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. 2020 Nov 21;7(2):465–473. doi: 10.1002/vms3.378

Identification of optimal assisted aspiration conditions of oocytes for use in porcine in vitro maturation: A re‐evaluation of the relationship between the cumulus oocyte complex and oocyte quality

Giuseppe Silvestri 1, Claudia C Rathje 1, Simon C Harvey 2, Rebecca L Gould 3, Grant A Walling 4, Peter JI Ellis 1, Katie E Harvey 2,✉, Darren K Griffin 1
PMCID: PMC8323607

Abstract

The quality of porcine oocytes for use in IVF is commonly graded according to the number of layers of cumulus cells (CCs) surrounding the oocyte; together these form the cumulus oocyte complex (COC). At least three compact layers of CCs is regarded as important for efficient IVP. To test this, oocytes were scored according to cumulus investment, with grade A representing COCs with three or more cumulus layers including granulosa cell‐cumulus oocyte complexes, grade B those with an intact corona radiata surrounded by another layer of cumulus cells and grades C and D representing COCs with lower CC investment. These oocytes were then monitored for in vitro maturation (IVM), as assessed by tubulin immunostaining for meiotic progression, the development of a cortical granule ring, and by glutathione levels. Results indicate that grading correlates closely with nuclear maturation and cytoplasmic maturation, suggesting that grading oocytes by cumulus investment is a reliable method to predict IVM success. Importantly, Grade A and B oocytes showed no significant differences in any measure and hence using a cut‐off of two or more CC layers may be optimal. We also determined the effect of assisted aspiration for oocyte retrieval, comparing the effect of needle size and applied pressure on the retrieval rate. These data indicated that both variables affected oocyte recovery rates and the quality of recovered oocytes. In combination, these experiments indicate that grade A and B oocytes have a similar developmental potential and that the recovery of oocytes of these grades is maximised by use of an 18‐gauge needle and 50 mmHg aspiration pressure.

Keywords: In vitro production, Morphology, Oocyte, Pig

The global population is expected to rise dramatically over the next few decades. This means that the demand for meat, including pig derived products will also increase, and that a more sustainable way to produce such products is necessary. The establishment of production scale pig embryo in vitro production (IVP) could be one answer to this problem, however some challenges still exist, one of which is that pig IVP requires the consistent isolation of large numbers of high‐quality oocytes. Here, ways of maximising both the yield and the developmental potential of oocytes from abattoir derived ovaries has been investigated.

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1. INTRODUCTION

Pig embryo in vitro production (IVP) often involves oocyte collection from the ovaries of abattoir animals. Usually, oocyte donors will not have undergone pharmacological treatments to regulate or induce the production of mature oocytes. As such, their gametes will be immature and require laboratory culture, a process known as in vitro maturation (IVM). Following slaughter, oocyte retrieval can be achieved using either follicular aspiration or ovary slicing. Aspiration methods can be either manual, using a syringe barrel and needle, or assisted, using a negative pressure aspiration pump and attached needle. Automated aspiration procedures generally allow for a more consistent collection environment when compared to manual aspiration, which is prone to inter‐operator variability (Marques et al., 2015). The oocytes retrieved for IVM are found in combination with cumulus cells (CCs), forming the cumulus oocyte complex (COC). CCs and oocytes share a complex network of interactions (Gilchrist et al., 2004) and there is a strong correlation between the number of CCs and the ability of an oocyte to complete both nuclear and cytoplasmic maturation (Dang‐Nguyen et al., 2011; Lin et al., 2016; Nagai et al., 1993).

The oocyte is maintained in meiotic arrest by the CCs, which supply it with stable levels of the meiotic progress inhibitor cyclic adenosine monophosphate (Anderson & Albertini, 1976; Racowsky, 1985) and of the phosphodiesterase inhibitor, cyclic guanosine monophosphate (Norris et al., 2009). Interestingly, the CCs also control meiosis resumption in response to high luteinising hormone levels (Mattioli & Barboni, 2000; Norris et al., 2008). Further to this, CCs promote the migration of cortical granules (CGs) towards the periphery of the oocyte (Galeati et al., 1991), a key element of cytoplasmic maturation. Cytoplasmic maturation has also been shown to be enhanced by high levels of glutathione (GSH) which CCs actively synthesise and transport to the oocyte (Maedomari et al., 2007; You et al., 2010), where it acts as a scavenger of reactive oxygen species (ROS) (Tatemoto et al., 2000) and increases amino acid transport and protein synthesis (Lafleur et al., 1994). Furthermore, high levels of GSH seem important for correct male pronuclear formation upon fertilisation (Niwa, 1993; Yoshida et al., 1993). Due to the many functional roles of CCs, the morphology of the COC is commonly used to determine candidates for IVM in pigs and in other farm animals (Alvarez et al., 2009; Nagano et al., 2006). Laboratory‐based retrieval methods and aspiration pressures used in ovum pickup from live animals have shown that aspiration pressure also has an impact on the morphology of the COC (Brüssow et al., 1997; Marques et al., 2015).

In pigs, full oocyte meiotic competence is achieved in follicles with a diameter of more than three mm and a positive correlation has been observed between follicle size and oocyte in vitro developmental competence (Marchal et al., 2002). Interestingly, oocytes from gilts display a reduced developmental potential when compared to oocytes from sows (Lechniak et al., 2007); this may be because the average follicle size is smaller in gilts (Bagg et al., 2007). Given these results, primary oocytes for pig IVP are usually recovered from follicles ranging between 3 and 8 mm (Bagg et al., 2007); it is however difficult for operators to judge follicle size accurately (Lin et al., 2016), and as such appropriate criteria for post retrieval oocyte selection are essential. Even though it is often recommended that only COCs formed of multiple compact layers of CCs should be selected for IVM (Bagg et al., 2007; Esaki et al., 2004; Fowler et al., 2018; Lee et al., 2012; Lin, Lee, Shin, Oqani, & Jin, 2015; Long et al., 1999; Rath et al., 1995; Sherrer et al., 2004), this practice results in wastage as COCs formed of three of more layers form only a proportion of the total yield (Lin et al., 2016). With this in mind, here we investigate how maturation, as assessed by meiotic progression, the development of a CG ring, and by GSH levels, varies in COCs with different numbers and morphologies of their CC layers. We have also investigated how specific combinations of aspiration pressure and needle gauge affect COC recovery and morphology. These analyses indicate that oocytes with three or more cumulus layers including granulosa cell‐cumulus oocyte complexes (Grade A) and those with intact corona radiata surrounded by another layer of CCs (Grade B) have similar developmental potentials and that the recovery of oocytes of these grades is maximised by use of an 18‐gauge needle and 50mmHg aspiration pressure.

2. MATERIALS AND METHODS

Ovaries were collected from unsynchronised animals on an abattoir line by trained staff from JSR Genetics Ltd. and were stored and transported to the laboratory in phosphate buffered saline (PBS) at 38.5°C within 6 hr of collection. PBS, and all other chemicals, were acquired from Sigma‐Aldrich (Gillingham, UK) except when stated otherwise. Upon arrival, ovaries were decanted into an autoclaved 500 ml beaker, and maintained at 38.5°C until use. For all experiments, follicles ranging between three and eight mm in diameter were aspirated (see below for details) to recover oocytes. The number of ovaries received in each batch varied depending on availability (between 30 and 50 in total).

Following aspiration, the collected follicular fluid was transferred into a pre‐warmed (38.5˚C) petri dish and all COCs were recovered using a stereomicroscope and an EZ‐grip pipette (Origio) equipped with a 290 µm EZ‐tip (Origio). Collected COCs were washed twice in tyrode lactate buffered 4‐(2‐hydroxyethyl)‐1‐piperazineethanesulfonic acid (TL‐HEPES) polyvinyl‐alcohol (PVA) medium and categorised as A, B, C or D (Figure 1) according to the number and morphology of their CC layers and ooplasm quality (granular patterns, colour and density): Grade A, three or more cumulus layers including granulosa cell‐cumulus oocyte complexes, even cytoplasm; Grade B, intact corona radiata surrounded by another layer of CCs, even cytoplasm; Grade C, incomplete corona radiata or partially denuded oocyte, uneven cytoplasm; Grade D: denuded oocyte, uneven cytoplasm. COC grading was replicated by two independent operators.

FIGURE 1.

FIGURE 1

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Representative examples of oocyte morphological categorisation. (a) grade A cumulus oocyte complex; several layers of CCs and even cytoplasm, (b) grade B cumulus oocyte complex; intact corona radiata surrounded by another layer of CCs and even cytoplasm, (c) grade C cumulus oocyte complex; incomplete corona radiata or partially denuded oocyte and uneven cytoplasm, and (d) grade D, denuded oocyte with an uneven cytoplasm. CC = cumulus cells; CR = corona radiata

2.1. Oocyte maturation

To assess how maturation varies in COCs that that differ in the number and morphology of their CC layers, oocytes were manually aspirated using a 5ml syringe and a 19‐gauge needle with the aid of a small volume of TL‐HEPES‐PVA medium. Comparisons of nuclear maturation potential were repeated using oocytes obtained from five separate deliveries, with assays of cytoplasmic maturation potential repeated on oocytes from six separate deliveries.

IVM of graded oocytes was undertaken in North Carolina State University (NCSU)‐23 media (Petters & Wells, 1993) supplemented with 10% porcine follicular fluid (pFF) that had been collected from a separate batch of abattoir‐derived ovaries. pFF was prepared by centrifugation for 30 min at 3,000rpm, and subsequent filter sterilisation of the supernatant using a Minisart single use filter (0.2 mm). pFF was stored at −20°C until use. Graded oocytes were matured separately in groups of 50 in 500 µl media under 6% CO2 at 37°C. For the first 22 hr, the culture medium was supplemented with hormones (1:100 PG600, Intervet, Milton Keys, UK), 0.8mM L‐cysteine, 10ng/ml EGF, 1mM db‐cAMP and 50 µM β‐mercaptoethanol. Oocytes were then transferred to fresh medium for a further 22 hr with the hormones and db‐cAMP excluded. The IVM oocytes produced were then tested as described below.

To assess the nuclear maturation potential of oocytes of different grades, the meiotic spindles of IVM oocytes were immunostained with anti α‐tubulin. IVM oocytes were denuded of CCs by incubation for 10 min in 0.2%w/v Hyaluronidase Type VIII from bovine testis and passing them several times through a 125‐µm tip. These oocytes were then fixed overnight at 4°C in 4% PFA in PHEM buffer (60 mM PIPES, 25 mM HEPES, 10 mM EGTA, 4 mM MgSO4, pH 7.0). Fixed oocytes were then rinsed three times in PBS for five minutes and permeabilised for ten minutes in 1% Triton X‐100 in PHEM buffer. After rinsing in PBS, the oocytes were blocked in 20% fetal bovine serum (FBS) PHEM buffer for one hour at room temperature. Oocytes were then stained for 30 min in 1:200 anti‐α tubulin‐Alexa 488 (ab195887, Abcam, UK), 5% FBS in PHEM buffer. After rinsing in PBS, oocytes were counterstained in 1 µg/ml Hoechst 33,342 in PBS, mounted on slides with an antifade agent (Fluoroshield). Meiotic stage was classified as previously published (Ma et al., 2003), with oocytes that had reached at least anaphase I considered to have achieved full nuclear maturation.

Two approaches were taken to assess the cytoplasmic maturation potential of oocytes of different grades. First, CGs in post IVM oocytes (N = 107, 64, 93 and 68 for Grades A, B, C and D, respectively) were stained with peanut agglutinin (PNA) lectin‐Alexa 488. Second, the levels of intrinsic GSH were measured in four replicates of 120 oocytes for each oocyte grade to investigate the capability of the oocytes to cope with oxidative stress.

PNA staining, using a modification of the method of Zhang and colleagues (Zhang et al., 2010) was undertaken on IVM oocytes that had been denuded and fixed as described above. Oocytes were rinsed three times in 0.3% bovine serum albumin (BSA) in PBS for five minutes, followed by permeabilisation for 5 min in 0.1% Triton X‐100 in PBS. Permeabilised oocytes were rinsed twice in PBS and stained in 100 µg/ml PNA lectin‐Alexa 488 (L21409, Life Technologies, Paisley, UK) in PBS. After three washes of 5 min in 0.3% BSA, 0.01% Triton X‐100 in PBS, oocytes were counterstained with 1 µg/ml Hoechst 33,342, mounted with Fluoroshield and observed. Oocytes were defined as cytoplasmically mature if they showed a clear, continuous ring of CGs close to their membrane rather than a homogeneously dispersed pattern throughout the cytoplasm.

GSH content measurement was undertaken on IVM oocytes that had been denuded as described above, then washed three times in PBS to eliminate any possible thiol carryover from the culture media (mainly from β‐mercaptoethanol and L‐Cysteine). Measurements were completed as described previously (Funahashi et al., 1994) in a final volume of 1 ml and using 0.25U GSH reductase from baker's yeast (G3664). A Biomate 3S spectrophotometer (ThermoScientific, Waltham, MA) was set for continuous reading at 412 nm and measurements were taken every 20 s for two minutes. The oocyte GSH content was then estimated from a linear calibration curve employing 1, 0.1 and 0.01 nmol reduced GSH per reaction.

All fluorescence observations were completed at x200 total magnification using an Olympus BX60 fluorescence microscope equipped with standard DAPI and Fluorescein isothiocyanate (FITC) bandpass filters. Images were captured using a Hamamatsu ORCA‐03G camera and processed through the software SmartCapture (version 3; Digital Scientific, Cambridge, UK), but CG stained oocyte images are not shown as the static images obtained were not as clear as observations seen by eye.

Rates of nuclear maturation and of cytoplasmic maturation as assessed by PNA staining were analysed by chi‐squared test, with differences between treatments determined by false discovery rate (FDR) corrected pairwise tests (Benjamini & Hochberg, 1995). The GSH content of matured oocytes of different grades were compared by Kruskal–Wallis, with pairwise Mann–Whitney U tests used for post hoc comparisons. Data analysis was performed in R version 3.3.4 (R Core Team, 2018).

2.2. Oocyte recovery

For analysis of how specific combinations of aspiration pressure and needle gauge affect COC recovery and morphology, oocytes were collected using an aspirator pump (Labotect Aspirator 3). The rubber tubing attaching the needle to the aspirant collection tube was primed with TL‐HEPES‐PVA (Funahashi et al., 1997). A needle of appropriate diameter (18, 19, 20, 21 and 23‐gauge) (BD Microlance 3) was attached to the aspirator pump, set to an appropriate pressure setting (25, 50, 75, 100, 125 or 150 mmHg), connected to a 50 ml aspirant collection tube sealed with a rubber bung. TL‐HEPES‐PVA media was flushed through the rubber tube prior to oocyte collection and subsequently periodically to ensure all COCs were recovered into the aspirant collection tube. All 25 combinations of needle and pressure could not be tested on each batch of ovaries as this would either limit sample size per treatment or make the collection period too long. Therefore, for each round of oocyte collection, between three and six different combinations of needle and pressure were tested, with follicles harvested from ten random ovaries from the shipment for each combination. The effects of needle gauge and aspiration pressure on the recovery of (a) oocytes, (b) grade A COCs and (c) grade A and B COCs, from the follicles aspirated was assessed by using a chi‐squared test using Bonferroni corrected p values for different aspiration pressures within each needle gauge. For significant tests, differences between aspiration pressures were then determined by false discovery rate (FDR) corrected pairwise tests (Benjamini & Hochberg, 1995). Differences between the standardised Pearson residuals were used to determine pressures associated with increased recovery rates. In total, 38,595 follicles were aspirated using a range of pressures and needle gauges, yielding 26,370 oocytes.

2.3. Ethical approval process

Oocytes were obtained from abattoir derived ovaries. No specific ethical approval was required.

3. RESULTS

3.1. Oocyte maturation

To assess how maturation varies in COCs that that differ in the number and morphology of their cumulus cell layers, quality and both nuclear and cytoplasmic maturation were assessed in COCs recovered by manual aspiration. These COCs were categorised as grade A, B, C or D according to the number and morphology of their CC layers: Grade A, three or more cumulus layers including granulosa cell‐cumulus oocyte complexes; Grade B, intact corona radiata surrounded by another layer of CCs; Grade C, incomplete corona radiata or partially denuded oocyte; Grade D: denuded oocyte (see Figure 1 for examples of these categories).

Nuclear maturation in IVM oocytes was evaluated by visualising the metaphasic spindles with α‐tubulin immunostaining (Figure 2). This showed that the standard diagnostic features of meiosis could be identified and that the oocytes were maturing. Analysis of these data indicates that IVM oocytes of different grades differ in their rates of nuclear maturation (χ2 = 10.05, df = 3, p = .018, and χ2 = 35.19, df = 3, p < .0001 for the percentage assessed as metaphase I and metaphase II, respectively), with higher grade oocytes displaying higher levels of nuclear maturation (Figure 3i and b). These analyses indicate that the maturation potential of Grade D oocytes is very limited, with only a small percentage (mean of 2.9%) reaching metaphase I (Figure 3i) and no oocytes of this grade assessed as having achieved full nuclear maturation (Figure 3ii). These analyses also indicate that, in terms of nuclear maturation, there is no significant difference between the development of Grade A and B oocytes (Figure 3i and 3ii).

FIGURE 2.

FIGURE 2

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Meiotic stage of pig oocytes visualised by α‐tubulin immunostaining. Shown are examples of immunostained oocytes, with tubulin shown in green (Alexa488) and DNA shown in blue (Hoechst 33,342). (a) Immature oocyte at the germinal vesicle stage, no spindle is detectable. (b) Prometaphase I, a spindle is forming. (c) Metaphase I, a clear tubulin spindle has assembled around the chromosomal compartment. (d) Anaphase I, two sets of chromosomes are separated by a tubulin bridge. (e) Metaphase II, two independent spindles can be detected belonging to either the oocyte or its polar body. Images captured by fluorescence microscopy at x200 total magnification

FIGURE 3.

FIGURE 3

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Oocyte grade affects potential for nuclear maturation. Comparison of maturation rates in IVM oocytes of different grades. Shown are the percentage of oocytes achieving a particular stage, with the error bars showing the 95% confidence intervals. This indicates that the percentage of oocytes developing to both i) metaphase I and to ii) metaphase II differ. Numbers associated with each oocyte grade relate to post hoc tests, with oocyte grades that do not share numbers being significantly different (FDR corrected pairwise tests, p < .05)

Cytoplasmic maturation of IVM oocytes as assessed by PNA lectin‐Alexa 488 staining of CGs indicated that maturation rates differed between oocyte grades (χ2 = 26.64, df = 3, p < .0001) (Figure 4i). Here, oocytes in complex with two or more complete layers of CCs (Grades A and B) displayed a continuous peripheral ring of CGs more often than partially or fully denuded oocytes (Grades C and D) (Figure 4i). Levels of GSH also differed between oocytes classes (Kruskal–Wallis H = 11.89, df = 3, p = .01), with higher levels of GSH seen in Grade A and Grade B oocytes (6.3 ± 0.5 pmol/oocyte and 5.8 ± 1.9 pmol/oocyte, respectively) than in Grade C and Grade D oocytes (2.2 ± 0.3 pmol/oocyte and 0.9 ± 0.3 pmol/oocyte, respectively) (Figure 4ii).

FIGURE 4.

FIGURE 4

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Oocyte grade affects potential for cytoplasmic maturation. i) The percentage, with the error bars showing the 95% confidence intervals, of oocytes assessed as mature by CG staining (N = 107, 64, 93 and 68 for Grades A, B, C and D, respectively). Numbers associated with each oocyte grade relate to post hoc tests, with oocyte grades that do not share numbers being significantly different (FDR corrected pairwise tests, p < .05. ii) Box plots showing the glutathione content per oocyte, with the central bars representing the median value, the lower and upper hinges corresponding to the first and third quartiles and the whiskers extending from the hinge to the largest value no further than 1.5 of the interquartile range from the hinge, numbers associated with each oocyte grade relate to post hoc tests, with oocyte grades that do not share numbers being significantly different (FDR corrected pairwise tests, p < .05)

3.2. Oocyte recovery

The aspiration of 38,595 follicles, using a range of pressures and needle gauges yielded a total of 26,370 oocytes, with the recovered oocytes unequally distributed across the quality grades (Grade A: 8.1%; Grade B: 24.1%; Grade C: 35.7%; Grade D: 32.1%) (Figure 5). These data were analysed to investigate three questions (Table 1). Specifically, which combinations of aspiration pressure and needle gauge recovered the maximum percentage of 1) oocytes, 2) grade A and B COCs, and 3) grade A COCs, from the follicles aspirated (Figure 5, Table 1). These analyses indicated that, for nearly all needle gauges, the aspiration pressure affects recovery and quality of oocyte (Table 1). The exception to this is for the 23‐gauge needle, where no difference between aspiration pressures in the percentages of grade A COCs recovered was identified (Table 1). However, oocyte recovery is maximised at higher aspiration pressures (Table 1) and the worst recovery is seen when using 23‐gauge needles (Table 1, Figure 5). We also observed that the recovery of high‐quality oocytes (either grade A only, or grades A and B combined) is the highest when low aspiration pressures are used (Table 1). For example, if grade A and B COCs are required, these data suggest that recovery is maximised using an 18‐gauge needle and an aspiration pressure of 50mmHg.

FIGURE 5.

FIGURE 5

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The effect of needle gauge and aspiration pressure on the quality of recovered oocytes. Shown is the percentage of aspirated follicles that yielded oocytes of different grades for the various combinations of needle gauges and aspiration pressures. In each column, the top of the D grade oocyte bar presents the total percentage of aspirated follicles from which an oocyte was recovered

TABLE 1.

The effect of needle gauge and aspiration pressure on the recovery rates and quality of recovered oocytes. Shown for comparisons of 1) oocyte recovery, 2) recovery of A and B grade oocytes, and 3) recovery of A grade oocytes only, are the χ2 values and Bonferroni corrected p values for tests of different aspiration pressures within each needle gauge, and results of post hoc comparisons, where aspiration pressures that differ significantly do not share a number. The aspiration pressures associated with recovery of increased numbers of oocytes, or of higher proportions of high‐quality oocytes (grades A and B in 2 and grade A only in 1), are also listed (as determined by inspection of the standardised Pearson residuals). Where multiple pressures are listed, the order indicates the strength of the association with the first pressure listed having the greatest effect

Comparison Needle χ2 Aspiration pressure Pressure(s) with
Gauge (p value) 25 50 75 100 125 150 highest recovery
1) Oocytes 18

603.5

(p < .001)

 1 2 3 2 4 5 125, 150
19

472.13

(p < .001)

 1 1 1 1 2 3 125, 150, 50, 100
20

323.1

(p < .001)

 1 2 3 3 4 5 150, 100, 75
21

177.3

(p < .001)

 1 2 3 2 4 1 100, 50, 75
23

282.3

(p < .001)

 1 2 2 2,4 3 4 150, 50, 100, 75
2) A and B grade 18

351.3

(p < .001)

 1 2 3 1 4 1 50, 75
19

141.3

(p < .001)

 1 1 2 3 3 2 50, 25
20

351.3

(p < .001)

 1 2 3 1 4 1 50, 75
21

98.4

(p < .001)

 1 1,3 2 2,3 2 4 25, 50, 100
23

100.0

(p < .001)

 1 2 2 2 3 1 75, 50, 100
3) A grade 18

195.1

(p < .001)

 1 2 2 1 2 1 50, 75
19

127.9

(p < .001)

 1 2 2 3 3 2 25, 50
20

53.0

(p < .001)

 1,2 2 1,3 3,4 4 4 50, 25, 75
21

97.4

(p < .001)

 1 2 2 2 3 3 25, 50, 75
23

25.5

(p = .03)

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4. DISCUSSION

The global population is predicted to rise to 9.8 billion by the year 2050 (Bruinsma, 2002). This growth, in combination with changes in dietary preferences, is increasing the global demand for animal protein, with the livestock sector challenged to find new ways of accommodating this need. The global shipment of genetically advanced breeding stock to upgrade or replace local genetic lines involve high production, environmental and logistical costs. In some circumstances movement of live animals is not permitted because of the perceived risk of the introduction of disease into the territory, for example, there are regulations around boar movement and pig trade because African swine fever virus infection risk (Taylor et al., 2020). Taken together, these factors have resulted in the in vitro production (IVP) of pig embryos being of increased interest to producers, given that great financial and environmental benefits could be made. Pig IVP has the potential to become a viable alternative to artificial insemination for agricultural and biomedical purposes, however for this to be achieved oocyte quality is paramount, and systematic comparisons between oocytes matured in vitro and in vivo are still needed. The acquisition of adequate numbers of competent oocytes is the first, critical step to successful subsequent in vitro embryo production; it is imperative that the maximum number of the highest quality oocytes can be recovered. Here we have investigated the both the number and quality of oocytes that can retrieved from abattoir derived ovaries, and have determined that CC investment can act as a predictor of oocyte developmental competence. Further to this, we have also shown that both needle size and applied pressure alters the retrieval rate of developmentally competent COCs.

Our results indicate that nuclear maturation and cytoplasmic maturation is variable across the four grades of oocytes (Figure 1). Grade A and B oocytes were observed to mature at high rates whilst fewer Grade C and D oocytes were assessed as having completed maturation (Figure 4). Interestingly, fully denuded oocytes (Grade D), which were never observed to complete nuclear maturation, were occasionally observed to have progressed to full cytoplasmic maturation, therefore indicating that these two processes are not necessarily linked. Alternatively, as the developmental capabilities of denuded oocytes can be rescued by co‐culture with COCs (Luciano et al., 2005), the limited development seen here may have been the result of the presence of a limited number of CCs in the wells in which Grade D oocytes were cultured.

The IVM system in this study included the use of both cysteine and β‐mercaptoethanol. It has been reported that the mean GSH content in pig oocytes increased from 7.9 ± 0.6 pmol/oocyte to 10.4 ± 2.8 pmol/oocyte (means ± standard errors) in the presence of 50 µM β‐mercaptoethanol (Abeydeera et al., 1998), while another found that supplementing the maturation medium with 0.57 mM cysteine increased oocyte GSH from 4.0 ± 0.8 pmol/oocyte to 15 ± 0.3 pmol/oocyte (means ± standard errors) (Yoshida et al., 1993). Our findings indicated that Grade A and B oocytes have levels of GSH comparable to these levels, while Grade C and D oocytes appear largely depleted of GSH (Figure 4). This shows the presence of cysteine and β‐mercaptoethanol alone during IVM is not sufficient to increase intracellular GSH levels in pig oocytes as CCs are required to complete the process (Tatemoto et al., 2000). Moreover the absence of enough GSH in Grade C and D oocytes could well explain their reduced cytoplasmic and nuclear maturation potentials.

In our investigation of the recovery rates of oocytes, we found that A and B oocytes formed only a minority of the total yield (8.1% and 24.1% of A and B, respectively, Figure 5), which is in line with previous findings (Lin et al., 2016). Our comparisons of oocytes recovered using different aspiration pressures and needle gauges (Figure 5, Table 1) indicate that both factors affect oocyte recovery; this supports previous work that identified a negative correlation between the size of the COCs retrieved and the aspiration pressure, with higher pressures increasing the incidence of denuded oocytes (Brüssow et al., 1997; Marques et al., 2015). Furthermore, these data suggest the existence of a compromise between recovery rates and oocyte quality, with quality maximised by lower pressures. Given that CCs support both nuclear and cytoplasmic oocyte maturation (Tanghe et al., 2002), it is desirable to minimise disruption or damage to the COC when retrieving oocytes from follicles.

5. CONCLUSIONS

Taken together, our previous observations on the maturational competence of oocytes concur with the well‐established link between COC investment and oocyte developmental competence (Alvarez et al., 2009; Bagg et al., 2007; Kim et al., 2010; Lin et al., 2016; Marchal et al., 2002; Nagano et al., 2006), and suggest that previous studies may have been too stringent in COC selection for subsequent IVP. Here, optimum results were achieved using an 18‐gauge needle and 50mmHg aspiration pressure (Figure 5). Given that Grade A and B COCs are equivalent in their developmental competence, it is possible to maximise both the number, and the quality of oocytes retrieved with this combination. If however other considerations in future investigations indicate that only Grade A oocytes are required for IVP, then there are a broader range of needle and pressure combinations that can be considered (Figure 5). Going forward, it would be interesting to determine whether the developmental potential of Grades C and D pig oocytes could be rescued using meiotic inhibition and an extended IVM protocol, in concordance with other work in both pigs and cattle (Li et al., 2016; Park et al., 2016; Sugimura et al., 2018). This would, in principle, afford these oocytes more time to grow and develop fully. It is important that future work aims to support these findings with subsequent in vitro embryo development data.

CONFLICTS OF INTEREST

The authors declare no competing interests.

AUTHOR CONTRIBUTION

Giuseppe Silvestri: Investigation; Writing‐original draft; Writing‐review & editing. Claudia C Rathje: Investigation; Writing‐original draft; Writing‐review & editing. Simon Harvey: Formal analysis; Writing‐original draft; Writing‐review & editing. Rebecca Louise Gould: Investigation; Writing‐review & editing. Grant A Walling: Conceptualization; Writing‐review & editing. Peter James Ivor Ellis: Conceptualization; Methodology; Writing‐review & editing. Katie Evelyn Harvey: Conceptualization; Investigation; Methodology; Supervision; Writing‐original draft; Writing‐review & editing. Darren Griffin: Conceptualization; Funding acquisition; Supervision; Writing‐review & editing.

PEER REVIEW

The peer review history for this article is available at https://publons.com/publon/10.1002/vms3.378.

ACKNOWLEDGEMENTS

This work was supported by the Technology Strategy Board (now called Innovate UK). The authors thank JSR Genetics for supplying the pig ovaries used in this project.

Silvestri G, Rathje CC, Harvey SC, et al. Identification of optimal assisted aspiration conditions of oocytes for use in porcine in vitro maturation: A re‐evaluation of the relationship between the cumulus oocyte complex and oocyte quality. Vet Med Sci.2021;7:465–473. 10.1002/vms3.378

Giuseppe Silvestri and Claudia Rathje are joint first authors

Katie Evelyn Harvey and Darren Karl Griffin are joint last authors

DATA AVAILABILITY STATEMENT

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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