Theriogenology 82 (2014) 1055–1067

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Seasonal variations in developmental competence and relative abundance of gene transcripts in buffalo (Bubalus bubalis) oocytes Ahmed S. Abdoon a, b, *, Christoph Gabler b, Christoph Holder b, Omaima M. Kandil a, Ralf Einspanier b a

Department of Animal Reproduction and Artificial Insemination, Veterinary Research Division, National Research Center, Cairo, Egypt b Institute of Veterinary Biochemistry, Freie Universität Berlin, Berlin, Germany

a r t i c l e i n f o

a b s t r a c t

Article history: Received 21 October 2013 Received in revised form 29 June 2014 Accepted 3 July 2014

Hot season is a major constraint to production and reproduction in buffaloes. The present work aimed to investigate the effect of season on ovarian function, developmental competence, and the relative abundance of gene expression in buffalo oocytes. Three experiments were conducted. In experiment 1, pairs of buffalo ovaries were collected during cold season (CS, autumn and winter) and hot season (HS, spring and summer), and the number of antral follicles was recorded. Cumulus oocyte complexes (COCs) were aspirated and evaluated according to their morphology into four Grades. In experiment 2, Grade A and B COCs collected during CS and HS were in vitro matured (IVM) for 24 hours under standard conditions at 38.5
C in a humidified air of 5% CO2. After IVM, cumulus cells were removed and oocytes were fixed, stained with 1% aceto-orcein, and evaluated for nuclear configuration. In vitro matured buffalo oocytes harvested during CS or HS were in vitro fertilized (IVF) using frozen-thawed buffalo semen and cultured in vitro to the blastocyst stage. In experiment 3, buffalo COCs and in vitro matured oocytes were collected during CS and HS, and then snap frozen in liquid nitrogen for gene expression analysis. Total RNA was extracted from COCs and in vitro matured oocytes, and complementary DNA was synthesized; quantitative Reverse Transcription-Polymerase Chain Reaction was performed for eight candidate genes including GAPDH, ACTB, B2M, GDF9, BMP15, HSP70, and SOD2. The results indicated that HS significantly (P < 0.01) decreased the number of antral follicles and the number of COCs recovered per ovary. The number of Grade A, B, and C COCs was lower (P < 0.05) during HS than CS. In vitro maturation of buffalo oocytes during HS significantly (P < 0.01) reduced the number of oocytes reaching the metaphase II stage and increased the percentage of degenerated oocytes compared with CS. Oocytes collected during HS also showed signs of cytoplasmic degeneration. After IVF, cleavage rate was lower (P < 0.01) for oocytes collected during HS, and the percentage of oocytes arrested at the two-cell stage was higher (P < 0.01) than oocytes IVF during CS. Oocytes matured during CS showed a higher (P < 0.01) blastocyst rate than those matured during HS. Also, COCs recovered in HS showed significant (P < 0.05) upregulation of HSP70 mRNA expression compared with those recovered in CS. For in vitro matured oocytes, CS down regulated the transcript abundance of ACTB and upregulated GAPDH and HSP70 mRNA levels compared with HS condition. In conclusion, HS could impair buffalo fertility by reducing the number of antral follicles and oocyte quality. In vitro maturation of buffalo oocytes during HS impairs their nuclear and cytoplasmic maturation, fertilization, and

Keywords: Buffalo Season Oocyte morphology Developmental competence Gene expression

* Corresponding author. Tel.: þ20 1221941292; fax: þ20 233370931. E-mail address: [email protected] (A.S. Abdoon). 0093-691X/$ – see front matter Ó 2014 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.theriogenology.2014.07.008

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subsequent embryo development to the morula and blastocyst stages. This could be in part because of the altered gene expression found in COCs and in vitro matured oocytes. Ó 2014 Elsevier Inc. All rights reserved.

1. Introduction High environmental temperature during summer is a major constraint for the production and fertility of agricultural animals worldwide. In USA, fertility in Holstein cows decreased from 52% to 32% as maximum air temperature increased from 23.9
C to 32.2
C during the summer [1]. In Brazil, pregnancy rates of Holstein cows housed in free stalls were reduced from 71.2% in winter to 45.7% in summer [2]. Moreover, buffaloes are poor thermoregulatory animals because of their morphologic and anatomic peculiarities. They are sensitive to heat stress, which affects their biological functions including reproduction [3]. Inactive ovaries represent 93% of summer infertility in buffaloes raised in Middle Egypt [4]. The decreased fertility associated with summer is a multifactorial problem in which hyperthermia affects cellular function in various tissues of the female reproductive tract [5]. In cattle, high ambient temperature affects the duration and intensity of the expression of estrus and increases the duration of anestrus and silent ovulation [6]. Hot summer compromises bovine follicular growth [7,8], hormonal secretion [9], uterine blood flow [10], and endometrial function [11]. Reduced fertility during summer was associated with irregular luteal forms, which may have a negative effect on bovine ovarian function and oocyte quality [12]. Bovine oocytes are more susceptible to summer heat stress during the preovulatory period [13]. Bovine oocytes recovered during summer season exhibited lower quality [14,15], and reduction in their developmental potential [14,16] and preimplantation embryonic development [17]. Blastocyst rate was compromised, and the fragmentation rate of repeat breeder Holstein cows’ blastocysts was enhanced during summer compared with winter. The association of repeat breeder fertility problems and HS may potentially impair oocyte quality [18]. In buffaloes, no differences were found in terms of oocyte recovery per ovary among seasons; however, the percentage of small oocytes was higher during spring and summer (0.9  0.1 and 0.9  0.2) compared with autumn and winter (0.3  0.1 and 0.2  0.1). Both cleavage and embryo rate increased during the period from October to December (71.7  3.1 and 26.5  2.1, respectively) compared with the period from April to June (58.0  2.4 and 18.8  1.6, respectively) [19]. The mechanisms underlying the disruption of oocyte maturational processes by season are likely to be complex and probably involve reduced intracellular protein synthesis [20], increased glutathione content [21], altered cortical granule types [22], disrupted cytoskeleton [23,24], microfilament, microtubule architecture, disorganized oocyte meiotic spindle [25], and induction of oocyte death through apoptosis [26], and consequently, the proportion of oocytes that completes nuclear maturation is reduced [27].

A better understanding of the molecular mechanisms underlying how summer could impair fertility may lead to the development of additional approaches to alleviate these effects. Exposing the ovarian pool of oocytes to environmental stress appears to impair maternal mRNA storage and/or the mechanism of transcription renewal, which in turn affects gene expression before and after embryonic genome activation [15]. The transcript levels of c-morpholino oligonucleotides (c-MOs), growth differentiation factor 9 (GDF9), the pluripotency markers (POU domain class 5 transcription factor 1 [POU5F1; formerly named OCT4]), and glyceraldehyde 3phosphate dehydrogenase (GAPDH) were lower in metaphase II (MII) in bovine oocytes collected during summer than winter [28]. Furthermore, heat stress during early embryonic development decreased caudal type homeobox 2 (CDX2), placenta-specific 8 (PLAC8), and prostaglandin-endoperoxide synthase 2 (PTGS2; formerly named cyclooxygenase 2) mRNA expressions in bovine embryos produced in vitro [29]. Also, heat-induced alterations in transcriptional levels of genes involved in cell growth, cell cycle, and programmed cell death have been documented in human oocytes [30]. Up to now, there is no literature available investigating the mechanisms by which season can cause such catastrophic effects on buffalo ovarian function. Therefore, it appears necessary to define the cellular, physiological, and molecular mechanisms that identify potential local effects of season on buffalo cumulus oocyte complexes (COCs) and oocytes’ developmental competence. Deeper knowledge is a prerequisite to allow design of more appropriate methods to alleviate such detrimental effects. Therefore, the present study was designed to (1) investigate the effects of season on ovarian follicular population and oocyte yield and morphology; (2) evaluate the ability of COCs recovered during cold season (CS) and hot season (HS) to mature and develop in vitro; and (3) study the relative mRNA expression of selected candidate genes in buffalo COCs and in vitro matured oocytes during CS and HS. 2. Materials and methods 2.1. Chemicals and reagents All reagents and media used in this study were obtained from Sigma–Aldrich (St. Louis, MO, USA) unless otherwise mentioned. 2.2. Experimental design The present work was conducted on buffalo ovaries collected during CS (from November 2010 to March 2011) and HS (from April 2011 to October 2011). The maximal and minimal temperatures and relative humidity are presented in Figure 1.

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Fig. 1. The average daily high (red) and low (blue) temperatures (A) and high (blue) and low (brown) relative humidity (B) (http://weatherspark.com/averages/ 29245/Cairo-Al-Qahirah-Egypt). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article).

2.2.1. Experiment 1: effects of season on ovarian follicular development and oocytes’ yield and morphology in buffalo 2.2.1.1. Determination of the number of antral follicles. During CS and HS, genital tracts of multiparous buffalo cows were obtained once a week at local abattoirs (EL-Moneib and ElWaraq) at Giza, Egypt. Pairs of ovaries were collected and transported to the laboratory within 2 hours in a thermos (at 32
C–35
C) containing physiological saline solution (0.9% wt/vol NaCl) supplemented with 100 IU/mL penicillin G sodium and 100 mg/mL streptomycin. At the laboratory, pairs of ovaries were washed separately in a warm saline solution and the number of antral follicles 2 to 10 mm in diameter was recorded for each animal. 2.2.1.2. Recovery of COCs. After counting the number of antral follicles during CS and HS, COCs were aspirated from follicles 3 to 8 mm in diameter by using an 18-gauge needle attached to a 10-mL sterile disposal syringe. Aspirated follicular fluid was transferred into 15 mL sterile tubes and kept in a water bath at 37
C for 15 minutes. The resulting sediment was transferred to a 9-cm culture dish (Nunclon, Rosalind, Denmark) containing 10-mL PBS (PAA, Pasching, Austria) supplemented with 4 mg/mL BSA and 50 mg/mL gentamicin. Cumulus oocyte complexes were selected under a stereomicroscope (Olympus, Tokyo, Japan) at a magnification of 20. After collection, COCs were washed at least three times in PBS and then classified under a stereomicroscope at a magnification of 50 according to Khurana and Niemann [31], based on their morphologic characters (number of cumulus cell layers and the homogeneity of the cytoplasm) into four Grades: oocytes with homogenous evenly granulated and many tight layers of cumulus cells (Grade A); oocytes with homogenous ooplasm and two to three cumulus cell layers (Grade B); oocytes with heterogeneous ooplasm surrounded by one cumulus cell layer (Grade C); and denuded oocytes (Grade D). 2.2.2. Experiment 2: effects of season on the developmental competence of buffalo oocytes 2.2.2.1. Effect of season on nuclear maturation of buffalo oocytes. Grade A, B, and C COCs were washed at least three

times in PBS and once in in vitro matured (IVM) medium (Medium 199 with Earl’s salt and bicarbonate buffer and supplemented with 10% fetal calf serum, 10 mg/mL FSH, 10 IU/mL equine chorionic gonadotropin (eCG, Folligon; Intervet, Boxmeer, The Netherlands), 10 IU/mL human chorionic gonadotropin (hCG, Pregnyl; El Nile Co., Cairo, Egypt), 20 ng/mL epidermal growth factor (EGF), and 50 mg/ mL gentamycin. During the CS and HS and under the same standard laboratory conditions of 26
C, 15 to 20 COCs were matured in 500 mL of maturation medium in a four-well culture dish (Nunclon, Rosalind, Denmark) for 22 to 24 hours at 38.5
C in a humidified air of 5% CO2. 2.2.2.2. Assessment of nuclear maturation. After IVM of buffalo oocytes during CS and HS, cumulus cells were removed by gentle pipetting. Matured oocytes were fixed for 48 hours in acetic:ethanol (1:3), and then stained with 1% aceto-orcein (Merck, Darmstadt, Germany). The stained oocytes were evaluated for nuclear maturation under a microscope at a magnification of 400 according to the method adopted by Hunter and Polge [32] into (1) immature oocytes, at either the germinal vesicle (GV) stage or germinal vesicle break down stage (GVBD); (2) mature oocytes in the anaphase, telophase, or MII (with first polar body) stage; and (3) degenerated oocytes without any chromatin structures. 2.2.2.3. Effect of season on cleavage rate and development of buffalo embryos in vitro. Buffalo COCs were in vitro matured during CS and HS as mentioned in the previous section. A single ejaculate of frozen semen from a buffalo bull of known fertilization rate (86%) in vitro was used for IVF. Frozen semen straws were thawed at 37
C for 30 seconds and then overlaid on the top of two layers of 1 mL (v:v) of 45% and 90% Percoll density gradient. Semen was centrifuged at 1800 rpm for 20 minutes; the supernatant was removed and the sperm pellet was resuspended in 5 mL of Sperm-Tyrode’s albumin lactate pyruvate (Sp-TALP) medium supplemented with 4 mg/mL BSA and 50 mg/mL gentamycin, and then centrifuged at 1800 rpm for 5 minutes. After two sperm wash steps, the sperm pellet was

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in vitro matured oocytes using the PicoPure RNA Isolation kit (MDS Analytical Technologies, Ismaning, Bayern, Germany) according to the manufacturer’s instructions. To generate single-stranded complementary DNA (cDNA), 100 ng total RNA was processed using 3.75 mM random hexamers (Amersham Biosciences, Freiburg, Germany) and 200 U M-MuLV Reverse Transcriptase (Fermentas, St. LeonRot, Germany) in a 60-mL reaction mixture as described earlier in detail [33]. The generated cDNA served as a template for quantitative real-time polymerase chain reaction and was stored at 20
C until use.

suspended in 0.5 mL of fresh Fert-TALP medium supplemented with 6 mg/mL BSA-FAF (fatty acid-free) and 50 mg/ mL gentamycin. Sperm concentration was adjusted to 1 to 2  106 spermatozoa/mL. Then, 600 mL of motile sperm suspension supplemented with 10 mL PHE (0.5 mM penicillamine, 0.25 mM hypotaurine, and 25 mM epinephrine in 0.9% [wt/vol] NaCl) was placed in a four-well culture plate (Nunc) and covered with 200 mL of mineral oil. In vitro matured oocytes during CS and HS were washed in FertTALP media at least three times, then transferred into the sperm suspension (25–30 oocytes per well) and incubated at 38.5
C in a humidified air of 5% CO2 for 18 to 20 hours. The presumptive zygotes were thoroughly washed in modified synthetic oviduct fluid (mSOFaa) supplemented with 5 mg/ mL BSA, 5 ng/mL insulin, and 50 mg/mL gentamycin. Ten to 15 zygotes were transferred into 50 mL droplets of mSOFaa medium covered with mineral oil and incubated at 38.5
C in a humidified air of 5% CO2. Cleavage rate and embryo development to the blastocyst stage were evaluated on Days 2, 5, and 7, and the medium was replaced every 48 hours.

2.2.3.3. Quantitative real-time polymerase chain reaction. Quantitative real-time polymerase chain reaction was performed on a Rotor-Gene 3000 (Corbett Research, Mortlake, Australia) using SYBR Green, following The Minimum Information for Publication of Quantitative Real-Time PCR Experiments guidelines [34] as described previously [33]. Briefly, a 10-mL reaction mixture was prepared for each sample, containing 1 mL of cDNA, 0.4 mM of each primer, and 1 SensiMixLow-ROX (Bioline, Luckenwalde, Germany). The specific primer pairs synthesized by Eurofins MWG Operon (Eurofins, Ebersberg, Germany) for the following genes of interest are listed in Table 1: GAPDH, beta-actin (ACTB), beta-2-microglobulin (B2M), GDF9, bone morphogenetic protein 15 (BMP15), PTGS2, superoxide dismutase 2 (SOD2), and heat shock protein 70 (HSP70). Table 1 also lists the primer pairs for potential housekeeping genes 18S rRNA, ribosomal protein L19 (RPL19), histone deacetylase 1 (HDAC1), peptidylprolyl isomerase A (PPIA), and ubiquitously expressed transcript (UXT). In preliminary experiments, designed primer pairs were evaluated to confirm the expected product size and the optimal annealing temperature, as described in detail in ref. [33]. In addition,

2.2.3. Experiment 3: effect of season on gene expression in buffalo COCs and in vitro matured oocytes 2.2.3.1. Sample collection. In experiments 1 and 2, 12 pools of buffalo COCs and in vitro matured oocytes were collected during CS and HS (n ¼ 15–20 COCs or in vitro matured oocytes per pool), then washed twice in PBS þ 0.1% polyvinyl alcohol and snap frozen in cryotubes in liquid nitrogen. After that, frozen COCs and in vitro matured oocytes were stored at 80
C until used for RNA isolation. 2.2.3.2. Ribonucleic acid isolation and complementary DNA synthesis. Total RNA was isolated from buffalo COCs and

Table 1 Primer sequences, annealing temperature (TM), and amplicon size of each gene target analyzed in buffalo cumulus oocyte complexes and in vitro matured oocytes by quantitative real-time PCR. Name

EMBL accession no.

Bovine BMP15

NM_001031752.1

Bovine SOD2

NM_201527.2

Bovine GDF9

GQ922451.1

Bovine HSP70

JN604432.1

Bovine 18S rRNA

AF176811.1

Bovine GAPDH

U85042

Bovine ACTB

AY141970

Bovine PTGS2

AF031698.1

Bovine B2M

NM_173893.2

Bovine RPL19

NM_001040516

Bovine HDAC1

NM_001037444

Bovine PPIA

NM_178320

Bovine UXT

NM_001037471

Sequence F R F R F R F R F R F R F R F R F R F R F R F R F R

0

0

5 -GGCACTTCATCATTGGACAC-3 50 -TTGAAAAGGGTGGAGGGAAC-3 50 -ACGTGAACAACCTCAACGTC-30 50 -AGTCACGTTTGATGGCTTCC-30 50 -TGTAAGATTGTCCCGTCACC-30 50 -CGAGGGTTGTATTTGTGTGG-30 50 -TTCGTGGAGGAGTTCAAGAG-30 50 -TGAAGATCTGCGTCTGCTTC-30 50 -GAGAAACGGCTACCACATCCAA-30 50 -GACACTCAGCTAAGAGCATCGA-30 50 -CCCAGAAGACTGTGGATGG-30 50 -AGTCGCAGGAGACAACCTG-30 50 -CGGTGCCCATCTATGAGG-30 50 -GATGGTGATGACCTGCCC-30 50 -CTCTTCCTCCTGTGCCTGAT-30 50 -CTGAGTATCTTTGACTGTGGGAG-30 50 -AGTAAGCCGCAGTGGAGGT-30 50 -CGCAAAACACCCTGAAGACT-30 50 -GGCAGGCATATGGGTATAGG-30 50 -CCTTGTCTGCCTTCAGCTTG-30 50 -CCAGTGCAGTTGTCTTGCAG-30 50 -TTAGGGATCTCCGTGTCCAG-30 50 -CTGAGCACTGGAGAGAAAGG-30 50 -TGCCATCCAACCACTCAGTC-30 50 -CGCTACGAGGCTTTCATCTC-30 50 -TGAAGTGTCTGGGACCACTG-30

Size (bp)

TM (
C)

228

66.0

201

61.0

207

61.0

565

66.0

337

61.0

306

62.0

266

58.0

359

60.4

108

60.0

232

60.0

217

60.0

259

60.0

207

61.0

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obtained amplicons of these preliminary approaches were cut from the gel, eluted using the Invisorb Spin DNA Extraction Kit (Stratec, Berlin, Germany), and used for generation of standard curves. The cycling conditions comprised an initial denaturation step at 95
C for 10 minutes, three-step amplification including denaturation at 95
C for 15 seconds, annealing at the temperature indicated in Table 1 for 20 seconds, and extension at 72
C for 30 seconds, a melting curve program (50
C–99
C) to confirm specific amplification, and a final cooling step to 4
C. Expression analysis of all samples (COCs and in vitro matured oocytes) was performed together in the same run for each and every reference gene and gene of interest, respectively. Quantities of the genes of interest were calculated in comparison with a simultaneously amplified standard curve of known quantities of the specific amplicon. Amplification efficiencies of the used primer pairs varied from 85% to 100%. The coefficient of determination (r2) varied from 0.97 to 0.99, revealing the linearity of the standard curve. As negative controls, reactions containing no template (H2O) or nonreverse transcribed total RNA were included to verify that obtained PCR products were not derived from contaminations or genomic DNA. Expression data were then normalized with the geNorm (http://medgen.ugent.be/wjvdesomp/genorm) tool [35], which calculates a normalization factor based on the geometric mean of expression levels of housekeeping genes. The optimal number of the housekeeping genes for normalization was calculated as the GeNormV value. The GeNormV value is the pairwise variation of the resultant normalization factors between n and n þ 1 housekeeping genes in each bar and should be below the critical limit of 0.15. The GeNormV value indicates the effect of including an additional gene in the calculation for normalization factors and is helpful in minimizing the number of required housekeeping genes. However, there was a tendency of significance between the groups. Therefore, the mentioned housekeeping genes were selected for COCs or in vitro matured oocytes to have a broader basis for normalization. The most stable housekeeping genes in these experiments were 18S rRNA, RPL19, HDAC1, PPIA, and UXT for the COCs and 18S rRNA, ACTB, and GAPDH for the in vitro matured oocytes. The obtained amplicons were checked for specificity by sequencing (GATC Biotech, Konstanz, Germany) and showed a 98 to 99% homology to the published bovine sequences.

2.3. Statistical analysis The paired t test and chi-square analysis (X2) were used to compare HS and CS. P values less than 0.05 and less than 0.01 were considered significant. Statistical evaluations for ovarian follicular population and oocyte yield, quality, maturation, and cleavage rate were performed by using Decision Analyst STATS2.0 (www.decisionanalyst.com). For analysis of mRNA expression, the Kruskal–Wallis test for independent samples was used. When this test detected significant differences between groups, it was followed by the Mann–Whitney U test. These statistical analyses were performed with SPSS Statistics for Windows Version 20 (IBM, Ehningen, Germany).

1059

Normalized values were used to generate box and whisker plots for data presentation with SPSS 20. Data in Figures 4 and 5 show median values with 50% of data within the box. Extreme values are shown as asterisks and outliers as circles. An outlier was defined as a value between 1.5- and 3.0-fold of the interquartile range, whereas an extreme value was a value beyond 3.0-fold of the interquartile range. 3. Results 3.1. Experiment 1 The effect of season on number of follicles and oocytes’ yield and morphology in buffalo ovaries is presented in Table 2. The average numbers of ovarian follicles and COCs recovered per ovary were significantly (P < 0.01) lower in HS than CS. Also, HS has affected the quality of COCs recovered per ovary. A significantly (P < 0.01) lower number of grade A, B, and C COCs were recovered during HS than CS (Table 2). In addition, oocytes collected during HS showed cytoplasmic degeneration and fragmentation, and degeneration of the surrounding cumulus cells compared with those recovered during CS. 3.2. Experiment 2 3.2.1. Effect of season on maturation rate of buffalo oocytes Buffalo oocytes matured during CS showed homogenous cytoplasm and healthy expanded cumulus cells (Fig. 2A). In contrast, oocytes matured during HS showed dark cytoplasm and degenerated cumulus cells (Fig. 2B). Metaphase II stage matured oocytes harvested during CS showed evenly granulated cytoplasm and the presence of first polar body (Fig. 2C). Not only nuclear maturation, but buffalo oocytes in vitro matured during HS showed a higher rate of cytoplasmic degeneration, as indicated by the presence of large lipid vacuoles (Fig. 2D). A significantly (P < 0.01) higher percentage of in vitro matured buffalo oocytes collected during HS were arrested at the GV breakdown stage, and the total number of immature oocytes was significantly (P < 0.01) higher in HS compared with CS (Table 3). Moreover, the total number of matured oocytes was significantly higher (P < 0.01) for oocytes recovered in CS than in HS. A Table 2 Effect of season on number of surface ovarian follicles, and oocytes’ yield and quality in buffalo ovaries. Item

Cold season

Hot season

Number of pairs of ovaries Number of antral follicles/ovary (mean  SD) Number of COCs ovary (mean  SD) Quality of COCs recovered (mean  SD) Grade A Grade B Grade C Grade D

316 9.8  1.1a

249 4.1  0.7c

6.2  1.4a

2.7  0.9c

2.8 1.5 1.2 0.7

   

0.4a 0.3a 0.2a 0.3

1.1 0.5 0.1 1.0

   

0.3c 0.2c 0.1c 0.3

Superscripts a and c within the same column differ significantly at P < 0.01.

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Fig. 2. In vitro matured buffalo oocytes during CS and HS. Oocytes matured during CS showed homogenous cytoplasm and healthy expanded cumulus cells (A), whereas oocytes matured during HS showed dark cytoplasm and degenerated cumulus cells (B). Metaphase II matured oocytes during CS, with evenly granulated cytoplasm and the presence of first polar body (C), and degenerated in vitro matured oocyte collected during HS, showing degenerated cytoplasm (D). CS, cold season; HS, hot season.

higher (P < 0.01) percentage of in vitro matured oocytes collected during CS reached the MII stage (Table 3) compared with those collected during HS (Fig. 3A). Also, the percentage of degenerated in vitro matured oocytes detected in HS was higher (P < 0.01) than that detected in CS (Table 3). In contrast, higher percentage of degenerated oocytes matured during HS showed the presence of large lipid droplets occupying the cytoplasm (Fig. 3B).

3.2.2. Effect of season on the developmental competence of in vitro matured buffalo oocytes In vitro maturation of buffalo COCs during HS had a detrimental effect on their developmental competence (Table 4). Cleavage rate was lower (P < 0.01) in oocytes matured in HS compared with those harvested during CS. The percentage of cleaved embryos arrested at the two-and four-cell stage was higher (P < 0.01) in matured/fertilized

Table 3 Effect of hot season on nuclear maturation of in vitro matured buffalo oocytes. Season

Cold season Hot season

No. of oocytes

Immature oocytes (%) GV

GVBD

Total

Mature oocytes (%) AI

TI

MII

Total

107 98

d 2 (2)

9 (8.4)a 33 (33.7)c

9 (8.4)a 35 (35.7)c

18 (16.8)a 9 (9.2)c

13 (12.1)c 8 (8.2)a

60 (56.1)a 25 (25.5)c

91 (85.0)a 42 (42.9)c

Degenerated oocytes (%) 7 (6.5)a 21 (21.4)c

Superscripts a and c differ significantly within the same column at P < 0.01. Abbreviations: GV, germinal vesicle; GVBD, germinal vesicle break down; AI, Anaphase I; TI, Telophase I; MII, metaphase II.

Fig. 3. In vitro matured buffalo oocytes collected during cold season and stained with aceto-orcein showed maturation to the metaphase II stage, and first polar body (1st pb) (A), and degenerated oocyte matured during hot season, showing the presence of large lipid droplets occupying the cytoplasm (B).

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Table 4 Effect of hot season on cleavage rate and the development of in vitro produced buffalo embryos. Season

No. of fertilized oocytes

Cleavage rate (%)

Cold season Hot season

286 237

84.3  1.3a 45.6  2.4c

Percentage of embryo developmental  SEM 2–4 Cells

8–16 Cells

Morula

Blastocyst

6.3  0.9c 34.6  1.6a

12.1  1.1a 7.6  1.9b

54.3  1.3a 39.8  1.6c

28.3  1.1a 13.0  2.1c

Superscripts a and b within the same column differ significantly at P < 0.05. Superscripts a and c within the same column differ significantly at P < 0.01.

during HS than in CS. Moreover, oocytes harvested during HS showed a decreased (P < 0.01) ability to develop to the morula and blastocyst stage compared with oocytes harvested during CS.

3.3. Experiment 3 3.3.1. Effect of season on gene expression in buffalo COCs The relative transcript abundance of GAPDH, ACTB, B2M, GDF9, BMP15, HSP70, PTGS2, and SOD2 in buffalo COCs is presented in Figure 4. The results showed high individual variance between the pools. The transcript abundance of HSP70 mRNA was significantly higher in the pools of COCs collected during HS than those collected during CS. The

A

other genes were obviously not affected in their expression pattern by season. 3.3.2. Effect of season on gene expression in in vitro matured buffalo oocytes In vitro maturation of buffalo oocytes harvested during HS showed a significant downregulation (P < 0.05) of the relative transcript abundance of GAPDH (Fig. 5A). The same tendency (P ¼ 0.093) was observed for HSP70 mRNA expression (Fig. 5H). In contrast, HS showed a tendency to upregulate mRNA expression of ACTB (P ¼ 0.115) in in vitro matured buffalo oocytes compared with those matured during CS (Fig. 5B). The expression pattern of the other investigated genes was obviously not affected by season.

B normal. ACTB expression

normal. GAPDH expression

150

100

50

0

120

80

40

0

Hot

Cold

Hot

Season

C

D normal. BMP15 expression

120

normal. B2M expression

Cold

Season

80

40

12

9

6

3

0

0

Hot

Cold

Season

Hot

Cold

Season

Fig. 4. Transcript abundance of GAPDH (A), ACTB (B), B2M (C), BMP15 (D), GDF9 (E), PTGS2 (F), SOD2 (G), and HSP70 (H) mRNA expressions in buffalo cumulus oocyte complexes during hot and cold seasons. Extreme values are shown as asterisks and outliers as circles. normal., normalized.

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E

F 1.5

normal. PTGS2 expression

normal. GDF9 expression

6

4

2

0

Hot

1.0

0.5

0

Cold

Hot

Season

G

H 24

normal. HSP70 expression

normal. SOD2 expression

45

30

15

0

Cold

Season

Hot

16

8

0

Cold

Season

Hot

Cold

Season Fig. 4. (continued).

4. Discussion Buffaloes are more susceptible to heat stress than cattle. Anestrus, silent heat, poor conception rate, high incidence of early embryonic death, and longer intercalving period are major constraints to the reproductive performance and productivity in buffaloes under heat stress conditions. However, the direct cellular effects of hyperthermia on specific buffalo reproductive functions are a challenge to determine in vivo. It is known that hyperthermia can affect cellular function in various tissues of the female reproductive tract [5]. In the present work, the number of antral follicles was lower (P < 0.01) during HS than CS. Similarly, summer season altered follicular population and oocyte quality in buffalo [36], decreased the number of bovine small ovarian follicles [37], or decreased bovine follicular growth and dynamics [38]. In published studies, early antral follicles of approximately 0.5 to 1.0 mm in diameter were shown to be sensitive to heat stress during summer [39]. Furthermore, heat stress during summer could act directly on bovine ovary to decrease its sensitivity to gonadotropin stimulation [40], or disrupt steroidogenesis [41]. On the contrary, no differences were found in terms of oocyte recovery per ovary among seasons, but the percentage of small oocytes was higher during spring and summer than autumn and winter [19]. In contrast, previous studies indicated that in

cattle, heat stress during summer either increased the number of small ovarian follicles [42] or had no effect on this [43]. Discrepancies between these studies could be related to differences in experimental design, time of heat stress exposure, the severity of temperatures during summer, localities, or differences in genetic susceptibility. Furthermore, HS has a compromising effect on buffalo oocytes’ yield and quality. As shown here, the total number of COCs harvested during HS was lower (P < 0.01) than that recovered during CS. Also, the percentage of grade A, B, and C COCs harvested during HS was lower (P < 0.01) than that harvested in CS. At the same time, the cytoplasm of the oocytes recovered during HS showed degenerative changes, as indicated by the presence of dark cytoplasmic batches, and a higher incidence of degenerated cumulus cells. These results are in accordance with previously published studies indicating that during HS buffaloes produced fewer goodquality oocytes than their unstressed counterparts [44]. Concomitantly, the quality of bovine oocytes recovered during summer was compromised compared with the quality of those recovered during wintertime [18]. It is known that heat stress has a detrimental effect from early stages of folliculogenesis, leading to carryover effects on ovulated oocytes of low quality [26,41]. Furthermore, heat stress during HS induced a decrease in oocyte function as a result of a series of cellular alterations affecting nuclear and cytoplasmic compartments of the bovine oocyte [45].

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Fig. 5. Relative abundance of transcripts of GAPDH (A), ACTB (B), B2M (C), BMP15 (D), GDF9 (E), PTGS2 (F), SOD2 (G), and HSP70 (H) in buffalo oocytes matured during hot and cold seasons. Extreme values are shown as asterisks and outliers as circles. normal., normalized.

However, Takuma et al. [38] reported that the quality of COCs recovered from pregnant cows did not differ between HS and CS. This discrepancy could be because of the differences in localities, species, or reproductive status, and is probably associated with the severity of environmental temperature during summer season. Furthermore, other effects such as body condition score, lactation period, age, feeding, and management cannot be excluded. The present study started with the high number of 560 cows to minimize individual differences. Therefore, these mentioned effects can rather be considered minor effects because of such a high number of included animals. It was reported that a higher number of follicles was observed and a higher number of oocytes were recovered in buffaloes during the peak breeding season compared with the low breeding season [46]. Although, we could show that HS has a detrimental effect on the IVM rate of buffalo oocytes. In the comparison of buffalo oocytes in vitro matured during CS and HS, the proportions of oocytes reaching the MII stage were significantly (P < 0.01) lower and the proportions of immature and degenerated oocytes were higher (P < 0.01) during HS than in CS. Not only nuclear maturation but also HS had deleterious effects on the cytoplasm of matured oocytes. The cytoplasm was faint and completely occupied by large

vacuoles. In previous studies, summer season had a detrimental effect on the maturation rate of buffalo [44] and bovine oocytes [14,15]. Reduced nuclear maturation is one of the changes caused by heat shock in bovine oocytes. Exposure of oocytes at GV-stage [39] or maturing oocytes [16,24] from crossbred Bos indicus to 41
C heat shock decreased the proportion of oocytes that reached MII stage after IVM. In these experiments, heat shock blocked meiotic progression by increasing the proportion of MI stage oocytes. Furthermore, there is evidence that oocyte cytoplasm is more susceptible to the adverse effects of increased temperature than the nucleus [47]. Heat stress impairs intracellular events associated with both nuclear and cytoplasmic maturation, such as translocation of cortical granules to the oolemma [39], cytoskeletal rearrangement [24], and spindle formation [23,24]. Recently, Maya-Soriano et al. [48] reported that when matured under heat shock conditions, bovine oocytes obtained in both HS and CS were similarly affected in terms of nuclear maturation, whereas a seasonal effect was observed on cytoplasmic maturation. Meanwhile, the present work illustrated that IVM and IVF of buffalo oocytes during HS significantly (P < 0.01) decreased cleavage rate and embryo development until the blastocyst stage. Similarly, Mishra et al. [49] have suggested that high ambient temperature during summer have

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Fig. 5. (continued).

influenced the competence of buffalo oocytes to cleave and develop to the blastocyst stage. Developmental competence of bovine GV-stage Holstein oocytes was affected by increased temperature [14,50,51]. The proportion of such GV-stage oocytes develops to the blastocyst stage after fertilization was less in the warm season compared with the cool season [14]. In a study that used ultrasound-guided follicular aspiration (i.e., oocyte pickup) to collect Holstein and Brahman oocytes during the CS and HS, the percentage of Holstein oocytes with normal morphology decreased from 80% in the CS to 25% in the HS. During the CS, embryo development was higher (44.4% eight-cell stage, 34.2% morula, and 29% blastocysts) than the HS (1.1% eight-cell stage, 0% morula, and 0% blastocyst) [50]. On the contrary, a series of in vivo [50,52] and in vitro studies [53] revealed that elevated temperature did not affect oocyte cleavage rate, but the blastocyst rate was lower than that under normal conditions [22,54]. Therefore, the reason for these discrepancies in embryo development may be genetic differences in susceptibility to elevated temperature between species or differences in the duration of exposure to or the severity of temperature. The mechanisms by which increased temperature affects oocyte physiology are not completely understood. Oocyte morphology associated with competence to undergo meiosis and development to the blastocyst stage can be positively

correlated with a greater relative abundance of known mRNA transcripts. quantitative Reverse Transcription-Polymerase Chain Reaction analysis showed that the important housekeeping gene GAPDH was differentially expressed in in vitro matured oocytes at different seasons, with a significant downregulation of in vitro matured oocytes during summer season. Similar results were recorded for bovine oocytes by Gendeleman and Roth [15], showing that GAPDH expression was lower in HS than CS. Glyceraldehyde 3-phosphate dehydrogenase plays a key role in energy metabolism and is involved in the glycolytic pathway [55]. In the present work, the transcript abundance of ACTB mRNA tended to be increased in in vitro matured oocytes during HS than CS. However, such an effect was not observed in COCs. The mechanism by which HS could impair ACTB mRNA expression is still unknown. Actin microfilaments are responsible for cortical granule translocation to the cortical region of the oocyte during maturation [56]. Exposure of GV-stage [39] and maturing oocytes [57] to 41
C heat shock increased the proportion of oocytes that had a type III cortical granule distribution, indicating that heat shock hastened cytoplasmic maturation kinetics and induced oocyte aging. Also, the percentage of mouse oocytes carrying incomplete cortical granule migration during IVM was greater in oocytes matured at 40
C than 37
C [58]. So, the higher expression of ACTB mRNA in in vitro matured

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oocytes during HS may indicate that high environmental temperature during summer may hasten cytoplasmic maturation kinetics and induce oocyte aging and consequently lead to the higher incidence of cytoplasmic degeneration in the present work. It has been reported that GDF9 and BMP15 are essential for the proliferation of granulosa cells during preantral and antral follicular development [59,60]. This would explain why the transcript abundance of the developmentally important genes GDF9 and BMP15 in our work was almost similar between buffalo COCs or in vitro matured oocytes collected during CS and HS. This is in contrast to results that showed a higher expression in bovine oocytes recovered during CS compared with those recovered during HS [15]. Seasonally induced alteration of GDF9 expression in the ovarian pool of follicles and/or their enclosed oocytes underlies the immediate and carryover effect of summer heat stress on follicular function and oocyte developmental competence [15], which might explain the reduced proportion of mature oocytes in the bovine [8,57]. However, our results are supported by Gendelman et al. [51], who found that the level of GDF9 mRNA in bovine embryos did not show any difference between HS and CS. Moreover, BMP15 may play a key role in the developmental competence of bovine oocytes. The lower developmental competence of calf oocytes may be partially explained by a deficiency of BMP15 in cumulus cells [61]. Bone morphogenetic protein 15 maintains a low incidence of cumulus cell apoptosis by establishing a localized gradient of BMPs [62]. Contradictions among the results may be because of the differences in the degree and duration of temperature treatment among studies and/or in the ability of different genetic lines to tolerate high environmental temperature. In the present work, no difference was found for PTGS2 expression in COCs and in vitro matured oocytes between seasons. This is supported by the finding that COC quality has no effect on the expression of PTGS2 [63]. Silva et al. [29] reported that heat stress decreased the expression of PTGS2 mRNA in bovine embryos. This is supported by a reported finding that PTGS2 was more abundant in fully competent blastocysts than in those resulting in abortion after transfer [64]. This difference could be because of difference in experimental design, the developmental stage, genetic differences between species, or severity of temperature. Furthermore, HSP70 mRNA expression was differentially regulated in buffalo COCs and in vitro matured oocytes during HS and CS. This gene has a proapoptotic function and its upregulation indicated signs of degeneration in buffalo COCs recovered during HS. Transcription of HSP70 may be increased by stressful conditions like heat shock [65]; its expression has been used as an indicator of stress in bovine embryos [66]. In cattle, immature Jersey oocytes were more sensitive to heat stress and contained higher HSP70 mRNA levels than Gyur oocytes [67], showing species or breed differences. Exposure of COCs to elevated temperature increased HSP70 mRNA in cumulus cells, and some of the negative effects of heat stress on oocytes were mediated through the cumulus cells [68]. Also, exposure of pig ovaries to elevated temperature (41.5
C) for 1 hour significantly disrupted oocyte maturation and resulted in upregulation of HSP70 mRNA expression [69].

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In the present data, the relative abundance of GAPDH, ACTB, GDF9, BMP15, PTGS2, and HSP70 mRNAs was higher in buffalo COCs than in in vitro matured oocytes both in HS and CS. It is known that maternal mRNAs of many genes accumulate in the oocyte and are progressively degraded during maturation and early cleavage [70]. It is possible that IVM might increase the sensitivity of maturing oocytes to heat stress. 4.1. Conclusions The present data show for the first time in the buffalo species that the lower developmental competence during HS is because of a poorer quality of COCs, decreased nuclear and cytoplasmic maturation, and consequently, reduced development to the blastocyst stage. These results could be in part because of the observed relative downregulation of developmentally important genes and upregulation of stress response genes during oocyte growth and maturation. Acknowledgments The authors greatly appreciated the financial support from the National Research Center of Egypt (in-house project 9040207) and the support by the Deutsche Forschungsgemeinschaft (DFG EI 296/15-1 and DFG GA1077/ 5-1). Ahmed S. Abdoon was financially supported by the Deutsche Forschungsgemeinschaft to work at Free University of Berlin. References [1] Badinga L, Collier RJ, Thatcher WW, Wilcox CJ. Effects of climatic and management factors on conception rate of dairy cattle in subtropical environment. J Dairy Sci 1985;68:78–85. [2] Pires MFA, Ferreira AM, Saturnino HM, Teodoro RL. Taxa de gestação em fêmeas da raça Holandesa confinadas em free stall, no verão e inverno. Arq Bras Med Vet Zootecn 2002;54:57–63. [3] Marai IF, Haeeb AA. Buffalo’s biological functions as affected by heat stressda review. Livest Sci 2010;127:89–109. [4] Ali A, Abdel-Razek AKh, Derar R, Abdel-Rheem HA, Shehata SH. Forms of reproductive disorders in cattle and buffaloes in Middle Egypt. Reprod Domest Anim 2009;44:580–6. [5] Hansen PJ, Drost M, Rivera RM, Paula-Lopes FF, Al-Katanani YM, Krininger CE. Adverse impact of heat stress on embryo production: causes and strategies for mitigation. Theriogenology 2001;55:91– 103. [6] Gwazdauskas FC, Thatcher WW, Kiddy CA, Paape MJ, Wilcox CJ. Hormonal patterns during heat stress following PGF2 alpha-tham salt induced luteal regression in heifers. Theriogenology 1981;16: 271–85. [7] Badinga L, Thatcher WW, Diaz T, Drost M, Wolfenson D. Effect of environmental heat stress on follicular development and steriodogenesis in lactating Holstein cows. Theriogenology 1993;39:797– 810. [8] Wolfenson D, Thatcher WW, Badinga L, Savio JD, Meidan R, Lew BJ, et al. Effect of heat stress on follicular development during the estrous cycle in lactating dairy cattle. Biol Reprod 1995;52:1106–13. [9] Roth Z, Meidan R, Braw-Tal R, Wolfenson D. Immediate and delayed effects of heat stress on follicular development and its association with plasma FSH and inhibin concentration in cows. J Reprod Fertil 2000;120:83–90. [10] Roman-Ponce H, Thatcher WW, Canton D, Barron DH, Wolcox CJ. Thermal stress effects on uterine blood flow in dairy cows. J Anim Sci 1978;46:175–80. [11] Malayer JR, Hansen PJ, Buhi WC. Effect of day of oestrus cycle, side of the reproductive tract and heat shock on in-vitro protein secretion by bovine endometrium. J Reprod Fertil 1988;84:567–78.

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