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research-article |
Program in Cellular, Molecular and Developmental Biology,3 Tufts University School of Medicine, Boston, Massachusetts 02111
Department of Molecular and Integrative Physiology,4 The Center for Reproductive Sciences,5 University of Kansas, Kansas City, Kansas 66160
Marine Biological Laboratory,6 Woods Hole, Massachusetts 02543
ABSTRACT
In oocytes, asymmetric cytokinesis represents a conserved strategy for karyokinesis during meiosis to retain ooplasmic maternal factors needed after fertilization. Given the role of gamma-tubulin in cell cycle progression and microtubule dynamics, this study focused on gamma-tubulin as a key regulator of asymmetric cytokinesis in mouse oocytes. Gamma-tubulin properties were studied using multiple-label digital imaging, Western blots, quantitative RT-PCR, and microinjection strategies in mouse oocytes matured in vivo (IVO) or in vitro (IVM). Quantitative image analysis established that IVO oocytes extrude smaller first polar bodies (PBs), contain smaller spindles, and have more cytoplasmic microtubule organizing centers (MTOCs) relative to IVM oocytes. Maturation in culture was shown to alter gamma-tubulin distribution, as evidenced by incorporation throughout the meiotic spindle and within the first PB. Western blot analysis confirmed that total gamma-tubulin content remained elevated in IVM oocytes compared with IVO oocytes. Analysis of gamma-tubulin mRNA during maturation revealed fluctuations in IVO oocytes, whereas IVM oocytes maintained relatively stable at lower levels for the time points examined (0–16 h). Selective reduction of gamma-tubulin mRNA by injection of siRNA diminished both spindle and PB size, whereas overexpression of enhanced green fluorescent protein gamma-tubulin had the opposite effect. Together, these studies reinforce the notion that limiting gamma-tubulin availability during meiotic maturation ensures coordination of karyokinesis and cytokinesis and conservation of gamma-tubulin as an embryonic reserve.
cumulus cells,, follicle-stimulating hormone, meiosis, oocyte development, ovum
Animal oocytes typically invoke asymmetric cytokinesis during extrusion of the first and second polar bodies (PBs) as they exit the first and second meiotic metaphases [1]. This extreme form of cytokinesis allows female germ cells to complete successive rounds of reductive karyokinesis without significantly misappropriating oocyte cytoplasmic contents. Hence, maternal resources are conserved for embryogenesis. In addition, meiotic spindle position marks the animal pole of the embryo in many species, and this specialized cortical domain may be key to the establishment of asymmetry in animal eggs exhibiting either determinative or regulative types of development [2, 3]. Delineating oocyte factors that regulate PB size and position during meiotic maturation is therefore needed to understand those events during oogenesis and ovulation that contribute to embryonic viability and developmental competence.
Among these factors, Longo and Chen [4] showed in mouse oocytes that the emergence of the microvillar-free domain of the oocyte cortex overlying the spindle was dependent upon actin-based cortical migration in the first meiotic spindle. Maro et al. [5] further demonstrated that remodeling of the actin cortex, which results in microvillar-free domains, is induced by chromatin resident factors in the absence of an intact spindle a finding reinforced recently by the demonstration of cortical reorganization after injection of condensed sperm heads into mouse oocytes [6, 7]. The importance of spindle/chromatin approximation to the oocyte cortex also was revealed by the phenotype of Fmn2–/– mice. These studies showed that failure to dock the first meiotic spindle at the cortex resulted in the absence of cytokinesis, even though karyokinesis proceeded during meiotic progression, yielding abnormal aneuploid embryos [8]. Thus, while chromatin locally influences cortical cytoskeletal remodeling for PB extrusion, other components within the meiotic spindle are likely required to effect the initiation and completion of asymmetric cytokinesis [2, 9].
In mitotic cells, the spindle harbors factors that mediate spindle positioning and cleavage furrow location [10]. For example, astral fiber growth at anaphase onset and microtubule plus end interaction with the cell cortex aid in centralization of the mitotic apparatus in cells that execute symmetric forms of cytokinesis [11, 12]. In Caenorhabditis elegans zygotes, asymmetries in spindle pole components and imbalanced patterns of astral fiber growth contribute to an asymmetric division plane [13]. Whether such asymmetries exist in the poles of oocyte meiotic spindles is not fully resolved, although some reports have suggested this with respect to the distribution of pericentrin, a key component of microtubule organizing centers (MTOCs) [14]. Typically, meiotic spindles of mouse oocytes are anastral, although extremes in the degree of spindle pole tapering or minus end focusing vary widely between species [3] and with respect to the conditions under which meiotic maturation occurs [15]. Moreover, cycles of astral fiber growth at anaphase I and II onset have been reported [16–18]. It is interesting to note that cultured mouse oocytes typically display large barrel-shaped anastral spindles that are more pronounced after various treatments [15, 19]. The significance of deviations in meiotic spindle shape and size with respect to the initiation and completion of PB formation has not been fully explored, even though reports of unusually large PBs in mouse oocytes displaying atypical spindles have been noted in certain strains such as LT/Sv [20], or Mos–/– and Fmn2–/– mice [8, 21]. Several studies have described patterns of 
-tubulin throughout maturation and early embryonic development in mice [18, 22–25]. However, few studies have directly addressed the relationship between microtubule remodeling and asymmetric cytokinesis during the course of maturation in vivo. Our studies in mice have suggested that the redistribution of -tubulin is regulated in a cell cycle stage-specific fashion [18], and further that the spatial allocation of -tubulin between the spindle and MTOCs is a determinant of spindle size [14, 15, 17, 26]. The recent findings of Yuba-Kubo et al. indicate that deletion of maternal -tubulin arrests early mouse development at the morulae stage and provide compelling evidence for this microtubule regulator in the expression of developmental competence [27]. Building on this evidence, our present results show that the patterns and levels of -tubulin expression directly influence PB position, size, and content. These results reinforce the idea that chromatin-based remodeling in the mouse oocyte cortex involves a balanced spatial segregation and conservation of -tubulin to specific subcellular locations that ensure that the mouse embryo inherits adequate stores of -tubulin.
Collection and Maturation of Oocytes
All experiments were performed using 21-day-old CF-1 mice (Charles River Laboratories). Animals were handled according to the Guide for Care and Use of Laboratory Animals (National Academy of Sciences, 1996) and were maintained in a 14L:10D photoperiod under a constant temperature and relative humidity. Food and water were provided ad libitum. Ovaries were removed from animals that were primed 48 h prior with 5 IU/100 µl eCG. Cumulus-enclosed oocytes (COCs) were collected in Hepes-buffered Eagle minimum essential medium (EMEM) with Hanks salts supplemented with 50 µg/ml gentamicin and 0.3% BSA. Spontaneous maturation of COCs was conducted in either 1) a basal maturation medium (IVMb) consisting of EMEM, 2 mM glutamine, 0.23 mM pyruvate, 0.3% BSA, and 50 µg/ml gentamicin, or 2) enriched IVM medium (IVM+) consisting of 2 mM glutamine, 0.23 mM pyruvate, 10% fetal bovine serum (FBS), 0.6 mM L-cysteine, 0.5 mg/ml D-glucosamine, 0.02 µM ascorbate, 1% insulin-transferring selenium (ITS), 50 µg/ml gentamicin, and 0.2 IU/µl recombinant human FSH (Serono Reproductive Biological Institute). IVM+ medium was chosen because it promotes cumulus expansion, unlike IVMb medium. Specifically, the inclusion of serum, glucosamine, and FSH is required to support cumulus expansion. COCs were cultured for 16 h (20–30 COCs per dish in 1 ml medium per organ culture dish) in a humidified atmosphere of 5% CO2 and air at 37°C. At the end of culture COCs were treated 2–3 min at 30°C–37°C with hyaulronidase (100 µg/ml; Sigma), and oocytes were fixed in microtubule stabilizing buffer-extraction fix for 30 min at 37°C [17]. In vivo-ovulated (IVO) COCs were collected from the oviductal ampulla from animals at T0 and 2, 4, 8 and 16 h after 5 IU hCG (Calbiochem), which was given 48 h following eCG (see above). All oocytes were either fixed and stored in wash solution containing 0.2% azide, 2% normal goat serum, 1% BSA, 0.1 M glycine, and 0.1% Triton X-100 at 4°C until further processing or until usage for quantitative RT-PCR or protein analysis.
Sample Preparation for Microscopy
Oocytes were processed for indirect immunolabeling to ascertain maturation state, centrosome number and position, spindle size, PB size, and -tubulin content. A total of 20–30 oocytes per treatment group (four replicates) were incubated in primary antibody for 1 h at 37°C with gentle agitation, followed by three10-min washes in wash buffer, followed by a 1-h incubation of secondary antibody at 37°C with agitation. Labeling pairs were 1) 
/ß-tubulin cocktail (1:100; mouse; Sigma) followed by Alexa 488 goat anti-mouse IgG (1:500; Molecular Probes) and rhodamine-phalloidin (1:200; Molecular Probes) and 2) YOL34 (1:100; rat; Harlan) and -tubulin (1:100; mouse; Sigma), followed by Alexa 488 goat anti-rat IgG (1:500; Molecular Probes) and Alexa 568 goat anti-mouse IgG (1:500; Molecular Probes). Green fluorescent protein (GFP) antibody (1:100; rabbit; Invitrogen) followed by Alexa goat anti-rabbit IgG (1:500; Molecular Probes) was used to detect pEGFP construct after microinjection. Oocytes were incubated in 1 µg/ml Hoechst 33258 (Molecular Probes) in wash solution for 3 min to label chromatin and were mounted in 2 µl of a 50% glycerol/PBS solution containing 25 mg/ml sodium azide and 1 µg/ml Hoechst 33258. Samples were analyzed on an inverted IM35 microscope (Zeiss, Thornwood, NY) equipped with a 100-W mercury arc lamp and were imaged using 40x and 63x Neofluor objectives (Zeiss). Digital images were collected as 300-ms exposures using an Orca ER digital camera (model C4742–95; Hamamatsu Corp., Bridgewater, NJ) interfaced with MetaMorph Imaging System (Universal Imaging Corp., Downington, PA). A triple band pass dichroic and automated excitation filter selection specific for fluorescein (Alexa 488), rhodamine (Alexa 568), and bisbenzimides (Hoechst 33258) permitted the collection of in-frame images with minimal magnification or spatial distortion. A Zeiss LSM 5 Pascal confocal microscope was used to collect z-axis data sets from rhodamine-phalloidin-stained uncompressed oocytes at 4-µm intervals with a 40x Plan-Apo objective (numerical aperture 1.0) using HeNe 543 laser excitation and LP560 detection. Volumes were computed from 3D reconstructions of single oocytes using LSM 5 software.
Fluorescence Intensity and Volume Measurements
Archived images of 16 bits were analyzed for oocyte, PB, spindle measurements, MTOC number, and -tubulin fluorescence intensity. Metamorph imaging software was manually calibrated using a micrometer for the 40x and 63x objectives. Measurements then were integrated into the Metamorph software's automated measurement tool. Relative fluorescence intensity measurements also were obtained for -tubulin. Each image in one complete experiment was analyzed with a standard threshold. Average intensity values were obtained for the whole oocyte, spindle, PB, and MTOCs as the sum of grey value arbitrary units (a.u.) for a region of interest. Thresholded grey values ranged from a minimum of 241 units to a maximum of 552 units for a 16-bit image. Fluorescence background grey values were determined by measuring intensity after staining samples with secondary antibody alone. The average background level approximated 50 greyscale units and was subtracted from the final thresholded value to give a standardized grey value for our analysis. Volume data were collected by analyzing uncompressed oocytes labeled with rhodamine-phalloidin. Three-dimensional images were built from Z-series collected on a Zeiss LSM 5 Pascal to obtain oocyte volume (see sample preparation). Ten oocytes per treatment group were examined, and each experiment was run in triplicate.
Oocytes were stripped of cumulus cells by incubation for 10 min in Earles buffered salt solution (EBSS) containing 1 mg/ml BSA, 6.8 mM EGTA, and 9 mM HEPES, followed by a 2-min incubation in EBSS containing 1.8 mM EGTA, 50 mM NaHCO3, 9 mM HEPES, and 245 mM sucrose. Ten oocytes per treatment group were added to 5 µl of 1x lysis buffer (Cell Signaling Technologies) containing 200 mM PMSF and 40 µM phenylarsine oxide. Lysates were boiled for 5 min with 1 µl of 10x sample buffer. The samples were separated using 40% acrylamide/bis-acrylamide (29:1) and were transferred onto a 0.2-µm polyvinylidene fluoride membrane using Genie electrophoretic transfer (IDEA Scientific). Membranes then were incubated in mouse anti--tubulin (1:1000; GTU-88; Sigma), and ZP1(M-20)-bound (1:1000; sc-23708; Santa Cruz Biotechnology) antibodies were detected by goat anti-mouse or rabbit anti-goat horseradish peroxidase followed by Supersignal West Dura Enhancer substrate (Pierce).
To determine the expression of -tubulin throughout oocyte maturation, RNA was isolated from 20 naked oocytes at 0, 2, 4, 8, and 16 h during maturation (IVM and IVO) using TRI reagent (Sigma) in four replicate experiments. Rabbit RNA (0.6 pg) was added to TRI reagent before chloroform extraction. RNA isolation was followed according to the TRI reagent protocol. Glycogen (1 µl) was added to each sample before the addition of isopropanol to aid in RNA precipitation. Reverse transcription was performed at 37°C for 2 h, followed by inactivation for 15 min at 67°C.
Gamma-tubulin and rabbit -globin primers and probes were generated using Primer Express 2.0 (5'GGCCAGTGCGGCAATC and 5'CGGGACTGATGCCATGCT; Applied Biosystems). We used SYBR Green detection system (Applied Biosystems) for -tubulin and Taqman probe system (Applied Biosystems) for -globin, and we performed the quantitative PCR reaction on an ABI Prism 7900HT Sequence Detection System (Applied Biosystems). Samples were compared to a standard curve that was generated by serial dilution of a sample containing elevated levels of the target amplicon.
Gamma-Tubulin Overexpresison and siRNA Knockdown Strategies
Germinal vesicle (GV)-stage oocytes were partially denuded by gentle pipetting and were maintained in a 30-µl drop of FHM (Clontech) containing 0.2 mM IBMX (Sigma) covered with sterile mineral oil (M8410; Sigma). Oocytes were either injected with 1) 
10 pl of injection buffer containing 5 mM Tris, pH 7.5, and 0.1 mM EDTA (EmbryoMax MR-09510F; Clontech) or 2) 10 pl linearized enhanced green fluorescent protein (pEGFP)--tubulin (5 ng/µl) [28] or 3) 10 pl -tubulin siRNA (10 µM; sc-29323; Santa Cruz Biotechnology) into the GV using Eppendorf Femtojet with EggJek injection needles (catalog no. EJ-01; MicroJek) using Precision Micro Devices holding pipets (catalog no. H2085-25-07; CellTram Air; Eppendorf). A total of 20–30 oocytes were injected per treatment group, and each experiment was run in triplicate (siRNA control, n = 71; siRNA, n = 83; plasmid control, n = 68; plasmid, n = 58). Injected oocytes were returned to 37°C for 1 h before being transferred to maturation medium (1.5 ml). Oocytes injected with -tubulin siRNA and the paired control-injected oocytes were put in IVM+ medium to ascertain the effects of selective knockdown under conditions that normally produce spindles and PBs of a reduced size. In contrast, oocytes injected with pEGFP--tubulin and the paired control-injected oocytes were cultured in IVMb, since these conditions typically promote spindles and PBs of increased size. Oocytes were cultured for 16 h in 37°C with 5% CO2, fixed for 30 min in 2% paraformaldehyde, treated for 30 min in wash solution containing 0.1% Triton, and processed for immunofluorescence labeling.
Oocyte and PB volume and spindle measurements, -tubulin intensity measurements, and quantitative PCR data were subjected to one-way analysis of variance followed by Bonferroni multiple comparison post-hoc test to determine the significance of each treatment group using Prism 4 (GraphPad Software Inc.). Calculated values are shown as ±SEM, with a significance level of P < 0.05 being considered significant.
Variations in Spindle Size, PB Size, and MTOC Number and Localization
The spindle, PB, and cytoplasmic characteristics of oocytes matured under three maturation conditions were analyzed using a triple-labeling strategy and digital imaging. Representative images of oocytes from each treatment group are shown in Figure 1. GV-stage mouse oocytes (four experiments, n = 130) matured in a basal medium (IVMb) typically contained broad, barrel-shaped spindles juxtaposed to large PBs (Fig. 1A) that were prominently labeled with -tubulin antibodies (Fig. 1D). Large amounts of -tubulin were distributed throughout the spindle that was stained with -tubulin (red), shown by the overlapping yellow signal. It is important to note that the -tubulin (green) was not focused at the poles (Fig. 1D, inset). Oocytes matured in vivo displayed small pointed spindles associated with the microvillar-free domain (red; Fig. 1B), as well as small -tubulin foci restricted to the spindle poles (Fig. 1E, inset). Metaphase II oocytes (MII) matured in IVM+ exhibited smaller and somewhat tapered spindles (compared with IVMb), with focused -tubulin staining at the poles and low-level staining through the spindle proper (Fig. 1F and inset). MTOCs were detected with -tubulin antibodies in all treatment groups. These were few in number, diffusely stained, and displaced from the oocyte cortex in oocytes of the IVMb group (Fig. 1D). In contrast, oocytes from IVO and IVM+ groups contained MTOCs that were consistently more numerous, focused, and cortically located based on through focus analysis (Z-series; Fig. 1, E and F; red spots seen in IVM+ in panel F are remnants of transzonal projections emanating from stripped cumulus cells). These variations of IVM and IVO oocytes with respect to PB and spindle size prompted a morphometric analysis in each experimental group.
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Volumetric Variations in the Oocyte, PB, and Spindle
A digital z-series database was created for individual uncompressed oocytes from each treatment group that included 1) oocyte and PB volumes by f-actin cortex staining, 2) -tubulin to monitor length and width of the MII spindle, and 3) -tubulin labeling to obtain MTOC numbers. From this database, mean oocyte and PB volumes (Table 1) and spindle dimensions (Fig. 2A) were extracted. The volume of IVMb oocytes was consistently 15% smaller on average than IVO and IVM+ oocytes, and it appeared to be due to prominent contraction during maturation, as documented by time lapse digital imaging (data not shown). PBs from IVMb oocytes were approximately 35%–40% larger than those from IVO oocytes. Oocytes from the IVO group displayed spindles with small pole lengths and overall lengths and widths compared with IVMb spindles (Fig. 2A). Interestingly, oocytes matured in IVM+ were similar to the IVO group with respect to all spindle parameters. Thus, these data suggest a relationship between spindle and PB volume when IVMb, IVO, and IVM+ groups are compared. Our studies also show an indirect relationship between spindle size and MTOC number. IVM+ and IVO oocytes exhibited a 2- to 3-fold (10.2 ± 1.7) to a 10-fold (22.9 ± 2.4) increase, respectively, in mean MTOC number relative to IVMb (3.65 ± 2.45; Fig. 2B), which is consistent with previous reports [26]. Because of the relationship of PB size to spindle size and its inverse relationship with MTOC number, we next examined the spatial allocation of -tubulin within these structures.
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Qualitative Differences in -Tubulin Allocation
Using digital imaging analysis, we first found that the amount of immunodetectable -tubulin in IVMb oocytes (7.8 x 106 a.u.) was an order of magnitude higher than that observed in IVO or IVM+ oocytes (4.7 x105 and 5.6 x105 a.u., respectively; Fig. 3), a finding consistent with the higher protein levels found in IVMb oocytes (Fig. 3, B and C). In IVO oocytes, 57.1% (2.7 ± 0.7 x 105 a.u.) of the immunodetectable -tubulin was rationed to the spindle, where it was confined specifically to spindle pole foci. By contrast, IVMb oocytes allocated 48% (3.8 ± 1.5 x 106 a.u.) of immunodetectable -tubulin to the MII spindle that was distributed diffusely. Variations in nonspindle compartments also were seen between groups. Whereas a substantial fraction of -tubulin (36.2%; 1.7 ± 0.2 x 105 a.u.) was present in MTOCs of IVO oocytes, MTOCs accounted for only 2.8% (2.2 ± 0.5 x 105 a.u.) of the total -tubulin in IVMb oocytes. Most importantly, little or no immunodetectable -tubulin was evident in the first PBs of IVO oocytes, whereas a significant fraction (23%; 1.8 ± 0.4 x 106 a.u.) of total oocyte -tubulin was detected in first PBs of IVMb oocytes. Moreover, -tubulin foci were often detected in the PBs of IVMb oocytes but not in PBs of IVO oocytes (data not shown). Whereas IVM+ oocytes, as shown earlier, resembled IVO oocytes in some respects, this was not the case for -tubulin allocation. Thus, while 46.4% (2.6 ± 0.9 x 105 a.u.) of -tubulin was localized to the spindle poles, there was a relative reduction to 21.4% (1.2 ± 0.3 x 105 a.u.) of total -tubulin associated with the MTOC compartment, suggesting that the mechanism involved with segregation of -tubulin during IVM may be compromised relative to IVO oocytes. These findings highlight discrete variations in -tubulin allocation associated with discernible alterations in the properties of PBs in maturing mouse oocytes. Because of the variations in total -tubulin protein and immunodetectable epitope found in IVMb oocytes, we wanted to next examine whether similar variations in -tubulin mRNA were related to the intergroup protein variance.
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Quantification of -Tubulin mRNA in Maturing Oocytes
We next examined the -tubulin mRNA expression levels in IVMb, IVM+, and IVO oocytes (n = 20 oocytes per group) at various time points during oocyte maturation using quantitative RT-PCR (Fig. 4). Data represent the mean for four replicate experiments. The T0 treatment samples exhibited similar -tubulin mRNA expression levels. Within 2 h (T2) of isolation, -tubulin mRNA levels dropped 3.5-fold in the IVMb and IVM+ groups, whereas only a 40% loss of mRNA in the IVO was seen. Between T4 and T16, IVMb and IVM+ -tubulin levels did not change appreciably nor significantly at each of the time points examined. However, IVO group -tubulin levels at T4 decreased appreciably, and significant variations in -tubulin mRNA levels were observed at both T8 (3-fold increase) and T16 (2- to 3-fold decrease).
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Experimental Manipulation of -Tubulin Levels Alters Meiotic Cytokinesis
To determine whether depletion of -tubulin mRNA would impact the meiotic spindle or PB extrusion, -tubulin siRNA was injected into GV-stage oocytes prior to in vitro maturation. Conversely, a linearized pEGFP--tubulin construct was injected to determine the effects of ectopic overexpression of -tubulin during in vitro maturation. The results are summarized in Table 2. A total of 87.3% and 85.3% of the controls (siRNA and pEGFP--tubulin groups, respectively) proceeded to metaphase of meiosis 2 (M2), establishing that the injection procedure itself had little effect on meiotic progression. In contrast, only 51.8% of the -tubulin siRNA-injected oocytes progressed to M2, indicating that downregulation of -tubulin mRNA levels adversely affected meiotic progression. In the case of siRNA-injected oocytes, some arrested at germinal vesicle breakdown (GVBD; 13.3%) or metaphase of meiosis 1 (M1; 24.1%) stages, whereas those that proceeded exhibited striking alterations. The most apparent effects of siRNA treatment were decreased cytoplasmic MTOC number, the elimination of immunodetectable -tubulin in the spindle (Fig. 5B), and the presence of tapered spindle poles and malaligned chromosomes on the spindle relative to controls (Fig. 5A). Importantly, PB size was significantly decreased in this group (Table 2). Oocytes injected with pEGFP--tubulin expressed high levels of immunodetectable GFP (Fig. 5D) relative to controls (Fig. 5C). Control oocytes (Fig. 5E) exhibited tapered spindles with -tubulin foci at each pole, as typically seen in IVMb conditions. Under these conditions, GFP was evident in the PB and throughout the spindle (Fig. 5D). Staining with tubulin and -tubulin (Fig. 5F) showed large, barrel-shaped spindles with prominent plaques of -tubulin at the poles and expression of -tubulin throughout the spindle (inset). Quantitation of the injected oocytes, as shown in Table 2, verified that the majority of pEGFP--tubulin-injected oocytes exhibited increases in MTOC numbers (means: 17.7 vs. 13) and PB size (102 µm vs. 80 µm) relative to controls. Thus, coupled with the result of decreased PB and spindle sizes after siRNA, these data provide direct evidence that -tubulin is a key regulator of PB size by virtue of its spindle-associated functions.
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Just before fertilization, oocytes resume meiosis in response to ovulatory gonadotropins and extrude the first PB, signifying completion of karyokinesis during MI. While the process of oocyte maturation occurs readily in culture in many mammalian species, little is known about the fidelity and regulation of this form of asymmetric cytokinesis. Preserving cytoplasmic volume and content in large cells such as oocytes is critical to embryogenesis following fertilization. This is an especially relevant subject of study, since effects of culture conditions on cytokinesis may affect the overall developmental potential of IVM oocytes. The present findings indicate that 1) the size, shape, and content of the first PBs and M2 spindles are directly influenced by IVM; 2) the variations observed between in vivo- and in vitro-matured oocytes link cytokinesis and karyokinesis based upon the availability and distribution of -tubulin; and 3) when experimentally perturbed, active synthesis and turnover of -tubulin cause an imbalance in the forces that dictate the shape and size of the spindle regulating the spatial parameters of chromatin-induced actin remodeling.
Relationship Between Spindle and PB Volume
The present study demonstrates dysregulation of PB and spindle size due to culture in IVMb oocytes when compared to oocytes undergoing this process in vivo, and it corroborates our previous studies on IVM oocytes exhibiting hypertrophy of MI spindles that was absent in IVO oocytes [15]. Here we extend these observations to show that spindle hypertrophy is linked to both increased PB size and decreased cortical MTOC number. This is consistent with previous studies suggesting that restricting MTOC access to the first meiotic spindle dually limits spindle size and available MTOCs for cortical stabilization and anaphase onset [26]. IVO MII spindles are small, pointed, and anastral, and these oocytes contain anastral cortical MTOCs (Fig. 1). Interestingly, IVMb oocytes display a phenotype remarkably similar to the Mos–/– [29–31] and LT mutants [32, 33]. In both of these murine models, oocytes extrude first PBs that are remarkably large. Moreover, Mos–/– and LT strains of mutant oocytes have hypertrophied spindles and few cortical MTOCs. PBs from these mutant strains also are known to persist and go through a round of cytokinesis, like IVMb oocytes, rather than degrade soon after emission, as seen in IVO oocytes [34, 35]. One final parallel to note is that IVMb, Mos–/–, and the LT strains of mutant COCs cannot undergo cumulus expansion, a process known to occur during in vivo maturation and ovulation. It is therefore relevant to point out that the addition of rhFSH to culture media (IVM+ oocytes) promotes partial reversion of IVMb oocytes to the IVO phenotype, suggesting that COC integrity and hormone-regulated expansion play pivotal roles in microtubule (MT) patterning during oocyte maturation.
Previous work has shown that PB formation is a microfilament-driven process [5], and conditions that lead to gross chromosome dispersion cause an expansion of chromatin-based remodeling of the actin cortex [5, 14]. Similarly, studies on somatic cells established that mitotic spindles dictate chromosome spatial positioning, thereby impacting both furrow location and volume equivalence of cytokinesis [36]. Here we show in the case of asymmetric cytokinesis in mouse oocytes that a similar principle holds. Specifically, two populations of MTOCs that associate with either spindle poles or the oocyte cortex [16, 17, 37–39] are present in mouse oocytes [17]. If spindle-associated MTOCs were limited in number or mass during spindle morphogenesis, then smaller, tapered spindles in oocytes and numerous cortical MTOCs would be expected, as we report here for IVO oocytes. One mechanism for limiting MTOC access to the forming spindle in vivo appears to be persistence of the nuclear lamina [26]. The delayed appearance of -tubulin within the nuclear lamina is consistent with this model [18]. We speculate that by excluding MTOCs from the microenvironment created by the nuclear lamina, a more chromatin-dominated process occurs to ensure rapid alignment and assembly of a -tubulin-free spindle. The fact that -tubulin is distributed throughout meiotic spindles after overexpression (Fig. 5F) or IVMb culture conditions further supports such a mechanism as the underlying link between the spindle interchromosomal spacing and PB size.
We took advantage of the consistent properties of in vivo-matured oocytes relative to cultured and spontaneously maturing oocytes (IVM) to evaluate compartmentalization of -tubulin within MII oocytes. Three distinguishing properties for PBs in IVMb oocytes relative to IVO oocytes were noted in this work. First, IVMb oocytes contained 23% more immunodetectable -tubulin. Second, IVM PBs contained intact MTOCs. Finally, IVMb PBs were more likely to undergo another round of cytokinesis compared with PBs from IVO oocytes (data not shown). The fact that IVO oocytes contained 36% of their total immunodetectable -tubulin in cortical MTOCs and typically yielded PBs unlikely to cleave further suggests that restricted positioning of MTOCs is regulated in vivo. Perhaps anchoring MTOCs to the cortex prevents their involvement in spindle assembly, fostering a chromatin bias (see above) that would limit PB size and the spurious appearance of MTOCs in the PB. That somatic signaling confers MTOC stabilization in the oocyte cortex is one mechanism that has been proposed for localizing assembled -tubulin complexes [40]. Whether regulation is further manifested at a transcriptional or translational level also was addressed.
Real-time PCR analysis of -tubulin mRNA levels revealed absolute and temporal variations between study groups. Although it is well known that global transcription is arrested during oocyte maturation [41], some transcription may occur, particularly from genes that are directly involved in this process. Interestingly, the increase in -tubulin mRNA between T4–T8 in IVO oocytes coincides with a critical time point when transzonal projections (TZPs) in the surrounding cumulus are being remodeled. TZPs, somatic cell-to-oocyte connections, are essential for transporting cAMP, a small molecule essential for arresting the oocyte in prophase of M1. Cyclic AMP also is essential in gene transcription and has been linked with the upregulation of tubulin mRNA in neuroblastomas [42]. In IVM oocytes, -tubulin mRNA drops initially from T0 to T2 and remains low throughout maturation, possibly because TZPs retract when COCs are released from the follicle into culture [34, 35, 43]. The decrease in -tubulin mRNA during IVM coincides temporally with the accelerated GVBD seen in IVM oocytes compared with GVBD in IVO oocytes [26]. It is interesting to consider whether fluctuations in mRNA levels in IVO oocytes reported here are related to classical studies on tubulin gene expression that showed how steady-state levels of unpolymerized (cytoplasmic) tubulin effect a negative feedback loop at both transcriptional and translational levels to suppress new mRNA or protein production [44]. The high levels of cytoplasmic (unpolymerized) -tubulin reported here in IVMb oocytes (Fig. 3A) contrasts with the relatively lower amounts of cytoplasmic -tubulin detected in IVO oocytes that is perhaps due to selective stabilization and/or decreased degradation of existing mRNAs. That maternal centrosomal constituents impact subsequent embryonic development in the mouse has already been elegantly demonstrated by the gene deletion studies of Yuba-Kubo et al. [27] cited earlier. Thus, some form of inheritable maternal -tubulin (Tubg1), is required for early embryonic cell cycles to proceed.
In that light, it was interesting that downregulation with -tubulin siRNA resulted in a paucity of cortical MTOCs and in M2 oocytes that lacked -tubulin in small, tapered spindles (Fig. 5B). On the one hand, it would be important to establish whether -tubulin-depleted oocytes lose their developmental competence, as was seen in the knockouts. On the other hand, it seems paradoxical that limiting -tubulin availability yields an IVO phenotype at the level of the spindle and PB. These observations again emphasize that it may be essential for oocytes to limit the involvement of -tubulin in meiotic cell cycle progression, since its precocious consumption and/or utilization would impact reserves designated for postfertilization mitotic progression. Interestingly, the majority of siRNA-injected oocytes were blocked in the cell cycle at points between GVBD and M1, which is consistent with the fact that -tubulin is an essential cell cycle protein [45]. Conversely, -tubulin overexpression resulted in large, barrel-shaped spindles and large PBs. This phenotype is consistent with our hypothesis that if -tubulin is not anchored to the cortex its incorporation into the forming spindle enlarges the domain of influence for chromatin-based remodeling of the actin cortex and extruding PB.
These findings draw specific attention to determinants of oocyte quality that are known to influence both clinical and agricultural programs in assisted reproductive technology and reinforces mounting concerns over the widespread use of clinical IVM.
ACKNOWLEDGMENTS
We thank past and present members of the Albertini Laboratory (Gloria Lee, Dr. Alexandra Sanfins, Dr. Susan Messinger, Paty Rodrigues, Lynda McGinnis, Darlene Limback, and Dr. Karla Hutt) and Dr. Lane Christenson for their support and comments on the manuscript. We thank Lynda McGinnis for all of her help with microinjection. Dr. William Kinsey provided assistance with Western blot studies, and the pEGFP--tubulin plasmid was kindly provided by Dr. Quingshen Gao. Dr. Steve Palmer and Dr. Daniel de Matos at Serono Reproductive Biological Institute provided guidance and support for this work. This work is being submitted as partial fulfillment for a Ph.D. at Tufts University Sackler School of Graduate Biomedical Sciences.
FOOTNOTES
1Supported by National Institutes of Health grant RO1-HD42076, the Eshe Fund, Serono Reproductive Biology Institute, and The Hall Family Foundation. 
Correspondence: 2David F. Albertini, 3901 Rainbow Blvd., University of Kansas Medical Center, Kansas City, KS 66160. FAX: 913 588 0456; e-mail: dalbertini{at}kumc.edu
Received: 5 September 2006.
First decision: 12 October 2006.
Accepted: 2 February 2007.
REFERENCES
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