification is strongly influenced by the equilibration time in the solution and that the vitrification solution should be selected based on the physiological characteristics of mammalian cells to achieve better results.

5.4. Lipid Content

Higher intracellular lipid levels are associated with lower cryotolerance [117]. Under low-temperature conditions, membrane lipids undergo a phase transition from the liquid-crystalline to the gel state [118]. This phase change directly disrupts membrane architecture, thereby interfering with functional contacts among lipid droplets, the endoplasmic reticulum, and mitochondria, ultimately compromising cellular metabolic energy homeostasis [119]. Moreover, the morphology of lipid droplets influences cold sensitivity. In bovine GV-stage oocytes, lipid droplets are structurally simple, appearing as dense and homogeneous spheroids that remain morphologically stable at low temperatures and help maintain membrane integrity, whereas porcine GV-stage oocytes contain structurally complex lipid droplets, including homogeneous dark droplets and grey droplets with electron-lucent striations involved in lipid metabolism, which makes them more susceptible to phase transition and peroxidation during cryopreservation, thereby increasing cryo-damage [120]. To reduce lipid content in oocytes, researchers have developed several strategies, including centrifugation, mechanical delipidation via micromanipulation, and chemical delipidation. A study revealed that mechanical delipidation of porcine oocytes significantly reduced mitochondrial distribution, decreased ROS levels and cleavage rates (delipidated group 21.4% vs. non-delipidated group 10.4%) after vitrification [121]. In addition to the aforementioned factors, selecting an appropriate cooling medium temperature is critical for improving the efficiency of mammalian oocyte vitrification. Liquid helium (LHe), which has a substantially lower temperature than liquid nitrogen (LN₂), has been reported to yield superior cryopreservation outcomes in immature bovine germinal vesicle (GV) oocytes compared with LN₂-based vitrification [50,88]. The volume of the vitrification droplet is another important determinant of vitrification efficiency. In conventional practice, the droplet volume is typically restricted to less than 1 μL to facilitate ultra-rapid cooling and to minimize ice crystal formation [89]. Similarly, the osmolarity of vitrification solutions plays a pivotal role. During the vitrification process, oocytes are exposed to dramatic osmotic changes, transitioning from approximately 280 mOsm (culture medium) to around 2700 mOsm (equilibration solution), and subsequently to about 5600 mOsm in the vitrification solution. The warming process involves a reverse osmotic shift [26]. These abrupt osmotic fluctuations can impose severe stress on the cell membrane, potentially leading to significant cellular damage.

Table 1.

Advantages of different vitrification devices.

Devices	Steps	Advantages	Reference
Cryoloop	Oocytes are transferred onto a thin film supported by a nylon loop, followed by storage in LN2.	This approach requires a minimal volume of cryoprotectant and facilitates rapid cooling.	[122]
open pulled straw (OPS)	Straws are heated-pulled to create a narrow tip, onto which multiple oocytes are loaded before storage in LN2.	Increases cooling rate and reduces cellular damage.	[123]
Cryotop	Under a stereomicroscope, oocytes are loaded onto a plastic strip tip, submerged in LN2, and stored long-term.	Enables ultra-rapid cooling and warming rates.	[86]
Hemi-straw	A 0.25 mL straw is cut into a 1 cm × 0.5 mm strip, onto which oocytes are aligned prior to storage in LN2.	Simple operation with a large contact surface area with LN2.	[86]
solid surface vitrification (SSV)	A solid medium is pre-cooled in LN2, onto which a droplet of CPA–oocyte mixture is deposited for long-term storage in LN2.	Easy to perform and cost-effective.	[124]
Electron microscopy grids	Oocytes are loaded onto electron microscopy grids, blotted to remove excess CPAs, and directly plunged into LN2 for storage.	High-throughput processing capability.	[125]
Automated vitrification	Precise regulation of CPA concentration around oocytes can be achieved through linear loading and removal protocols.	Cryoprotectant volume to automate freezing.	[107]
3D photopolymerized device	Place single cells/embryos in pods, place multiple pods in a Garage, repeat vitrification heating and cycling.	Minimal volume of cryoprotectant required.	[106]

6. Strategies to Improve Oocyte Vitrification Efficiency

Improving the quality of vitrified oocytes is key to advancing the rapid advancement in assisted reproductive technology (ART) and the efficient in vitro production of embryos. Currently, the main strategies to enhance vitrification efficiency include reducing oxidative stress, inhibiting ice crystal formation, suppressing apoptosis, delipidation or lipid content reduction, and stabilizing the cytoskeleton, as well as the application of novel biosynthetic materials and the development and application of novel vitrification devices.

6.1. Reducing Oxidative Stress

During vitrification, excessive ROS generation induces oxidative stress. Numerous studies have reported that supplementing antioxidants such as ascorbic acid, melatonin, L-carnitine, mitoquinone (MtQ), chlorogenic acid, resveratrol, astaxanthin and spermidine can mitigate cryo-induced damage. L-carnitine is a class of small-molecule substances that can promote fatty acid β-oxidation and ATP synthesis. Fatty acids in the cytoplasm cannot directly enter mitochondria for oxidative decomposition and must rely on carnitine, a small-molecule carrier, to translocate into mitochondria [126,127]. MtQ is an antioxidant that can deliver ubiquinone into mitochondria. It alleviates cellular oxidative stress by reducing lipid peroxidation, prevents apoptosis induced by leakage of superoxide radicals from the mitochondrial respiratory chain, and increases intracellular glutathione levels [128,129]. Thus, mtQ acts as an antioxidant during vitrification and plays a critical role in mitigating oxidative damage in cells. Resveratrol and astaxanthin are both common antioxidants; resveratrol can act on multiple molecular targets simultaneously to effectively reduce ROS production in cells, while astaxanthin scavenges ROS and oxygen free radicals and inhibits the process of lipid peroxidation [130,131,132]. The applications of the above-discussed antioxidants in mammalian oocyte vitrification are summarized in Table 2.

6.2. Inhibiting Ice Crystal Formation

AFPs, originally identified in deep-sea fish and subsequently isolated from fungi and plants, have been extensively studied for their ability to modify ice crystal structure and inhibit ice formation [133,134]. AFPs have been successfully used for the cryopreservation of oocytes and embryos across different species. For example, Lee et al. [135] demonstrated that supplementing the vitrification solution with 0.05–0.1 mg/mL of various antifreeze proteins, namely, Flavobacterium frigoris ice-binding proteins (FfIBP), Glaciozyma sp. ice-binding proteins (LeIBP), and AFP-III from bacteria, yeast, and fish during mouse oocyte cryopreservation significantly improved oocyte survival (85.0% vs. 75.0%), cleavage (81.2% vs. 58.7%), and blastocyst formation rates (76.8% vs. 58.7%). Li et al. [136] compared the effects of the straw vitrification device and programmed freezer on the vitrification of ovine oocytes and found that supplementation of 10 μg/mL antifreeze proteins from Anatolia polita (ApAFP914) into the vitrification solution could significantly improve the survival rate of in vitro fertilized ovine embryos (97.17% in the vitrification group vs. 72.47% in the slow freezing group). Leal et al. [137] vitrified feline oocytes, and supplementation with 0.1–1 μg/mL AFP-I increased the cooling rate to 1700 °C/min; compared with the control group, this treatment improved the survival rate of feline oocytes (75% in the control group vs. 89–90% in the AFP I-supplemented groups), enhanced mitochondrial activity, and reduced intracellular ROS levels. However, high production costs limit their widespread use. Notably, synthetic ice crystal inhibitors have also come into view. For instance, Santos et al. [138] found that the synthetic ice blockers Supercool X-1000 (SC) and carboxylated ε-poly-L-lysine (COOH-PLL) exerted beneficial effects in the vitrification of porcine oocytes.

6.3. Suppressing Apoptosis

Apoptosis induced during vitrification can be mitigated using apoptosis inhibitors, thereby improving oocyte survival and developmental competence. Treatment with the pan-caspase inhibitor benzyloxycarbonyl-Val-Ala-Asp -fluoromethyl ketone (Z-VAD-FMK) reduced DNA damage and caspase activity in vitrified feline oocytes, yielding a maturation rate (53.13%) comparable to fresh controls (65.38%) and similar cleavage rates (34.38% vs. 31.78%) [139]. These findings substantiate that excessive caspase activation is a key mediator of cryo-induced injury. However, it should be noted that caspase inhibition primarily targets downstream apoptotic execution and may not fully prevent upstream cellular stress triggered by vitrification. Pre-treatment of bovine oocytes with a 10 μM Rho-associated coiled-coil kinase (ROCK) inhibitor (R)-(+)-trans-N-(4-pyridyl)-4-(1-aminoethyl)cyclohexanecarboxamide (Y-27632) for 2 h significantly improved survival, cleavage, and blastocyst rates after vitrification [140]. Thus, adding apoptosis inhibitors during vitrification and warming reduces activation of mitochondrial and extrinsic apoptotic pathways, minimizes cellular damage, and enhances oocyte cryosurvival and developmental potential.

6.4. Delipidation or Lipid Reduction

Given the high lipid content and extreme cold sensitivity of oocytes, reducing cytoplasmic lipid levels is an effective strategy to improve cryotolerance. Although mechanical delipidation via microinjection through the zona pellucida effectively reduces lipid content, the procedure poses a significant risk to cellular structural integrity [141]. Alternatively, adding lipolytic agents to oocyte culture media can safely reduce lipid content and improve freezing tolerance. Fu et al. [142] demonstrated that supplementing maturation media with 10 μM forskolin significantly decreased lipid levels and increased post-vitrification survival. During the vitrification of feline oocytes, supplementation with 2 mg/mL levocarnitine can significantly improve the survival rate and ATP levels of vitrified oocytes and reduce intracellular ROS levels [143]. During the vitrification of bovine oocytes using the OPS device, supplementing with 1 μM β-nicotinamide mononucleotide (NMN), 2.5 μM berberine (BER), or 1 μM cordycepin reduced lipid content, downregulated lipid synthesis-related genes, and decreased ROS and apoptosis levels [15]. During the vitrification of feline oocytes, reducing lipid content using lipid modulators such as conjugated linoleic acid, forskolin, and L-carnitine improved oocyte viability (74% in the experimental group vs. 53% in the control group) [144]. Forskolin acts as a lipolytic agent by stimulating adenylate cyclase, thereby reducing intracellular lipid levels. In contrast, carnitine functions as a lipid metabolism enhancer, facilitating the transport of long-chain fatty acids into mitochondria for β-oxidation and promoting mitochondrial energy metabolism [145,146]. However, several studies have also indicated that supplementation with fatty acids during the vitrification of mouse oocytes or 4-cell embryos can increase neutral lipid content in mouse oocytes, improve the blastocyst development rate, and upregulate the expression of genes related to fatty acid β-oxidation in mouse embryos [147]. Fatty acid β-oxidation is a metabolic process that occurs within mitochondria, in which fatty acids are broken down into acetyl-CoA. This process promotes ATP production via the electron transport chain [148,149], thereby supporting cellular energy metabolism after thawing. Therefore, the use of small-molecule lipid-reducing agents (Forskolin, levocarnitine) is a promising approach to enhance vitrification outcomes.

Table 2.

The effects of antioxidant supplementation on the vitrification of mammalian oocytes or embryos.

Antioxidants	Research Subject	Dose	Research Findings	Reference
Ascorbic acid	Mouse 2-cell embryos or blastocysts	0.1 mM	Ascorbic acid reduces hydrogen peroxide levels in embryos, enhances inner cell mass development, and decreases lactate dehydrogenase activity.	[150]
Melatonin	Human oocytes	10−9 M	Addition of 10−9 M melatonin during human oocyte vitrification significantly reduces intracellular ROS and Ca2+ levels, maintains membrane integrity, and mitigates oxidative stress.	[151]
Astaxanthin	Porcine oocytes	2.5 μM	Astaxanthin improves the survival rate of vitrified oocytes, reduces ROS levels, increases glutathione levels, and enhances lysosomal fluorescence intensity.	[152]
mitoquinone (MtQ)	Mouse Oocytes	0.02 μM	MtQ improves post-warming survival, enhances MMP, and reduces the Bax/Bcl2 ratio and caspase3 expression.	[153]
L-carnitine	Porcine oocytes	10 mM	L-carnitine significantly reduces lipid droplet content and restores SOD1 expression in vitrified oocytes; however, it does not improve overall cryopreservation outcome.	[154]
Resveratrol	porcine oocytes	2 μM	Resveratrol improves the quality of vitrified porcine oocytes and regulates apoptosis.	[11]
Chlorogenic Acid	Ovine oocytes	40 μM	Chlorogenic acid mitigates oxidative stress, enhances mitochondrial function, and downregulates the expression of apoptosis and antioxidant-related genes in vitrified oocytes.	[17]
Resveratrol	Bovine oocytes	1 μM	Resveratrol reduces the incidence of abnormal spindles and lowers ROS levels in vitrified oocytes to physiological ranges, thereby enhancing cryopreservation efficiency.	[155]
Spermidine	Mouse oocytes	50 μM	Spermidine improves the survival and blastocyst formation rates of vitrified mouse oocytes and restores the expression levels of 43.3% of dysregulated genes.	[23]

6.5. Stabilizing the Cytoskeleton

The addition of cytoskeletal stabilizers during oocyte maturation has been widely reported to improve cryosurvival by maintaining cytoskeletal integrity. Commonly used stabilizers include cytochalasin B (CB), cytochalasin D, and paclitaxel. Pretreating buffalo oocytes with 8 μg/mL cytochalasin B before vitrification attenuated the reduction in tubulin expression and improved development to the 8-cell and blastocyst stages [156]. Moawad et al. [157] found that treating sheep GV oocytes with 7.5 μg/mL CB prior to vitrification improved post-warming cleavage rates.

Based on the cryotop vitrification of mouse oocytes, supplementation with docetaxel alleviates vitrification-induced damage by modulating the expression of apoptosis-related genes [158]. Existing studies have demonstrated that the addition of cytoskeletal stabilizers (e.g., cytochalasin B, docetaxel, etc.) during oocyte maturation can improve their survival rate and subsequent developmental potential by maintaining cytoskeletal integrity and regulating the expression of apoptosis-related genes [159], with the efficacy varying across different species and stabilizer concentrations. Future research should prioritize the development of novel, low-toxicity cytoskeletal stabilizers and integrate multi-omics frameworks to delineate their underlying molecular mechanisms. Such efforts are essential for the precise optimization of oocyte cryopreservation protocols across diverse species.

6.6. Application of Novel Biosynthetic Materials

Over the years, novel biosynthetic materials, such as nanomaterials, have been widely applied in oocyte vitrification. Importantly, novel ice-inhibiting nanomaterials, including hydroxyapatite (HA) nanoparticles, possess excellent biocompatibility, antioxidant properties, thermal conductivity, and membrane permeability. Studies have found that the addition of HA nanoparticles to vitrification solutions can significantly improve the cryosurvival rate of porcine MII oocytes reported that the survival rate increased from 14.7% to 25.9–35.4% [160]; its protective mechanism does not rely on enhancing cooling rates but rather on inhibiting ice crystal formation and recrystallization, thereby alleviating mechanical damage to cells during the freezing–thawing process [160,161]. Liu et al. [162] reported that during the vitrification of ovine oocytes using cryotop devices, supplementation with HA nanoparticles reduced ROS and apoptosis levels, increased cleavage rates, and restored MMP post-vitrification. Studies have shown that nanoparticles can scavenge intracellular oxygen-free radicals, reduce intracellular ROS levels, modulate MMP levels, and alleviate oxidative stress [163,164]. In addition, antioxidant nanomaterials, such as Fe₃O₄ magnetic nanoparticles and nanocarriers loaded with resveratrol [13] or melatonin [165,166] have been shown to alleviate oxidative stress during cryopreservation. Fe₃O₄ nanoparticles significantly improved nuclear maturation and embryonic development in mouse immature oocytes [167]. These findings collectively indicate that novel biomaterials have extremely broad application prospects in the vitrification of mammalian oocytes and are highly beneficial for improving oocyte vitrification efficiency.

6.7. Development of Novel Vitrification Devices

With the continuous advancement in science and technology, emerging vitrification devices have been increasingly developed and applied to oocyte vitrification preservation. Researchers have successively developed automated devices for oocyte or embryo vitrification that help mitigate osmotic damage during CPA loading [168,169]. Secondly, the gradual progression of ultra-rapid vitrification has opened up a novel avenue for oocyte vitrification. Ultra-rapid vitrification minimizes CPA volume and attenuates its cytotoxicity [56], thereby improving the post-vitrification survival rate of oocytes.

The integrated application of these strategies significantly improves the efficiency of oocyte vitrification in mammalian species (Figure 3). Furthermore, the implementation of fully automated vitrification systems helps to minimize cellular or embryonic damage induced by manual manipulation. Concurrently, as artificial intelligence continues to advance, vitrification robots specifically engineered for mammalian oocytes or embryos and powered by large language models are anticipated to catalyze an industrial transformation in cryobiology.

Figure 3.

A schematic diagram of combined strategies for enhancing the developmental potential of vitrified oocytes. The integrated approach combines fully automated vitrification devices with targeted intracellular and extracellular protective strategies to mitigate cryodamage, ultimately enhancing oocyte viability and post-thaw embryonic development rates following in vitro fertilization/intracytoplasmic sperm injection.

7. Comparative Analysis of Oocyte Vitrification Across Different Species

The vitrification process for oocytes from different species shares a common procedural framework consisting of three key steps: equilibration, vitrification, and warming [108,140]. During equilibration, oocytes are briefly exposed to an equilibration solution (ES) containing permeating CPAs (typically EG and DMSO), then transferred to a vitrification solution (VS) with higher concentrations of these permeating CPAs combined with non-permeating CPAs such as sucrose or trehalose to induce rapid dehydration [86,112]. Subsequently, oocytes are loaded into an ultra-minimal volume (<1 μL) on a vitrification device and immediately plunged into liquid nitrogen for storage; warming typically involves stepwise sucrose dilution to alleviate osmotic stress [86]. Due to interspecies variations in oocyte physiology, the optimal CPA composition and equilibration duration differ markedly. In mice, oocytes are briefly equilibrated 30 s in 10% (v/v) EG and 10% (v/v) DMSO, followed by a short exposure of 25 s to 15% (v/v) of EG and DMSO with sucrose [13,170]. In humans, equilibration is markedly longer, typically around 15 min in 7.5% EG and 7.5% propanediol (PROH), prior to transfer into a VS supplemented with sucrose or trehalose and serum substitutes, with stepwise sucrose dilution during warming [25]. In livestock species, protocol parameters are further adjusted. Ovine oocytes typically require intermediate equilibration of about 1 min and exposure to higher cryoprotectant concentrations [171], whereas bovine oocytes show protocol variability depending on the carrier system (e.g., Cryotop vs. OPS), with corresponding differences in VS composition and exposure time [16]. Porcine oocytes, characterized by large size and high lipid content, require the longest equilibration time, typically 5 to 15 min, and more complex warming procedures involving higher initial sucrose concentrations and gradual dilution [11]. The general workflow for the vitrification and recovery of oocytes across diverse species is illustrated in Figure 4. These interspecies discrepancies are predominantly attributable to variations in oocyte lipid profiles and membrane composition. Phospholipids, particularly phosphatidylcholine and sphingomyelin, modulate membrane fluidity and permeability [172], thereby governing water and cryoprotectant exchange [119]. Substantial intracellular lipid levels, particularly in porcine oocytes, reduce membrane permeability and prolong equilibration requirements.

Figure 4.

Operational procedure of mammalian oocyte vitrification and thawing. Following in vitro maturation, oocytes are generally equilibrated in equilibration solution (ES) for 30 s ~15 min, and then exposed to vitrification solution (VS) for 20~60 s, loaded onto a cryotop, and rapidly plunged into liquid nitrogen. During warming, oocytes are sequentially transferred to a trehalose/sucrose (TS) solution for 1~3 min, followed by a dilution solution (DS) for 3~5 min, and a wash solution (WS) for 1~10 min, before being placed into recovery medium for 1~3 h.

Due to differences in lipid content among oocytes of different species, their vitrification efficiency also varies [37]. With a lipid content 3–5 times that of cattle and sheep [144], porcine oocytes undergo lipid phase transitions during the freezing process, resulting in a vitrification survival rate of only 58–65%, with cleavage and blastocyst rates of 35–40% and 2–5%, respectively [18,19,20]. Mouse and human oocytes have low lipid content and smaller volumes, resulting in higher vitrification survival rates. Studies have indicated that mouse oocytes exhibit vitrification survival rates of 82–90%, cleavage rates above 59–68%, and blastocyst rates of 20–37% [22,23,24]. The survival rate, good-quality embryo rate, and blastocyst rate of vitrified human oocytes were 94–100%, 54–71%, and 31.7–34.9%, respectively [25,173]. The primary reason for these differences lies in the substantial variation in lipid composition and content among oocytes; the higher the cellular lipid content, the lower the tolerance to low temperatures. During vitrification, lipid phase transitions compromise cell membrane integrity, inducing apoptosis and cell death [118]. Additionally, the types of lipids that constitute the cell membrane influence its tolerance to low temperatures. Research has found that membranes with high cholesterol and low phospholipid content are less sensitive to temperature changes, and that the higher the cellular lipid content, the lower the tolerance to low temperatures [174]. This may account for some of the differences in vitrification efficiency across species.

8. Discussion

Vitrification currently represents the preferred strategy for oocyte cryopreservation, surpassing slow-cooling protocols by effectively mitigating cryoinjury and maintaining cellular architecture. This transition has solidified its role as a pivotal tool in both clinical reproductive medicine and the conservation of biodiversity via endangered species germplasm banks [1,175]. Although vitrification has been widely accepted, several controversial issues persist in its application. On one hand, the relative merits of open versus closed vitrification systems remain debated: some studies suggest that closed systems prevent liquid nitrogen-mediated pathogen contamination and offer greater safety [176], whereas others report no significant differences between the two systems in terms of post-warming survival rates or developmental competence [177]. On the other hand, there is disagreement regarding the optimal developmental stage for oocyte vitrification. The mature MII stage allows direct use for in vitro fertilization with a straightforward workflow, while the immature GV stage theoretically reduces the risk of aneuploidy by avoiding spindle depolymerization induced by cryopreservation [92,93]. These discrepancies primarily stem from heterogeneity in experimental protocols, such as the type and concentration of cryoprotectants and cooling rates, as well as species-specific factors [37,178]. Current literature lacks sufficient longitudinal data on offspring health and epigenetic risks. Moreover, the precise mechanisms underlying cryoinjury-induced ultrastructural defects, such as mitochondrial and cytoskeletal abnormalities, require further elucidation [12]. Addressing these limitations will be essential for developing species-tailored vitrification strategies and improving reproductive outcomes across diverse mammalian systems.

In the future, the adoption of vitrification instruments and automated vitrification devices will enable the precise addition and removal of cryoprotectants at the nanoscale, while uniform warming processes will facilitate the standardization of vitrification techniques. Cryoprotectant-free strategies or non-permeable cryoprotectants such as trehalose and antifreeze proteins may gradually replace sucrose, enabling cell cryopreservation without chemical toxicity and eliminating cytotoxic effects. This technology will extend beyond assisted reproduction to broader applications, including establishing “germplasm banks” for endangered species, thereby supporting biodiversity conservation. In summary, through interdisciplinary integration, oocyte vitrification is evolving toward safer, more efficient, and more intelligent approaches, opening new frontiers in reproductive medicine and species conservation.

9. Conclusions

Oocyte vitrification is the primary method for preserving germplasm in female animals and has been successfully applied in multiple mammalian species. It is widely recognized that the vitrification process induces osmotic stress and cryoprotectant toxicity, leading to cellular damage that compromises developmental competence. However, the molecular mechanisms underlying these injuries, particularly their long-term epigenetic consequences, remain largely unknown. Whether oxidative stress, mitochondrial dysfunction, and apoptosis occur independently or interact to cause cellular damage remains unclear. It also remains unexplored whether epigenetic alterations induced by vitrification can be transmitted to offspring and affect their health. Key priorities include decoding the molecular and epigenetic impacts of cryoinjury, standardizing oocyte vitrification through automation, and developing biocompatible cryoprotectants to ensure precise delivery and minimal toxicity. Addressing these challenges is essential for improving the efficiency of oocyte vitrification in mammals, particularly in large animals such as pigs, cattle, and sheep, for accelerating genetic improvement in farm animals, and for facilitating the preservation of germplasm resources from endangered species.

Abbreviations

The following abbreviations are used in this manuscript:

CPAs	Cryoprotectants
EG	Ethylene glycol
DMSO	Dimethyl sulfoxide
GV	Germinal vesicle
MII	Metaphase II
OPS	Open pulled straw
NADPH	Reduced nicotinamide adenine dinucleotide phosphate
MMP	Mitochondrial membrane potential
ATP	Adenosine triphosphate
ICSI	Intracytoplasmic sperm injection
ROS	Reactive oxygen species
SSV	Solid surface vitrification
ZP	Zona pellucida
MtQ	Mitoquinone
IGF2R	Insulin-like growth factor 2 receptor
PPP1R9A	Protein phosphatase 1 regulatory subunit 9A
PEG3	Paternally expressed gene 3
H3K9ac	Histone H3 lysine 9 acetylation
H4K5ac	Histone H4 lysine 5 acetylation
H4K21ac	Histone H4 lysine 21 acetylation
H3K9me3	H3 lysine 9 trimethylation
LDHA	Lactate dehydrogenase A
LDHA	Lactate dehydrogenase B
EP300	E1A binding protein p300
SIRT1	Sirtuin1
Gtl2	Gene trap locus 2
FfIBP	Flavobacterium frigoris ice-binding proteins
LeIBP	Glaciozyma sp. ice-binding proteins
ApAFP914	Antifreeze proteins from Anatolia polita
CB	Cytochalasin B
HA	Hydroxyapatite
Z-VAD-FMK	Benzyloxycarbonyl-Val-Ala-Asp -fluoromethyl ketone
Y-27632	(R)-(+)-trans-N-(4-pyridyl)-4-(1-aminoethyl)cyclohexanecarboxamide

Author Contributions

Conceptualization, X.L. and T.Y.; Writing—original draft preparation, K.S.; Project administration, X.H. and R.Q.; Visualization, L.W., F.Y., Q.M. and J.B.; Investigation, L.W. and S.G.; Resources, H.Z. and Q.M.; Supervision, F.Y. and R.Q.; Formal analysis, H.Z., X.H. and J.B.; Funding acquisition, T.Y.; Writing—review & editing, S.G., X.L. and T.Y. All authors have read and agreed to the published version of the manuscript.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available in this article.

Conflicts of Interest

The co-author Hongqian Zhu is an employee of Muyuan Foods Co., Ltd. The authors declare that there are no other conflicts of interest.

Funding Statement

This research was funded by National Key R&D Program of China (2024YFD1301002), National Natural Science Foundation of China (32202671), China Postdoctoral Science Foundation (2023M730999), Key Science and Technology Research Projects in Henan Province (242103810023), Henan Province Higher Education Institutions Key Scientific Research Project Service Industry Specialization (25CY014).