Biol Reprod Lalor Postdoctoral Fellowships -- Application Deadline January 15, 2009
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

BOR - Papers in Press, published online ahead of print June 2, 2004.
Biol Reprod 2004, 10.1095/biolreprod.104.029264
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
71/4/1150    most recent
biolreprod.104.029264v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Tatemoto, H.
Right arrow Articles by Nakada, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Tatemoto, H.
Right arrow Articles by Nakada, T.
Agricola
Right arrow Articles by Tatemoto, H.
Right arrow Articles by Nakada, T.
BIOLOGY OF REPRODUCTION 71, 1150–1157 (2004)
DOI: 10.1095/biolreprod.104.029264
© 2004 by the Society for the Study of Reproduction, Inc.

Gamete Biology

Protection of Porcine Oocytes Against Cell Damage Caused by Oxidative Stress During In Vitro Maturation: Role of Superoxide Dismutase Activity in Porcine Follicular Fluid1

Hideki Tatemoto2,3, Norio Muto4, Ikuko Sunagawa3, Akihisa Shinjo3, and Tadashi Nakada3

Department of Bioproduction,3 Faculty of Agriculture, University of the Ryukyus, Nishihara-cho, Okinawa 903-0213, Japan School of Bioresources,4 Hiroshima Prefectural University, Shobara, Hiroshima 727-0023, Japan


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
To elucidate the beneficial effects of porcine follicular fluid (pFF) added to maturation medium on the sustenance of cytoplasmic maturation responsible for the subsequent developmental competence after in vitro fertilization (IVF) of porcine oocytes, we focused on the antioxidative role of pFF in its function of protecting oocytes from reactive oxygen species (ROS)-induced cell damage. Porcine follicular fluid collected from small (2–6 mm) follicles had about 7.2-fold higher levels of superoxide dismutase (SOD) activity than that of fetal bovine serum (FBS), and this activity was markedly blocked by the CuZn-SOD inhibitor, diethyldithiocarbamate (DETC). The interruption of meiotic progression and the increasing intracellular glutathione (GSH) content throughout the maturation period, as well as an outbreak of DNA damage in oocytes and cumulus cells were difficult to detect in oocytes cultured in a medium supplemented with 10% pFF, even in the presence of ROS generated by the hypoxanthine-xanthine oxidase system, whereas cell damage encompassed by ROS was prominent in oocytes cultured with 10% FBS and 10% pFF plus 100 µM DETC. Similarly, significant enhancement to the degree of transformation of the sperm nucleus into the male pronucleus (MPN) after in vitro fertilization was shown by the addition of pFF to the maturation medium. The presence of DETC during in vitro maturation reduced the ability of oocytes to promote MPN formation to the same extent as oocytes matured with FBS. The proportion developing to the blastocyst stage was increased in oocytes that matured with pFF, but this developmental competence was significantly lowered by treatment with DETC (P < 0.05). These findings suggest that pFF plays a critical role in protecting oocytes from oxidative stress through a higher level of radical scavenging activity elicited from SOD isoenzymes, resulting in the enhancement of cytoplasmic maturation responsible for developmental competence postfertilization.

apoptosis, early development, gamete biology, in vitro fertilization, ovum


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
It has been a longstanding objective to identify optimal conditions for in vitro maturation (IVM) of follicular oocytes from domestic animals to enhance basic agricultural research and biotechnology. Although porcine oocytes can develop to the blastocyst stage following maturation and fertilization in vitro [1–4], their developmental potential is lower than that of oocytes that mature in vivo [5–7]. Oocyte maturation includes both nuclear and cytoplasmic maturation. The poor developmental competence might be caused by the lack of cytoplasmic maturation in porcine oocytes maturing in vitro, even though they undergo normal nuclear maturation [4, 8–10]. Usually, the deficiency of cytoplasmic maturation during IVM is reflected in an oocyte's low ability to form a male pronucleus (MPN) and to develop to the blastocyst stage after in vitro insemination [11].

In vitro cultures are maintained under higher concentrations of O2 than those that occur in vivo, resulting in increased production of reactive oxygen species (ROS). In our previous study [12], cumulus cells efficiently protected porcine oocytes against cell damage caused by oxidative stress during IVM. After IVM, the oocytes surrounded by cumulus cells (cumulus-oocyte complexes; COCs) had significantly increased concentrations of intracellular glutathione (GSH), whereas the GSH content in cumulus-denuded oocytes was markedly decreased, and exposure of denuded oocytes to ROS resulted in an increase in the frequency of apoptotic cell death. Moreover, the protection of porcine oocytes from oxidative stress during IVM efficiently enhanced the acquisition of developmental competence after fertilization [13]. However, no data exist on the intrinsic functions or factors relating to the meiotic progression accompanying cytoplasmic maturation in porcine oocytes.

In vivo, oocytes are nourished by follicular fluid (FF) and undergo meiotic maturation. The effects of adding FF to maturation media on the developmental competence after in vitro fertilization (IVF) have been investigated in bovine [14–16] and porcine [17–20] oocytes. As reported by Naito et al. [17] and Rath et al. [19], porcine FF (pFF) added to IVM medium had beneficial effects on the resumption of meiosis and on MPN formation in combination with FSH. Yoshida et al. [18] also reported that the addition of pFF to IVM medium significantly increased the rate of nuclear maturation, normal fertilization, and normal cleavage of porcine oocytes after IVF, and one or more heat-labile (56°C) acidic factors with a molecular mass between 10 and 200 kDa in pFF is responsible for oocyte maturation and developmental capacity. A variety of peptide growth factors and steroid hormones have been demonstrated to be present in FF [15, 21–25]. From these findings, it seems that pFF is highly effective in enhancing the developmental competence of porcine oocytes in vitro.

The presence of the superoxide dismutase (SOD) isoenzyme has been reported in human FF [26–28]. SOD activity and CuZn-SOD isoenzymes are localized in developing follicles, membranes of granulosa cells of Graafian follicles, and postovulatory follicles [29]. Further studies on rat and human ovaries suggest that ROS and SOD isoenzymes may play a role in the ovulation process and in the development of oocytes [26–28, 30]. Depending on the site of formation and on the tension of oxygen, ROS such as superoxide anions (O2–), hydroxyl radicals (OH–), and hydrogen peroxide (H2O2) can cause lipid peroxidation and enzyme inactivation, resulting in tissue injury in humans and animals [31]. Therefore, the present study was conducted to examine the protective effects of SOD activity in pFF on oxidative stress caused by the hypoxanthine-xanthine oxidase (XOD) system during IVM of porcine oocytes. Additionally, to clarify whether the addition of pFF to IVM media supported the cytoplasmic maturation responsible for the subsequent developmental competence, the desirable effects of the SOD isoenzyme in pFF through the scavenging ability to protect porcine oocytes from the detrimental effects of ROS during IVM were compared with SOD activity-abolished pFF by the SOD inhibitor, diethyldithiocarbamate (DETC), and fetal bovine serum (FBS).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
All chemicals used in the study were purchased from Sigma Chemical Company (St. Louis, MO) unless otherwise stated.

Collection of Oocytes and pFF

Ovaries were collected from peripubertal gilts at a local slaughterhouse and transported to the laboratory in 0.9% (w:v) NaCl containing 100 mg/L kanamycin sulfate (Meiji Seika, Tokyo, Japan) at 30°C. Within 2 h postslaughter, the follicular contents were recovered by excising the visible small antral follicles (about 2–6 mm in diameter) on the ovarian surface by using a razor, and by scraping the inner surface of the follicle walls with a disposable surgical blade. Only COCs with a uniform ooplasm and a compact cumulus cell mass were collected and washed three times with Hepes-buffered Tyrode medium containing 0.01% (w:v) polyvinyl alcohol (H-TL-PVA). Porcine FF was aspirated from follicles of 2- to 6-mm in diameter by using a 10-ml syringe fitted with an 18-gauge needle, centrifuged at 10 000 x g for 15 min at 4°C to remove cellular debris, and stored at –30°C until use.

Maturation Culture of Oocytes

The basic medium for maturation culture of oocytes was BSA-free North Carolina State University 37 (NCSU37; [6]) supplemented with 0.6 mM cysteine, 2% (v:v) modified Eagle medium (MEM) amino acids solution (Invitrogen, Carlsbad, CA), 1% (v:v) MEM nonessential amino acids solution (Invitrogen), 0.04 units/ml ovine FSH, 0.02 units/ml ovine LH, and 1 mM hypoxanthine. After washing in the basic medium, groups of 20 COCs were transferred into 100-µl droplets of culture medium that had been previously equilibrated in a CO2 incubator. The culture medium was supplemented with 10% (v:v) pFF, 10% pFF and 100 µM DETC (pFF + DETC), or 10% (v:v) FBS. To precisely clarify the cell damage caused by oxidative stress, some portions of the COCs were cultured for the entire period in the medium containing 1 mU/ml XOD to expose them to ROS during IVM. All media containing oocytes were covered with mineral oil and cultured at 39°C in an atmosphere of 5% CO2 in air. After 44 h of maturation culture, the COCs were treated to strip their cumulus cells by pipetting through a narrow-bore pipette in H-TL-PVA containing 0.1% (w:v) hyaluronidase.

Determination of SOD Activity

To determine SOD activity, a novel tetrazolium salt, 4-[3-(4-iodophenyl)-2-(4-nitrophenyl)-2H-5-tetrazolio]-1,3-benzene disulfonate sodium salt (WST-1; Dojindo Laboratories, Kumamoto, Japan), which forms a highly water-soluble formazan after reduction, was used as described previously [32] with some modifications. The reaction mixture consisted of 700 µl of reaction buffer (50 mM sodium carbonate buffer containing 0.15 mM ethylenediamine tetraacetic acid [EDTA] and 0.15 mM hypoxanthine pH 9.4), 100 µl of sample solution, 100 µl of inhibitor solution, and 50 µl of 1 mM WST-1 solution. The reaction was initiated by adding 50 µl of 200 mU/ml XOD solution and the absorbance was monitored continuously at 438 nm with a spectrophotometer (JASCO Co., Tokyo, Japan) for 3 min, with readings recorded every 0.5 min. The SOD activity of each sample was determined based on the standard curve of CuZn-SOD (S-2515; Sigma) with a defined specific activity. The inhibition of SOD activity was tested by using KCN and DETC as specific CuZn-SOD inhibitors [33].

Analysis of DNA Damage

To precisely clarify the DNA damage in oocytes caused by oxidative stress, the COCs were cultured in the absence or presence of 1 mU/ml XOD during IVM. These COCs as well as oocytes freshly isolated from their follicles (freshly isolated oocytes) were transferred into 0.1% (w:v) protease solution in H-TL-PVA at room temperature to remove the zona pellucida, and washed quickly in PBS (Invitrogen) containing 0.3% (w:v) BSA. DNA damage in each oocyte was detected by the single-cell microgel electrophoresis (COMET) assay reported by Singh et al. [34] and Tatemoto et al. [12] with the following modifications. Fifteen to 20 zona-free oocytes were mixed with 10-µl drops of 2% (w:v) low-melting agarose (SeaPlaque GTG agarose; FMC BioProducts, Rockland, ME) at 39°C on a glass slide, and the cell suspension was immediately covered with a coverslip. The space between the coverslip and the glass slide was filled with 2% low-melting agarose, and the slides were then kept at 4°C for 20 min to allow solidification of the agarose. After gently removing the coverslip, the slides stuck with the oocyte-embedded agarose were immersed in a lysing solution (1% N-lauroyl-sarcosine, 2.5 M NaCl, 20 mM EDTA, 10 mM Tris pH 10, 20 µg/ml proteinase K, and 1% Triton X-100) for 1 h to lyse the cells and permit DNA unfolding. The slides were then placed on a horizontal gel electrophoresis unit and equilibrated for 20 min in tris-borate-EDTA (TBE) electrophoresis buffer. Electrophoresis was conducted for 20 min at 50 V. After electrophoresis, the slides were stained with 10 µg/ml of bisbenzimide Hoechst 33342 for 10 min, washed with distilled water, and then covered with a coverslip. The slides were sealed with clear nail polish and examined using a fluorescent microscope. The length of the streak of the DNA COMET tail between the edge of the plasma membrane and the end of the visible COMET tail was individually measured in 40–45 oocytes of each experimental group by using a micrometer.

To detect DNA damage of cumulus cells, briefly, 1.0 x 106 cumulus cells were separated from COCs and centrifuged at 750 x g for 10 min. Cell pellets were resuspended in 200 µl of TE buffer (50 mM Tris-HCl pH 7.5, and 20 mM EDTA) containing 1.0% Triton X-100, and allowed to stand for 10 min at 4°C with gentle pipetting. After centrifugation at 2000 x g for 10 min, the supernatant was transferred to a new tube and treated with 2.5 µl of 2mg/ml ribonuclease A solution. After 60 min of incubation at 37°C, 5 µl of 20 mg/ml proteinase K solution was added and the extract was incubated for an additional 45 min at 50°C. After incubation, 250 µl of 2-propanol and 50 µl of 5 M NaCl solution were added, and the tube was left overnight at –30°C. The sample was centrifuged at 12 000 x g for 10 min (4°C). After the supernatant was completely removed, 7 µl of TBE electrophoresis buffer, 2 µl of 5x loading buffer (40% sucrose and 0.25% bromophenol blue), and 1 µl of 0.1 mg/ ml ethidium bromide were added. DNA was analyzed by horizontal 2% agarose gel electrophoresis. The intensity of DNA ladders was quantified by using an Image PC (Scion Co., Frederick, MD).

Assay of Intracellular GSH Content

COCs treated with or without XOD, in addition to freshly isolated oocytes, were assayed for intracellular GSH content. Denuded oocytes were washed three times with the stock buffer (0.2 M sodium phosphate buffer pH 7.2 containing 10 mM EDTA), and groups of 40 oocytes in 5 µl of the stock buffer were transferred to 1.5-ml microtubes, and 5 µl of 1.25 M H3PO4 was added. Samples were stored at –80°C until assay. The concentration of intracellular GSH in oocytes was determined using the 5,5-dithio-bis(2-nitrobenzoic acid)-glutathione disulfide (DTNB-GSSG) reductase recycling assay as described previously [12], which detects both GSH and GSSG in the oocyte.

In Vitro Fertilization

Denuded oocytes were washed three times with modified Tris-buffered medium (mTBM [1]), designated as IVF medium supplemented with 2 mM caffeine sodium benzoate and 0.1% (w:v) BSA. After washing, 25– 30 oocytes were transferred to 50-µl droplets of IVF medium that had been covered with warm mineral oil. The droplets containing oocytes were kept in an incubator for 30 to 45 min until spermatozoa were added for fertilization. For sperm preparation, frozen-ejaculated boar spermatozoa were thawed (39°C) and washed two times by centrifugation at 400 x g for 4 min in PBS supplemented with 0.1% (w:v) polyvinyl alcohol at pH 7.2. At the end of the washing procedure, the sperm pellets were resuspended at 4 x 108 cells/ml in mTBM supplemented with 2 mM caffeine sodium benzoate, 0.1% (w:v) BSA, and 0.5% (v:v) pFF, and were then incubated for 90 min at 39°C. After sperm preincubation, 50 µl of diluted sperm suspension in IVF medium was added to a droplet containing oocytes for a final sperm concentration of 1 x 106 cells/ml. Oocytes were coincubated with spermatozoa for 7 h at 39°C in an atmosphere of 5% CO2 in air.

Embryo Culture

After insemination, oocytes were removed from fertilization drops, washed three times, and cultured in 50 µl of NCSU37 medium at 39°C in an atmosphere of 5% CO2 in air. At 48 and 168 h after IVF, cleavage and blastocyst formation, respectively, were evaluated under a stereomicroscope. The percentages of cleavage and development to the blastocyst stage were determined from the number of oocytes that were placed into the maturation media.

Assessment of Meiotic Maturation and Fertilization Parameters

After IVM culture or 10 h after IVF, groups of 30–40 oocytes were mounted, fixed in acetic acid-ethanol (1:3, v:v) for 72 h, stained with 1% (w:v) lacmoid in 45% (v:v) acetic acid, and examined for nuclear maturation or fertilization parameters, respectively, under a phase-contrast microscope at 400x magnification. Germinal vesicle breakdown (GVBD), metaphase I (MI), maturation to metaphase II (MII), sperm penetration, polyspermy, and MPN formation were assessed. Oocytes were considered to be penetrated by spermatozoa at the MII stage when they had two polar bodies, one or more swollen sperm heads, MPN (or a combination of these), and the corresponding sperm tail.

Statistical Analysis

Values are presented as the mean of four independent experimental replicates. Variation between experiments is illustrated using SEM. For evaluation of the differences between groups, data on the percentage of meiotic maturation, fertilization parameters, and embryo development were checked for homogeneity, pooled, and then subjected to contingency table analysis followed by a Tukey test for nonparametric multiple comparisons [35]. Statistical analyses of data on SOD activity, migration of DNA, GSH content, and mean number of spermatozoa per penetrated oocyte were carried out by a Shapiro-Wilk normality test and analysis of variance followed by a Tukey-Kramer test. All analyses were carried out using the Statistical Analysis System R software package (Cary, NC). A probability of P < 0.05 was considered statistically significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Comparison of SOD Activity Between pFF and FBS

In pFF, 16.0 ± 0.6 U/ml of SOD activity was detected and was about 7.2-fold higher than that of FBS (2.2 ± 0.1 U/ml; Fig. 1). However, SOD activity in pFF and FBS was remarkably decreased to 0.4 ± 0.1 U/ml and 0.3 ± 0.1 U/ ml, respectively, by the addition of 1 mM KCN, indicating that a higher level of radical scavenging activity in pFF is mainly addressed by CuZn-SOD isoenzymes.



View larger version (13K):
[in this window]
[in a new window]
  		FIG. 1. SOD activities of pFF and FBS in the presence or absence of 1 mM KCN. Values are expressed as the mean ± SEM. Values with different superscripts are significantly different (P < 0.05)

As shown in Figure 2, SOD activity in the IVM medium supplemented with 10% pFF was inhibited by the addition of DETC in a concentration-dependent manner. The addition of 100 µM DETC to the IVM medium reduced SOD activity to 0.19 ± 0.02 U/ml, which was similar to that of the IVM medium supplemented with 10% FBS (0.20 ± 0.05 U/ml).



View larger version (15K):
[in this window]
[in a new window]
  		FIG. 2. Inhibitory effects of DETC on SOD activity in maturation medium supplemented with 10% pFF. Values are expressed as the mean ± SEM. Values with different superscripts are significantly different (P < 0.05)

Nuclear Maturation

In the absence of XOD, there was no significant difference in the percentages of oocytes undergoing GVBD and reaching the MII stage among COCs cultured with pFF, pFF + DETC, or FBS (Table 1). The maturation rate at the MII stage of COCs cultured with pFF slightly decreased to 65% by treatment with XOD compared with that of untreated oocytes (78%). However, the harmful effect of XOD on meiotic maturation was more markedly observed in COCs cultured with pFF + DETC and FBS, and the maturation rates of these oocytes treated with XOD was significantly suppressed to 48% and 33%, respectively, compared with that of oocytes treated without XOD (69% and 75%, respectively) (P < 0.05). Nonetheless, the percentages of GVBD and degeneration in these oocytes did not differ significantly compared with those of oocytes cultured with pFF. It is interesting that treatment with XOD in oocytes cultured with pFF + DETC and FBS inhibited emission of the first polar body and significantly blocked meiotic progression at the MI stage (30% and 39%, respectively), compared with that of untreated oocytes (12% and 10%, respectively) (P < 0.05).


View this table:
[in this window]
[in a new window]
  		TABLE 1. Effects of oxidative stress caused by the hypoxanthine-XOD system on the meiotic progression of porcine oocytes cultured for 44 h in media with supplements

DNA Damage in Oocytes and Cumulus Cells

To confirm the effect of SOD activity in pFF on DNA damage caused by oxidative stress, the extent of DNA migration of a single oocyte was measured by the COMET assay following treatment with or without XOD for 44 h. As shown in Figure 3A, the length of DNA migration of oocytes cultured with pFF was not significantly affected by the absence or presence of XOD (74.4 ± 4.4 µm or 96.5 ± 9.0 µm, respectively), compared with that of freshly isolated oocytes (73.4 ± 4.7 µm). In contrast, the addition of XOD during IVM of oocytes cultured with pFF + DETC and FBS caused irreversible damage to oocyte DNA, and the length of DNA migration in these oocytes increased remarkably to 161.0 ± 7.7 µm and 152.8 ± 8.3 µm, respectively, compared with that of oocytes cultured with pFF (P < 0.05).



View larger version (35K):
[in this window]
[in a new window]
  		FIG. 3. Effects of oxidative stress caused by the hypoxanthine-XOD system on the length of DNA migration (A) and the concentration of intracellular GSH (B) in porcine oocytes cultured for 44 h in media supplemented with pFF, pFF + DETC, and FBS. Values are expressed as the mean ± SEM. Values with different superscripts are significantly different (P < 0.05)

Inhibitory effects of SOD activity in pFF on DNA damage caused by ROS was also found in cumulus cells. The degree of DNA damage in cumulus cells removed from COCs was analyzed by agarose gel electrophoresis. DNA laddering was almost not detectable in cumulus cells at the start of IVM (Fig. 4, lane 1), whereas it was clearly visible after 44 h of IVM culture, irrespective of the culture condition (Fig. 4, lanes 2–4). In particular, upon exposure to ROS generated by XOD, the intensities of the DNA ladders in cumulus cells collected from COCs cultured with pFF + DETC and FBS were 1.5-fold to 1.7-fold higher than those in COCs treated without XOD (Fig. 4, lanes 6 and 7). However, the obvious increase in intensity (1.1-fold) was not observed in cumulus cells cultured with pFF despite treatment with XOD (Fig. 4, lane 5).



View larger version (80K):
[in this window]
[in a new window]
  		FIG. 4. DNA fragmentation in cumulus cells cultured for 44 h in the absence (lanes 2–4) or presence (lanes 5–7) of ROS generated by the hypoxanthine-XOD system. Cumulus cells were separated from COCs freshly isolated from their follicles (lane 1), cultured in media supplemented with pFF (lanes 2 and 5), pFF + DETC (lanes 3 and 6), and FBS (lanes 4 and 7). M, 100-base pair DNA marker

Intracellular GSH Content in Oocytes

As shown in Figure 3B, GSH contents were significantly higher in oocytes after maturation culture with pFF, pFF + DETC, and FBS (9.6 ± 0.4 pmol/oocyte, 7.6 ± 0.4 pmol/ oocyte, and 8.3 ± 0.7 pmol/oocyte, respectively) without the addition of XOD, compared with that of freshly isolated oocytes (5.0 ± 0.3 pmol/oocyte), and a significantly lower increasing level of GSH was found in oocytes cultured with pFF + DETC (P < 0.05). Although increasing GSH content with the advancement of oocyte maturation was not interrupted by oxidative stress in oocytes cultured with pFF (7.3 ± 0.3 pmol/oocyte), treatment with XOD significantly inhibited the increase of intracellular GSH levels in oocytes cultured with pFF + DETC and FBS (5.2 ± 0.4 pmol/ oocyte and 5.1 ± 0.4 pmol/oocyte, respectively) (P < 0.05).

IVF and Embryo Development

After in vitro insemination of oocytes matured in media containing pFF, pFF + DETC, and FBS, there was no difference in the rates of penetration (71–79%; Table 2). In the absence of XOD during IVM culture, the proportion of MPN formation in oocytes matured with pFF (70%) was significantly higher than that of oocytes matured with FBS (50%) (P < 0.05). However, the positive effect of pFF on MPN formation was abated by the addition of DETC, and this rate was reduced to 47%, or to the same extent as oocytes matured with FBS. In the presence of XOD, the incidence of polyspermic fertilization in oocytes matured with pFF (61%) was significantly lower than that of oocytes matured with pFF + DETC and FBS (84% and 80%, respectively); consequently, the mean number of spermatozoa per oocyte was significantly lower in oocytes matured with pFF (P < 0.05). Furthermore, the proportions of MPN formation in oocytes matured with pFF + DETC and FBS were significantly reduced to 16% and 16%, respectively, compared with that of oocytes cultured with pFF (44%) (P < 0.05).


View this table:
[in this window]
[in a new window]
  		TABLE 2. Effects of added supplements to the maturation medium in the absence or presence of XOD on the fertilization parameters of porcine oocytes

As shown in Figure 5, the cleavage rate after 48 h of in vitro insemination was significantly decreased to 43% in oocytes matured with FBS in the presence of XOD compared with that in the absence of XOD (61%) (P < 0.05). In the absence of XOD during IVM, significantly lower proportions of oocytes matured in the medium supplemented with pFF + DETC developed to the blastocyst stage (7%) compared with that of oocytes cultured with pFF and FBS (18% and 13%, respectively) (P < 0.05). Although the addition of XOD during IVM of oocytes cultured with pFF caused a slight decrease in the potential for development to the blastocyst stage (9%), the developmental ability of oocytes matured with pFF + DETC and FBS were drastically abolished by treatment with XOD (0% and 1%, respectively).



View larger version (30K):
[in this window]
[in a new window]
  		FIG. 5. Cleavage and blastocyst formation rates after IVF of porcine oocytes matured in media supplemented with pFF, pFF + DETC, and FBS in the absence or presence of XOD. Findings are expressed as the mean ± SEM. Approximately 170 oocytes were examined in each group. Within the same category, values with different superscripts are significantly different (P < 0.05)


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The present study assessed the effects of SOD activity in pFF added to maturation medium in porcine oocytes on the protection of cell damage caused by ROS and the improvement of MPN formation and developmental competence after IVF. The main finding was that pFF during IVM culture efficiently protected oocytes against oxidative stress through its radical scavenging activity elicited from one or more SOD isoenzymes. The poor developmental competence of porcine oocytes matured in vitro might reflect deficient cytoplasmic maturation of oocytes, and many studies have indicated that the addition of pFF to maturation media can improve cytoplasmic maturation [17–20]. Although the desirable effects of pFF on the cytoplasmic maturation responsible for the subsequent developmental competence is affirmed in these reports, there are still unclear points with respect to factor identification. In the present study, pFF had about 7.2-fold higher levels of superoxide scavenging activity than that of FBS, and this activity was overridden by treatment with KCN and the CuZn-SOD inhibitor, DETC (Figs. 1 and 2), indicating that pFF possesses a higher level of the CuZn-SOD isoenzyme compared with FBS. Similarly, the presence of CuZn-SOD isoenzymes and a higher level of SOD activity were also detected in human FF. It is possible to speculate that there may be a secretion or concentration mechanism of CuZn-SOD isoenzymes into the FF during follicular maturation [26, 27].

The dismutation of O2– to H2O2 is a primary step in the antioxidative pathway during both normal cellular metabolism and various pathological processes [36, 37]. Different molecules take part in this process, with the major part played by a family of three SOD isoenzymes. The two intracellular isoenzymes, CuZn-SOD [38] and Mn-SOD [39], are both synthesized by virtually all cell types in the body. The secretory extracellular SOD (EC-SOD) is the major isoenzyme in extracellular fluids, such as plasma, lymph, and synovial fluid, but has also been demonstrated in all mammalian species [40, 41]. EC-SOD is evolutionarily related to CuZn-SOD, because the middle protein of the EC-SOD sequence shows strong homology with the part of the CuZn-SOD sequence that defines the active site [42]. The present study could not identify which isoenzyme of SOD (CuZn-SOD or EC-SOD) participates in a higher level of radical scavenging activity in pFF. However, Tilly and Tilly [43] reported that gonadotropin-mediated rat follicular development coincided with enhanced expression of EC-SOD and Mn-SOD messenger RNA, but not CuZn-SOD messenger RNA, in granulosa cells. Therefore, EC-SOD may be responsible for radical scavenging activity in pFF.

Several studies of mouse embryos indicate that the two-cell block occurring in vitro could be alleviated by protection against oxidative stress mediated by a potent scavenger of O2–, CuZn-SOD [44–46]. Matsuoka et al. [47] reported that when CuZn-SOD and catalase were added to a serum-free culture medium of mouse oocytes, fertilization and cleavage rates were improved. In bovine oocytes, the addition of CuZn-SOD during IVM apparently improved the developmental potential to the blastocyst stage after IVF [48]. In contrast, when bovine oocytes were matured in vitro in the presence of CuZn-SOD, the developmental competence of the oocytes after IVF was not significantly improved [49]. It seems that these conflicting data may be attributed to different antioxidative defense mechanisms between CuZn-SOD and EC-SOD. CuZn-SOD and Mn-SOD are found primarily in the cytosol and the mitochondria, respectively, of eukaryotes. On the other hand, EC-SOD, an extracellular form of CuZn-SOD, is localized predominantly in the extracellular matrix of tissues, as well as in the extracellular fluids [50]. It has been reported that EC-SOD is an important antioxidant molecule in the testis and is under germ cell regulation [51]. Taken together, further studies are required to investigate the differences in the functional mechanisms between CuZn-SOD and EC-SOD in regulating and supporting the developmental competence of oocytes during maturation.

We previously demonstrated that oxidative stress during IVM culture provoked porcine oocytes into undergoing apoptotic cell death, judging from the findings that DNA cleavage by TUNEL analysis, the increased length of DNA migration by COMET assay, and the activation of caspase-3 were prominently detected in degenerated oocytes by treatment with hypoxanthine and XOD [12, 13, 52]. In the present study, the maturation rate was slightly reduced under ROS conditions generated by the hypoxanthine-XOD system (Table 1), but the addition of pFF to the IVM medium efficiently precluded an increase in the length of DNA migration in oocytes and the intensity of DNA ladders in cumulus cells (Figs. 3A and 4). However, this fully protective potency against DNA damage caused by ROS was not found in oocytes cultured with pFF + DETC, nor in FBS, in accordance with their lower levels of SOD activity; consequently, a large number of those oocytes blocked the meiotic progression at the MI stage compared with oocytes cultured with pFF. Thus, it is likely that the addition of pFF to the IVM medium may function to protect porcine oocytes and cumulus cells from cell damage caused by oxidative stress.

It is interesting that the apoptotic DNA fragments in cumulus cells occurred during IVM culture, irrespective of the addition of pFF and treatment with XOD. Very recently, this spontaneous apoptosis in cumulus cells during IVM was also observed in bovine COCs, suggesting that the degree of apoptosis in cumulus cells may be correlated with the developmental competence of oocytes after IVF [53]. In humans, a low degree of apoptosis in cumulus cells has been an indicator of a higher developmental competence of oocytes after IVF or an intracytoplasmic sperm injection program [54, 55]. We previously reported that cumulus cells play a critical role in protecting oocytes against oxidative stress-induced apoptosis by enhancing the intracellular GSH content in oocytes [12]. In the present study, an increased intensity of DNA ladders in cumulus cells was detected in COCs cultured with pFF + DETC and with FBS in the presence of XOD. Furthermore, the developmental competence to the blastocyst stage was hard to observe in these oocytes, based on results of significantly lower levels of intracellular GSH, and lower abilities to promote MPN formation, and to prevent the frequency of polyspermic fertilization than oocytes cultured with pFF (P < 0.05) (Figs. 3B and 5, and Table 2). It has been found that intracellular GSH plays an important role in protecting the cell from oxidative damage [56], and is a molecular marker to predict cytoplasmic maturation in porcine oocytes [2, 9, 57, 58]. To our knowledge, the present study is the first that directly shows the involvement of pFF in protection of porcine oocytes against cell damage caused by ROS during IVM through its radical scavenging activity of SOD isoenzymes, resulting in supporting cytoplasmic maturation.

With regard to the effect of SOD activity in pFF on the cytoplasmic maturation, the present study demonstrated that the higher proportions of MPN formation and development to the blastocyst stage were recorded for oocytes matured with pFF compared with oocytes matured with FBS, whereas the abilities of porcine oocytes to promote transformation of sperm nuclei into MPN and the subsequent embryo development were strongly reduced by the further addition of DETC (Fig. 5). It was also reported that the addition of pFF collected from medium-size (5–7 mm in diameter) follicles to porcine IVM medium can improve meiotic maturation, reduce polyspermy, and increase normal fertilization rates in vitro [20]. In humans, oocytes successfully fertilized in vitro corresponded closely to a higher level of antioxidant capacity in FF obtained at the same time of follicle aspiration [28]. Although it has been reported that besides SOD isoenzymes, pFF possesses various factors associated with enhancement of cytoplasmic maturation in comparison with FBS [15, 21–25], the reason why MPN formation ability is compromised in oocytes cultured with pFF + DETC despite the absence of XOD is still a matter of discussion. The inhibition of SOD activity by 2-methoxyoestradiol causes accumulation of cellular O2–, and leads to free-radical-mediated damage to mitochondrial membranes, the release of cytochrome c from mitochondria, and apoptosis of human leukemia cells [59]. It is thus speculated that inhibition of SOD activity during IVM may give rise to the accumulation of cellular O2–, resulting in disturbances of physiological processes that are essential for cytoplasmic maturation, because the intracellular GSH content in oocytes cultured with pFF (9.6 ± 0.5 pmol/oocyte) significantly decreased to 7.6 ± 0.4 pmol/oocyte with further addition of DETC (P < 0.05) (Fig. 3B).

From these findings, it becomes clear that the protection of porcine oocytes against oxidative stress during IVM results in an improvement in the potential for acquiring the cytoplasmic maturation responsible for developmental competence after IVF, and it seems that SOD activity in pFF is closely associated with the defense mechanisms from the harmful effect of ROS generated by critical cellular processes, such as oxidative respiration, in oocytes matured not only in vitro but also in vivo.

In summary, the data reported here demonstrate that a higher level of SOD activity in pFF efficiently blocked the decreasing intracellular GSH content and DNA damage caused by oxidative stress in porcine oocytes and cumulus cells, resulting in sustenance of the ability to undergo MPN formation and development to the blastocyst stage after in vitro insemination; however, these abilities were interrupted by the SOD inhibitor. Therefore, it is suggested that the radical scavenging activity of SOD isoenzymes, which is one of the desirable effects of pFF, can support cytoplasmic maturation relating to developmental competence postfertilization by alleviating oxidative stress during oocyte maturation.


   ACKNOWLEDGMENTS
 
We are grateful to the staff of the Meat Inspection Office of the City of Oosato, Japan, for supplying the porcine ovaries.


   FOOTNOTES
 
1 This study was partially supported by a Grant-in-Aid for Young Scientists (B) 14760181 to H.T. from the Ministry of Education, Culture, Sports, Science, and Technology of Japan. Back

2 Correspondence. FAX: 81 98 895 8757; hidettmt{at}agr.u-ryukyu.ac.jp Back

Received: 5 March 2004.

First decision: 30 March 2004.

Accepted: 27 May 2004.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Abeydeera LR, Day BN. Fertilization and subsequent development in vitro of pig oocytes inseminated in a modified tris-buffered medium with frozen-thawed ejaculated spermatozoa. Biol Reprod 1997 57:729-734[Abstract]
  2. Wang WH, Abeydeera LR, Cantley TC, Day BN. Effects of oocyte maturation media on development of pig embryos produced by in vitro fertilization. J Reprod Fertil 1997 111:101-108
  3. Kano K, Miyano T, Kato S. Effects of glycosaminoglycans on the development of in vitro-matured and -fertilized porcine oocytes to the blastocyst stage in vitro. Biol Reprod 1998 58:1226-1232[Abstract/Free Full Text]
  4. Marchal R, Caillaud M, Martoriati A, Gerard N, Mermillod P, Goudet G. Effect of growth hormone (GH) on in vitro nuclear and cytoplasmic oocyte maturation, cumulus expansion, hyaluronan synthases, and connexins 32 and 43 expression, and GH receptor messenger RNA expression in equine and porcine species. Biol Reprod 2003 69:1013-1022[Abstract/Free Full Text]
  5. Beckmann LS, Day BN. Effects of media NaCl concentration and osmolarity on the culture of early-stage pig embryos and the viability of embryos cultured in a selected superior medium. Theriogenology 1993 39:611-622
  6. Petters RM, Wells KD. Culture of pig embryos. J Reprod Fertil 1993 48:suppl61-73
  7. Dobrinsky JR, Johnson LA, Rath D. Development of a culture medium (BECM3) for pig embryos: effects of bovine serum albumin and fetal bovine serum on embryo development. Biol Reprod 1996 55:1069-1074[Abstract]
  8. Ka H-H, Sawai K, Wang W-H, Im K-S, Niwa K. Amino acids in maturation medium and presence of cumulus cells at fertilization promote male pronuclear formation in porcine oocytes matured and penetrated in vitro. Biol Reprod 1997 57:1478-1483[Abstract]
  9. Yamauchi N, Nagai T. Male pronuclear formation in denuded porcine oocytes after in vitro maturation in the presence of cysteamine. Biol Reprod 1999 61:823-833[Abstract/Free Full Text]
  10. Marchal R, Tomanek M, Terqui M, Mermillod P. Effects of cell cycle dependent kinases inhibitor on nuclear and cytoplasmic maturation of porcine oocytes. Mol Reprod Dev 2001 60:65-73[CrossRef][Medline]
  11. Niwa K. Effectiveness of in vitro maturation and in vitro fertilization techniques in pig. J Reprod Fertil 1993 48:suppl49-59
  12. Tatemoto H, Sakurai N, Muto N. Protection of porcine oocytes against apoptotic cell death caused by oxidative stress during in vitro maturation: role of cumulus cells. Biol Reprod 2000 63:805-810[Abstract/Free Full Text]
  13. Tatemoto H, Ootaki K, Shigeta K, Muto N. Enhancement of developmental competence after in vitro fertilization of porcine oocytes by treatment with ascorbic acid 2-O-{alpha}-glucoside during in vitro maturation. Biol Reprod 2001 65:1800-1806[Abstract/Free Full Text]
  14. Elmileik AMA, Maeda T, Terada T. Higher rates of development into blastocyst following the in vitro fertilization of bovine oocytes matured in a medium supplemented with the fluid from large bovine follicles. Anim Reprod Sci 1995 38:85-96[CrossRef]
  15. Khatir H, Carolan C, Lonergan P, Mermillod P. Characterization of calf follicular fluid and its ability to support cytoplasmic maturation of cow and calf oocytes. J Reprod Fertil 1997 111:267-275
  16. Ikeda S, Azuma T, Hashimoto S, Yamada M. In vitro maturation of bovine oocytes with fractions of bovine follicular fluid separated by heparin affinity chromatography. J Reprod Dev 1999 45:397-404[CrossRef]
  17. Naito K, Fukuda Y, Toyoda Y. Effects of porcine follicular fluid on male pronucleus formation in porcine oocytes maturation in vitro. Gamete Res 1988 21:289-295[CrossRef][Medline]
  18. Yoshida M, Ishizaki Y, Kawagishi H, Bamba K, Kojima Y. Effects of pig follicular fluid on maturation of pig oocytes in vitro and on their subsequent fertilizing and developmental capacity in vitro. J Reprod Fertil 1992 95:481-488
  19. Rath D, Niemann H, Tao T. In vitro maturation of porcine oocytes in follicular fluid with subsequent effects on fertilization and embryo yield in vitro. Theriogenology 1995 44:529-538
  20. Vatzias G, Hagen DR. Effects of porcine follicular fluid and oviduct-conditioned media on maturation and fertilization of porcine oocytes in vitro. Biol Reprod 1999 60:42-48[Abstract/Free Full Text]
  21. Spicer LJ, Echternkamp SE, Canning SF, Hammond JM. Relationship between concentrations of immunoreactive insulin-like growth factor-I in follicular fluid and various biochemical markers of differentiation in bovine antral follicles. Biol Reprod 1988 39:573-580[Abstract]
  22. Hofmann GE, Scott RT, Brzyski RG, Jones HW. Immunoreactive epidermal growth factor concentrations in follicular fluid obtained from in vitro fertilization. Fertil Steril 1990 54:303-307[Medline]
  23. Knight PG, Muttukrishna S, Groome NP. Development and application of a two-site enzyme immunoassay for the determination of ‘total’ activin-A concentration in serum and follicular fluid. J Endocrinol 1996 148:267-279[Abstract/Free Full Text]
  24. Seli E, Zeyneloglu HB, Senturk LM, Bahtiyar OM, Olive DL, Arici A. Basic fibroblast growth factor: peritoneal and follicular fluid levels and its effect on early embryonic development. Fertil Steril 1998 69:1145-1148[CrossRef][Medline]
  25. Liu R-H, Li Y-H, Jiao L-H, Wang X-N, Wang H, Wang W-H. Extracellular and intracellular factors affecting nuclear and cytoplasmic maturation of porcine oocytes collected from different sizes of follicles. Zygote 2002 10:253-260[CrossRef][Medline]
  26. Shiotani M, Noda Y, Narimoto K, Imai K, Mori T, Fujimoto K, Ogawa K. Immunohistochemical localization of superoxide dismutase in the human ovary. Human Reprod 1991 6:1349-1353[Abstract/Free Full Text]
  27. Sabatini L, Wilson C, Lower A, Al-Shawaf T, Grudzinskas JG. Superoxide dismutase activity in human follicular fluid after controlled ovarian hyperstimulation in women undergoing in vitro fertilization. Fertil Steril 1999 72:1027-1034[CrossRef][Medline]
  28. Oyawoye O, Gadir AA, Garner A, Constantinovici N, Perrett C, Hardiman P. Antioxidants and reactive oxygen species in follicular fluid of women undergoing IVF: relationship to outcome. Human Reprod 2003 18:2270-2274[Abstract/Free Full Text]
  29. Laloraya M, Kumar GP, Laloraya MM. Histochemical study of superoxide dismutase in the ovary of the rat during the oestrous cycle. J Reprod Fertil 1989 86:583-587
  30. Sato EF, Kobuchi H, Edashige K, Takahashi M, Yoshioka T, Utsumi K, Inoue M. Dynamic aspects of ovarian superoxide dismutase isozymes during the ovulatory process in the rat. FEBS Lett 1992 303:121-125[CrossRef][Medline]
  31. Ribarov SR, Benov LC. Relationship between the hemolytic action of heavy metals and lipid peroxidation. Biochem Biophys Acta 1981 640:721-726[Medline]
  32. Ukeda H, Kawana D, Maeda S, Sawamura M. Spectrophotometric assay for superoxide dismutase based on the reduction of highly water-soluble tetrazolium salts by xanthine-xanthine oxidase. Biosci Biotechnol Biochem 1999 63:485-488[CrossRef]
  33. Van Camp W, Bowler C, Villarroel R, Tsang EW, Van Montagu M, Inze D. Characterization of iron superoxide dismutase cDNAs from plants obtained by genetic complementation in Escherichia coli. Proc Natl Acad Sci U S A 1990 87:9903-9907[Abstract/Free Full Text]
  34. Singh NP, McCoy MT, Tice RR, Schneider EL. A simple technique for quantitation of low level of DNA damage in individual cells. Exp Cell Res 1988 175:184-191[CrossRef][Medline]
  35. Ryan TA. Significance tests for multiple comparison of proportions, variances, and other statistics. Psychol Bull 1960 57:318-328[CrossRef][Medline]
  36. Ames BN, Shiganaga MK, Hagen TM. Oxidant, antioxidant and the degenerative diseases of the aging. Proc Natl Acad Sci U S A 1993 90:7915-7922[Abstract/Free Full Text]
  37. Repetto M, Reides C, Gomez Carretero ML, Griemberg MCG, Llesuy S. Oxidative stress in blood of HIV infected patients. Clin Chim Acta 1996 255:107-117[CrossRef][Medline]
  38. McCord JM, Fridovich I. Superoxide dismutase. An enzymic function for erythrocuprein (hemocuprein). J Biol Chem 1969 244:6049-6055[Abstract/Free Full Text]
  39. Weisiger RA, Fridovich I. Mitochondrial superoxide simutase: site of synthesis and intramitochondrial localization. J Biol Chem 1973 248:4793-4796[Abstract/Free Full Text]
  40. Marklund SL. Extracellular superoxide dismutase and other superoxide dismutase isoenzymes in tissues from nine mammalian species. Biochem J 1984 222:649-655[Medline]
  41. Marklund SL. Expression of extracellular superoxide dismutase by human cell lines. Biochem J 1990 266:213-219[Medline]
  42. Hjalmarsson K, Marklund SL, Engstrom A, Edlund T. Isolation and sequence of complementary DNA encoding human extracellular superoxide dismutase. Proc Natl Acad Sci U S A 1987 84:6340-6344[Abstract/Free Full Text]
  43. Tilly JL, Tilly KI. Inhibitors of oxidative stress mimic the ability of follicle-stimulating hormone to suppress apoptosis in cultured rat ovarian follicles. Endocrinology 1995 136:242-252[Abstract]
  44. Noda Y, Matsumoto H, Umaoka Y, Tatsumi K, Kishi J, Mori T. Involvement of superoxide radicals in the mouse two-cell block. Mol Reprod Dev 1991 28:356-360[CrossRef][Medline]
  45. Nonogaki T, Noda Y, Narimoto K, Umaoka Y, Mori T. Effects of superoxide dismutase on mouse in vitro fertilization and embryo culture system. J Assist Reprod Genet 1992 9:274-280[CrossRef][Medline]
  46. Chun YS, Kim JH, Lee HT, Chung KS. Effect of superoxide dismutase on the development of preimplantation mouse embryos. Theriogenology 1994 41:511-520
  47. Matsuoka I, Fujino Y, Ogita S, Inoue M. Impact of erythrocytes on mouse embryonal development in vitro. FEBS Lett 1995 371:297-299[CrossRef][Medline]
  48. Luvoni GC, Keskintepe L, Brackett BG. Improvement in bovine embryo production in vitro by glutathione-containing culture media. Mol Reprod Dev 1996 43:437-443[CrossRef][Medline]
  49. Blondin P, Coenen K, Sirard M-A. The impact of reactive oxygen species on bovine sperm fertilizing ability and oocyte maturation. J Androl 1997 18:454-460[Abstract/Free Full Text]
  50. Oury TD, Day BJ, Crapo JD. Extracellular superoxide dismutase in vessels and airways of humans and baboons. Free Radic Biol Med 1996 20:957-965[CrossRef][Medline]
  51. Mruk DD, Silvestrini B, Mo M-Y, Cheng CY. Antioxidant superoxide dismutase—a review: its function, regulation in the testis, and role in male fertility. Contraception 2002 65:305-311[CrossRef][Medline]
  52. Tatemoto H, Okuda T, Sogo N, Muto N. Male pronuclear formation and blastocyst formation are improved by supplementation of ascorbic acid 2-O-{alpha}-glucoside during in vitro maturation cultured of denuded porcine oocytes. J Reprod Dev 2001 47:329-339[CrossRef]
  53. Ikeda S, Imai H, Yamada M. Apoptosis in cumulus cells during in vitro maturation of bovine cumulus-enclosed oocytes. Reproduction 2003 125:369-376[Abstract]
  54. Nakahara K, Sakai N, Saito H, Tezuka N, Saito T, Hiroi M, Ito M, Watanabe H, Ohta N. Incidence of apoptotic bodies in membrana granulosa of the patients participating in an in vitro fertilization program. Fertil Steril 1997 67:302-308[CrossRef][Medline]
  55. Host E, Mikkelsen AL, Lindenberg S, Smidt-Jensen S. Apoptosis in human cumulus cells in relation to maturation stage and cleavage of the corresponding oocyte. Acta Obstet Gynecol Scand 2000 79:936-940[CrossRef][Medline]
  56. Meister A. Selective modification of glutathione metabolism. Science 1983 220:472-477[Abstract/Free Full Text]
  57. Abeydeera LR, Wang WH, Cantley TC, Prather RS, Day BN. Presence of ß-mercaptoethanol can increase the glutathione content of pig oocytes matured in vitro and the rate of blastocyst development after in vitro fertilization. Theriogenology 1998 50:747-756[CrossRef][Medline]
  58. Mori T, Amano T, Shimizu H. Roles of gap junctional communication of cumulus cells in cytoplasmic maturation of porcine oocytes cultured in vitro. Biol Reprod 2000 62:913-919[Abstract/Free Full Text]
  59. Huang P, Feng L, Oldham EA, Keating MJ, Plunkett W. Superoxide dismutase as a target for the selective killing of cancer cells. Nature 2000 407:390-395[CrossRef][Medline]



This article has been cited by other articles:


Home page
 Exp. Biol. Med.Home page
A. M. Benoit, H. A. LaVoie, G. L. McCoy, and C. A. Blake
Expression of Sperm Protein 22 (SP22) in the Rat Ovary During Different Reproductive States
Experimental Biology and Medicine, July 1, 2007; 232(7): 910 - 920.
[Abstract] [Full Text] [PDF]

Home page
 ReproductionHome page
D. Nestler, M. Risch, B. Fischer, and P. Pocar
Regulation of aryl hydrocarbon receptor activity in porcine cumulus-oocyte complexes in physiological and toxicological conditions: the role of follicular fluid
Reproduction, May 1, 2007; 133(5): 887 - 897.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
71/4/1150    most recent
biolreprod.104.029264v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Tatemoto, H.
Right arrow Articles by Nakada, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Tatemoto, H.
Right arrow Articles by Nakada, T.
Agricola
Right arrow Articles by Tatemoto, H.
Right arrow Articles by Nakada, T.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS