Biol Reprod Track the topics, authors and articles important to you
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
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 My Folders
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 Mori, T.
Right arrow Articles by Shimizu, H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Mori, T.
Right arrow Articles by Shimizu, H.
Agricola
Right arrow Articles by Mori, T.
Right arrow Articles by Shimizu, H.
Biology of Reproduction 62, 913-919 (2000)
© 2000 Society for the Study of Reproduction, Inc.

Articles

Roles of Gap Junctional Communication of Cumulus Cells in Cytoplasmic Maturation of Porcine Oocytes Cultured In Vitro

Tadashi Mori1,a, Tomoko Amano2,a, and Hiroshi Shimizua

a Animal Breeding and Reproduction, Division of Bioresources and Product Science, Graduate School of Agriculture, Hokkaido University, Sapporo 060-8589, Japan


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cumulus cells of the oocyte play important roles in in vitro maturation and subsequent development. One of the routes by which the factors are transmitted from cumulus cells to the oocyte is gap junctional communication (GJC). The function of cumulus cells in in vitro maturation of porcine oocytes was investigated by using a gap junction inhibitor, heptanol. Cumulus-oocyte complexes (COCs) were collected from the ovaries of slaughtered gilts by aspiration. After selection of COCs with intact cumulus cell layers and uniform cytoplasm, they were cultured in a medium with 0, 1, 5, or 10 mM of heptanol for 48 h. After culture in vitro, one group of oocytes was assessed for nuclear maturation and glutathione (GSH) content, and another group was assigned to in vitro fertilization and assessed for the penetrability of oocytes and the degree of progression to male pronuclei (MPN) of penetrated spermatozoa.

At the end of in vitro maturation, the oocytes reached metaphase II at a high rate (about 80%) regardless of the presence of heptanol at various concentrations. Cumulus cell expansion and the morphology of oocytes cultured in the medium with heptanol were similar to those of control COCs matured without heptanol. The amount of GSH in cultured oocytes tended to decrease as the concentration of heptanol in the medium was increased. Although there was no difference in the rates of penetrated oocytes cultured in media with different concentrations of heptanol, the proportion of oocytes forming MPN after insemination decreased significantly (P < 0.01) at all concentrations tested. A higher rate of sperm (P < 0.01) failed to degrade their nuclear envelopes after penetration into the oocytes that were treated with heptanol.

GJC between the oocyte and cumulus cells might play an important role in regulating the cytoplasmic factor(s) responsible for the removal of sperm nuclear envelopes as well as GSH inflow from cumulus cells.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cumulus cells play important roles in in vitro maturation and subsequent development of the oocyte. The ability of the oocyte to form male pronuclei (MPN) after sperm penetration strongly depends on the presence of cumulus cells during maturation [1–5] and fertilization [6, 7]. One of the routes by which the cumulus cells transmit the factors to the oocyte is gap junctional communication (GJC) [8, 9]. Low molecular substrates (<1000 Mr) such as ions, nucleotides, and amino acids can also traverse from cumulus cells to the oocyte via GJC [10, 11].

A significant relationship between MPN formation and glutathione (GSH) content in the oocyte has been reported [12, 13]. GSH is synthesized from glutamine and cysteine in the cell. Since direct supplementation of GSH to a maturation medium can not promote MPN formation [14], it has been thought that exogenous substances of GSH are essential for the synthesis of GSH in the oocyte [14, 15]. Since the GSH content of an oocyte is highly correlated with the presence of cumulus cells [15–17], GJC is thought to contribute to the increase in GSH content in the oocyte through transportation of GSH (Mr 307.33) or its substrates, cysteine (Mr 121.16) and glutamine (Mr 146.15).

Cumulus cells have been removed to void their effect on the maturation of an oocyte. However, the removal of cumulus cells during maturation induces a premature migration of cortical granules, leading to an increase in exocytotic events and a decrease in penetrability compared with those in cumulus-enclosed oocytes [18]. Also, the presence of cumulus cells affects the penetrability of spermatozoa because the expanded cumulus cell mass induces an acrosome reaction and promotes penetration into the oocyte [19].

Heptanol, a seven-carbon alcohol, reduces gap junctional coupling between cumulus cells and the oocyte [20] and is a possible alternative for inhibiting the function of cumulus cells in the maturation of the oocyte without their removal. Coskun and Lin [9] showed that transforming growth factor {alpha} induced signaling from cumulus cells to the oocyte via the gap junction by using heptanol. However, there has been no report on the effect of heptanol on penetrability and subsequent MPN formation.

The present study was designed to clarify the roles of GJC between the oocyte and cumulus cells in nuclear maturation, penetration of spermatozoa, and formation of the MPN of the oocyte cultured in vitro by using heptanol instead of removing the cumulus cells.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Medium

The medium used in the handling of oocytes and spermatozoa was M2 [21]. The basic culture medium used for the maturation of oocytes was tissue culture medium (TCM)-199 supplemented with 25 mM Hepes and 26.2 mM sodium hydrogen-carbonate (pH 7.4). After in vitro fertilization in Brackett and Oliphant (BO) solution [22], oocytes were cultured in NCSU23 [23].

Preparation and In Vitro Maturation of Oocytes

Ovaries from prepubertal gilts were collected at a local slaughterhouse and transported to the laboratory within 1 h in a warm 0.85% sterile saline solution. Cumulus-oocyte complexes (COCs) were aspirated from antral follicles 2–5 mm in diameter using an 18-gauge needle fixed to a 5-ml disposable syringe and then pooled in 10-ml test tubes placed in a water bath (37°C) for settling. The COCs were collected and washed twice with M2, and only COCs with intact cumulus cell layers and uniform cytoplasm were selected. After COCs were washed with the maturation medium, TCM-199 containing 10 IU/ml hCG, 10 IU/ml eCG, 1 mM L-glutamine, 10% fetal calf serum (Filtron, Brooklyn, NSW, Australia), and 10% pig follicular fluid, 20 COCs were transferred to 200 ml of the maturation medium under mineral oil (M 8410; Sigma Chemical Co., St. Louis, MO) in a 35-mm Petri dish (Falcon 1008; Becton & Dickinson, Lincoln Park, NJ) and cultured in a CO2 incubator (5% CO2 in air at 39°C) for 42–48 h. The cumulus cells were removed from some of the oocytes by the use of a pipette of narrow bore size (about 100 µm) prior to the culture for maturation.

Sperm Preparation and In Vitro Fertilization

The sperm-rich fraction of ejaculate was obtained from a Large White boar using the gloved-hand method and kept overnight at 15°C. The COCs were transferred into fertilization medium droplets (90 µl) under mineral oil. Semen samples were washed with M2 once and then with BO solution equilibrated in 5% CO2 in air at 37°C. The spermatozoa were then suspended in equilibrated BO solution to a concentration of 2 x 106 cells/ml. Ten microliters of the sperm suspension was introduced into the fertilization medium containing COCs, and the final concentration of spermatozoa was 2 x 105 cells/ml. After 6 h of coincubation, oocytes were transferred to the embryo culture medium, and culturing was continued for a further 2 h.

Assessment of Pronuclear Formation

After the removal of cumulus cells by brief treatment with 0.1% hyaluronidase, oocytes (8 h postinsemination) were fixed with acid-alcohol (ethanol and acetic acid, 3:1, v:v) and stained with 1% aceto-orcein for phase-contrast microscopy. Oocytes with a female pronucleus and 2 polar bodies were considered to have been penetrated by spermatozoa at metaphase II. Spermatozoa that had penetrated into oocytes and progressed to MPN were classified into the following 5 stages (Fig. 1) according to Poccia and Collas [24]: fusion of spermatozoa with the oolemma (stage 1); vesiculation of the sperm nuclear membrane at the equatorial segment and the start of decondensation (stage 2); disassembly of the sperm nuclear membrane and full decondensation of the chromatin (stage 3); commencement of stranding of the sperm chromatin and apparent absence of the nuclear membrane (stage 4); and reassembly of the sperm nuclear membrane and formation, expansion, and development of MPN (stage 5).



View larger version (126K):
[in this window]
[in a new window]
  		FIG. 1. Microphotographs of oocytes fixed after 8 h of coincubation with spermatozoa. Spermatozoa that had penetrated into oocytes and progressed to MPN were classified into 5 stages. In stage 1, spermatozoa fused with the oolemma (arrow), and these spermatozoa stained darker than unfused spermatozoa (arrowhead) (a). In stage 2, the nuclear envelope of the spermatozoa vesiculated at the equatorial segment and started to decondense (b). In stage 3, the nuclear membrane disassembled and the chromatin fully decondensed (arrows) (c). In stage 4, the chromatin started to strand and the nuclear membrane appeared to be absent (arrow) (d). In stage 5, the nuclear membrane reassembled, and the MPN formed (arrow) (e) and expanded and developed (arrow) (f). All photographs were taken at the same magnification. Bar = 5 µm

Assay of GSH Content in the Oocyte

After 48 h of maturation, the amount of GSH in the oocytes was assayed. Cumulus cells were removed from the oocytes by treatment with 0.1% hyaluronidase and agitation using a narrow-bore pipette. Denuded oocytes were washed twice with PBS containing 1 mg/ml polyvinyl alcohol (P8136; Sigma) (PBS-PVA). Then 20–30 oocytes were put into polypropylene microcentrifuge tubes, and they were washed 3 times with PBS-PVA by centrifugation. After the supernatant was removed, 5 µl of distilled water was added to each tube, and the samples were frozen at -20°C. The frozen samples were then thawed at room temperature, and 5 µl of 1.25 M phosphoric acid solution was added to each tube. The samples were refrozen for storage at -20°C until use for the assay of GSH content. GSH content was determined by the enzymatic cycling assay according to the method described by Tietze [25]. Each sample was thawed and diluted with 190 µl of distilled water, and then transferred to a cuvette (Microcuvettes; Kartell, Milan, Italy). The following solutions were subsequently added to the cuvette: 10 µl of 200 IU/ml glutathione reductase (Sigma) and 100 µl of 6 mM 5,5-dithiobis (2-nitrobenzonic acid; Sigma). Immediately after the addition of 700 µl of 0.33 mg/ml NADPH dissolved in stock buffer (0.2 M sodium phosphate, 10 mM EDTA, pH 7.2), the absorbance was monitored at 412 nm by use of a spectrophotometer (Hitachi U-3210; Tokyo, Japan). Blank solution (900 µl of PVA-PBS and 100 µl of stock buffer) and GSH (oxidative form of GSH; Wako Pure Chemical Industry, Osaka, Japan) standards were also assayed under the same conditions. The amount of GSH in each sample was determined from these standards. The amount of GSH was calculated as that per 30 oocytes.

Experiment 1

To clarify the effect of heptanol on the GSH content in oocytes, heptanol was supplemented to the maturation medium at a concentration of 1 mM, 5 mM, or 10 mM. At the end of maturation, the amount of GSH in oocytes was measured. The oocytes matured in the medium without heptanol (0 mM) were treated as a control. Oocytes cultured without cumulus cells were also treated as a negative control.

Experiment 2

To clarify the effect of inhibition of GJC during culture on maturation, oocytes matured for 48 h in medium containing 0 mM (control), 1 mM, 5 mM, or 10 mM heptanol were fertilized in vitro; then nuclear maturation of oocytes, sperm penetration, and MPN formation were evaluated.

Experiment 3

To evaluate the significance of GSH in the oocyte for MPN formation, 100 µM cysteamine (Sigma) was added to a maturation medium in the presence or absence of 10 mM heptanol; the rate of MPN formation and GSH contents in oocytes were then analyzed.

Statistical Analysis of Data

Data were analyzed for statistically significant differences (P < 0.05) by Fisher's protected least significant differences (Statview ver.5; Abacus Concepts, Berkeley, CA).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Experiment 1

The amount of GSH in oocytes immediately before culture was 0.231 nmol/30 oocytes, and this amount remained constant until the end of maturation when oocytes were cultured in the absence of heptanol (0.219 nmol/30 oocytes). However, the amount of GSH decreased as the concentration of heptanol added to the medium was increased (0.163 and 0.142 nmol/30 oocytes cultured with 1 and 5 mM heptanol, respectively) (Fig. 2). The amount of GSH in oocytes cultured in the presence of 10 mM heptanol was significantly lower than that in the control (0.124 vs. 0.219 nmol/30 oocytes, P < 0.05). The amount of GSH in oocytes cultured in the absence of cumulus cells (0.125 nmol/30 oocytes) was the same as that in oocytes treated with 10 mM heptanol.



View larger version (50K):
[in this window]
[in a new window]
  		FIG. 2. Effect of heptanol on the content of GSH in oocytes matured in vitro. COCs were cultured in media with 0, 1, 5, or 10 mM heptanol; at the end of culture the amounts of GSH were measured by the enzymatic cycling assay after removal of cumulus cells. The amounts of GSH in oocytes at the start of culture (IVM 0h) and denuded oocytes (DO) cultured without heptanol were also measured. The results represent the mean ± SEM. The numbers in parentheses are the total numbers of samples containing 25–30 oocytes/numbers of replicates. *Significantly different from 0 mM treatment (P < 0.05)

Experiment 2

At the end of in vitro maturation, most of the oocytes reached metaphase II in the presence of heptanol at any concentration tested (Table 1). Cumulus cell expansion and the morphology of oocytes cultured in the medium with heptanol were almost the same as those of the control without heptanol. There was also no difference between the rates of penetrated oocytes cultured in the presence of heptanol at various concentrations (P > 0.1) (Table 1). However, the proportion of oocytes that formed MPN (stage 5) decreased significantly (P < 0.01) at all concentrations of heptanol tested. Close observation of the morphology of penetrated spermatozoa showed that a higher concentration of heptanol added to the maturation medium resulted in a higher rate of oocytes that had only sperm of stage 1 (P < 0.01) (Fig. 3) and a higher rate of failure of spermatozoa to degrade their nuclear envelopes (P < 0.01) (Fig. 4).


View this table:
[in this window]
[in a new window]
  		TABLE 1. Effects of heptanol added to the maturation medium on maturation and penetrability of porcine oocytes



View larger version (89K):
[in this window]
[in a new window]
  		FIG. 3. Effect of heptanol on the ability of oocytes to decondense the sperm head and to form the MPN. Of the oocytes cultured in media with various concentrations of heptanol and penetrated by spermatozoa (Table 1), the percentages of oocytes with the most advanced stage of the sperm head (see Fig. 1) have been plotted. **, *: Significantly different from 0 mM treatment in each stage at P < 0.01 and P < 0.05, respectively



View larger version (59K):
[in this window]
[in a new window]
  		FIG. 4. Effect of heptanol on decondensation of the sperm head and male pronuclear formation. Percentages of spermatozoa that penetrated and progressed to each stage (see Fig. 1) in each treatment have been plotted. A total of 565–676 spermatozoa were analyzed in each treatment. **, *: Significantly different from 0 mM treatment in each stage at P < 0.01 and P < 0.05, respectively

Experiment 3

When 100 µM cysteamine was added to the medium, the rate of MPN formation was improved (63% vs. 35% in experiment 2). An effect of cysteamine on MPN formation was observed even if the medium contained 10 mM heptanol, and a significantly higher proportion of oocytes formed MPN (P < 0.05) (Table 2). The amount of GSH in oocytes cultured with both cysteamine and heptanol (0.277 ± 0.017 pmol/30 oocytes, mean ± SEM from 9 samples in 5 replicates) was the same as that for oocytes cultured with cysteamine only (0.245 ± 0.018 pmol/30 oocytes, mean ± SEM from 4 samples in 3 replicates); and these amounts were significantly higher (P < 0.05) than the amount in oocytes cultured with heptanol only (0.168 ± 0.016 pmol/30 oocytes, mean ± SEM from 7 samples in 6 replicates).


View this table:
[in this window]
[in a new window]
  		TABLE 2. Effect of cysteamine on male pronuclear formation of porcine oocytes cultured with or without heptanol


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The gap junctional channel is composed of two symmetrical structures: connexons that create an intracellular channel and allow passage of nutrients, ions, and regulatory molecules of less than about 1 kDa between contacting cells [26]. Connexons are composed of a hexamer of membrane proteins, termed connexins, that constitute a large family [27]. Gap junctions are ubiquitous in multicellular organisms and play an important role in the regulation of the proliferation, differentiation, and homeostasis of the cell [28]. In the antral follicle, the oocyte and somatic cells have the communication via gap junctions that is essential for normal oocyte and follicular development [29]. It has been proposed that mural granulosa cells generate a meiosis-arresting substance and transport it to the oocyte through granulosa-cumulus and cumulus-oocyte GJCs [30, 31]. Once the COC becomes separate from the granulosa cells by a reduction in granulosa-cumulus cell coupling [31–35] or by the isolation of the COC from the follicle [30, 31, 36, 37], the oocyte undergoes germinal vesicle breakdown. It has been reported that the disruption of GJC between the oocyte and the cumulus cells induces meiotic resumption [11, 38, 39]. On the other hand, Fagbohun and Downs [20] reported that gap junctional coupling between the cumulus cells and the oocyte is necessary to mediate the action of Concanavalin A and FSH, which induce meiotic resumption in the mouse. Other studies have demonstrated that the meiotic resumption precedes loss in GJCs [40, 41]. Considering previous studies, both mural granulosa cells and cumulus cells must take part in the regulation of oocyte maturation by generating inhibitory or stimulatory effects with complex modes. In the present study there were no statistically significant differences from control values in the rates of oocytes that were arrested at germinal vesicle stage or reached metaphase II at the end of culture in the presence of heptanol (Table 1), and heptanol did not inhibit nuclear maturation. This finding is consistent with the result reported by Coskun and Lin [9]. To elucidate the regulatory action of cumulus cells in mitotic resumption more precisely, the timing of the progression of nuclear maturation of oocytes cultured in the medium with or without heptanol should be investigated.

Fagbohun and Downs [20] reported that 1 mM heptanol reduced gap junctional coupling between oocytes and the cumulus cells by 13%. Gap junction channels are embedded in relatively cholesterol-rich domains of the membrane, and heptanol decreases the gap junctional conductance by decreasing the fluidity of these cholesterol-rich domains [42]. This results in a decrease in probability that gap junction channels will open [42, 43]. Lars Bastiaanse et al. [42] suggested that more heptanol is needed to achieve efficient uncoupling of gap junctions in cholesterol-rich lipid domains. In the present study, a relatively high concentration of heptanol (10 mM) was needed to reduce the amount of GSH to the level in cumulus-free, denuded oocytes (experiment 1). Therefore we used 10 mM heptanol thereafter.

Our results showed that there was no difference between the numbers of spermatozoa that penetrated the oocytes in each of the media and that heptanol does not disturb the penetrability of oocytes. Galeati et al. [18] reported that the removal of cumulus cells during maturation induces a premature migration of cortical granules, leading to an increase in exocytotic events and a decrease in penetrability compared with values in cumulus-enclosed oocytes. In contrast to their experiments, cumulus cells were not removed and only GJC was inhibited in our experiments. Although localization of the cortical granules was not investigated in the present study, it was thought that heptanol did not induce the premature migration of cortical granules in the oocytes because there was no difference from the control in the proportion of penetrated oocytes and the mean number of penetrated spermatozoa.

In contrast to the penetrability of oocytes, heptanol influenced the oocyte's ability to shed the nuclear envelope of spermatozoa. Many spermatozoa that penetrated into the heptanol-treated oocytes were arrested at stage 1, and significantly more spermatozoa failed to proceed to stages 4–5. According to observations in the hamster by Usui and Yanagimachi [44], nuclear envelope removal and the subsequent chromatin decondensation are under different ooplasmic regulations. If this is the case in porcine oocytes, the oocytes treated with heptanol might lack the ability to remove the nuclear envelope, because the stage 1 spermatozoa seemed to fuse with oolemma but failed to proceed to the next step. Berrios and Bedford [45] and Szollosi et al. [46] also reported that the nuclear envelopes of spermatozoa that had penetrated into immature oocytes remained intact. It is likely that mammalian oocytes obtain the ability to break down the sperm nuclear envelope during cytoplasmic maturation. Therefore, the arrest of spermatozoa at stage 1 may be related to the lower ability of the oocyte to remove the nuclear envelope, and it is possible that the function of cumulus cells via GJC should be involved in the development of this ability.

Porcine follicular oocytes and cumulus cells cultured in vitro have GJC during the first 24–32 h of culture [47], and the factors related to cytoplasmic maturation might be transported from cumulus cells to the oocyte in this duration. These factors were not determined in the present study, but GSH may be a candidate, because our study showed that heptanol added to the medium decreased the GSH content in oocytes and inhibited the formation of MPNs. Yoshida [13] also reported the significance of GSH in oocytes during the initial and mid phases of maturation for MPN formation. Recently, Yamauchi and Nagai [48] demonstrated that cysteamine increased the content of GSH and promoted MPN formation in cumulus-free oocytes. Consistent with their result, our results showed that cysteamine increased the GSH content and the rate of MPN formation even though the oocyte was treated with 10 mM heptanol. However, the possibility of involvement of other factors, such as intracellular pH and intracellular [Ca], that are regulated via GJC cannot be excluded.

In summary, this study showed that heptanol, a gap junction inhibitor, is a useful reagent for investigating the function of cumulus cells during oocyte maturation without disturbing the penetrability of oocytes. The GJC between the oocyte and the cumulus cell is thought to play an important role in regulating ooplasmic factor(s) involved in the removal of sperm nuclear envelopes as well as in GSH transportation.


   ACKNOWLEDGMENTS
 
The authors would like to thank Dr. M. Yamashita, Division of Biological Sciences, Graduate School of Science, Hokkaido University, for his critical advice. We are also grateful to the staff of the Ebetsu Meat Inspection Office and the staff of the Experimental Farm, Faculty of Agriculture, Hokkaido University, for their help in the collection of porcine ovaries and semen, respectively.


   FOOTNOTES
 
First decision: 27 September 1999.

1 Correspondence. FAX: 81 11 716 0879; tamori{at}anim.agr.hokudai.ac.jp Back

2 Current address: Central Research Institute, Itoham Food Inc., Moriya, Kitasoma, Ibaraki 302-0104, Japan. Back

Accepted: November 16, 1999.

Received: August 18, 1999.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Mattioli M, Galeati G, Seren E. Effect of follicle somatic cells during pig oocyte maturation on egg penetrability and male pronuclear formation. Gamete Res 1988; 20:177–183.[CrossRef][Medline]
  2. Mattioli M, Galeati G, Bacci ML, Seren E. Follicular factors influence oocyte fertilizability by modulating the intercellular co-operation between cumulus cells and oocytes. Gamete Res 1988; 21:223–232.[CrossRef][Medline]
  3. Moor RM, Mattioli M, Ding J, Nagai T. Maturation of pig oocytes in vivo and in vitro. J Reprod Fertil Suppl 1990; 40:197–210.[Medline]
  4. Fukui Y. Effect of follicular cells on the acrosome reaction, fertilization, and developmental competence of bovine oocytes matured in vitro. Mol Reprod Dev 1990; 26:40–46.[CrossRef][Medline]
  5. Vanderhyden BC, Armstrong DT. Role of cumulus cells and serum on the in vitro maturation, fertilization, and subsequent development of rat oocytes. Biol Reprod 1989; 40:720–728.[Abstract]
  6. Kikuchi K, Nagai T, Motlik J, Shioya Y, Izaike Y. Effect of follicular cells on in vitro fertilization of pig follicular oocytes. Theriogenology 1993; 39:593–599.
  7. Ka HH, Sawai K, Wang WH, Im KS, 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]
  8. Gilula NB, Epstein ML, Beers WH. Cell-to-cell communication and ovulation: a study of the cumulus-oocyte complexes. J Cell Biol 1978; 78:58–75.[Abstract/Free Full Text]
  9. Coskun S, Lin YC. Effect of transforming growth factors and activin-A on in vitro porcine oocyte maturation. Mol Reprod Dev 1994; 38:153–159.[CrossRef][Medline]
  10. Anderson E, Albertini DF. Gap junctions between the oocytes and companion follicle cells in the mammalian ovary. J Cell Biol 1976; 71:680–686.[Abstract/Free Full Text]
  11. Dekel N, Beers WH. Development of rat oocytes in vitro: inhibition and induction of maturation in the presence or absence of cumulus-oophorus. Dev Biol 1980; 75:247–254.[CrossRef][Medline]
  12. Perreault SD, Barbee RR, Slott VL. Importance of glutathione in the acquisition and maintenance of sperm nuclear decondensing activity in maturing hamster oocytes. Dev Biol 1988; 125:181–186.[CrossRef][Medline]
  13. Yoshida M. Role of glutathione in the maturation and fertilization of pig oocytes in vitro. Mol Reprod Dev 1993; 35:76–81.[CrossRef][Medline]
  14. Yoshida M, Ishigaki K, Persel VG. Effect of maturation media on male pronucleus formation in pig oocytes matured in vitro. Mol Reprod Dev 1992; 31:68–71.[CrossRef][Medline]
  15. Sawai K, Funahashi H, Niwa K. Stage-specific requirement of cysteine during in vitro maturation of porcine oocytes for glutathione synthesis associated with male pronuclear formation. Biol Reprod 1997; 57:1–6.[Abstract]
  16. Zuelka KA, Parreault SD. Hamster oocyte and cumulus cell glutathione concentrations increase rapidly during in vitro maturation. Biol Reprod 1994; 50(suppl):144 (abstract).
  17. Funahashi H, Day BN. Effects of cumulus cells on glutathione content of porcine oocyte during in vitro maturation. J Anim Sci 1995; 73(suppl 1):90 (abstract).
  18. Galeati G, Modina S, Lauria A, Mattioli M. Follicle somatic cells influence pig oocyte penetrability and cortical granule distribution. Mol Reprod Dev 1991; 29:40–46.[CrossRef][Medline]
  19. Mattioli M, Lucidi P, Barboni B. Expanded cumuli induce acrosome reaction in boar sperm. Mol Reprod Dev 1998; 51:445–453.[CrossRef][Medline]
  20. Fagbohun CF, Downs SM. Metabolic and ligand-stimulated meiotic maturation in the mouse oocyte-cumulus cell complex. Biol Reprod 1991; 45:851–859.[Abstract]
  21. Quinn P, Barros C, Whittingham DG. Preservation of hamster oocytes to assay the fertilizing capacity of human spermatozoa. J Reprod Fertil 1982; 66:161–168.[Abstract/Free Full Text]
  22. Brackett BG, Oliphant G. Capacitation of rabbit spermatozoa in vitro. Biol Reprod 1975; 12:260–274.[Abstract]
  23. Petters RM, Wells KD. Culture of pig embryos. J Reprod Fertil Suppl 1994; 48:61–73.
  24. Poccia D, Collas P. Nuclear envelop dynamics during male pronuclear development. Dev Growth Differ 1997; 39:541–550.[CrossRef][Medline]
  25. Tietze F. Enzymic method for quantitative determination of nanogram amounts of total and oxidized glutathione: applications to mammalian blood and other tissues. Anal Biochem 1969; 27:502–522.[CrossRef][Medline]
  26. Barhoumi R, Bowen JA, Stein LS, Echols J, Burghardt RC. Concurrent analysis of intracellular glutathione content and gap junctional intercellular communication. Cytometry 1993; 14:747–756.[CrossRef][Medline]
  27. Haefliger JA, Bruzzone R, Jenkins NA, Gilbert DA, Copeland NG, Paul D. Four novel members of the connexin family of gap junction proteins. J Biol Chem 1992; 267:2057–2064.[Abstract/Free Full Text]
  28. Grazul-Bilska AT, Reynolds LP, Redmer DA. Gap junctions in the ovaries. Biol Reprod 1997; 57:947–957.[CrossRef][Medline]
  29. Buccione R, Schroeder AC, Eppig JJ. Interactions between somatic cells and germ cells throughout mammalian oogenesis. Biol Reprod 1990; 43:543–547.[Abstract]
  30. De Loos FAM, Zeinstra E, Bevers MM. Follicular wall maintains meiotic arrest in bovine oocytes cultured in vitro. Mol Reprod Dev 1994; 39:162–165.[CrossRef][Medline]
  31. Kalous J, Sutovsky P, Rimkevicova Z, Shioya Y, Lie B, Motlik J. Pig membrana granulosa cells prevent resumption of meiosis in cattle oocytes. Mol Reprod Dev 1993; 34:58–64.[CrossRef][Medline]
  32. Lartsen WJ, Tung HN, Polking C. Response of granulosa cell gap junctions to human chorionic gonadotropin (hCG) at ovulation. Biol Reprod 1981; 25:1119–1134.[Abstract]
  33. Larsen WJ, Wert SE, Brunner GD. Differential modulation of rat follicle cell gap junction populations at ovulation. Dev Biol 1987; 122:61–71.[CrossRef][Medline]
  34. Larse WJ, Wert SE, Brunner GD. A dramatic loss of cumulus cell gap junctions is correlated with germinal vesicle breakdown in rat oocytes. Dev Biol 1986; 113:517–521.[CrossRef][Medline]
  35. Racowsky C, Baldwin KV, Larabell CA, DeMarais AA, Kazilek CJ. Down-regulation of membrana granulosa cell gap junctions is correlated with irreversible commitment to resume meiosis in golden Syrian hamster oocytes. Eur J Cell Biol 1989; 49:244–251.[Medline]
  36. Pincus G, Saunders B. The comparative behavior of mammalian eggs in vivo and in vitro. I. The activation of ovarian eggs. J Exp Med 1935; 62:655–675.
  37. Edwards RG. Maturation in vitro of mouse, sheep, cow, pig, rhesus monkey and human ovarian oocytes. Nature 1965; 208:349–351.[CrossRef][Medline]
  38. Dekel N, Lawrence TS, Gilua NB, Beers WH. Modulation of cell-cell communication in the cumulus-oocyte complex and the regulation of oocyte maturation by LH. Dev Biol 1981; 86:356–362.[CrossRef][Medline]
  39. Isobe N, Maeda T, Terada T. Involvement of meiotic resumption in the disruption of gap junctions between cumulus cells attached to pig oocytes. J Reprod Fertil 1998; 113:167–172.[Abstract/Free Full Text]
  40. Moor RM, Smith MW, Dawson RMC. Measurement of intercellular coupling between oocytes and cumulus cells using intercellular markers. Exp Cell Res 1980; 126:15–29.[CrossRef][Medline]
  41. Eppig JJ. The relationship between cumulus cell-oocyte coupling, oocyte maturation, and cumulus expansion. Dev Biol 1982; 89:268–272.[CrossRef][Medline]
  42. Lars Bastiaanse EM, Jongsma HJ, van der Laarse A, Takens-Kwak BR. Heptanol-induced decrease in cardiac gap junctional conductance is mediated by a decrease in the fluidity of membranous cholesterol-rich domains. J Membr Biol 1993; 136:135–145.[CrossRef][Medline]
  43. Meda P. Gap junctional coupling and secretion in endocrine and exocrine pancreas. In: Sperelakis N, Cole WC (eds.), Cell Interactions and Gap Junctions, vol. 1. Boca Raton, FL: CRC Press; 1989: 59–83.
  44. Usui N, Yanagimachi Y. Behavior of hamster sperm nuclei incorporated into eggs at various stages of maturation, fertilization and early development. J Ultrastruct Res 1976; 57:276–288.[CrossRef][Medline]
  45. Berrios M, Bedford JM. Oocyte maturation: aberrant post-fusion responses of the rabbit primary oocyte to penetrating spermatozoa. J Cell Sci 1976; 39:1–12.[Abstract/Free Full Text]
  46. Szollosi D, Szollosi MS, Czolowska R, Tarkowski AK. Sperm penetration into immature mouse oocytes and nuclear changes during maturation: an EM study. Biol Cell 1986; 42:140–151.
  47. Motlik J, Fulka J, Flechon JE. Changes in intercellular coupling between pig oocytes and cumulus cells during maturation in vivo and in vitro. J Reprod Fertil 1986; 76:31–37.[Abstract/Free Full Text]
  48. Yamauchi N, Nagai T. Male pronuclear formation in denuded porcine oocytes after in vitro maturation in the presence of cysteamine. Biol Reprod 1999; 61:828–833.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 Biol. Reprod.Home page
A. Bettegowda, O. V. Patel, K.-B. Lee, K.-E. Park, M. Salem, J. Yao, J. J. Ireland, and G. W. Smith
Identification of Novel Bovine Cumulus Cell Molecular Markers Predictive of Oocyte Competence: Functional and Diagnostic Implications
Biol Reprod, August 1, 2008; 79(2): 301 - 309.
[Abstract] [Full Text] [PDF]

Home page
 ReproductionHome page
P. Pocar, D. Nestler, M. Risch, and B. Fischer
Apoptosis in bovine cumulus-oocyte complexes after exposure to polychlorinated biphenyl mixtures during in vitro maturation
Reproduction, December 1, 2005; 130(6): 857 - 868.
[Abstract] [Full Text] [PDF]

Home page
 Hum ReprodHome page
J. Hasegawa, A. Yanaihara, S. Iwasaki, Y. Otsuka, M. Negishi, T. Akahane, and T. Okai
Reduction of progesterone receptor expression in human cumulus cells at the time of oocyte collection during IVF is associated with good embryo quality
Hum. Reprod., August 1, 2005; 20(8): 2194 - 2200.
[Abstract] [Full Text] [PDF]

Home page
 Biol. Reprod.Home page
H. Tatemoto, N. Muto, I. Sunagawa, A. Shinjo, and T. Nakada
Protection of Porcine Oocytes Against Cell Damage Caused by Oxidative Stress During In Vitro Maturation: Role of Superoxide Dismutase Activity in Porcine Follicular Fluid
Biol Reprod, October 1, 2004; 71(4): 1150 - 1157.
[Abstract] [Full Text] [PDF]

Home page
 Biol. Reprod.Home page
C. Campagna, M.-A. Sirard, P. Ayotte, and J. L. Bailey
Impaired Maturation, Fertilization, and Embryonic Development of Porcine Oocytes Following Exposure to an Environmentally Relevant Organochlorine Mixture
Biol Reprod, August 1, 2001; 65(2): 554 - 560.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
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 My Folders
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 Mori, T.
Right arrow Articles by Shimizu, H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Mori, T.
Right arrow Articles by Shimizu, H.
Agricola
Right arrow Articles by Mori, T.
Right arrow Articles by Shimizu, H.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS