Morphometric Assessment of the Bovine Ovary for in vitro Matured Oocyte Quality to Determine Developmental Competence

Embryo production in vitro requires three steps: in vitro maturation (IVM), in vitro fertilization (IVF) and in vitro culture (IVC). Oocyte developmental competence is a determining factor that influences the outcomes of an IVF cycle regarding the ability of a female gamete to reach maturation, be fertilized and uphold embryonic development until the blastocyst stage (Sirait et al., 2021). The first step in the in vitro maturation of oocytes is germinal vesicle breakdown (GVBD), followed by completion of the 1^st meiotic division and formation of the 1^st polar body. The most important step of in vitro production (IVF) is selecting good-quality oocytes that can undergo nuclear and cytoplasmic maturation, which occurs during follicle growth (Sirard, 2001).
       
Nuclear maturation is the ability to resume meiotic division up to metaphase II by initiating the breakdown of germinal vesicles (GVBDs) and formation of the 1^st polar body. Secondary oocytes are then extrused from Graafian follicles and the second metaphase plate of the oocytes appear (Watson, 2007).
       
Cytoplasmic maturation includes a succession of transformations, essentially of mitochondria, cortical granules and smooth and rough endoplasmic reticulum (Hyttel et al., 1997), all of which are necessary for the maturation and blockade of polysperm. These events indicate that new protein synthesis occurs, such as the “factor of the growth of masculine pronucleus” MPGF and maturation-promoting factor MPF.
       
In addition to these phenomena, the mucification and expansion of the cells surrounding the oocyte and reduction in the number of intercellular junctions between the granulosa cells and oocytes begin, interrupting ionic transport between the cells of the cumulus and oocytes (Hyttel et al., 1997). Cytoskeletal proteins (Sun and Schatten, 2006) and organelle distribution (Brevini et al., 2007) are crucial for oocyte maturation and may affect oocyte competence. Kim et al., (1996, 2000) demonstrated the close relationship between nuclear maturation and cytoskeleton dynamics in pig and cattle oocytes. Cytoskeleton dynamics are also related to oocyte developmental competence (Brevini et al., 2007). Correct organelle positioning is also critical because low-quality bovine oocytes fail to translocate to the mitochondria from the cytoplasm periphery to the center (Stojkovic et al., 2001). Bilodeau-Goeseels, (2006) reported that glucose, pyruvate, lactate and glutamine were stimulatory to nuclear maturation in bovine oocytes. Insufficient cytoplasmic maturation of M II oocytes may be one reason for the low rate of embryo production in vitro (Marchal et al., 2001).
       
Dairy cattle have a variety of health issues, the most prevalent of which are mastitis, which may be either obviously or subclinical (Chouhan et al., 2021) and cystic ovarian disease, which is a widespread and economically important ailment in dairy cattle that affects fertility (Chauhan et al., 2021) and do not forget the role of the season on dairy cows and their production (Kumar et al., 2021).
       
Therefore, the crucial step in selecting bovine oocytes requires careful morphological measurement to predict subsequent developmental competence, particularly when the available oocytes are limited. The morphometric selection of in vitro matured oocytes has been reported in humans and several animal species (Kitagawa and Nimura, 2006; Yousaf and Chohan, 2003; Lasiene et al., 2011). Therefore, this study aimed to determine morphometric dimensions of bovine ovaries and oocytes, such as the ovary weight and diameter and different types of follicles, including the layer of follicle cells surrounding the oocytes and the thickness of the zona pellucida around the oocytes. Additionally, we evaluated the ooplasm and perivitelline space with or without the first polar body to increase the probability of oocytes that could support in vitro embryonic development.

Unless otherwise specified, all the chemicals and hormones used in this study were obtained from Sigma-Aldrich Corp. (St. Louis, MO, USA).
       
Cow ovaries were obtained from a native slaughterhouse in Riyadh and were transported in 0.9% (v/v) NaCl solution at 30-33°C to the laboratory within 2-3 h. The contents of antral follicles were aspirated using a 19-gauge needle attached to a 10 mL disposable syringe containing 0.5 ml of tissue culture medium (TCM-199 with Hank’s salts, L-glutamine, 25 mM HEPES and L-amino acids supplemented with 10% fetal calf serum (FCS), 0.3 mM sodium pyruvate, 25 mg/ml of gentamycin sulfate and 143 mg/ml of heparin sodium salt (Caisson Lab. Inc., Smithfield, UT, USA). Cumulus-oocyte complexes (COCs) with uniformly granulated cytoplasm and more than three layers of compact cumulus cells were chosen and evaluated (Al mutary et al., 2020).
       
COCs with more than 3 layers of cumulus cells and uniform cytoplasm (n ¼ 2641) were matured for 24 h in TCM-199 Earle’s salts supplemented with 10% fetal bovine serum (FBS), 0.3 mM sodium pyruvate, 1 mg/ml of estradiol-17b, 0.023 IU/ml of luteinizing hormone (LH), 0.02 IU/ml of follicular stimulating hormone (FSH) and 25 mg/ml of gentamycin sulfate at 38.5°C and exposed to 5% carbon dioxide (CO₂) in air at high humidity. The oocytes (10-15 per droplet) were cultured in 60 mm Petri dishes containing 10 drops, each 50 ml in size and were covered with embryo-tested mineral oil (Al mutary et al., 2020).
       
The cumulus cells were removed from the oocytes after IVM. Trituration was performed in HEPES-buffered TCM-199 with 0.1% (v/v) hyaluronidase and two washes in TCM-199 augmented with 10% (v/v) FBS. The morphometric parameters of the oocytes were assessed according to previous studies (Otoi et al., 1997; Griffin et al., 2006). Images were captured using a camera and the software of the microscope (Leica Application Suite, Version 4.0) and the morphometric parameters of the captured images were analyzed using Image 1.50i software (NIH, USA) and the scale bar as an arbitrary scale for pixel analysis by the software in the same oocyte image and all the results were recorded. The morphometric parameters described are as follows: oocyte outer diameter (ZPO), zona pellucida thickness (ZPT), inner oocyte diameter (ZPI) and ooplasm diameter (OD).

The collected ovaries were divided into normal and abnormal ovary types, according to their morphological structure with a corpus luteum or large vacuole follicles (Fig 1a). Statistical analysis of the mean ovary weight showed no significant difference (13.4±1.73 g and 18.45±1.38 g) between the normal and abnormal ovaries, respectively (Table 1). Additionally, the volume, length and width of the ovaries were not significantly different between the ovary types (Table 1). Furthermore, the dominant follicle diameter in both ovary types showed no significant differences (8.41±0.88 vs. 8.52±0.99 mm). Additionally, the subordinate follicles in both ovary types showed significant differences in the diameter (1.65±0.16 and 0.16 mm) and their number per ovary (9.5±1.4 vs. 8.55±1.58 mm) of abnormal subordinate follicles (Table 1).  
 

 

       
Before maturation of the cumulus oocytes, the complex showed no significant differences in the diameter (normal: 52.73±7.23 mm; abnormal: 43.015±5.41 mm). However, after maturation, a highly significant difference (P<0.001) was found in the diameter between the normal (178.10±15.36 µm) and abnormal ovaries (10.45±7.99 µm). Additionally, maturation between the normal and abnormal conditions (P<0.001) (Table 2) will affect post maturation steps such as IVF and later embryo development in vitro. Data analysis of the oocyte parameters of normal and abnormal ovaries revealed similar findings, such as those regarding the outer oocyte diameter of normal (189.016±2.24 µm) and abnormal (188.91±1.77 µm) ovaries. The oocyte zona pellucida thicknesses in normal and abnormal oocytes were 17.017±0.59 µm and 17.245±0.73 µm, respectively. The ooplasm diameters in normal and abnormal oocytes were 134.82±1.56 µm and 137.63±1.51 µm, respectively. The sizes of the perivitelline space in normal and abnormal oocytes were 55.09±2.26 µm and 51.22±1.30 µm, respectively (Table 2). Data analysis of the oocytes with or without 1^st polar bodies showed a highly significant difference (P<0.001) between the oocytes with (32.15±4.19) or without (10.95±1.59) 1^st polar bodies in normal ovaries and oocytes with (7.0±078) or without (3.3±0.32) 1^st polar bodies in abnormal ovaries.
 

       
The best selection of oocytes in vitro from normal good ovaries will affect the subsequent steps in in vitro maturation, in vitro fertilization or cloning and in vitro embryo culture and increase embryo development quality. Some studies have shown that the corpus luteum in an ovary enhances the quality and developmental competence of oocytes derived from that ovary (Reis et al., 2006; Moreno et al., 1993; Penitente-Filhe et al., 2015). However, our study findings contrast those of other studies (Shabankareh et al., 2015; Quezada-Casasola et al., 2018), as well as subsequent embryonic development (Gonzalez-Bulnes et al., 2005; Manjunatha et al., 2008; Boediono et al., 1995; Pirestani et al., 2011). Additionally, several studies have reported no effects (de Wit et al., 2000; Sungulle, 2008). Bovine oocytes with a diameter of 110 mm can achieve complete meiotic competence, while smaller oocytes have significantly less transcriptional activity, indicating that they are still in the growing phase (Fair et al., 1995).
       
Additionally, oocytes with diameters of 110-120 and 120 µm reached MII in similar proportions (76 and 81%, respectively) (Fair et al., 1995). Otoi et al., (1997) indicated that meiotic competence was attained once oocytes reached 115 µm in diameter, while full developmental capacity was obtained when the diameter was at least 120 µm. Bovine oocytes with an inside-zona diameter smaller than 95 μm cannot resume meiosis in vitro; however, a high proportion of bovine oocytes can resume meiosis. Otoi et al., (1997) showed that the oocyte must measure 110 μm or higher to reach the MII stage. In the present study, all mature oocytes from both normal and abnormal ovaries had ZP thicknesses of 17.017±0.59 and 17.245 μm, respectively, values similar to those reported by Cavelia et al., (2008). Additionally, bovine oocytes with an inside-zone diameter smaller than 95 mm could not resume meiosis in vitro Sungulle et al., (2008).
       
No significant differences were found in the outer oocyte diameter, ooplasm diameter or oocyte periventricular space between bovine oocytes collected from normal and abnormal ovaries. Other studies have indicated that nuclear maturation, such as that in pigs (Luca et al., 2002), buffalos (Yousaf and Cohan, 2003) and camels (Saadeldin et al., 2017) and blastocyst production in cows were positively correlated with the oocyte diameter (Otoi et al., 1997; de Wit and Kruip, 2001; Arlotto et al., 1996). The same parameters employed in evaluating oocyte morphology in human oocytes by Lasiene et al., (2011) include the appearance of the structure of the cumulus-oocyte complex, oocyte cytoplasm, perivitelline space, zona pellucida, polar body and meiotic spindle.
       
The lower rate of in vitro fertilization and in vitro embryo development could be because most oocytes derived from the ovary for in vitro maturation originate from subordinate or growing follicles that are at least viewed days away from any possible ovulation. Although most of these oocytes complete their nuclear maturation, few develop to the blastocyst stage.

The critical point at which the normal ovary produces better in vitro matured follicles with good oocyte quality and produces the 1st polar body determines developmental competence. Therefore, the best selection of normal ovaries will enhance in vitro maturation for subsequent experiments, such as in vitro fertilization or cloning and in vitro embryo development.

The authors thank the Researchers Supporting Project for funding this work (number RSP-2021/232) at the King Saud University, Riyadh, Saudi Arabia.

None.

This research was funded by the Researchers Supporting Project (number RSP-2021/232), King Saud University, Riyadh, Saudi Arabia.