Confocal fluorescence assessment of bioenergy/redox status of dromedary camel (Camelus dromedarius) oocytes before and after in vitromaturation
© Russo et al.; licensee BioMed Central Ltd. 2014
Received: 22 October 2013
Accepted: 12 February 2014
Published: 18 February 2014
Reproductive biotechnologies in dromedary camel (Camelus dromedarius) are less developed than in other livestock species. The in vitro maturation (IVM) technology is a fundamental step for in vitro embryo production (IVP), and its optimization could represent a way to increase the success rate of IVP. The aim of the present study was to investigate the bioenergy/oxidative status of dromedary camel oocytes before and after IVM by confocal microscopy 3D imaging.
Oocytes were retrieved by slicing ovaries collected at local slaughterhouses. Recovered oocytes were examined before and after IVM culture for nuclear chromatin configuration and bioenergy/oxidative status, expressed as mitochondria (mt) distribution and activity, intracellular Reactive Oxygen Species (ROS) levels and distribution and mt/ROS colocalization.
The mean recovery rate was 6 oocytes/ovary. After IVM, 61% of oocytes resumed meiosis and 36% reached the Metaphase II stage (MII). Oocyte bioenergy/redox confocal characterization revealed changes upon meiosis progression. Immature oocytes at the germinal vesicle (GV) stage were characterised by prevailing homogeneous mt distribution in small aggregates while MI and MII oocytes showed significantly higher rates of pericortical mt distribution organized in tubular networks (P < 0.05). Increased mt activity in MI (P < 0.001) and MII (P < 0.01) oocytes compared to GV stage oocytes was also observed. At any meiotic stage, homogeneous distribution of intracellular ROS was observed. Intracellular ROS levels also increased in MI (P < 0.01) and MII (P < 0.05) oocytes compared to GV stage oocytes. The mt/ROS colocalization signal increased in MI oocytes (P < 0.05).
This study provides indications that qualitative and quantitative indicators of bioenergy and oxidative status in dromedary camel oocytes are modified in relation with oocyte meiotic stage. These data may increase the knowledge of camel oocyte physiology, in order to enhance the efficiency of IVP procedures.
The dromedary camel (Camelus dromedarius) is a seasonal polyestrous breeder and reproductive activity is limited to the coolest month of the year . Local environmental factors such as rainfall, external temperature, and nutrition, rather than photoperiod, are supposed to influence the moment of initiation of sexual activity and length of breeding season . Like other camelids, dromedary camels are induced ovulators  but spontaneous ovulation sometimes occurs . The breeding season occurs during winter (November/December to March/April), though there are conflicting reports about its beginning and length . Outside the breeding season, mating activity ceases and the ovaries are inactive or have a few small follicles .
The interest in dromedary camel breeding is due to its role as a food resource in semi-arid areas and also to the commercial interest in the racing sector. The total number of camels in the world is reckoned to be around 25 million heads; though this number is probably underestimated, a steady increase has been recorded in the last 20 years . In order to optimize breeding programs in this species, monitoring and increasing efficiency plans of the reproductive activity are desirable. To this end, application of assisted reproductive technologies (ARTs), such as artificial insemination, oocyte in vitro maturation (IVM) and fertilization, oocyte and embryo cryopreservation, embryo transfer and cloning could provide a substantial opportunity to improve reproductive efficiency and genetic performance and to better understand factors regulating reproductive activity in this species.
The successful development of ARTs relies on obtaining functionally competent oocytes which are a mandatory requisite for successful embryo production.
IVM is a fundamental step in in vitro embryo production (IVP) and, in any species, its optimization could represent a way to increase the success rate of IVP. The regulation of oocyte maturation not only affects the proportion of oocytes capable of undergoing maturation, but also their subsequent fertilization and embryo development . There is very little information on oocyte physiology in camelids, although the kinetics of oocyte nuclear maturation has been studied [8, 9]. The complex process of cytoplasmic maturation has not been widely investigated in this species. To date, only one study reported offspring in camelids obtained by transfer of embryos produced by IVP . These authors recovered oocytes from abattoir-derived ovaries, cultured them in vitro for 30 h in TCM-199 medium supplemented with fetal calf serum (FCS), epidermal growth factor (EGF), follicle stimulating hormone (FSH), estradiol and cysteamine. After maturation, oocytes underwent in vitro fertilization with motile spermatozoa obtained by centrifugation of fresh semen on a Percoll discontinuous gradient and putative embryos were cultured in a semi-defined medium (mKSOMaa supplemented after 48 h with 10% FCS). The same authors reported 23% blastocyst formation rate and 13 normal dromedary camel offspring produced by IVP . Other studies have been reported on IVM technology in dromedary camel oocytes [5, 9, 11–13]. These studies analysed the role of reproductive season and the effects of different culture media on in vitro oocyte meiotic development.
Increasing evidence shows the role of mitochondria (mt) as determinant for developmental competence of human and mammalian oocytes [14–16]. Regular mitochondrial bioenergetic activities within the oocyte include mt adenosine triphosphate (ATP) generation, roles in Ca2+ homeostasis, regulation of cytoplasmic redox state, and signal transduction [17, 18]. It has been reported that oocyte competence is directly related to cytoplasmic bioenergetic capacity, and that ATP supply/demand imbalances, associated with suboptimal levels of mitochondrial ATP generation, may be a common cause of several developmental defects . Mitochondrial dysfunctions or abnormalities may compromise developmental processes by inducing chromosomal segregation disorders, maturation and fertilization failures, or oocyte/embryo fragmentation resulting in mt-driven apoptosis [15, 17]. Mitochondria are the major source of reactive oxygen species (ROS), which are produced during oxidative phosphorylation . Under physiologic conditions, ROS are neutralized by an elaborate defence system . If disequilibrium between ROS production and antioxidative capacity of the cell takes place, oxidative stress occurs . Although the impact of oxidative stress on oocyte and reproductive function remains unclear, it has been shown that oxidative stress affects oocyte Ca2+ homeostasis, and thus also oocyte maturation and oocyte-sperm interaction .
To our knowledge, only one study has been reported to date on mt complement distribution and activity in dromedary camel oocytes as assessed by confocal microscopy . These authors investigated mt distribution and activity, ATP, glutathione (GSH) content and calcium oscillation during IVM. They observed a highly polarized mt distribution pattern in MII oocytes compared to immature ones. In the same study, higher ATP content was observed in MII oocytes than in immature ones.
The aim of the present study was to investigate the bioenergy/oxidative status of dromedary camel oocytes in relation to their meiotic stage by simultaneous analysis of oocyte nuclear chromatin configuration and qualitative and quantitative aspects of bioenergy/oxidative status. A confocal laser scanning microscopy (CLSM)-based multiparametric assessment of oocyte bioenergy/redox status, expressed as mitochondrial distribution pattern, mitochondrial activity, intracellular ROS levels and localization and mitochondria/ROS colocalization, was performed. Increasing the knowledge of camel oocyte physiology could be useful to enhance the efficiency of IVM procedures, for a species in which reproductive biotechnologies would have wide application.
The study was performed in accordance with the ethical standards laid down in the 1964 Declaration of Helsinki and its subsequent amendments. All the procedures with animals were performed following good veterinary practice for animal welfare according to the Italian law (D.Lgs 116/92).
Chemicals and drugs
All chemicals for in vitro cultures and analyses were purchased from Sigma-Aldrich (Milan, Italy) unless otherwise indicated.
Oocyte collection and maturation in vitro
Oocyte collection and in vitro culture were performed in Egypt. Five replicates were performed during the early non breeding season (May-June 2012 and 2013). Ovaries were collected at a local abattoir, transported to the laboratory in physiological saline solution at 30–35°C and washed once in 70% ethanol and twice with warm (37°C) phosphate buffered saline (PBS) containing 100 IU/mL penicillin G sodium and 100 μg/mL streptomycin. Cumulus-oocyte complexes (COCs) were recovered by slicing ovaries as previously reported . Oocytes with uniform cytoplasm and multilayered cumulus cells were selected, washed twice in HEPES-TCM medium: TCM 199 supplemented with 25 mM HEPES, 2 mM sodium bicarbonate, 2 mM sodium pyruvate, 1 mM L-glutamine and 10% FCS and destined to either immediate evaluation or in vitro maturation (IVM) culture. The IVM culture was performed in TCM199 (Gibco 22340) supplemented with 15% FCS, 0.25 mg/mL pyruvic acid, 20 ng/mL EGF, 50 μg/mL gentamicin, 10 μg/mL FSH (Sigma-Aldrich, F2293), 10 μg/mL luteinizing hormone (LH; Sigma-Aldrich, L6420), 1 μg/mL estradiol and 0.3 mM cystine. Groups of 25 COCs were matured in 400 μl of IVM medium, covered with 400 μl of mineral oil, in four well culture plates (Nunc, Roskilde, Denmark). On the basis of previously reported studies, oocytes were cultured for 40–42 h at 38.5°C under 5% CO2 in air with 95% relative humidity [12, 24]. Oocytes were denuded by gentle pipetting in HEPES-TCM 199 with 80 IU of hyaluronidase and underwent epifluorescence and confocal microscopy evaluation.
Oocyte mitochondria and ROS staining
To investigate mitochondrial distribution and apparent energy status, oocytes were washed three times in PBS with 3% bovine serum albumin (BSA) and incubated for 30 min in the same medium containing 280 nM MitoTracker Orange CMTM Ros (Molecular Probes M-7510, Oregon, USA) at 38.5°C under 5% CO2[25, 26]. The cell-permeant probe contains a thiol-reactive chloromethyl moiety. Once the MitoTracker probe accumulates in the mitochondria, it can react with accessible thiol groups on peptides and proteins to form an aldehyde-fixable conjugate. This cell-permeant probe is readily sequestered only by active mitochondria [27, 28]. The organelle-specificity of the probe was assessed, as reported by Valentini et al. . Control oocytes were imaged after incubation in MitoTracker Orange and further incubation for 5 min in the presence of 5 μM of the mt membrane potential (Delta Psi)-collapsing uncoupler carbonyl cyanide 3-chloro phenyl hydrazone (CCCP; Molecular Probes; Monza, Italy). This molecule inhibits mt respiratory activity, thus reducing fluorescence intensity. After incubation with mt probe, oocytes were washed three times in PBS with 0.3% BSA and incubated for 15 min in the same media containing 10 μM 2′,7′-dichlorodihydrofluorescein diacetate (H2DCF-DA) [30–32] in order to detect and localize intracellular sources of ROS. The principle underlying this procedure may be described briefly as follows: non-ionized H2DCF-DA is membrane-permeant and is therefore able to diffuse readily into cells. Once within the cell, the acetate groups are hydrolysed by intracellular esterase activity forming 2′,7′-dichlorodihydrofluorescein (H2DCF) which is polar and thus trapped within the cell. H2DCF fluoresces when it is oxidized by H2O2 or lipid peroxides to yield 2′,7′-dichlorofluorescein (DCF). The level of DCF produced within the cells is related linearly to that of peroxides present and thus its fluorescent emission provides a measure of the peroxide levels . After incubation, oocytes were washed three times in pre-warmed PBS without BSA and fixed overnight at 4°C with 2% paraformaldehyde solution in PBS.
Nuclear chromatin evaluation of oocytes
To evaluate nuclear chromatin, oocytes were stained with 2.5 μg/mL Hoechst 33258 in 3:1 (v/v) glycerol/PBS and mounted on microscope slides covered with cover slips, sealed with nail polish and kept at 4°C in the dark until observation. Oocytes were evaluated in relation to their meiotic stage under an epifluorescence microscope (Nikon Eclipse 600; Nikon Instruments, Amsterdam, The Netherlands; 400x magnification) equipped with B-2 A (346 nm excitation/460 nm emission) filter, as germinal vesicle (GV), metaphase I (MI) and metaphase II (MII) with 1st polar body extruded. Oocytes with multipolar spindle, irregular chromatin distribution or absence of chromatin were classified as degenerated and were excluded from further analysis.
Assessment of oocyte mitochondrial distribution pattern and intracellular ROS localization
For evaluation of mt distribution pattern, oocytes were selected among those having homogeneous cytoplasmic texture. Oocytes were observed at 600x magnification in oil immersion with Nikon C1/TE2000-U laser scanning confocal microscope (Nikon Instruments, Amsterdam, The Netherlands). A helium/neon laser ray at 543 nm and the G-2 A filter (551 nm exposure and 576 nm emission) were used to point out the MitoTracker Orange CMTM Ros. An argon-ion laser ray at 488 nm and the B-2 A filter (495 nm exposure and 519 nm emission) were used to point out the DCF. For each oocyte, scanning was conducted with 25 optical series from the top to the bottom of the oocyte with a step size of 0.45 μm to allow three-dimensional distribution analysis. General criteria for definition of oocyte mt patterns were adopted on the basis of previous studies in other species [25, 26, 29, 32]. Homogeneous/even distribution of small mt aggregates (SA) throughout the cytoplasm was considered as indication of immature cytoplasmic condition, whereas heterogeneous/uneven distribution of small and/or large aggregates within the cytoplasm indicated a metabolically active ooplasm. In particular, tubular mt networks spread throughout the cytoplasm (diffused tubular network, DTN) or their accumulation in the peripheral cytoplasm (pericortical tubular network, PCTN) or in the periphery and around the nucleus (perinuclear/pericortical tubular network, PN/PCTN) were considered as sequential aspects of the developmental program of cytoplasmic maturation . Oocytes showing irregular distribution of large mt clusters unrelated to the specific cell compartments were classified as abnormal and were excluded from further quantification analysis.
Quantification of MitoTracker orange CMTM Ros and DCF fluorescence intensity
In each individual oocyte, MitoTracker Orange CMTM Ros and DCF fluorescence intensities were measured at the equatorial plane, as in previous studies in human  and animal oocytes [25, 26, 29, 32], with the aid of the EZ-C1 Gold Version 3.70 image analysis software platform for Nikon C1 (Nikon Instruments) confocal microscope. A circle of area = 100 in diameter (arbitrary value) was drawn to measure only the cytoplasmic area (512 by 512 pixels). The fluorescence intensity encountered within the programmed scan area was recorded and plotted against the conventional pixel unit scale (0–255). Fluorescence intensity was expressed as Arbitrary Densitometric Units (ADU). Parameters related to fluorescence intensity were maintained at constant values for all measurements. In detail, images were taken under fixed scanning conditions with respect to laser energy, signal detection (gain) and pinhole size.
Oocyte mitochondria/ROS colocalization analysis
Colocalization analysis of mitochondria and ROS was performed by using the EZC1 Gold Version 3.70 software. Degree of colocalization was reported as a Pearson’s correlation coefficient quantifying the overlap degree between MitoTracker Orange CMTM Ros and DCF fluorescence signals [26, 35].
Nuclear maturation rates and mt distribution patterns were compared by chi-square analysis. For confocal quantification analysis of mt activity and intracellular ROS levels, the least-square means of the dependent variable (MitoTracker Orange CMTM Ros and DCF fluorescence intensity) were calculated in examined samples and the statistical significance of the least-square means between treated and control groups was calculated by one-way ANOVA followed by Multiple Comparison Dunn’s method (SigmaPlot software). For mt/ROS colocalization, mean values of Pearson’s correlation coefficient were compared between treated and control groups by one-way ANOVA followed by Multiple Comparison Dunn’s method (SigmaPlot software). Differences with P < 0.05 were considered as being statistically significant.
Mitochondrial distribution pattern of dromedary camel oocytes before (*) and after IVM
Nuclear chromatin configuration
Mitochondrial distribution pattern
Results of mt distribution pattern analysis are presented in Table 1. The rate of oocytes showing PCTN mt distribution pattern was significantly higher in oocytes cultured in vitro and reaching the MI and MII stages compared with uncultured GV stage oocytes (31% vs 9% for MII and GV; 33% vs 9% for MI and GV, respectively, P < 0.05). The rate of oocytes showing PN/PCTN also increased with maturation, though not statistically significant. Likewise, the rate of oocytes showing mt distribution in SA was significantly reduced both in MI and MII oocytes compared to GV oocytes (41% versus 78% for MI and GV, respectively, P < 0.02; 38% vs 78% for MII and GV, respectively, P < 0.01).
Bioenergy/redox status analysis
IVM culture duration could have a significant impact on oocyte metabolic/oxidative status, as expressed by examined confocal microscopy parameters. Wani and Nowshari  investigated the optimum time and the kinetics of IVM in the dromedary camel. These authors found that dromedary camel oocytes required 32–44 h of in vitro culture to get an optimum number of oocytes in the MII stage. On this basis, we performed our experiments after 40 h of maturation, which is reported by Wani and Nowshari  as the optimum IVM time leading to the highest maturation rate. We analyzed the basal bioenergy/oxidative status of IVM oocytes and how it changes during meiosis progression. Our results on the maturation rate are in line with those reported by other authors [5, 12]. Zeidan et al.  reported 68.2% maturation rate when using TCM199 in oocytes recovered during the breeding season. In the same study, the maturation rate decreased to 44.4% in oocytes recovered in the non-breeding season. Thus, due to our experimental period (early non-breeding season), our maturation rate (36%) are in line with those reported by these authors for oocytes recovered during the non-breeding season. Other studies reported similar  or higher maturation rates (84%,  and 81%, ) as related to the use of different culture media. Abdoon et al.  reported a MII rate of 37.5% by using TCM-199 supplemented with 10% FCS, 10 μg/mL FSH, 10 ng/mL EGF, 10 IU/mL equine chorionic gonadotropin (eCG), and 10 IU/mL human chorionic gonadotropin (hCG). Khatir et al.  obtained a maturation rate of 84% by using TCM-199 supplemented with 10% FCS, 10 ng/ml EGF and 250 μM cysteamine. Wani and Wernery  reported that 81% of in vitro cultured oocytes reached the MII stage by using TCM-199 supplemented with 0.4% bovine serum albumin, 10 μg/mL FSH, 10 μg/mL LH, and 1 μg/mL estradiol.
Regular mt bioenergetic activities within the oocyte play a crucial role as determinants for developmental competence for human and mammalian oocytes [15, 16]. During oocyte maturation and fertilization, nuclear changes are coordinated with movements of organelles, in particular mitochondria, and with biochemical changes in the cytoplasm to ensure normal embryo development . Of the numerous cytoplasmic changes occurring during oocyte maturation, the positioning of mitochondria may be involved in concentrating ATP or calcium or in regulating pH in specific regions of the oocyte to support normal developmental processes. A mt distribution pattern polarized in the peri-spindle (perinuclear) and in the pericortical region of the oocyte has been regarded as one aspect of cytoplasmic maturation in different species, as reported in previous studies from our unit in equine, ovine, canine and human oocytes [25, 26, 29, 32, 34]. In the present study, qualitative analysis of mt distribution in dromedary camel oocytes, performed before or after in vitro maturation, revealed that mt distribution patterns within the oocytes were changed in relation to the meiotic stage. Oocytes at the GV stage showed a predominantly uniform mt distribution, organized in SA. In oocytes at the MI and MII stages, a reduction in oocytes showing SA mt distribution pattern and an increase in oocytes with a heterogeneous PCTN mt distribution pattern were observed. These results are in line with those reported in previous studies. Using confocal analysis, Abdoon et al.  reported mt clustering in MI oocytes (referred as matured oocytes without PB in their study). These authors also reported highly polarized mt distribution pattern in the subcortical region of MII oocytes showing the first PB extruded. Using ultrastructural TEM analysis, Kafi et al.  reported arrangement of most mitochondria in the oocyte periphery at 24–36 hours IVM culture, which is demonstrated as being the time of MI to MII transition.
To our knowledge, this is the first study jointly evaluating the nuclear chromatin configuration and the bioenergy and redox status of dromedary camel oocytes, with a triple staining procedure and an epifluorescence/3D confocal evaluation of individual oocytes, in relation with meiotic progression. Moreover, the quantification study was performed by objective multiparametric analysis. MitoTracker Orange CMTM Ros fluorescence intensity analysis revealed an increase of mt activity in oocytes at MI and MII stage compared with those at GV stage. This finding indicated that the achievement of nuclear maturation was related to an increase in the number of actively respiring mitochondria within the ooplasm and/or to an increase in the number of active sites per each mitochondrion, as well as to a change in mt distribution. These data are in line with observations by Abdoon et al. , concerning increased ATP content in MII compared with GV oocytes. DCF fluorescence intensity also revealed an increase in intracellular ROS levels in oocytes at MI and MII compared to GV stage. This result could be explained in terms of increased oxidative phosphorylation which may be translated into an increase in ROS production.
Mitochondria/ROS colocalization, objectively expressed as Pearson’s correlation coefficient, was higher in oocytes at MI stage compared with GV stage oocytes. It also tended to remain at higher values in MII compared with GV stage oocytes, though not significantly different. Colocalization of ROS and actively respiring mitochondria was reported to be indicative of higher ATP turnover resulting from a more intense mt activity and thus indicative of healthy cell conditions, as demonstrated in hepatocytes . Another study, in mouse oocytes, reported that the regions within the oocyte producing high levels of ROS colocalized with the active mitochondria, as visually assessed by using confocal microscopy, in the majority of in vivo MII ovulated oocytes . Moreover, higher levels of mt/ROS colocalization were reported as a reliable marker of in vivo matured MII ovine oocytes .
In conclusion, this study provides simultaneous information on bioenergy and redox status of dromedary camel oocytes as related to their meiotic stage before and after in vitro maturation and contributes to knowledge of camel oocyte physiology, which may enhance the efficiency of IVM procedures and in vitro production of embryos in this species.
This document has been produced with the financial assistance of the European Union: ENPI-CBC- MED “PROCAMED” Project: Promotion des systèmes camelins innovants et des filières locales pour une gestion durable des territoires saharienne; reference number I.B/1.1/493. The contents of this document are the sole responsibility of “Section of Clinics and Animal Productions, Department of Emergency and Organ Trasplantation - University of Bari Aldo Moro, Italy” and can under no circumstances be regarded as reflecting the position of the European Union. Authors are grateful to the Direction of CURP for the logistic support and help during the implementation of this work. Additional supports were received from CEGBA project, University of Bari Aldo Moro Italy and ONEV MIUR PONa3_00134 project n.254/R&C 18/05/2011. The authors thank Anthony Green of the Department of Veterinary Medicine, University of Bari Aldo Moro for language revision and assistance.
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