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Calmodulin synthesis and accumulation during oogenesis and maturation of Xenopus laevis oocytes
DEVELOPMENTAL
BIOLOGY
113,174-181
(1986)
Calmodulin Synthesis and Accumulation during Oogenesis and Maturation of Xenopus Levis Oocytes F.
MICHAEL Department
of Biological Received
May
CICIRELLI*
Sciences,
AND
L.
Purdue
University,
6, 1985; accepted
in revised
DENNIS
SMITH
West Lafayette, form
August
Indiana
47907
3, 19X5
The calmodulin levels in stage 6 Xenopus oocytes averaged 89 f 24 (SD) ng/oocyte and had largely accumulated by stage 3 of oogenesis. From stage 3 to early stage 6, calmodulin levels did not increase further. However, in large stage 6 oocytes (>1.25 mm diam) calmodulin levels again rose to a level as high as 121 ng/oocyte. Calmodulin levels did not change during the maturation of stage 6 oocytes and the results of measurements on animal and vegetal oocyte halves from control and mature oocytes showed no evidence of a redistribution of calmodulin during maturation. Measurements of calmodulin synthesis in stages 1 and 2 oocytes, stage 4 oocytes, and stage 6 oocytes indicated that calmodulin was being synthesized continuously during oogenesis and that the rate of synthesis increased during oogenesis. In stage 1 and 2 oocytes (combined), the synthesis rate was 3.5 pg/hr/oocyte; in stage 4 oocytes it was 48 pg/hr/oocyte, and in large stage 6 oocytes the rate had increased to 160 pg/hr/oocyte. These changes in the rates of synthesis were discussed as they relate to the pattern of calmodulin accumulation during oogenesis. o 1986 Academic press, IX INTRODUCTION
During the period of oogenesis, growing oocytes synthesize and accumulate a variety of macromolecules for use during embryonic development (reviews Davidson, 1976; Smith and Richter, 1985). Considerable attention has focused on RNA, especially putative mRNA, and it is clear that the various species of RNA are not synthesized and accumulated coordinately. For example in Xenopus oocytes, mRNA accumulation is completed during Dumont (1972) stage 2 (Golden et al., 1980) while rRNA accumulation continues throughout oogenesis (Davidson, 1976). Many diverse proteins also are known to be stored in Xenopus and Rana oocytes for use after fertilization. These include histones (Woodland, 1980), nucleoplasmin (Laskey and Earnshaw, 1980), DNA polymerases (Zierler et al., 1985), RNA polymerases (Roeder, 1974), tubulins (Pestell, 1975), actin (Clark and Merriam, 1977), proteins which bind to snRNAs (Zeller et al., 1983), proteins which bind to 5s RNA (Shastry et al., 1984) and proteins associated with cytoplasmic mRNAs (Darnbrough and Ford, 1981; Richter and Smith, 1983). Less attention has been devoted to the kinetics of synthesis and accumulation of such proteins. Some proteins appear to increase in amount as a function of oocyte size, maintaining the same level on a volume basis. Examples of this include tubulin: tyrosine ligase (Preston et al, 1981), RNA polymerase (Roeder, 1974), and tubulin * Current address: Laboratories, University Wash. 98195. 0012-1606/86
Howard Hughes of Washington
$3.00
Copyright 8 1986 by Academic Press. Inc. All rights of reproduction in any form reserved.
Medical School
Institute, of Medicine,
Research Seattle,
174
(Pestell, 1975). Qualitatively, the majority of proteins identified by electrophoretic techniques are synthesized at all stages of oogenesis, increasing only in amount as growth occurs (Ruderman and Pardue, 1977; Harsa-King et al., 1979), and the same protein species may appear after fertilization. Thus, it would appear that for many proteins, accumulation during oogenesis reflects the maintenance of a constant proportionality, perhaps similar to that found in somatic cells. On the other hand, clearly there are examples of proteins which do not follow this pattern of accumulation. Proteins such as the 5S RNA binding protein (Shastry et al., 1984) and poly(A)+ RNA binding proteins (Darnbrough and Ford, 1981; Richter and Smith, 1983) reach a maximal content in small oocytes and then decrease during oogenesis. Histones are synthesized in growing oocytes, but are reported to show a marked 20- to 50-fold increase during the relatively brief period of oocyte maturation (review by Woodland, 1980); an increase not requiring transcription. The developmental significance of different patterns of protein synthesis and accumulation during oogenesis and oocyte maturation is not clear. To understand this significance, more information concerning the synthesis and accumulation of proteins with known functions is required. This paper is concerned with studies on the synthesis and accumulation of a protein, calmodulin, known to be involved in a variety of cellular control mechanisms (reviews by Means and Dedman, 1980; Cheung, 1980). Furthermore, calmodulin levels are reported to change rather dramatically during the period of oocyte maturation, reaching a value 70% greater than
CICIRELLI
AND
SMITH
in controls (Cartaud et al., 1980). We have examined the accumulation of calmodulin during oogenesis and oocyte maturation, and have measured the rate of calmodulin synthesis during the same times. We find that calmodulin levels do not increase during oocyte maturation. Further, calmodulin shows a pattern of accumulation different than any of the other proteins studied thus far. METHODS
AND
MATERIALS
Animals. Adult female Xenops laevis females were purchased from South Africa (Snake Farm, Fish Hoek, Cape Province) or from Nasco (Fort Atkinson, Wis.). The females were maintained in the laboratory as previously described (Webb et al., 1975) and their ovaries were surgically removed after hypothermic anesthesia. Different stage oocytes were manually defolliculated with watchmaker’s forceps and cultured in OR-2 (Wallace et al., 1973). Alternatively, in some experiments, oocytes were obtained by collagenase digestion. Small ovarian sections were incubated at room temperature in 0.15% collagenase (type II, Sigma) in OR-2 (minus CaC&) with gentle swirling. As oocytes were released, they were removed and washed extensively with regular OR-2. Measurement of calmodulin content. Oocytes, or animal and vegetal halves, were homogenized in Buffer C (10 mM imidazole pH 7.5,lO mM MgClz, 0.15 MNaCl, 1 mM EGTA, 1 mM2-mercaptoethanol; Chafouleas et ah, 1979) and the homogenate was placed in a boiling water bath for 5 min. After heating, the homogenate was cooled on ice for 5 min and centrifuged at 12,000g for 15 min (O5°C). Aliquots (50 ~1) of the supernatant were used for radioimmunoassay (RIA Kit; New England Nuclear). The validity of the RIA procedure was checked by adding increasing amounts of authentic calmodulin to the extract from both control and mature oocytes; the assay accurately measured each addition. For the determination of recovery, propionylated calmodulin (N-propionate-2,3-3H; New England Nuclear) was added at the time of homogenization. The marker was used only if it migrated as a single band on SDSpolyacrylamide gels and was greater than 90% TCA precipitable. To measure radioactivity attributable to the marker, the supernatant (see above) was put on ice, made 1 mg/ml BSA, and 10% TCA, and allowed to precipitate for lo-30 min. The precipitate was collected on Whatman GF/C glass fiber filters and was washed 3 times with 2 ml ice cold 10% TCA. The filters were dried at 6O”C, incubated with 0.5 ml NCS (Amersham) for 2 hr at 60°C and counted in 10 ml PPO/toluene (4 g/liter) after chemiluminescence had disappeared. Measurement of calmodulin synthesis. Rates of calmodulin synthesis were estimated for stage 1 and 2, stage
Calmdulin
during
Oogenvsis
175
4, and stage 6 oocytes. In the first case, approximately 50,000 stage 1 and 2 oocytes were incubated in a dish with L-[3H]leucine (1.6 mCi/2 ml, 55 Ci/mmole) in OR2. Incorporation into the acid soluble pool was determined over a 60-min span by homogenizing groups of oocytes (30 per time point) in 20% TCA and measuring acid soluble radioactivity; radioactivity had reached steady state within 20 min. Pool specific activity was calculated from the steady state level of radioactivity, assuming the leucine pool (average of stage 1 and 2 oocytes) to be 1.7 pmol/oocyte (Taylor and Smith, 1985). The oocytes remaining in the leucine after 1 hr were used to determine radioactivity incorporated into calmodulin. This single incorporation time point was divided by the pool specific activity to estimate the rate of calmodulin synthesis, assuming the weight percentage of leucine in ovarian calmodulin to be 6% (Cartaud et al., 1980). For the determination of calmodulin synthesis in stage 4 and 6 oocytes, 200 oocytes were rapidly injected with [35S]methionine (about 5 X lo6 cpm/51.8 pmole/l5 nl) and allowed to incubate for 1 hr. The endogenous methionine pool size for stage 4 and 6 oocytes is 4 and 12 pmol per oocyte, respectively (Taylor and Smith, unpublished data). Hence, the injected methionine expanded the endogenous pool sufficiently to generate linear incorporation kinetics over the 60-min period (see Shih et al., 1978). The rate of calmodulin synthesis was determined by dividing the radioactivity incorporated into calmodulin by pool specific activity (radioactivity injected divided by expanded pool size) and assuming that the weight percentage methionine in calmodulin is 7.6% (Cartaud et al., 1980). To determine incorporation into calmodulin, oocytes were homogenized in 40 mM Tris pH 7.5, 5 mM EGTA, and the supernatant from centrifugation at 12,000g for 15 min. was brought to 90°C in a microwave oven, then rapidly cooled in an ethanol-ice bath (-10°C). Denatured protein was removed (12,000g for 15 min) and the supernatant was made 6 mMCaC1, and stored frozen overnight. The sample was then thawed, centrifuged again (12,OOOg,15 min) and the supernatant was loaded directly onto a phenothiazine column (Bio Rad, 1 ml vol) equilibrated with binding buffer (0.3 M NaCl, 100 pM CaC&, 1 mM 2-mercaptoethanol, 50 mM Tris, pH 7.5). The column was washed with 4 bed vol of binding buffer and calmodulin was eluted with 10 ml of buffer E (5 mM EGTA, 50 mMTris, pH 7.5). The calmodulin was dialyzed against 3.5 liters of 5 mM NaCl, 4 mM Tris, pH 7.5, twice, lyophilized, and resuspended in 1 ml distilled water. This sample was divided into two parts, each was precipitated by addition of 4 vol of ethanol (-20°C). The ethanol precipitates were collected and dried. The isolated calmodulin was subjected to electropho-
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resis on 15% SDS-polyacrylamide gels (Laemmli, 1970). One pellet (half the original sample) was resuspended in electrophoresis sample buffer containing 4 mMEDTA, while the other pellet was resuspended in sample buffer containing 10 mM CaClz, 1 mM EDTA. After electrophoresis, the gels were stained with Coomassie blue and fluorographed using Enhance (New England Nuclear). Calmodulin bands were either cut out of the gel and oxidized using a Packard Tri-carb oxidizer (3H label) or incubated with NCS in a scintillation vial for 2 hr at 60°C prior to adding scintillation fluid (35Slabel). In separate experiments, tritiated calmodulin prepared from stage 1 and 2 oocytes, as above, was used as a marker for the overall recovery. The marker was added at the time of homogenization and recovery was monitored through the gel electrophoresis step; recovery was less than 10%. RESULTS
Calmodulin
Content during
Oogenesis
To determine the calmodulin content of oocytes at different stages of oogenesis, and during oocyte maturation, we first performed experiments to determine the extraction efficiency of calmodulin from different sized oocytes. The recovery of marker calmodulin, added at the time of homogenization and monitored through the extraction procedure to the point at which the RlA was performed, is shown in Fig. 1. With stage 1 oocytes, or in a procedure carried out with no added oocytes, recovery of marker calmodulin was quite high, averaging about 80% .The 20% loss in these cases is mainly attributed to the 5-min heat treatment during extraction. Recovery clearly had decreased by the time oocytes had progressed to stage 3, and in stage 6 oocytes, recovery
STAGES OF OOGENESIS 100
r ‘12131
“I: 0
.2
.4
OOCYTE
4
.6
.0
DIAMETER
1516
1.0
1.2
1.4
(mm)
FIG. 1. Calmodulin recovery as a function of oocyte size. Groups of 10 oocytes ranging in size from stage 1 to stage 6 were homogenized with added recovery marker and processed as described under Methods and Materials. The best fit through the points was determined by second-order regression analysis.
OOCYTE
10 80 60 40 20 0r.2 .4 .6 .8 11 VOL. (~1)
FIG. 2. Calmodulin levels during oogenesis. Groups of 10 oocytes ranging in size from stage 1 to stage 6 were homogenized and assayed as described under Methods and Materials. The results from 5 different females are shown. In two cases (A and D), average values for stage 6 oocytes are given (KSEM).
of the marker was only 38% .Presumably, this additional loss is due to binding and/or trapping of calmodulin to components which pellet during the initial centrifugation. In this context, the progressive decrease in recovery during oogenesis coincides approximately with the increase in yolk protein during oogenesis (Wallace, 1983), and the yolk protein lipovitellin is known to bind calmodulin (Molla et al., 1983). The actual calmodulin levels measured in oocytes of different stages, corrected for recovery using the data in Fig. 1, are shown in Fig. 2 which shows independent measurements on oocytes from 5 different females. In all cases, calmodulin content reached an apparent plateau by about the completion of stage 3. While there was some variation in this level in oocytes from different females, (42-94 ng/oocyte), the plateau level for any given group of oocytes remained relatively constant until stage 6; oocyte volume increased about lo-fold during this time. In stage 6 oocytes, rather striking differences were observed. In oocytes from 3 of the females (Figs. 2A-C), calmodulin levels had increased dramatically,
CICIRELLI
AND
SMITH
Calnzodulin
reaching values about twofold greater than in stage 5 oocytes. In oocytes from the other 2 females, little or no increase was seen in calmodulin levels in stage 6 versus stage 5 oocytes. Calmodulin
Synthesis during Oogenesis
The observation that levels of calmodulin approach a plateau during stage 3 could be explained by assuming that synthesis of the protein ceases at about this time and the protein remains stable. Alternatively, the plateau could represent a steady state level, maintained by constant rates of synthesis and degradation. The significant increase in calmodulin levels in stage 6 oocytes from some females is more difficult to explain. However, in those cases, in which calmodulin levels were highest, the stage 6 oocytes in those particular females were larger, hence older, than in those instances in which little or no increase in calmodulin levels was seen. Thus, either oocytes would have to increase (or initiate) calmodulin synthesis at a certain time after reaching stage 6, with no change in turnover, or the turnover of calmodulin would have to decrease. To test these possibilities, we carried out experiments to determine the rate 2
during
177
of calmodulin synthesis in oocytes at different stages of oogenesis. As shown for stage 1 and 2 oocytes (Fig. 3), oocytes at all stages examined incorporated radioactive precursors into calmodulin. Radioactive calmodulin appeared as a doublet on the gels, and underwent the characteristic shift in mobility in the presence of calcium. Furthermore, in these “purified” preparations, other radioactive proteins did not appear to seriously contaminate the region of the gel containing calmodulin. Thus, calmodulin bands could be cut out and counted directly. In practice, both bands (with and without calcium) were cut from the gels and the counts in both were combined for rate calculations. When expressed on a volume basis, none of the calculated rates vary by more than threefold between the different stages of oogenesis. However, the rate of calmodulin synthesis expressed on a per oocyte basis in stage 1 and 2 oocytes was 3.5 pg/hr/oocyte, in stage 1 and 2 oocytes (combined), in stage 4 oocytes it was 48 pg/hr/oocyte, and in stage 6 oocytes the rate was 160 pg/hr/oocyte. Clearly, calmodulin is synthesized continuously throughout oogenesis and the rate of synthesis increases rather substantially comparing stage 1 and 2 with stage 6 oocytes. This increase approximately parallels the increase in total protein synthesis (Taylor and Smith, 1985) and therefore requires that increasing amounts of calmodulin mRNA must be recruited for translation during oogenesis. Calmodulin
FIG. 3. Calmodulin synthesis in stage 1 and 2 oocytes. Approximately lO,OOO-20,000 stage 1 and 2 oocytes (combined) were incubated in 2 ml OR-2 containing 1.37 mCi r3H]1eucine (55 Ci/mmole) for 7 hr. Two groups of approximately 20 oocytes were then removed, homogenized in 1 ml of 50 mM NaCl, and centrifuged at 12,OOOg for 15 min. The supernatants were precipitated with 4 vol of ethanol (-20°C overnight), centrifuged for 10 min at 25OOg, and the resulting pellets were dried. One of the pellet residues was resuspended in sample buffer containing 4 mM EDTA (lane l), while the other was resuspended in sample buffer containing 1 mM EDTA and 10 mM CaCl, (lane 2). The remaining oocytes in the incubation dish were homogenized and calmodulin was isolated as described in synthesis (lane 3, calmodulin in sample buffer containing 4 mM EDTA; lane 4, calmodulin in sample buffer containing 1 mM EDTA and 10 mM CaC12). A 15% SDS polyacrylamide gel was run as described under Methods and Materials.
Oqpmxis
Levels during Oocyte Maturation
Cartaud et al. (1980) reported that calmodulin levels increased as much as 70% during oocyte maturation. Since the level of calmodulin in stage 6 oocytes from different females varied from 58 to 121 ng/oocyte in our experiments (Fig. 2), a 70% increase would involve increases of 41 to 85 ng in calmodulin levels during maturation. By the same reasoning, since the total protein synthetic rate increases from about 20 ng/oocyte/hr to 40 ng/oocyte/hr during the approximate 12-hr period of maturation (Wasserman et al., 1982), and the rate increase occurs about midway through maturation, calmodulin synthesis would have to average at least 11% of the average synthetic rate throughout maturation to account for such a dramatic increase. However, neither an increase in calmodulin levels, nor in calmodulin synthesis, was observed in our experiments. In the latter case, our first experiments involved oocytes injected with [3H]leucine and homogenized in 50 mM NaCl; the soluble proteins were electrophoresed on SDS-polyacrylamide gels. Radioactivity in the region of the gel representing calmodulin was less than 0.8% of the total radioactivity for control oocytes and less than 1% of the total for mature oocytes (data not shown). In this par-
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titular experiment, the total rate of protein synthesis in control oocytes was 23.8 ng/oocyte/hr, while the rate in mature oocytes (after GVBD) was 56.2 ng/hr/oocyte. Using these figures, the rate of calmodulin synthesis in control and mature oocytes would be maximally 190 pg/ oocyte/hr and 562 pg/oocyte/hr, respectively. The rate of calmodulin synthesis in control oocytes estimated in this way (without purification) is remarkably similar to that obtained in the experiment in which purified calmodulin was measured (160 pg/oocyte/hr). More importantly, if we assume that calmodulin synthesis during the entire period of maturation (about 12 hr) is the average of the two values listed above (the rate increase occurs about midway during maturation), the predicted increase in calmodulin content of 4.5 ng would be insignificant. The point made above is confirmed directly by measurements of the actual level of calmodulin during maturation. The size of stage 6 oocytes may vary considerably from one female to another, and even among stage 6 oocytes from the same female. Thus, we isolated stage 6 oocytes of approximately the same size from a given female and then normalized the data from different females by expressing paired sets of data from control and progesterone treated oocytes at each time point as percentage of control value. As shown in Fig. 4, the results using oocytes from 3 different females demonstrate that no significant changes in calmodulin levels occur during maturation. It should be pointed out that this result was obtained using oocytes from a female in which calmodulin levels in stage 6 controls had increased minimally compared to stage 5 (Fig. 2D) as well as oocytes in which calmodulin levels at stage 6 were 1.8-fold greater than at stage 5 (Fig. 2A). The results described above are in direct opposition to those reported by Cartaud et al. (1980). The reason for this discrepancy is unclear, but may relate to differences in the assay procedures. Cartaud et al. (1980)
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quantitated calmodulin levels based on the ability of calmodulin to activate myosin light chain kinase in a Ca’+-dependent manner. It is conceivable that modulators of this enzyme activity may have increased (or decreased) in maturing oocytes and interfered with the ability of extracted calmodulin to stimulate the enzyme. Other explanations such as differential recovery of calmodulin from control and maturing oocytes (Wasserman and Smith, 1981) would not seem to be appropriate, since recovery was no different, at least in our hands, when control and mature oocytes were compared (data not shown). Intracellular
Localization
of Calmodulin
Although calmodulin levels do not change during maturation it is conceivable that calmodulin could be localized in the cell and redistribute during maturation. This possibility is suggested by the findings that calmodulin binds to the yolk protein lipovitellin (Molla et ab, 1983) and that yolk platelets are believed to redistribute during maturation, becoming more densely packed in the vegetal hemisphere (Lau et al., 1984). The discovery that calmodulin becomes localized in the mitotic apparatus at metaphase in eukaryotic cells (Welsh et al., 1978) also suggests that an animal-vegetal shift in calmodulin could occur during maturation. To test this possibility, calmodulin levels were measured in the animal and vegetal halves of control and mature oocytes from 4 different females; oocytes frozen on dry ice were cut in half at the midpoint of the animal-vegetal axis. Calmodulin values were corrected for recovery as in previous studies. Calmodulin in the animal halves of control and mature oocytes averaged 63 + 5.6 ng/oocyte (SEM) and 64.1 + 3.5 ng/oocyte, respectively, while the comparable values for vegetal halves were 46.3 f 10.2 and 50.8 + 13.9. Clearly, no major animal-vegetal redistribution of calmodulin occurs during oocyte maturation. DISCUSSION
160
T J
is
120
5
8
-
60
_-
L
IY.
$‘.i 0
60
--
__________
------------
------------
-A
120
100 TIME
240
360
420
(mln)
FIG. 4. Calmodulin levels during oocyte maturation. Oocytes were incubated in 5 ml OR-2 with 1 fig/ml progesterone or 5 ml OR-2 with an equivalent concentration of ethanol carrier (4.3 mM). At various times after the addition of oocytes to the solutions, 10 oocytes per time point were removed and assayed as described under Methods and Materials. The dashed line equals the control value, set at 100%.
During oogenesis the rate of total protein synthesis per oocyte increases from about 0.2 ng/hr in stage 1 oocytes to about 23 ng/hr in stage 6 oocytes (Taylor and Smith, 1985). The rate of calmodulin synthesis averaged less than 1% (0.6-0.8%) of the total protein synthetic rate at all stages of oogenesis tested. Thus, calmodulin is synthesized continuously throughout oogenesis at progressively increasing rates. However, levels of calmodulin clearly do not continue to accumulate throughout oogenesis (Fig. 2). Thus, calmodulin levels must be regulated in part by continually changing rates of degradation. The significance of this conclusion is better discussed in terms of the time required to progress through the various stages of oogenesis.
CICIRELLI
AND
SMITH
Calmodul
In growing females, Callen et al. (1980) have described four growth phases, each with distinctive kinetics. In the first phase, oocytes progress to early stage 1 within a few weeks after metamorphosis and then enter a second phase which may last several months. During this time, late stage l-early stage 2 oocytes appear and this phase is characterized by the synthesis of most, if not all, of the maternal poly(A)+ RNA; calmodulin also accumulates primarily at this time. Oocytes then enter a period of more rapid growth (stages 2-4) coincident with active vitellogenesis and then reach the last phase (stages 5-6) in which growth again is very slow. However, in adult females fewer oocytes representative of the rapid growth phases are apparent. Rather, ovaries contain predominantly stage 5 and 6 oocytes and late stage 1 oocytes which appear to represent a holding stage, perhaps of years duration (Callan et uh, 1980). The stimulus to leave the holding stage is hormonal. The induction of ovulation with gonadotropins not only causes release of stage 6 oocytes from their follicles but also stimulates the growth of smaller oocytes, resulting in the reappearance of stage 6 oocytes (Keem et al., 1979; Wallace, 1983). The rapid growth phase, charaterized by vitellogenesis, is known to be stimulated by estrogen (Wallace, 1983) and follicles during these stages actively synthesize estradiol, while estradiol is no longer synthesized by stage 6 follicles (Fortune, 1983). The possibility exists, therefore, that changes in the level of calmodulin synthesis and degradation throughout oogenesis may be directly (or indirectly) under hormonal control. Considering the above, we suggest the following sequence of events to explain the data (Fig. 2) on calmodulin synthesis and accumulation during oogenesis. In stage 1 and 2 oocytes from the adult females used in our studies, the very low rate of calmodulin synthesis is sufficient merely to maintain the levels of calmodulin synthesized prior to the time at which oocytes entered a holding phase. For example, assuming the 40 ng/oocyte actually measured represents a steady state, the estimated tI12 of the calmodulin would be 330 days, given a synthesis rate of 3.5 pg/oocyte/hr. As oocytes are selected to leave the holding stage, presumably in response to steriod hormones, synthetic activity would be accelerated. According to the data in Fig. 2, oocytes accumulate about 20 ng of additional calmodulin (average of 5 experiments) as they progress through stage 3. We have not measured the rate of synthesis of calmodulin directly in stage 3 oocytes. However, assuming it represents about 1% of total protein synthetic rate, stage 3 oocytes would synthesize calmodulin at a rate of 30 pg/hr. Thus, 28 days would be required to accumulate the additional calmodulin, assuming no turnover. This is less than twice the minimum time estimated by Callen
in during
0oqmesi.s
179
et ul. (1980) for oocytes to progress from stage 2 through stage 3. The data in Fig. 2 indicate that calmodulin levels remain constant at about 60 ng/oocytes through the completion of stages 4 and 5. However, we estimate that calmodulin is synthesized at a rate of 48 pg/oocyte/hr in stage 4 oocytes, and the rate might be expected to increase even further in stage 5 oocytes. Since calmodulin content does not increase, turnover must be occurring in this case. Assuming that the 60-ng level represents a steady state, and using a rate of synthesis of 48 pg/oocyte/hr, the tIjz for calmodulin in stage 4 oocytes would be about 36 days. This would decrease to 13 days in stage 5 oocytes, assuming that calmodulin synthesis continued at about 1% of the total level of protein synthesis. The pattern of calmodulin accumulation discussed above is characterized by continually increasing rates of synthesis during oogenesis matched by continually increasing rates of degradation, at least from stage 3 through stage 5. If this pattern were to continue as oocytes progressed to stage 6, we would predict that degradation would increase even more, since the rate of calmodulin synthesis approximately doubles between stage 5 and stage 6. This may explain the observation that calmodulin levels increase minimally, or not at all, in stage 6 oocytes from 2 of the 5 females examined. However, in the other three cases (Fig. 2) the rate of degradation would either have to decrease at some point after oocytes reach stage 6, the rate of calmodulin synthesis would have to increase even further, or some combination of the two events would have to occur in order to account for the additional calmodulin accumulation. In any case, a switch in the rate of degradation and/or synthesis would be required. Perhaps such a switch could result from a change in the type of steriod hormone synthesized in the stage 6 follicle (Fortune, 1983). At any rate, the idea of such a change in the largest stage 6 oocytes (necessarily the oldest), compared to smaller ones, is not novel. Wallace and Steinhardt (1977) reported a marked difference in membrane potential in “large” versus “small” stage 6 oocytes after removal from their follicles. It is rather surprising that growing oocytes would regulate the amount of an endogenous protein such as calmodulin at a post-translational level rather than at the level of recruitment and initiation of mRNA for translation. However, Dolecki and Smith (1979) suggested a similar metabolic pattern to explain accumulation of poly(A)+ RNA during oogenesis. In that case, synthesis of poly(A)+ RNA was observed throughout oogenesis in spite of the fact that the maximal content of poly(A)+ RNA was reached already in stage 2 oocytes. They also suggested that the constant level of poly(A)+
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DEVELOPMENTAL
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RNA was maintained by continually changing rates of synthesis and degradation during oogenesis. Thus, in both kinds of situations, the kinetics of synthesis and degradation are a reflection of the fact that progress through the stages of oogenesis is not a smooth and continuous process. The pattern of calmodulin accumulation during oogenesis clearly is different from that of other known proteins studied thus far. Since calmodulin is of obvious importance in regulating a variety of cellular functions (Means and Dedman, 1980; Cheung, 1980), one might suggest that the accumulation pattern reflects a fundamental role of the protein in events necessary to oogenesis or early development. If this is so, the nature of those events and the putative role of calmodulin in their regulation remain a mystery. For example, the concentration of calmodulin in stage 6 oocytes, even in those oocytes containing over 120 ng (Fig. 2), would only be about 14 pM, assuming a water volume of about 0.5 ~1 (Cicirelli et al., 1983). This value is not unusually high and does not suggest that oocytes accumulate a store of the protein for use later in development. Viewed differently, the concentration of free calcium in stage 6 oocytes is reported to be 0.14 pM (Robinson, 1985). Assuming this calcium pool is at equilibrium with all four calcium binding sites on calmodulin, only about 0.5% of the calmodulin would exist in the active form. On the other hand, assuming further that the free calcium concentration in stage 2-3 oocytes is the same as at stage 6, the concentration of calmodulin would be greater by several fold (Fig. 2). This suggests the possibility that the concentration of the Ca-calmodulin complex would be greater in small versus large oocytes. Perhaps, then, calmodulin-dependent events would become less important as oogenesis approaches completion.
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CLARK, T. G., and MERRIAM, R. W. (1977). Diffusible and bound actin in nuclei of Xenqrus laevis oocytes. Cell 12,883-891. DARNBROUGH, C., and FORD, P. J. (1981). Identification in Xenv luevis of ovary specific proteins which are bound to messenger RNA. Eur. J. Biochem.
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DAVIDSON, E. H. (1976). “Gene Activity in Early Development,” 2nd ed. New York, Academic Press. DUMONT, J. N. (1972). Oogenesis in Xenopus laevis (Daudin) I. Stages of oocyte development in laboratory maintained animals. J. Mmyhol 136,153-180. FORTUNE, J. E. (1983). Steroid production by Xenoms ovarian follicles at different stages. Dev. Biol. 99, 502-509. GOLDEN, L., SCHAFER, U., and ROSBASH, M. (1980). Accumulation of individual pA +RNAs during oogenesis of Xenopus laevis. Cell 22, 835-844.
HARSA-KING, M. L., BENDER, A., and LODISH, H. F. (1979). Stage specific changes in protein synthesis during Xenoyus oogenesis. Zn “Eucaryotic Gene Regulation. ICN-UCLA Symposia on Molecular and Cellular Biology” (R. Axel, T. Maniatis, C. F. Fox, eds.) Vol. XIV, pp. 239-250. KALT, M. R., and GALL, J. G. (1974). Observations on early germ cell development and premeiotic ribosomal DNA amplification in Xenopus laevis. J. Cell Biol. 62, 460-472. KEEM, K., SMITH, L. D., WALLACE, R. A., and WOLF, D. (1979). Growth rate of oocytes in laboratory-maintained Xenoyus laevis. Gam. Res. 2, 125-135.
LASKEY, R. A. and EARNSHAW, W. C. (1980). Nucleosome assembly. Nature (London) 286, 763-767. LAEMMLI, U. K. (1970). Cleavage of structural proteins during the assembly of the head bacteriophage T4. Nature (London) 227,680-685. LAU, Y. T., REYNHOUT, J. K., and HOROWITZ, S. B. (1984). Regional water changes during oocyte meiotic maturation: evidence of ooplasmic segregation. Dev. BioL 104, 106-110. MEANS, A. R., and DEDMAN, J. R. (1980). Calmodulin-an intracellular calcium receptor. Nuture (London) 285.73-77. MOLLA, A., CARTAUD, A., LAZARO, R., and OZON, R. (1983). Xenojuus lipovitellin, a new target protein for calmodulin. FEBS Letf. 154, 101-104. PELHAM, H. R. B., WORMINGTON, W. M., and BROWN, D. D. (1980). Similar 5s RNA transcription factors in Xenopus oocytes und somatic cells. Proc. Nutl. Acad Sci. USA 78, 1760-1764. PESTELL, R. Q. W. (1975). Microtubule protein synthesis during oogenesis and early embryogenesis in Xenows laevis. Biochem. J. 145, 527-534.
We thank Mrs. Marcia Kremer was supported by a grant from (HD04229) awarded to L.D.S.
for technical the National
assistance. Institutes
This work of Health
REFERENCES CALLEN, J.-C., DENNEBOUY, N., and MOUNOLOU, J. C. (1980a). analysis of entire oogenesis in Xenopus laevis. Dev. Growth
Kinetic Dzfer.
22,831-840.
CARTAUD, A., OZON, R., WALSH, M. P., HAIECH, J., and DEMAILLE, J. G. (1980).Xenom lae-uisoocyte calmodulin in the process of meiotic maturation. J Biol. Chem. 255,9404-9408. CHAFOULEAS, J. G., DEDMAN, J. R., MUNJAAL, R. P., and MEANS, A. R. (1979). Calmodulin. Development and application of a sensitive radioimmunoassay. J. Biol. Chem. 254.10262-10267. CHEUNG, W. Y. (1980). Calmodulin plays a pivotal role in cellular regulation. Science 207, 19-27. CICIRELLI, M. F., ROBINSON, K. R., and SMITH, L. D. (1983). Internal pH of Xenopus oocytes: A study of the Mechanism and role of pH changes during meiotic maturation. Dev. Biol. 100, 133-146.
PRESTON, S. F., DEANIN, G. G., HANSON, R. K., and GORDON, M. W. (1981). Tubulin: Tyrosine ligase in oocytes and embryos of Xeno;uus laevis. Dell. Biol. 81, 36-42. REECK, G. (1976). In “CRC Handbook of Biochemistry and Molecular Biology.” (G. D. Fasman and D. M. Kirschenbaum, eds.), 3rd ed., Vol. III, pp. 505-514. CRC Press, Cleveland, Ohio. RICHTER, J. D., and SMITH, L. D. (1983). Developmentally regulated RNA binding proteins during oogenesis in Xenopus laevis. J. BioL Chem. 258,4864-4869. ROBINSON, K. R. (1985). Maturation of Xenopus oocytes is not accompanied by electrode-detectable calcium changes. Dev. Biol. 109,504508.
ROEDER, R. G. (1974). Multiple forms of deoxyribonucleic acid-dependent ribonucleic acid polymerase in Xenmq laeuis. Levels of activity during oocyte and embryonic development. J. Biol. Chem. 249, 249256.
RUDERMAN, J. V., and PARDUE, M. L. (1977). Cell free translation analysis of messenger RNA in echinoderm and amphibian early development. Dev. Biol. 60, 48-68. SHASTRY, B. S., HONDA, B. M., and ROEDER, R. G. (1984). Altered levels of a 5s gene-specific transcription factor (TFIIIA) during oogenesis
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and embryonic development of Xenoms Inevis. J. Biol. Chem 259, 11,373-11,382. SHIH, R. J., O’CONNOR, C. M., KEEM, K., and SMITH, L. D. (1978). Kinetic analysis of amino acid pools and protein synthesis in amphibian oocytes and embryos. Dev. Biol. 66,1’72-182. SMITH, L. D., and RICHTER, J. D. (1985). Synthesis, accumulation, and utilization of maternal macromolecules during oogenesis and oocyte maturation. 171 “Biology of Fertilization” (C. B. Metz and A. Monroy, eds.), Vol. I., pp. 141-188. Academic Press, New York. TAYI.OR, M. A., and SMITH, L. D. (1985). Q uantitative changes in protein synthesis during oogenesis in Xenows laevis. Dev. Bid. 110, 230237. WALLACE, R. A. (1983). Interactions between somatic cells and the growing oocyte of Xenopus lnevis. In “Current Problems in Germ Cell Differentiation,” British Symp. for Dev. Biol. Symp. 7 (A. Mcharen and C. C. Wylie, eds.), pp. 285-306. Cambridge Univ. Press, London. WALLACE, R. A., JARED, D. W., DUMONT, J. N., and SEGA, M. W. (1973). Protein incorporation by isolated amphibian oocytes. III. Optimum incubation conditions. J. Exp 2001. 18, 321-334. WALI,ACE, R. A., and STEINHARDT, R. A. (1977). Maturation of Xenops
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durin.y
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oocgtes. II. Observations 316. WASSERMAN, W. J., and resumption of meiosis 394.
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potential.
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SMITH, L. D. (1981). Calmodulin triggers the in amphibian oocytes. J. Cell Bid. 89, 389-
WEBB, A. C., LAMARCA, M. J., and SMITH, L. D. (1975). Synthesis of mitochondrial RNA by full-grown and maturing oocytes of Rana pipiens and Xenopus laevis. Dev. Bid. 45, 44-45. WELSH, M. J., DEDMAN, Calcium-dependent paratus of eukaryotic WOODLAND, of Xequus.
J. R., BRINKLEY, B. R., and MEANS, A. R. (1978). regulator protein: Localization in mitotic apcells. Proc. N&l. Acud. Sci. USA 75,1867-1871.
H. R. (1980). FEBS Lett.
Histone 121, l-7.
synthesis
during
the development
ZELLER, R., NYFFENEGGER, T., and DEROBERTIS, E. (1983). Nucleocytoplasmic distribution of snRNPs and stockpiled snRNA binding protein during oogenesis and early development in Xenopus lnevis. Cdl 32, 425-434. ZIERLER, M. K., MARINI, N. J., STOWERS, (1985). Stockpiling of DNA polymerases bryogenesis in the frog, Xe7loFs laevis.
D. J., and BENBOW, R. M. during oogenesis and emJ. Biol. Chrm 260, 974-981.
BIOLOGY
113,174-181
(1986)
Calmodulin Synthesis and Accumulation during Oogenesis and Maturation of Xenopus Levis Oocytes F.
MICHAEL Department
of Biological Received
May
CICIRELLI*
Sciences,
AND
L.
Purdue
University,
6, 1985; accepted
in revised
DENNIS
SMITH
West Lafayette, form
August
Indiana
47907
3, 19X5
The calmodulin levels in stage 6 Xenopus oocytes averaged 89 f 24 (SD) ng/oocyte and had largely accumulated by stage 3 of oogenesis. From stage 3 to early stage 6, calmodulin levels did not increase further. However, in large stage 6 oocytes (>1.25 mm diam) calmodulin levels again rose to a level as high as 121 ng/oocyte. Calmodulin levels did not change during the maturation of stage 6 oocytes and the results of measurements on animal and vegetal oocyte halves from control and mature oocytes showed no evidence of a redistribution of calmodulin during maturation. Measurements of calmodulin synthesis in stages 1 and 2 oocytes, stage 4 oocytes, and stage 6 oocytes indicated that calmodulin was being synthesized continuously during oogenesis and that the rate of synthesis increased during oogenesis. In stage 1 and 2 oocytes (combined), the synthesis rate was 3.5 pg/hr/oocyte; in stage 4 oocytes it was 48 pg/hr/oocyte, and in large stage 6 oocytes the rate had increased to 160 pg/hr/oocyte. These changes in the rates of synthesis were discussed as they relate to the pattern of calmodulin accumulation during oogenesis. o 1986 Academic press, IX INTRODUCTION
During the period of oogenesis, growing oocytes synthesize and accumulate a variety of macromolecules for use during embryonic development (reviews Davidson, 1976; Smith and Richter, 1985). Considerable attention has focused on RNA, especially putative mRNA, and it is clear that the various species of RNA are not synthesized and accumulated coordinately. For example in Xenopus oocytes, mRNA accumulation is completed during Dumont (1972) stage 2 (Golden et al., 1980) while rRNA accumulation continues throughout oogenesis (Davidson, 1976). Many diverse proteins also are known to be stored in Xenopus and Rana oocytes for use after fertilization. These include histones (Woodland, 1980), nucleoplasmin (Laskey and Earnshaw, 1980), DNA polymerases (Zierler et al., 1985), RNA polymerases (Roeder, 1974), tubulins (Pestell, 1975), actin (Clark and Merriam, 1977), proteins which bind to snRNAs (Zeller et al., 1983), proteins which bind to 5s RNA (Shastry et al., 1984) and proteins associated with cytoplasmic mRNAs (Darnbrough and Ford, 1981; Richter and Smith, 1983). Less attention has been devoted to the kinetics of synthesis and accumulation of such proteins. Some proteins appear to increase in amount as a function of oocyte size, maintaining the same level on a volume basis. Examples of this include tubulin: tyrosine ligase (Preston et al, 1981), RNA polymerase (Roeder, 1974), and tubulin * Current address: Laboratories, University Wash. 98195. 0012-1606/86
Howard Hughes of Washington
$3.00
Copyright 8 1986 by Academic Press. Inc. All rights of reproduction in any form reserved.
Medical School
Institute, of Medicine,
Research Seattle,
174
(Pestell, 1975). Qualitatively, the majority of proteins identified by electrophoretic techniques are synthesized at all stages of oogenesis, increasing only in amount as growth occurs (Ruderman and Pardue, 1977; Harsa-King et al., 1979), and the same protein species may appear after fertilization. Thus, it would appear that for many proteins, accumulation during oogenesis reflects the maintenance of a constant proportionality, perhaps similar to that found in somatic cells. On the other hand, clearly there are examples of proteins which do not follow this pattern of accumulation. Proteins such as the 5S RNA binding protein (Shastry et al., 1984) and poly(A)+ RNA binding proteins (Darnbrough and Ford, 1981; Richter and Smith, 1983) reach a maximal content in small oocytes and then decrease during oogenesis. Histones are synthesized in growing oocytes, but are reported to show a marked 20- to 50-fold increase during the relatively brief period of oocyte maturation (review by Woodland, 1980); an increase not requiring transcription. The developmental significance of different patterns of protein synthesis and accumulation during oogenesis and oocyte maturation is not clear. To understand this significance, more information concerning the synthesis and accumulation of proteins with known functions is required. This paper is concerned with studies on the synthesis and accumulation of a protein, calmodulin, known to be involved in a variety of cellular control mechanisms (reviews by Means and Dedman, 1980; Cheung, 1980). Furthermore, calmodulin levels are reported to change rather dramatically during the period of oocyte maturation, reaching a value 70% greater than
CICIRELLI
AND
SMITH
in controls (Cartaud et al., 1980). We have examined the accumulation of calmodulin during oogenesis and oocyte maturation, and have measured the rate of calmodulin synthesis during the same times. We find that calmodulin levels do not increase during oocyte maturation. Further, calmodulin shows a pattern of accumulation different than any of the other proteins studied thus far. METHODS
AND
MATERIALS
Animals. Adult female Xenops laevis females were purchased from South Africa (Snake Farm, Fish Hoek, Cape Province) or from Nasco (Fort Atkinson, Wis.). The females were maintained in the laboratory as previously described (Webb et al., 1975) and their ovaries were surgically removed after hypothermic anesthesia. Different stage oocytes were manually defolliculated with watchmaker’s forceps and cultured in OR-2 (Wallace et al., 1973). Alternatively, in some experiments, oocytes were obtained by collagenase digestion. Small ovarian sections were incubated at room temperature in 0.15% collagenase (type II, Sigma) in OR-2 (minus CaC&) with gentle swirling. As oocytes were released, they were removed and washed extensively with regular OR-2. Measurement of calmodulin content. Oocytes, or animal and vegetal halves, were homogenized in Buffer C (10 mM imidazole pH 7.5,lO mM MgClz, 0.15 MNaCl, 1 mM EGTA, 1 mM2-mercaptoethanol; Chafouleas et ah, 1979) and the homogenate was placed in a boiling water bath for 5 min. After heating, the homogenate was cooled on ice for 5 min and centrifuged at 12,000g for 15 min (O5°C). Aliquots (50 ~1) of the supernatant were used for radioimmunoassay (RIA Kit; New England Nuclear). The validity of the RIA procedure was checked by adding increasing amounts of authentic calmodulin to the extract from both control and mature oocytes; the assay accurately measured each addition. For the determination of recovery, propionylated calmodulin (N-propionate-2,3-3H; New England Nuclear) was added at the time of homogenization. The marker was used only if it migrated as a single band on SDSpolyacrylamide gels and was greater than 90% TCA precipitable. To measure radioactivity attributable to the marker, the supernatant (see above) was put on ice, made 1 mg/ml BSA, and 10% TCA, and allowed to precipitate for lo-30 min. The precipitate was collected on Whatman GF/C glass fiber filters and was washed 3 times with 2 ml ice cold 10% TCA. The filters were dried at 6O”C, incubated with 0.5 ml NCS (Amersham) for 2 hr at 60°C and counted in 10 ml PPO/toluene (4 g/liter) after chemiluminescence had disappeared. Measurement of calmodulin synthesis. Rates of calmodulin synthesis were estimated for stage 1 and 2, stage
Calmdulin
during
Oogenvsis
175
4, and stage 6 oocytes. In the first case, approximately 50,000 stage 1 and 2 oocytes were incubated in a dish with L-[3H]leucine (1.6 mCi/2 ml, 55 Ci/mmole) in OR2. Incorporation into the acid soluble pool was determined over a 60-min span by homogenizing groups of oocytes (30 per time point) in 20% TCA and measuring acid soluble radioactivity; radioactivity had reached steady state within 20 min. Pool specific activity was calculated from the steady state level of radioactivity, assuming the leucine pool (average of stage 1 and 2 oocytes) to be 1.7 pmol/oocyte (Taylor and Smith, 1985). The oocytes remaining in the leucine after 1 hr were used to determine radioactivity incorporated into calmodulin. This single incorporation time point was divided by the pool specific activity to estimate the rate of calmodulin synthesis, assuming the weight percentage of leucine in ovarian calmodulin to be 6% (Cartaud et al., 1980). For the determination of calmodulin synthesis in stage 4 and 6 oocytes, 200 oocytes were rapidly injected with [35S]methionine (about 5 X lo6 cpm/51.8 pmole/l5 nl) and allowed to incubate for 1 hr. The endogenous methionine pool size for stage 4 and 6 oocytes is 4 and 12 pmol per oocyte, respectively (Taylor and Smith, unpublished data). Hence, the injected methionine expanded the endogenous pool sufficiently to generate linear incorporation kinetics over the 60-min period (see Shih et al., 1978). The rate of calmodulin synthesis was determined by dividing the radioactivity incorporated into calmodulin by pool specific activity (radioactivity injected divided by expanded pool size) and assuming that the weight percentage methionine in calmodulin is 7.6% (Cartaud et al., 1980). To determine incorporation into calmodulin, oocytes were homogenized in 40 mM Tris pH 7.5, 5 mM EGTA, and the supernatant from centrifugation at 12,000g for 15 min. was brought to 90°C in a microwave oven, then rapidly cooled in an ethanol-ice bath (-10°C). Denatured protein was removed (12,000g for 15 min) and the supernatant was made 6 mMCaC1, and stored frozen overnight. The sample was then thawed, centrifuged again (12,OOOg,15 min) and the supernatant was loaded directly onto a phenothiazine column (Bio Rad, 1 ml vol) equilibrated with binding buffer (0.3 M NaCl, 100 pM CaC&, 1 mM 2-mercaptoethanol, 50 mM Tris, pH 7.5). The column was washed with 4 bed vol of binding buffer and calmodulin was eluted with 10 ml of buffer E (5 mM EGTA, 50 mMTris, pH 7.5). The calmodulin was dialyzed against 3.5 liters of 5 mM NaCl, 4 mM Tris, pH 7.5, twice, lyophilized, and resuspended in 1 ml distilled water. This sample was divided into two parts, each was precipitated by addition of 4 vol of ethanol (-20°C). The ethanol precipitates were collected and dried. The isolated calmodulin was subjected to electropho-
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113. 1986
resis on 15% SDS-polyacrylamide gels (Laemmli, 1970). One pellet (half the original sample) was resuspended in electrophoresis sample buffer containing 4 mMEDTA, while the other pellet was resuspended in sample buffer containing 10 mM CaClz, 1 mM EDTA. After electrophoresis, the gels were stained with Coomassie blue and fluorographed using Enhance (New England Nuclear). Calmodulin bands were either cut out of the gel and oxidized using a Packard Tri-carb oxidizer (3H label) or incubated with NCS in a scintillation vial for 2 hr at 60°C prior to adding scintillation fluid (35Slabel). In separate experiments, tritiated calmodulin prepared from stage 1 and 2 oocytes, as above, was used as a marker for the overall recovery. The marker was added at the time of homogenization and recovery was monitored through the gel electrophoresis step; recovery was less than 10%. RESULTS
Calmodulin
Content during
Oogenesis
To determine the calmodulin content of oocytes at different stages of oogenesis, and during oocyte maturation, we first performed experiments to determine the extraction efficiency of calmodulin from different sized oocytes. The recovery of marker calmodulin, added at the time of homogenization and monitored through the extraction procedure to the point at which the RlA was performed, is shown in Fig. 1. With stage 1 oocytes, or in a procedure carried out with no added oocytes, recovery of marker calmodulin was quite high, averaging about 80% .The 20% loss in these cases is mainly attributed to the 5-min heat treatment during extraction. Recovery clearly had decreased by the time oocytes had progressed to stage 3, and in stage 6 oocytes, recovery
STAGES OF OOGENESIS 100
r ‘12131
“I: 0
.2
.4
OOCYTE
4
.6
.0
DIAMETER
1516
1.0
1.2
1.4
(mm)
FIG. 1. Calmodulin recovery as a function of oocyte size. Groups of 10 oocytes ranging in size from stage 1 to stage 6 were homogenized with added recovery marker and processed as described under Methods and Materials. The best fit through the points was determined by second-order regression analysis.
OOCYTE
10 80 60 40 20 0r.2 .4 .6 .8 11 VOL. (~1)
FIG. 2. Calmodulin levels during oogenesis. Groups of 10 oocytes ranging in size from stage 1 to stage 6 were homogenized and assayed as described under Methods and Materials. The results from 5 different females are shown. In two cases (A and D), average values for stage 6 oocytes are given (KSEM).
of the marker was only 38% .Presumably, this additional loss is due to binding and/or trapping of calmodulin to components which pellet during the initial centrifugation. In this context, the progressive decrease in recovery during oogenesis coincides approximately with the increase in yolk protein during oogenesis (Wallace, 1983), and the yolk protein lipovitellin is known to bind calmodulin (Molla et al., 1983). The actual calmodulin levels measured in oocytes of different stages, corrected for recovery using the data in Fig. 1, are shown in Fig. 2 which shows independent measurements on oocytes from 5 different females. In all cases, calmodulin content reached an apparent plateau by about the completion of stage 3. While there was some variation in this level in oocytes from different females, (42-94 ng/oocyte), the plateau level for any given group of oocytes remained relatively constant until stage 6; oocyte volume increased about lo-fold during this time. In stage 6 oocytes, rather striking differences were observed. In oocytes from 3 of the females (Figs. 2A-C), calmodulin levels had increased dramatically,
CICIRELLI
AND
SMITH
Calnzodulin
reaching values about twofold greater than in stage 5 oocytes. In oocytes from the other 2 females, little or no increase was seen in calmodulin levels in stage 6 versus stage 5 oocytes. Calmodulin
Synthesis during Oogenesis
The observation that levels of calmodulin approach a plateau during stage 3 could be explained by assuming that synthesis of the protein ceases at about this time and the protein remains stable. Alternatively, the plateau could represent a steady state level, maintained by constant rates of synthesis and degradation. The significant increase in calmodulin levels in stage 6 oocytes from some females is more difficult to explain. However, in those cases, in which calmodulin levels were highest, the stage 6 oocytes in those particular females were larger, hence older, than in those instances in which little or no increase in calmodulin levels was seen. Thus, either oocytes would have to increase (or initiate) calmodulin synthesis at a certain time after reaching stage 6, with no change in turnover, or the turnover of calmodulin would have to decrease. To test these possibilities, we carried out experiments to determine the rate 2
during
177
of calmodulin synthesis in oocytes at different stages of oogenesis. As shown for stage 1 and 2 oocytes (Fig. 3), oocytes at all stages examined incorporated radioactive precursors into calmodulin. Radioactive calmodulin appeared as a doublet on the gels, and underwent the characteristic shift in mobility in the presence of calcium. Furthermore, in these “purified” preparations, other radioactive proteins did not appear to seriously contaminate the region of the gel containing calmodulin. Thus, calmodulin bands could be cut out and counted directly. In practice, both bands (with and without calcium) were cut from the gels and the counts in both were combined for rate calculations. When expressed on a volume basis, none of the calculated rates vary by more than threefold between the different stages of oogenesis. However, the rate of calmodulin synthesis expressed on a per oocyte basis in stage 1 and 2 oocytes was 3.5 pg/hr/oocyte, in stage 1 and 2 oocytes (combined), in stage 4 oocytes it was 48 pg/hr/oocyte, and in stage 6 oocytes the rate was 160 pg/hr/oocyte. Clearly, calmodulin is synthesized continuously throughout oogenesis and the rate of synthesis increases rather substantially comparing stage 1 and 2 with stage 6 oocytes. This increase approximately parallels the increase in total protein synthesis (Taylor and Smith, 1985) and therefore requires that increasing amounts of calmodulin mRNA must be recruited for translation during oogenesis. Calmodulin
FIG. 3. Calmodulin synthesis in stage 1 and 2 oocytes. Approximately lO,OOO-20,000 stage 1 and 2 oocytes (combined) were incubated in 2 ml OR-2 containing 1.37 mCi r3H]1eucine (55 Ci/mmole) for 7 hr. Two groups of approximately 20 oocytes were then removed, homogenized in 1 ml of 50 mM NaCl, and centrifuged at 12,OOOg for 15 min. The supernatants were precipitated with 4 vol of ethanol (-20°C overnight), centrifuged for 10 min at 25OOg, and the resulting pellets were dried. One of the pellet residues was resuspended in sample buffer containing 4 mM EDTA (lane l), while the other was resuspended in sample buffer containing 1 mM EDTA and 10 mM CaCl, (lane 2). The remaining oocytes in the incubation dish were homogenized and calmodulin was isolated as described in synthesis (lane 3, calmodulin in sample buffer containing 4 mM EDTA; lane 4, calmodulin in sample buffer containing 1 mM EDTA and 10 mM CaC12). A 15% SDS polyacrylamide gel was run as described under Methods and Materials.
Oqpmxis
Levels during Oocyte Maturation
Cartaud et al. (1980) reported that calmodulin levels increased as much as 70% during oocyte maturation. Since the level of calmodulin in stage 6 oocytes from different females varied from 58 to 121 ng/oocyte in our experiments (Fig. 2), a 70% increase would involve increases of 41 to 85 ng in calmodulin levels during maturation. By the same reasoning, since the total protein synthetic rate increases from about 20 ng/oocyte/hr to 40 ng/oocyte/hr during the approximate 12-hr period of maturation (Wasserman et al., 1982), and the rate increase occurs about midway through maturation, calmodulin synthesis would have to average at least 11% of the average synthetic rate throughout maturation to account for such a dramatic increase. However, neither an increase in calmodulin levels, nor in calmodulin synthesis, was observed in our experiments. In the latter case, our first experiments involved oocytes injected with [3H]leucine and homogenized in 50 mM NaCl; the soluble proteins were electrophoresed on SDS-polyacrylamide gels. Radioactivity in the region of the gel representing calmodulin was less than 0.8% of the total radioactivity for control oocytes and less than 1% of the total for mature oocytes (data not shown). In this par-
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titular experiment, the total rate of protein synthesis in control oocytes was 23.8 ng/oocyte/hr, while the rate in mature oocytes (after GVBD) was 56.2 ng/hr/oocyte. Using these figures, the rate of calmodulin synthesis in control and mature oocytes would be maximally 190 pg/ oocyte/hr and 562 pg/oocyte/hr, respectively. The rate of calmodulin synthesis in control oocytes estimated in this way (without purification) is remarkably similar to that obtained in the experiment in which purified calmodulin was measured (160 pg/oocyte/hr). More importantly, if we assume that calmodulin synthesis during the entire period of maturation (about 12 hr) is the average of the two values listed above (the rate increase occurs about midway during maturation), the predicted increase in calmodulin content of 4.5 ng would be insignificant. The point made above is confirmed directly by measurements of the actual level of calmodulin during maturation. The size of stage 6 oocytes may vary considerably from one female to another, and even among stage 6 oocytes from the same female. Thus, we isolated stage 6 oocytes of approximately the same size from a given female and then normalized the data from different females by expressing paired sets of data from control and progesterone treated oocytes at each time point as percentage of control value. As shown in Fig. 4, the results using oocytes from 3 different females demonstrate that no significant changes in calmodulin levels occur during maturation. It should be pointed out that this result was obtained using oocytes from a female in which calmodulin levels in stage 6 controls had increased minimally compared to stage 5 (Fig. 2D) as well as oocytes in which calmodulin levels at stage 6 were 1.8-fold greater than at stage 5 (Fig. 2A). The results described above are in direct opposition to those reported by Cartaud et al. (1980). The reason for this discrepancy is unclear, but may relate to differences in the assay procedures. Cartaud et al. (1980)
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113. 1986
quantitated calmodulin levels based on the ability of calmodulin to activate myosin light chain kinase in a Ca’+-dependent manner. It is conceivable that modulators of this enzyme activity may have increased (or decreased) in maturing oocytes and interfered with the ability of extracted calmodulin to stimulate the enzyme. Other explanations such as differential recovery of calmodulin from control and maturing oocytes (Wasserman and Smith, 1981) would not seem to be appropriate, since recovery was no different, at least in our hands, when control and mature oocytes were compared (data not shown). Intracellular
Localization
of Calmodulin
Although calmodulin levels do not change during maturation it is conceivable that calmodulin could be localized in the cell and redistribute during maturation. This possibility is suggested by the findings that calmodulin binds to the yolk protein lipovitellin (Molla et ab, 1983) and that yolk platelets are believed to redistribute during maturation, becoming more densely packed in the vegetal hemisphere (Lau et al., 1984). The discovery that calmodulin becomes localized in the mitotic apparatus at metaphase in eukaryotic cells (Welsh et al., 1978) also suggests that an animal-vegetal shift in calmodulin could occur during maturation. To test this possibility, calmodulin levels were measured in the animal and vegetal halves of control and mature oocytes from 4 different females; oocytes frozen on dry ice were cut in half at the midpoint of the animal-vegetal axis. Calmodulin values were corrected for recovery as in previous studies. Calmodulin in the animal halves of control and mature oocytes averaged 63 + 5.6 ng/oocyte (SEM) and 64.1 + 3.5 ng/oocyte, respectively, while the comparable values for vegetal halves were 46.3 f 10.2 and 50.8 + 13.9. Clearly, no major animal-vegetal redistribution of calmodulin occurs during oocyte maturation. DISCUSSION
160
T J
is
120
5
8
-
60
_-
L
IY.
$‘.i 0
60
--
__________
------------
------------
-A
120
100 TIME
240
360
420
(mln)
FIG. 4. Calmodulin levels during oocyte maturation. Oocytes were incubated in 5 ml OR-2 with 1 fig/ml progesterone or 5 ml OR-2 with an equivalent concentration of ethanol carrier (4.3 mM). At various times after the addition of oocytes to the solutions, 10 oocytes per time point were removed and assayed as described under Methods and Materials. The dashed line equals the control value, set at 100%.
During oogenesis the rate of total protein synthesis per oocyte increases from about 0.2 ng/hr in stage 1 oocytes to about 23 ng/hr in stage 6 oocytes (Taylor and Smith, 1985). The rate of calmodulin synthesis averaged less than 1% (0.6-0.8%) of the total protein synthetic rate at all stages of oogenesis tested. Thus, calmodulin is synthesized continuously throughout oogenesis at progressively increasing rates. However, levels of calmodulin clearly do not continue to accumulate throughout oogenesis (Fig. 2). Thus, calmodulin levels must be regulated in part by continually changing rates of degradation. The significance of this conclusion is better discussed in terms of the time required to progress through the various stages of oogenesis.
CICIRELLI
AND
SMITH
Calmodul
In growing females, Callen et al. (1980) have described four growth phases, each with distinctive kinetics. In the first phase, oocytes progress to early stage 1 within a few weeks after metamorphosis and then enter a second phase which may last several months. During this time, late stage l-early stage 2 oocytes appear and this phase is characterized by the synthesis of most, if not all, of the maternal poly(A)+ RNA; calmodulin also accumulates primarily at this time. Oocytes then enter a period of more rapid growth (stages 2-4) coincident with active vitellogenesis and then reach the last phase (stages 5-6) in which growth again is very slow. However, in adult females fewer oocytes representative of the rapid growth phases are apparent. Rather, ovaries contain predominantly stage 5 and 6 oocytes and late stage 1 oocytes which appear to represent a holding stage, perhaps of years duration (Callan et uh, 1980). The stimulus to leave the holding stage is hormonal. The induction of ovulation with gonadotropins not only causes release of stage 6 oocytes from their follicles but also stimulates the growth of smaller oocytes, resulting in the reappearance of stage 6 oocytes (Keem et al., 1979; Wallace, 1983). The rapid growth phase, charaterized by vitellogenesis, is known to be stimulated by estrogen (Wallace, 1983) and follicles during these stages actively synthesize estradiol, while estradiol is no longer synthesized by stage 6 follicles (Fortune, 1983). The possibility exists, therefore, that changes in the level of calmodulin synthesis and degradation throughout oogenesis may be directly (or indirectly) under hormonal control. Considering the above, we suggest the following sequence of events to explain the data (Fig. 2) on calmodulin synthesis and accumulation during oogenesis. In stage 1 and 2 oocytes from the adult females used in our studies, the very low rate of calmodulin synthesis is sufficient merely to maintain the levels of calmodulin synthesized prior to the time at which oocytes entered a holding phase. For example, assuming the 40 ng/oocyte actually measured represents a steady state, the estimated tI12 of the calmodulin would be 330 days, given a synthesis rate of 3.5 pg/oocyte/hr. As oocytes are selected to leave the holding stage, presumably in response to steriod hormones, synthetic activity would be accelerated. According to the data in Fig. 2, oocytes accumulate about 20 ng of additional calmodulin (average of 5 experiments) as they progress through stage 3. We have not measured the rate of synthesis of calmodulin directly in stage 3 oocytes. However, assuming it represents about 1% of total protein synthetic rate, stage 3 oocytes would synthesize calmodulin at a rate of 30 pg/hr. Thus, 28 days would be required to accumulate the additional calmodulin, assuming no turnover. This is less than twice the minimum time estimated by Callen
in during
0oqmesi.s
179
et ul. (1980) for oocytes to progress from stage 2 through stage 3. The data in Fig. 2 indicate that calmodulin levels remain constant at about 60 ng/oocytes through the completion of stages 4 and 5. However, we estimate that calmodulin is synthesized at a rate of 48 pg/oocyte/hr in stage 4 oocytes, and the rate might be expected to increase even further in stage 5 oocytes. Since calmodulin content does not increase, turnover must be occurring in this case. Assuming that the 60-ng level represents a steady state, and using a rate of synthesis of 48 pg/oocyte/hr, the tIjz for calmodulin in stage 4 oocytes would be about 36 days. This would decrease to 13 days in stage 5 oocytes, assuming that calmodulin synthesis continued at about 1% of the total level of protein synthesis. The pattern of calmodulin accumulation discussed above is characterized by continually increasing rates of synthesis during oogenesis matched by continually increasing rates of degradation, at least from stage 3 through stage 5. If this pattern were to continue as oocytes progressed to stage 6, we would predict that degradation would increase even more, since the rate of calmodulin synthesis approximately doubles between stage 5 and stage 6. This may explain the observation that calmodulin levels increase minimally, or not at all, in stage 6 oocytes from 2 of the 5 females examined. However, in the other three cases (Fig. 2) the rate of degradation would either have to decrease at some point after oocytes reach stage 6, the rate of calmodulin synthesis would have to increase even further, or some combination of the two events would have to occur in order to account for the additional calmodulin accumulation. In any case, a switch in the rate of degradation and/or synthesis would be required. Perhaps such a switch could result from a change in the type of steriod hormone synthesized in the stage 6 follicle (Fortune, 1983). At any rate, the idea of such a change in the largest stage 6 oocytes (necessarily the oldest), compared to smaller ones, is not novel. Wallace and Steinhardt (1977) reported a marked difference in membrane potential in “large” versus “small” stage 6 oocytes after removal from their follicles. It is rather surprising that growing oocytes would regulate the amount of an endogenous protein such as calmodulin at a post-translational level rather than at the level of recruitment and initiation of mRNA for translation. However, Dolecki and Smith (1979) suggested a similar metabolic pattern to explain accumulation of poly(A)+ RNA during oogenesis. In that case, synthesis of poly(A)+ RNA was observed throughout oogenesis in spite of the fact that the maximal content of poly(A)+ RNA was reached already in stage 2 oocytes. They also suggested that the constant level of poly(A)+
180
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BIOLOGY
RNA was maintained by continually changing rates of synthesis and degradation during oogenesis. Thus, in both kinds of situations, the kinetics of synthesis and degradation are a reflection of the fact that progress through the stages of oogenesis is not a smooth and continuous process. The pattern of calmodulin accumulation during oogenesis clearly is different from that of other known proteins studied thus far. Since calmodulin is of obvious importance in regulating a variety of cellular functions (Means and Dedman, 1980; Cheung, 1980), one might suggest that the accumulation pattern reflects a fundamental role of the protein in events necessary to oogenesis or early development. If this is so, the nature of those events and the putative role of calmodulin in their regulation remain a mystery. For example, the concentration of calmodulin in stage 6 oocytes, even in those oocytes containing over 120 ng (Fig. 2), would only be about 14 pM, assuming a water volume of about 0.5 ~1 (Cicirelli et al., 1983). This value is not unusually high and does not suggest that oocytes accumulate a store of the protein for use later in development. Viewed differently, the concentration of free calcium in stage 6 oocytes is reported to be 0.14 pM (Robinson, 1985). Assuming this calcium pool is at equilibrium with all four calcium binding sites on calmodulin, only about 0.5% of the calmodulin would exist in the active form. On the other hand, assuming further that the free calcium concentration in stage 2-3 oocytes is the same as at stage 6, the concentration of calmodulin would be greater by several fold (Fig. 2). This suggests the possibility that the concentration of the Ca-calmodulin complex would be greater in small versus large oocytes. Perhaps, then, calmodulin-dependent events would become less important as oogenesis approaches completion.
VOLUME
113, 1986
CLARK, T. G., and MERRIAM, R. W. (1977). Diffusible and bound actin in nuclei of Xenqrus laevis oocytes. Cell 12,883-891. DARNBROUGH, C., and FORD, P. J. (1981). Identification in Xenv luevis of ovary specific proteins which are bound to messenger RNA. Eur. J. Biochem.
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We thank Mrs. Marcia Kremer was supported by a grant from (HD04229) awarded to L.D.S.
for technical the National
assistance. Institutes
This work of Health
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