1 Department of Anatomy and Cellular Biology, Tufts University School of Medicine, Boston, MA 02111, USA, 2 Instituto de Medicina Molecular, Unidade de Biologia da Reprodução, Faculdade de Medicina de Lisboa, 1649-028 Lisboa, Portugal and 3 Department of Biomedical Sciences, Tufts University School of Veterinary Medicine, Grafton, 01536 MA, USA
6 To whom correspondence should be addressed at: Department of Molecular and Integrative Physiology, University of Kansas Medical Center, 3901 Rainbow Boulevard, Kansas City, KS 66160-7401, USA. Email: dalbertini{at}kumc.edu
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Abstract |
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Key words: centrosome/in vivo or in vitro maturation/microtubules/MTOCs/nuclear lamina
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Introduction |
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Another aspect of oocyte maturation that may impinge upon oocyte quality and developmental competence is cytoskeletal remodeling. In mice, variations in microtubule remodeling during spindle morphogenesis were previously shown to underlie the distinct organization of meiotic spindles evident after either in vivo or in vitro maturation (Sanfins et al., 2003). Earlier studies of IVM oocytes suggested that once removed from the follicle, mouse oocytes undergo dramatic changes in microtubule patterning and centrosome positioning during spindle morphogenesis (Messinger and Albertini, 1991
; Combelles and Albertini, 2001
). Although cell cycle specific rearrangements in oocyte microtubule organizing centers (MTOCs) and
-tubulin have been defined during IVM in mouse oocytes (Messinger and Albertini, 1991
; Combelles and Albertini, 2001
), the overall significance of these cytoplasmic remodeling events to cell cycle control and oocyte nuclear maturation and developmental potential remains unknown. Accordingly, in an effort to better understand the importance of cell cycle markers during meiotic resumption and M-II arrest we have (i) analyzed cell cycle progression markers during the initial cytoskeletal remodeling in IVO and IVM oocytes, namely centrosome proteins (
-tubulin and pericentrin), nuclear lamin integrity (lamin B) and microtubule patterning; (ii) assessed the in vivo window of oocytes removed from ovulatory follicles at specific times after exposure to hCG and (iii) evaluated the effect of imposing a G2 cell cycle delay with the maturation-promoting factor (MPF) inhibitor, roscovitine, on the overall quality of cytoplasm and spindles. Our results indicate that the timing and spatial patterning of spindle morphogenesis influences the allocation of maternal
-tubulin stores during maturation such that both the physical properties of the meiotic spindle and the positioning of MTOCs are predictably distinct in IVM and IVO oocytes.
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Materials and methods |
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The influence of intrafollicular residence time post-hCG on M-II phenotype
To address the influence of the follicular environment during early stages of oocyte maturation, mice were injected with 5 IU of eCG followed 46 h later by 5 IU of hCG. At 1.5 and 5 h post-hCG injection (GV-stage and prometaphase/M-I stage, respectively), mice were sacrificed, COCs collected by follicular puncture from Graffian follicles and immediately cultured in IVM medium up to 16 h. Therefore, for the group collected 1.5 h post-hCG, oocytes were cultured for 14.5 h in IVM medium; and for the group collected 5 h post-hCG, oocytes were cultured for 11 h in IVM medium. Following removal of cumulus cells by gentle pipetting (1.5 h group) or brief treatment with hyaluronidase (5 h group), oocytes were immediately fixed and stored at 4°C until further processing. IVO and IVM matured oocytes, obtained 16 h post-hCG or post-culture, respectively, were used as controls for these experiments.
Roscovitine treatment of GV-stage oocytes
COCs were collected from 78-week old CF-1 mice 4648 h after injection with 5 IU eCG as described above. Oocytes were either exposed to 1% DMSO (control) or to 50 µM roscovitine in 1% DMSO (Biomol, Pennsylvania, PA) in maturation medium for 3 and 6 h. Fifty micromolar roscovitine was determined to be the optimal concentration to use based on reversibility experiments. Following treatment, oocytes were washed three times in 100x volume of maturation medium without roscovitine and were subsequently matured in vitro for 16 h without drug. Oocytes were immediately fixed and stored at 4°C until further processing.
Processing of oocytes for immunofluorescence
All oocytes were processed in parallel for fluorescence microscopy in specific combinations to assess meiotic status (Hoechst 33258) relative to microtubule (MT) patterning (tubulin), nuclear lamina stability (lamin B), and MTOCs bearing constitutive (pericentrin) or regulative (-tubulin) markers. Oocytes were incubated sequentially with primary and secondary antibodies. Oocytes were separately processed for
-tubulin (mouse) followed by anti-mouse secondary, then
-tubulin (rat) and anti-rat secondary. This sequence was followed for pericentrin (rabbit) and
-tubulin (rat); and
-tubulin (mouse) and lamin B (rabbit), as shown in Table I. Each antibody was incubated for 1 h at 37°C with shaking followed by three 15 min washes in blocking solution. Oocytes were mounted using 1.5 µl of a 50% glycerol/PBS containing sodium azide and Hoechst 33258 (1 µg/ml Polysciences Inc., Warrington, PA) to label chromatin. Incubation of oocytes in secondary antibodies alone failed to yield detectable staining either singly or in repeat sequence as above. Labeled oocytes were analyzed using a Zeiss IM-35 inverted microscope and a 50 W mercury arc lamp using 40x and 63x Neofluor objectives. Digital images were collected with a Hamamatsu Orca ER digital camera (model #C4742-95) interfaced with a Meta Morph Imaging System. A triple band pass dichroic and automated excitation filter selection permitted collection of in-frame images with minimal magnification or spatial distortion.
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Statistical analysis
The number of MTOCs was represented by notched box and whisker plots (Slide Write Plus for Windows, Version 5.01; Advanced Graphics Software Inc., Encinitas, CA). Notched box plots display order statistics, and the notches of the box plots correspond to median confidence limits. Statistically, two medians are considered significantly different at the 0.05 level if their confidence limits do not overlap. Additionally, data were analyzed using SPSS 10.0 (Statistics Package for Social Sciences, Chicago, IL). Comparison of cytoplasmic MTOC numbers for each category of oocytes was evaluated using a nonparametric KruskalWallis test followed by MannWhitney tests for two independent samples. Differences were considered significant at P<0.05.
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Results |
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These studies uncover basic distinctions in nuclear lamina integrity and MTOC behavior in IVM and IVO oocytes. Coupled with our earlier reports, it seems likely that containment of chromatin and limiting access to centrosome proteins may be linked to the smaller spindles and multiple MTOCs seen in IVO oocytes (Sanfins et al., 2003).
Influence of the intrafollicular environment on the M-II phenotype
The above results suggest that events soon after hCG reception in the follicle may specify properties in the M-II oocyte previously shown to affect spindle shape, size and cytoplasmic MTOC number. How soon after hCG would oocytes be committed to the expression of an IVO character? To determine if early signaling affected the properties of oocytes at M-II, experiments were designed using COCs recovered from pre-ovulatory Graffian follicles at 1.5 and 5 h post-hCG. Meiotic progression of the oocytes to M-II was not affected by removal at either time (80% from 1.5 h versus 72.7% from 5 h reached M-II stage). In oocytes retrieved 1.5 h post-hCG, M-II matured oocytes displayed a barrel shaped spindle (Figure 5C) with -tubulin distributed throughout the spindle proper (Figure 5B), similar to the IVM control group (Figure 5J). Typically, the 1.5 h group exhibited large spindles, lacking bundled microtubules and was characterized by a diffuse
-tubulin throughout (Figure 5C). In contrast, oocytes retrieved after 5 h exhibited pointed spindles containing distinct microtubule bundles (Figure 5F) and
-tubulin staining was only seen at the spindle poles (Figure 5E). The spindles were more cortically positioned in oocytes retrieved at 5 h versus 1.5 h (30/33 versus 10/36). Consistent with this, as shown in Figure 6, the number of MTOCs was greater when the intrafollicular residence time was longer, although significantly different from the IVO group (P<0.05). IVO oocytes (control) show the highest mean number of MTOCs/oocyte followed by oocytes collected 5 h post-hCG (27.3±6.3 and 21.9±6.48, respectively). Moreover, M-II oocytes collected 1.5 h post-hCG also exhibit a higher number of MTOCs when compared to the IVM group, but these were considerably reduced in MTOC mean number when compared to the 5 h post-hCG group (16.7±5.4, 10.8±3.1 and 21.9±6.48, respectively). Thus, between 1.5 and 5 h post-hCG, events have occurred that impact properties of M-II oocytes following retrieval and subsequent culture. These findings further support the idea that an initial delay in meiotic progression in vivo is due to resistance to effect a G2/M cell cycle transition.
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Discussion |
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The distribution of centrosomal proteins (-tubulin and pericentrin) and the microtubule cytoskeleton during the early stages of meiotic maturation is clearly distinct between IVO and IVM oocytes. IVO oocytes retain centrosomal proteins subcortically within multiple MTOCs of enhanced microtubule nucleation capacity that increase in number from diakinesis to M-I. In contrast, fewer cytoplasmic MTOCs are present in IVM oocytes over the same cell cycle stages, presumably as a result of the dramatic redistribution of
-tubulin and pericentrin to the nuclear lamina during spindle morphogenesis (Messinger and Albertini, 1991
; Combelles and Albertini, 2001
). While Can et al. (2004)
proposed that the availability of
-tubulin and pericentrin to the nuclear lamina during early stages of maturation in IVM oocytes reveals a shift of microtubules from stabilization of the cortex to morphogenesis of the spindle (Can et al., 2004
), our results on IVO oocytes add further insight. Centrosome positioning in somatic cells requires radial or astral arrays of cytoplasmic microtubules so that pushing and pulling forces involving MT mediated by motors can be anchored to and stabilize the overlying cortex (Burakov et al., 2003
). We suggest that IVM oocytes similarly exhibit more rapid and complete nuclear lamina disruption because of the rapid and nearly total accumulation of centrosomes upon GVBD. Interestingly enough, the minus end focusing motor dynein was also shown to be a key factor during nuclear envelope breakdown in mitosis due to M-phase dynamics of perinuclear centrosomes and their associated MTs (Salina et al., 2002
). Conversely, the delayed GVBD and persistence of the nuclear lamina in IVO oocytes is consistent with the fact that increased number of MTOCs support microtubule nucleation, preventing cytoplasmic MTOCs from participating in nuclear lamina disruption. Thus, while preserving integrity of the cortical microtubule cytoskeleton, spindle morphogenesis will proceed with minimal centrosome involvement within a constrained space. While admittedly speculative, the potential advantages of compartmentalizing spindle morphogenesis might include favoring chromosome directed over centrosome forces to achieve proper metaphase alignment and limiting the utilization of maternal molecules like
-tubulin in meiotic spindles so that adequate stores remain in the fertilized egg. Why would it be advantageous to limit centrosome access to the forming spindle? Some insight into this problem comes from recent studies on the regulation of the G2/M cell cycle transition in somatic cells.
Unlike what we report here and elsewhere in mouse oocytes (Combelles and Alberitini, 2001), somatic cells rapidly condense chromatin and depolymerize the nuclear lamina upon entry into M-phase. Moreover, it has been shown that active cyclin B1Cdk1 first appears on centrosomes in prophase (Jackman et al., 2003). The spatial segregation of cell cycle components appears to be central to achieving the correct temporal readout of checkpoint controls that allow somatic cells to reach and engage the chromosome alignment checkpoint at metaphase. While centrosomes can participate in mitotic spindle assembly, chromatin alone is necessary and sufficient to build a bipolar spindle as long as a minus end motor like dynein is present (Heald et al., 1997
). In fact, centrosomeless spindles function during mitosis and excess centrosomes can be deleterious to somatic cells, since they favor the formation of multipolar spindles that contribute to aneuploidy (Khodjakov and Rieder, 2001
; Sluder and Nordberg, 2004
). Given the large volume of the oocyte, and the requirement to retain maternal stores that likely include MTOC components needed for mitoses during embryogenesis, restricting MTOC access to the GV may serve to engage cortical anchoring of the spindle prior to first polar body extrusion and launch nuclear maturation within a nuclear lamina compartment favoring chromatin-directed spindle morphogenesis as noted above. Might this interplay between nuclear and cytoplasmic maturation also influence cell cycle progression?
The spatial distribution of cell cycle factors has been proposed to be a key factor for the coordination of nuclear and cytoplasmic maturation in mouse oocytes (Mitra and Schultz, 1996). As in somatic cells (Pines, 1999
), relative protein and mRNA levels of key cell cycle factors cannot account for differences in the ability of mouse oocytes to initiate a G2 (incompetent) to M (competent) cell cycle transition (Kanatsu-Shinohara et al., 2000
). Also, since centrosome phosphorylation has been associated with increased microtubule dynamics (Messinger and Albertini, 1991
) and meiotic competence acquisition (Wickramasinghe et al., 1991
), maintaining cortical centrosomes may stratify cell cycle machinery between the nucleus and cytoplasm to prevent precocious meiotic M-phase entry (Albertini and Carabatsos, 1998
). Thus, the synchrony in the rate of meiotic progression observed in IVO oocytes may be due to the selective retention of centrosomal material or MPF to the oocyte cortex whilst restricting recruitment of centrosomes/MPF for spindle morphogenesis. During IVM, it appears that oocytes liberate these factors leading to precocious MPF activation, the consequences of which would include rapid and more extensive dissolution of the nuclear membrane and lamina, resulting in formation of larger spindles and fewer cytoplasmic MTOCs. Asynchrony in the processes of nuclear and cytoplasmic maturation during IVM may then lead to compromised oocyte quality that might be ameliorated by better management of the G2/M cell cycle transition. With this in mind, the specific MPF inhibitor, roscovitine, was used to impose a delay on GVBD onset. Surprisingly, oocytes treated with roscovitine and allowed to re-initiate M-phase exhibited spindle and cytoplasmic properties resembling IVO oocytes (Figure 7). It has been reported that maintaining oocytes in a G2 cell cycle stage with roscovitine increases the developmental potential of porcine (Marchal et al., 2001
), bovine (Ponderato et al., 2001
; Lagutina et al., 2002
) and horse (Franz et al., 2003
) oocytes. Further studies are needed to elucidate the basis for oocyte quality improvement when a cell cycle delay is imposed prior to GVBD but our data with roscovitine imply that delaying activation of MPF and concurrent with microtubule stabilization may enhance the coordination of nuclear and cytoplasmic maturation and/or optimize oocyte metabolism as meiosis proceeds.
The metabolic and physiological complexity of the intrafollicular environment during in vivo ovulation must be addressed in any comparison with in vitro conditions (Sutton et al., 2003). Specific changes occur within the follicular milieu, mediated by oocytegranulosa cell contact and/or growth factors that coordinate oogenesis with folliculogenesis through ovulation (Albertini and Carabatsos, 1998
; Carabatsos et al., 2000
; Richards et al., 2002
). Not surprisingly then IVO oocytes exhibit elevated ATP content (Combelles and Albertini, 2003
), mitogen-activated protein (MAP) kinase activity (Su et al., 2003
) and characteristic mitochondrial distributions (Van Blerkom, 1991
; Sun et al., 2001
; Nishi et al., 2003
) compared to IVM counterparts. In addition, EGF-related growth factors (Park et al., 2004
) and specific transcriptional profiles responsible for cumulus expansion (Su et al., 2003
) and presumably oocyte quality determination are different in IVM oocytes. We addressed this directly by collecting IVO oocytes at 1.5 and 5 h post-hCG injection and culturing up to 16 h in IVM medium that did not recapitulate cumulus expansion conditions. Despite the fact that basal conditions yield IVM properties, oocytes removed from the follicles at 5 h post-hCG exhibited striking similarities to IVO oocytes in spindle shape and a slightly increased MTOC number (although MTOC number is significantly different to the IVO oocytes, P<0.05); oocytes removed from follicles at 1.5 h post-hCG more closely resembled IVM oocytes. These findings suggest that cumulus signaling and events occurring within the follicle in the first 5 h are critical to establish properties characteristic of IVO oocytes. This 5 h time interval is of interest because it coincides with the activation of cumulus MAPK and the production of EGF-like paracrine signals that mediate LH induced maturation (Su et al., 2003
; Park et al., 2004
).
Future efforts to optimize IVM should seek to recreate follicular changes in both cumulus and oocyte during the early events of oocyte maturation. Collectively these findings re-emphasize the importance of the early effects of LH on the process of oocyte maturation. Using markers for oocyte quality as have been identified here should help to understand the importance of maternal resource conservation mechanisms on embryonic developmental potential and give guidance to new protocols for in vitro maturation in humans.
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Acknowledgements |
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Notes |
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5Present address: Department of Biology and Biotechnology, Worcester Polytechnic Institute, 100 Institute Road, Worcester, MA 01609-2280, USA
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References |
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Submitted on April 13, 2004; accepted on August 26, 2004.