J. Embryol. exp. Morph. Vol. 61, pp. 347-365, 1981
Printed in Great Britain <S Company of Biologists Limited 1981
2>A1
Selective effect of
gonadotrophins on cell coupling, nuclear
maturation and protein synthesis in
mammalian oocytes
By R. M. MOOR1, J. C. OSBORN1, D. G. CRAN 1
AND D. E. WALTERS 2
From the Agricultural Research Council Institute of Animal Physiology, Cambridge and A.R.C. Statistics Group, Department of Applied Biology, Cambridge
SUMMARY
Individual gonadotrophic hormones were used to examine the; degree to which changes in
intercellular coupling between somatic and germ cells initiate rneiotic maturation, regulate
protein synthesis or alter the ultrastructure of the ovine oocyte. Follicle Stimulating Hormone
(FSH; SOngmh1) suppressed intercellular coupling to the same extent as that observed
during oocyte maturation in vivo. At low concentrations FSH did not, however, initiate
resumption of meiosis. By contrast, luteinising hormone (LH; 100-500 ng ml"1) invariably
initiated meiosis in oocytes cultured within the follicle but did not disrupt intercellular
coupling. We conclude that nuclear maturation is not dependent upon the disruption of cell
contact between the oocyte and the surrounding follicle cells.
The profile of proteins synthesized by untreated oocytes differed greatly from that of
oocytes matured for 18 h in follicles treated with a combination of FSH and LH. Pretreatment of follicles with either FSH or LH at low concentrations resulted in the synthesis of an
intermediate and more variable pattern of proteins. No correlation was found between
changes in protein synthesis and the extent of junctional communication between the cumulus
cells and oocyte.
Membrane vesiculation and lysosomal change in the transzonal processes are early structural
changes associated with the suppression of intercellular coupling in oocytes. These changes in
coupling probably result in the relocation of intracellular organelles in the final stages of
oocyte maturation.
INTRODUCTION
A variety of different cellular components in the oocyte undergo change after
the resumption of meiosis. Apart from the characteristic structural changes in
the nucleus, alterations also occur in cell metabolism., protein synthesis, membrane transport and in the localization of organelles within the cytoplasm (com1
Authors' address: Agricultural Research Council Institute of Animal Physiology,
307 Huntingdon Road, Cambridge, CB3 OJQ, U.K.
2
Author's address: ARC Statistics Group, Department of Applied Biology, Pembroke
Street, Cambridge, CB2 3DX.
348
R. M. MOOR AND OTHERS
prehensively reviewed by Masui & Clarke, 1979). To gain a clear appreciation
of the functional aspects of the maturational process it is, however, necessary
to move from the study of individual events to studies on interactions and interdependency between different compartments within the oocyte. The existing
data, although limited, nevertheless already shows that while some changes are
interdependent, others can occur in an apparently independent manner. Thus,
nuclear maturation can be induced under certain*conditions without inducing
all the changes in protein synthesis that characterize the fully mature oocyte
(Thibault, 1977; Warnes, Moor & Johnson, 1977). Similarly, amino-acid
fluxes across the oocyte membrane can be increased to those of the mature
oocyte without inducing comparable changes in the nucleus or in protein
synthesis (Moor & Smith, 1979).
Recent experiments have shown that low molecular-weight compounds
pass freely between the cytoplasm of follicle cells and that of the oocyte (Gilula,
Epstein & Beers, 1978; Moor, Smith & Dawson, 1980). These compounds
enter the oocyte through permeable junctions formed at points of contact
between the membranes of the somatic and germ cells. The term cell coupling
refers in this paper to passage of molecules from the follicle cells into the oocyte
through permeable junctions. This intercellular passage of small molecules is
sharply reduced after the resumption of meiosis. A favoured current hypothesis
suggests that the decline in intercellular coupling acts as the stimulus for the
breakdown of the germinal vesicle (Dekel & Beers, 1978). In this paper we test
this hypothesis and report on the relationship that exists between cell-coupling,
nuclear maturation and protein synthesis in ovine oocytes.
METHODS
Tissue preparation
Four separate experimental studies were undertaken (see Results). The
follicles used in each study were obtained from the ovaries of sheep that had
been injected on day 10-12 of the cycle with 1250 i.u. pregnant mare serum
gonadotrophin (PMSG) and slaughtered 40 h later. Follicles that had responded
to the exogenous gonadotrophin were dissected from the ovaries and cultured
for 9-18 h using similar techniques and culture media to those described previously (Moor & Trounson, 1977).
At the end of the culture period follicles were either fixed for structural studies
(expts 2 and 3) or were opened, washed to remove follicular fluid and then
further dissected to remove the entire cumulus-oocyte cell complex (expts 1, 3
and 4). These preparative procedures were carried out at 37 °C in an incubation
medium consisting of Dulbecco's phosphate-buffered saline supplemented with
bovine serum albumin (4 mg ml"1), pyruvate (0-36 HIM), lactate (23-8 HIM) and
glucose (5-5 HIM).
Hormones and compartmental change in oocytes
349
Table 1. Entry of choline via permeable junctions and the associated degree of
meiotic maturation in oocytes obtained from follicles cultured for 15 h in the
presence of different gonadotrophins
Percentage nuclear
(G.V.) change in
oocytes
Mean (±S.E.M.) choline
concentration in oocytes
OM)
Intact
(G.V.)
nuclear
breakdown
(GVBD)
88
45
516±46
160±46
86
11
14
89
48
42
216±32
87
38
13
62
Gonadotrophin
supplementation
No. of
oocytes
Untreated controls
FSH (5 fig ml"1) plus
LH (3 /tg ml"1)
FSH (100 ng ml"1)
LH (100 ng ml-1)
573 ±74
Measurement of intercellular coupling in oocytes
Intercellular coupling was quantified in oocytes obtained from follicles
cultured for 15 h in medium containing purified hormones (see Table 1).
Cumulus-oocyte complexes were placed in the wells of microtitre test plates and
maintained at 37 °C for 1 h in incubation medium containing 36 fiM methyl-[3H]
choline chloride, 5-15 Ci/m-mole, Radiochemical Centre, Amersham. Incubations were terminated by transferring the cell complexes into unlabelled medium
at 4 °C. Thereafter, oocytes were rapidly denuded of cumulus cells using finely
graded pipettes. Individual denuded oocytes were transferred to coverslips,
disrupted using 10/d sodium dodecyl sulphate (SDS) buffer and counted with
appropriate corrections for quenching and counting efficiency. Results are
expressed for convenience as the concentration of labelled choline in the
oocyte; it is, however, clear that most of the choline in oocytes is present as
phosphocholine or lecithin (Moor et al., 1980). The validity of using [3H]choline
for the measurement of intercellular coupling between cumulus cells and oocytes
has previously been discussed in detail (Moor et al., 1980).
Structural studies in oocytes
Groups of follicles (n = 8), removed 9, 12 or 15 h after culture in medium
containing either NIH-FSH-S9 (100 fig ml" 1 or NIH-LH-S18 (100 fig ml- 1 ), were
fixed in 4 % glutaraldehyde in collidine buffer (pH 7-2). The portion of follicle
wall bearing the cumulus-oocyte complex was removed and processed for
electron microscopy as described previously (Hay, Cran & Moor, 1976).
Hormonally induced changes in nuclear structure were examined in follicles
cultured for 15 h in medium containing different gonadotrophins (see Table 1).
The follicles were fixed and processed as for electron microscopy and 1 fim serial
12
EMB 6l
350
R. M. MOOR AND OTHERS
sections cut through the nucleus. Sections were stained in 1 % toludine blue in
1 % borax.
Electrophoretic separation of polypeptides in oocytes
Protein profiles were examined in oocytes obtained from follicles cultured for
18 h with different purified hormones. Groups of six to ten dissected and washed
oocyte-cumulus complexes were placed in microtitre plates and incubated at
37 °C for 3 h in 50 /*1 incubation medium containing 1 mCi ml" 1 of L-[ 3 5 S]
methionine (1000 Ci/m-mole; Radiochemical Centre, Amersham). After incubation, the oocyte-cumulus complexes were washed and denuded of cumulus cells.
Groups of three to five denuded oocytes were then briefly washed in 0-01 MTris-HCl, pH 7-4, collected in a small volume of Tris buffer (< 5 /*1) and frozen
at — 75 °C until required for electrophoresis.
Labelled oocytes were prepared for electrophoresis by adding 25-30 fi\ SDS
buffer (O'Farrell, 1975) and then freezing and thawing the samples twice. After
heating for one minute at 100 °C, a 5 / t l aliquot of each sample was used to
determine incorporation of radioactivity into TCA-precipitable material as
described by Van Blerkom (1978). A part of the remainder of each sample was
applied to an 8-15 % linear gradient SDS polyacrylamide slab gel (14 cm wide,
10 cm long and 0-15 cm thick) such that an equal number of TCA-precipitable
cpm (40000) was placed in each well.
Electrophoresis was carried out in the standard manner (Van Blerkom, 1978)
using the discontinuous SDS-glycine-Tris buffer system of Laemmli (1970).
Polypeptides were separated for 3 h at a constant current of 20 mA per gel.
Fluorography was carried out using the technique of Bonner & Laskey (1974).
Gels were dried on a Hoefer gel dryer and exposed to preflashed Kodak X-Omat
H film at - 7 0 °C for 48 h (Laskey & Mills, 1975). Molecular weight determinations were made using as standards: phosphorylase b (94K), albumin (67K),
ovalbumin (43K), carbonic anhydrase (30K), trypsin inhibitor (20-IK) and
lactalbumin (14-4K).
Sixteen polypeptide bands were selected for the quantitative analysis of
changes in protein synthesis by oocytes exposed to different hormone treatments.
Microdensitometer scans were made of each fluorogram and the relative amount
of protein in each marker band was obtained from the densitometer tracings
by determining the area under each peak using planimetric integration. The
results are presented as the amount of protein in each band expressed as a
percentage of the total amount of protein in the fluorogram.
Statistical analysis
An analysis was made of the pattern of proteins synthesized by oocytes after
treatment with one of four hormonal regimes (expt 4). The data used for analysis
consisted of 16 variates, being the percentage of protein in 16 individual marker
bands, obtained from each of the six or seven groups of oocytes (five oocytes/
Hormones and compartmental change in oocytes
351
Table 2. Relative amount of labelled protein in each of 16 marker bands expressed
as a percentage of the total protein synthesis in oocytes obtained from untreated
and gonadotrophin-treated groups of follicles. Each value represents the mean of
six or seven groups of oocytes (five oocytes/group) incubated in [355] methionine
for3h
Control
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
FSH
LH
FSH/LH
Std error*
of mean
(pooled)
2-37
3-61
211
3-36
2-87
3 07
2-97
2-61
1-77
104
2-65
1 99
0
0-55
0-78
1-39
4-72
4-41
5-48
3-54
0-79
113
0
0-35
3-60
3-60
3-81
3-53
4-99
6-81
6-50
604
3-26
2-46
301
3 04
7-66
6-25
6-95
6-79
7-72
7-12
6-66
6-98
2-77
4-45
3-75
316
1-42
2-20
2-23
2-26
208
1-54
1-88
2-24
3-20
1-83
2-23
2-38
1-28
1-41
1-75
1-46
* Statistics obtained from an analysis of variance for each
0-25
018
0-23
019
0-35
017
016
0-40
019
0-30
0-39
0-44
0-27
0-20
0-26
0-20
marker band
*
F ratio
8-25
1-17
816
919
5-32
7-99
0-54
3-89
319
3-65
1-31
2-82
2-27
2-36
4-68
105
group) that comprised the separate experimental treatments. The appropriate
mean values and standard errors for each of the individual marker bands are
shown in Table 2. The associated ' F ' ratios were calculated on each variate by
univariate analysis.
The data was subjected to multivariate analysis for a more thorough statistical
investigation. The purpose was to condense the original data, which in complete
diagrammatic representation would be in 16-dimensional space, into as few
dimensions as possible so that any underlying pattern could be recognized.
The canonical variate analysis proved particularly useful in this regard (Rao,
1952). This analysis selects the axis along which the inter-group differences are
greatest and then another axis, perpendicular to the first, which contains the
maximum amount of the remaining inter-group variation and so on to as many
variates as the data contains. This technique is, therefore, no more than a transformation of the multiple variates onto a scale which maximizes the discrimination between groups. The selection of the transformation takes account of the
variability of variates and the co-variation between each pair of variates.
Since a high proportion of the inter-group variation in our data was accounted
for by the first two canonical variates, a two-dimensional plot of the canonical
352
R. M. MOOR AND OTHERS
variate values provided a fairly accurate picture of the relative positions of the
treatment groups. A measure of the 'distance' between the treatment groups
was calculated as the Mahalanobis D statistic, which is the standardized difference between the group mean values, on the variate taken collectively. It is these
'distances' which are approximated by the inter-group differences in the canonical variate plot.
Due to irregularities in the error structure of the variates we found that a
logarithmic transformation of the data gave a more regular pattern in the
results. The data in Table 2, is for purposes of clarity, nevertheless still presented
in the original variable, whilst the canonical variate, analysis and the distance
statistics have been calculated on the log transformed variable.
RESULTS
(1) Measurement of intercellular coupling in oocytes
Studies in this paper are based upon the finding that FSH suppresses cell
coupling between oocytes and cumulus cells whilst LH is ineffective in this
respect (Moor et al. 1980). Tritiated choline has been used to measure changes
in the amount of intercellular coupling between these two cell types. This compound is rapidly taken up and phosporylated by follicle cells but cannot enter
the oocyte except through permeable junctions (Moore/ al., 1980). The rate at
which labelled choline metabolite enters the oocyte therefore provides a
functional measurement of the extent of cell coupling between the somatic and
germ cells.
The object of the experiments in this first section has been to define the lowest
concentration of FSH which consistently depresses intercellular passage of
labelled marker into oocytes. To establish this minimal hormonal requirement,
groups of follicles were cultured for 15 h in 50, 100 or 500 ng of FSH ml" 1
culture medium. The mean concentration of labelled choline in oocytes from these
groups of cultured follicles was 287 ± 82, 212 ±43 and 138 ± 34 JLIM respectively.
These concentrations were statistically indistinguishable from those of 175 ±26
and 160 + 46 /ui found in oocytes obtained (i) 15 h after the initiation of maturation in vivo (Moore/ al., 1980) or (ii) after 15 h culture in follicles treated with a
combination of high concentrations of FSH (5 /tg ml"1) and LH (3 /.ig ml"1).
In a parallel series of experiments, follicles were cultured for 15 h in medium
containing 100 or 500 ng LH ml"1. The mean concentration of choline in oocytes
from these two treatments was 541 ± 78 and 675 ± 188 jtm. These concentrations
were similar to those observed in oocytes cultured in the absence of hormones
(516±46/tM) but differed significantly from the concentrations in FSH-treated
oocytes (P < 001).
These results demonstrate that FSH, even at low concentrations, selectively
suppresses intercellular passage of low molecular weight compounds into
oocytes. On the basis of these findings it was decided to use FSH and LH at
Hormones and compartmental change in oocytes
353
1
concentrations of 100 ng ml" medium in all subsequent experiments described
in this paper.
(2) Time-dependent effects of FSH and LH on the structure of intercellular
junctions in oocytes
Intercellular transmission of compounds into oocytes occurs through slender
extensions of the cumulus cells which traverse the zona pellucida and become
enlarged on contact with the oolemma (Zamboni, 1972). Before gonadotrophin
treatment, each process contains numerous 6 nm microfilaments orientated
along its axis. Contact between the process and oolemma is maintained by
prominent intermediate junctions (Fig. 1). Occasional non-lysosomal vesicular
structures are found within the bulbous end of the process.
Addition of low levels of FSH to the culture medium resulted in the progressive degeneration of the cumulus cell processes. Within 9 h of FSH treatment lysosomes were observed within some of the processes (Fig. 5) and clusters
of vesicles, some 50 nm in diameter, became visible on the surface of some processes (Fig. 2). The electron micrographs showed clearly that these vesicles were
derived directly from the membrane of the cumulus cell process (Fig. 3). By
12 h after the addition of FSH most of the cumulus cell processes showed clear
signs of degeneration and after 15 h no intact processes were present. The
compact arrangement of cumulus cells at 9 h after FSH was similar to that
observed in untreated oocytes. However, by 12 h all of the cumulus cells except
for those abutting onto the zona pellucida (corona radiata) had undergone
marked dispersal (Fig. 4). Dispersal of the corona cells and separation from
the zona pellucida by 10-20 jam had occurred by 15 h after FSH treatment.
Nevertheless, these cells remained connected to the zona by cytoplasmic extensions containing abundant microfilaments. At this stage, few processes could
be detected in the zona pellucida at either the light or electron microscope level.
The addition of LH to the medium had no effect on the processes in most
oocytes examined 9-12 h later. The terminal ends of a small proportion of
processes, while retaining normal junctional contact with the oolemma, contained lysosome-like structures (Fig. 6). After 15 h exposure to LH, about half
the oocytes showed signs of process degeneration, involving the appearance of
lysosomes, loss of junctional integrity and the retraction of processes from the
surface, of the oocyte. In addition, the corona cells were more dispersed than at
12 h, and although difficult to quantify, there appeared to be fewer processes
traversing the zona than in the untreated group. It is noteworthy that the
membrane vesicles which regularly formed after FSH treatment were seldom
seen after treatment with LH.
An additional feature seen at 15 h after treatment with either FSH or LH was
an increase in the surface area of the oocyte membrane resulting from invaginations which varied from simple clefts (Fig. 7) to complex anastomosing networks
apparently isolating small regions of the oocyte cytoplasm (Fig. 8). These
354
R. M. MOOR AND OTHERS
' • • " * "
• • • • , . .
•
^ " -
'
.
;
;
;
«
.
Hormones and compartmental change in oocytes
355
specializations of the oocyte membrane, which were more apparent after LH
than FSH treatment, were readily penetrated with lanthanum which emphasized
their discontinuous nature (Fig. 9). In addition, the oocyte cytoplasm adjacent
to these areas changed structurally, containing a fine fibrillar material which
was particularly evident in lanthanum-impregnated oocytes (Fig. 10).
We conclude that the electron microscopical observations provide a structural
basis for the biochemical results which showed that FSH selectively depressed
junctional passage of molecules between cumulus cells and oocytes. From
the ultrastructure it is clear that the degenerative changes induced by FSH
are not only more rapid but are also more widespread and involve more
components of the processes than those induced by LH. By contrast, LH
appears to act most effectively upon the nonjunctional regions of the oocyte
membrane.
(3) Intercellular coupling and nuclear maturation in oocytes
In the first section of this paper we showed that the intercellular passage of
small molecules between cumulus cells and oocytes is disrupted by low levels of
FSH but is unaffected by LH. The purpose of the experiments in the present
section was to relate these changes in cell coupling to those obtained in parallel
studies on the morphology of the oocyte nucleus. The most important observations made in the study are summarized in Table 1. After 15 h culture with no
hormonal additions, intercellular passage of molecules was high and the great
majority of oocytes had intact germinal vesicles. Breakdown of cell coupling
and of the germinal vesicles occurred in oocytes cultured in follicles exposed to
high levels of FSH plus LH. That the two events are unrelated was, however,
clearly seen in follicles cultured with single hormones. Despite the suppression
of cellular coupling, FSH at low levels did not initiate germinal vesicle breakdown and nuclear maturation. By contrast, LH initiated nuclear maturation in
oocytes in which the degree of intercellular coupling remained at the same high
level as that recorded in untreated cells.
(4) Intercellular coupling and protein synthesis
There is almost no information available at present about the mechanisms
which induce changes in protein synthesis in mammalian oocytes during maturaFIGURES
1-3
Fig. 1. Terminal regions of cumulus cell processes (C) in an oocyte of a follicle
cultured without hormonal supplementation. Contact between the two cell types
is characterized by prominent intermediate junctions (arrows). Note tangentially
sectioned processes (P) in the zona pellucida. x 23500; bar = 1 //m.
Fig. 2. Cumulus cell process (C) after 9 h culture with FSH. It is surrounded by
numerous small vesicles and intermediate junctions are absent, x 57000; bar =
0-5 fim.
Fig. 3. Part of a process as in Fig. 2. Continuity between a vesicle and the process
plasma membrane is visible (arrow), x 70000; bar = 0-5/<m.
356
R. M. MOOR AND OTHERS
Hormones and compartmental change in oocytes
357
tion. It has, therefore, been of interest to examine the patterns of protein synthesis in oocytes in which selective changes have been induced in some of the
other intracellular compartments.
The effect of hormones on the incorporation of labelled methionine into TCAinsoluble material in oocytes was examined in the first part of the study.
Oocytes obtained from untreated follicles incorporated [35S]methionine at a mean
level (±S.E.M.) of 6-44 ± 0-85 f-mole methionine per oocyte h"1. The level of
incorporation was significantly increased (P < 0-0:5) in oocytes obtained from
follicles treated with FSH and LH to 8-62 + 0-44 f-imole methionine per oocyte
h~\ There was, however, no significant difference in the level of incorporation
of methionine between untreated oocytes and those obtained from follicles after
treatment with low levels of FSH (6-30 ± 1-4 f-mole per oocyte h"1) or LH
(4-91 ±0-89 f-mole per oocyte hr 1 ).
Illustrated in Fig. 11A is the pattern of proteins synthesized by oocytes
obtained from follicles cultured in the absence (untreated) or presence of
hormones.
Protein synthesis in untreated and FSH/LH-treated oocytes. A visual comparison of the fluorograms from untreated oocytes and oocytes obtained after
culture with high levels of gonadotrophin (FSH/LH group) showed numerous
changes in their protein profiles. Sixteen prominent bands (identified in Fig. 11 A),
which showed consistent changes in each of the six replicate experiments in the
study, were selected as markers of protein change during maturation. The effect
of hormones on these marker bands was analysed quantitatively by scanning
densitometry. Typical densitometer tracings from fluorograph tracks of untreated and FSH/LH-treated oocytes are shown in Fig. 11B. The numerically
labelled densitometer peaks correspond to the 16 bands selected as markers of
protein change (see Fig. 11 A). It should be noted that in a few instances the
resolution of the densitometer has been insufficient to separate a very dense
band from an immediately adjacent faint band. Where this has occurred the
area of the equivalent band combinations has been determined in both the
control and treated oocyte groups.
The rates of protein change in untreated and FSH/LH-treated oocytes are
shown in Table 2. When compared with the rates of change in untreated oocytes,
addition of gonadotrophin significantly altered the proportion of label inFIGURES
4-6
Fig. 4. Part of cumulus after treatment with FSH for 12 h. Those cells abutting
upon the zona pellucida (Z) are closely packed while the remainder are dispersed.
x2500; bar = 10 fim.
Fig. 5. Cumulus cell process (C) after treatment with FSH for 9 h. The end of the process contains prominent lysosomes (L). Z, zona pellucida. x 20500; bar = 1 /im.
Fig. 6. Process (C) within the cortical cytoplasm of an oocyte after 12 h LH. Although
a lysosome (L) is present, intermediate junctions (arrows) are intact, x 37000;
bar = 0-5/tm.
358
R. M. MOOR AND OTHERS
j*^'
>*-
Hormones and compartmental change in oocytes
359
corporated into a number of bands (P < 001). The gonadotrophin-induced
changes involved a substantial increase of incorporation into bands 1, 4, 6, 10
and 15 and a substantial reduction of incorporation into bands 3, 5, 8 and 12.
The canonical variate analysis provided a more rigorous examination of the
differences in synthesis between untreated and FSH/LH-treated oocytes. Since
the first two canonical variates accounted for 91-2 % of the intergroup variation,
the two-dimensional plot of these variates in Fig. 12 represents a fairly accurate
picture of the relative position of the two groups. The table of 'differences'
included in the figure shows clearly that the pattern of proteins in oocytes
treated with FSH and LH differed very substantially from that in untreated
oocytes.
Individual gonadotrophins and protein synthesis. The rates of protein change in
oocytes obtained from follicles treated with low concentrations (100 ng ml"1) of
either FSH or LH are shown in Table 2. Direct comparisons between individual
bands did not show the same large differences as seen between untreated and
FSH/LH treated oocytes. Nevertheless the ' F ' ratio statistic indicated particularly marked heterogeneity between the four treatment groups in bands 1, 3,
4 and 6.
The canonical variate analysis showed that oocytes treated with either FSH or
LH formed fairly compact and discrete clusters; there was no overlap with either
the untreated or FSH/LH-treated groups (Fig. 12). Moreover, the table of
'distances' confirmed that both the FSH and LH groups occupied positions
roughly equidistant from the untreated and FSH/LH-treated groups. The
individual gonadotrophins nevertheless altered protein synthesis in substantially
different ways, thereby accounting for the relatively large 'distance' between
the centroids of the FSH and LH groups.
DISCUSSION
This study is based upon the observation that signals generated in the follicle
cells are transmitted to the oocyte and form an integral part of the regulatory
system during maturation. Steroids and cyclic AMP are the best known examples
of these regulatory substances and probably influence the maturation of both
amphibian and mammalian oocytes (Masui, 1967; Mailer & Krebs, 1977; Cho,
FIGURES
7-10
Fig. 7. Plasma membrane of an oocyte after 15 h LH. It has invaginated to form deep
clefts (arrows), x 54500; bar = 0-5/«n.
Fig. 8. Complex surface foldings after LH. Note the close apposition of cortical
granules to the plasma membrane (arrows), x 12000; bar = 1 /tm.
Fig. 9. Surface foldings (arrows) impregnated with lanthanum demonstrating their
discontinuous nature. x7000; bar = 1 /tm.
Fig. 10. Surface invaginations after lanthanum impregnation. The associated cytoplasm contains finely filamentous material (arrows), x 18000; bar = 1 /*m.
360
R. M. M O O R
AND
OTHERS
Molecular weight X 10
Hormones and compartmental change in oocytes
361
Stern & Biggers, 1974; Moor, Polge & Willadsen, 1980). Amongst the other
suggested regulators is a low molecular weight inhibitor protein, secreted by
mammalian granulosa cells which possibly maintains oocytes in the dictyate
state in non-activated follicles (Chang, 1955; Foote & Thibault, 1969; Tsafriri,
Pomerantz & Channing, 1976). It has been our interest to examine the
mechanisms involved in the transmission of such putative signal molecules into
the oocyte (Moor & Smith, 1978; Moor et al, 1980).
The passage of small molecules through permeable junctions in most somatic
cells is thought to regulate and synchronize cell function in many multicellular
systems (reviewed by Loewenstein, 1979). However, it was not until the elegant
experiments of Lawrence, Beers & Gilula (1978) that unequivocal evidence was
obtained to show that hormonally induced signals are rapidly transmitted between somatic cells by contact-dependent mechanisms. That a similar signal
transmission system may operate between follicle cells and oocytes is suggested
by the high degree of coupling that exists between these heterologous cells
(Gi\u\a'et al., 1978; Moor et al., 1980). The experiments described in this paper
were designed to examine further this form of signal transmission and to relate
changes in the transmission system with intracellular events in the oocyte. The
finding that purified FSH,but not LH, selectively depresses intercellular coupling
in oocytes provided the means by which the different relationships have been
investigated. Our present results indicate that FSH levels as low as 50 ng ml" 1
effectively suppress intercellular coupling whilst 10-20 times higher LH levels
are ineffective in this regard. By using relatively low levels (100 ng ml"1) of the
individual gonadotrophins, the problems of cross-contamination with other
hormones has been minimized. The upper limit of contamination of NIH-LH-
S18 with FSH is less than 5 % while the LH contamination of NIH-FSH-S9 is
under 3 %.
The potential interpretative difficulties that could arise in the use of choline
as a marker of intercellular coupling in oocytes have been exhaustively discussed
in earlier papers (Moor et al., 1980). However, it is highly improbable that
difficulties in interpretation have been experienced in this study because the
marker uptake measurements have been confirmed in all cases by the results of
Fig. 11 A. Fluorograph of [35S]methionine-labelled proteins synthesized by oocytes
previously cultured for 18 h within follicles in the presence of the following
gonadotrophins (i) untreated controls, (ii) FSH, 5 fig ml ~x plus LH, 3 /<g ml"1, (iii)
FSH; lOOngml-1 (iv) LH, lOOngml-1. Polypeptide separation was by SDS
gradient acrylamide gel electrophoresis. The 16 marker bands selected for analysis
are indicated and numbered sequentially from the low to high molecular weight
regions.
Fig. 11 B. Densitometer tracers of fluorographs of [35S]methionine-labelled proteins
from oocytes obtained from follicles after 18 h culture in medium devoid of gonadotrophins (
; untreated controls) or in the presence of FSH 5/igml"1 plus LH
1
3 /<g mh (
FSH/LH group). The numbering of the 16 selected marker bands
corresponds to that shown in Fig. 11 a.
362
R. M. MOOR AND OTHERS
--3
LH
Control
FSH + LH
-5
-4
-3
-2
-1
1
2
3
4 •-•
5
—1
FSH
---2
Distance statistic
61
FSH
-
LH
FSH
LH
4-3
61
—
FSH/LH
61
6-9
5-4
Cont.
---3
x -- - 4
—
-L-5
Fig. 12. Analysis of the effect of hormones on protein profiles in oocytes obtained
from follicles cultured without gonadogrophins (untreated controls), with either
FSH or LH alone or with both gonadotrophins (FSH/LH group). The plot represents
the first two canonical variates for 130 oocytes in the four treatments. The table
gives the standardized 'distance' calculated as the Mahalanobis D statistic (Rao,
1952), between the centroids (*) of the treatment groups.
parallel ultrastructural studies. The differential effects of FSH and LH on the
cumulus cell processes and junctions are clearly discernable in the electron
microscope. Exposure to FSH causes rapid and widespread changes both within
the processes and on the membrane of the cell processes. By contrast LH induces a slow activation of lysosomes within some of the processes and does not
induce membrane vesiculation. It is however interesting that LH is far more
effective than FSH at inducing structural and functional changes in the nonjunctional regions of the oocyte membrane (Moor & Smith, 1978).
Our ultrastructural studies indicate that the suppression of functional coupling
between cumulus cells and oocytes by FSH involves two distinct morphological
events, namely degeneration and withdrawal of processes. The degenerative
process is characterized by the localized formation of lysosomal bodies in the
ends of processes which initially remain in junctional contact with the oolemma.
The disappearance of the intermediate junctions and the withdrawal of the
processes occur about 6 h after the initiation of the early degenerative changes.
Tt is possible that the disruption of these intermediate junctions is a necessary
Hormones and compartmental change in oocytes
363
precondition for the dispersal of the corona-radiata cells and the associated
retraction of their transzonal processes. The requirement for junctional disruption could explain why the large mass of cumulus cells undergo dispersal
about 3 h before the corona cells.
Szollosi (1980) has postulated that the disruption of intercellular coupling
in oocytes from the mouse, rabbit, pig and cow provides the stimulus for the
intracellular migration of cortical granules to a position directly beneath the
oolemma. In the sheep, cortical granules move towards the periphery of the
oocyte during follicular growth (Cran, Moor & Hay, 1980). However, between
15 and 18 h after the resumption of meiosis, or 3 to 5 h after the suppression of
cell coupling, the cortical granules in this species also became very closely
aligned beneath the plasma membrane. While a causal relationship between the
two events has still to be determined, it is possible that the disruption of cell
coupling may initiate changes in the cortical cytoplasm of the oocyte which
result in the redistribution of the organelles in this zone.
The protein results show that while both FSH and LH induce substantial
changes in the profile of labelled polypeptides synthesized by oocytes, only FSH
has the capacity to suppress intercellular coupling. Our overall conclusion from
these results is that the majority of changes in protein synthesis are not regulated by changes within the permeable junctions. However, our experiments do
not exclude the possibility that subtle variations in synthesis may be induced
by the disruption of cell coupling during maturation. Difficulties in detecting
and correctly interpreting minor changes in protein synthesis limit the degree to
which this possibility can be fully explored. Variability in synthetic activity
between different oocytes provides the first difficulty in detecting subtle intracellular changes. This variability arises from differences in the hormonal status
of donor animals at ovariectomy, differences in size, degree of differentiation
and extent of atresia of individual follicles and the conditions chosen for the
culture and labelling of the oocytes. For example, the variability in biosynthetic
activity in individual oocytes removed from follicles and cultured in an extrafollicular environment is directly influenced both by hormones in the medium
and by cell interactions with surrounding cumulus cells (I. M. Crosby, J. C.
Osborn & R. M. Moor, unpublished observations), ft is therefore evident that
pooling of oocytes for electrophoretic or other biochemical studies is acceptable
only if cellular homogeneity can first be demonstrated. We have found that
acceptable homogeneity (Fig. 12) can be obtained by using oocytes from intact
cultured follicles which, at explanation, were non-atretic, 3-0 to 5-0 mm in
diameter, and from animals that had been treated previously with a standardised
hormonal regime. Additional factors which limit the capacity to identify subtle
changes in synthesis are the technical requirements of the different protein
separation systems. More homogenous oocytes are required for a valid analysis
of both the basic and acid proteins using the highly sensitive two-dimensional
method of polypeptide separation than in the one-dimensional system. Despite
364
R. M. MOOR AND OTHERS
its reduced sensitivity, we have chosen the single dimensional form of separation
because of its advantages in providing, from very small numbers of oocytes,
quantitative data on both the acid and basic proteins. The problems of making
statistical comparisons between protein profiles containing large numbers of
different bands have been overcome by using the canonical variate analysis.
Dekel & Beers (1978) have proposed a model for oocyte regulation in mammals in which the dictyate state is maintained by elevated levels of cyclic AMP.
This putative meiotic inhibitor, after synthesis in the follicle cells, is thought to
enter the oocyte through contact-dependent mechanisms. The abolition of intercellular coupling between the cumulus cells and oocyte before ovulation would,
according to the model, result in a fall in the intracellular cyclic nucleotide
levels in the oocyte and the consequent resumption of meiosis. The use of low
levels of individual gonadotrophins provides a direct means of testing this model.
It is clear from the results presented in section 1 and 3 of the present paper that
FSH suppresses intercellular coupling between the cumulus cells and oocytes
without inducing nuclear maturation. Conversely, LH induces nuclear maturation in oocytes in which full intercellular coupling with the cumulus cells is
retained. We are therefore unable to support that part of the model which
postulates that the resumption of meiosis in mammalian oocytes is initiated by
the disruption of cell contact between the cumulus cells and oocyte. We presently favour the view that during maturation gonadotrophins act primarily to
alter the nature or intensity of the signals from the follicle cells. Changes in the
intercellular transmission system appear to be secondary to changes in signal
generation.
We thank Mrs Linda Musk and Mr Ian Crosby for valuable assistance in many aspects of this
work. The gonadotrophins were generously donated by the National Institute of Arthritis.
Metabolism and Digestive Diseases, National Institutes of Health, Bethesda, Maryland. One
of us (J.C.O.) is indebted to the Medical Research Council for financial support.
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(Received 25 July 1980, revised 15 September 1980)