Key Points
-
The growth and the onset of meiotic maturation of the mammalian oocyte are controlled by bidirectional interactions between the oocyte and the surrounding somatic cells.
-
Junctional complexes and transzonal projections (TZPs) form the structural basis for the passage of signalling molecules and metabolic substrates that support oocyte growth.
-
The meiotic spindle forms through self-organization of microtubules and motor proteins in response to a RAN GTPase-mediated chromatin signal in the absence of centriole-containing centrosomes.
-
Meiotic chromatin provides a signal for the establishment of oocyte cortical polarity, which required for asymmetric meiotic cell divisions and leads to polar body extrusion.
-
Asymmetric positioning of the meiosis I spindle is established through actin-based forces that are regulated by actin nucleating factors, including a formin-family protein and the actin-related protein 2/3 (ARP2/3) complex.
-
Actin-driven cytoplasmic streaming contributes to the establishment and maintenance of oocyte polarity, and the parameters of post-fertilization streaming may be prognostic of the developmental potential of the embryo.
Abstract
Mammalian oocytes go through a long and complex developmental process while acquiring the competencies that are required for fertilization and embryogenesis. Recent advances in molecular genetics and quantitative live imaging reveal new insights into the molecular basis of the communication between the oocyte and ovarian somatic cells as well as the dynamic cytoskeleton-based events that drive each step along the pathway to maturity. Whereas self-organization of microtubules and motor proteins direct meiotic spindle assembly for achieving genome reduction, actin filaments are instrumental for spindle positioning and the establishment of oocyte polarity needed for extrusion of polar bodies. Meiotic chromatin provides key instructive signals while being 'chauffeured' by both cytoskeletal systems.
This is a preview of subscription content, access via your institution
Access options
Subscribe to this journal
Receive 12 print issues and online access
$189.00 per year
only $15.75 per issue
Buy this article
- Purchase on Springer Link
- Instant access to full article PDF
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
References
Albertini, D. F. Regulation of meiotic maturation in the mammalian oocyte: inteplay between exogenous cues and the microtubule cytoskeleton. BioEssays 14, 97–103 (1992).
Burgoyne, P. S., Mahadevaiah, S. K. & Turner, J. M. The consequences of asynapsis for mammalian meiosis. Nature Rev. Genet. 10, 207–216 (2009).
Downs, S. M. Regulation of the G2/M transition in rodent oocytes. Mol. Reprod. Dev. 77, 566–585 (2010).
Watanabe, Y. Geometry and force behind kinetochore orientation: lessions from meiosis. Nature Rev. Mol. Cell Biol. 13, 370–382 (2012).
Pan, H. et al. Transcript profiling during mouse oocyte development and the effect of gonadotropin priming and development in vitro. Dev. Biol. 286, 493–506 (2005).
Park, J. Y. et al. EGF-like growth factors as mediators of LH action in the ovulatory follicle. Science 30, 682–684 (2004). Demonstrates that the production of EGF-like proteins by mural granulosa cells is essential to trigger the events within the cumulus cells–oocyte complex that will initiate the resumption of meiosis in oocytes.
Robinson, J. W. et al. Luteinizing hormone reduces the activity of the NPR2 guanylyl cyclase in mouse ovarian follicles, contributing to the cyclic GMP decrease that promotes resumption of meiosis in oocytes. Dev. Biol. 366, 308–316 (2012).
Simon, A. M., Goodenough, D. A., Li, E. & Paul, D. L. Female infertility in mice lacking connexin 37. Nature 385, 525–529 (1997).
Carabatsos, M. J., Sellitto, C., Goodenough, D. A. & Albertini, D. F. Oocyte–granulosa cell heterologous gap junctions are required for the coordination of nuclear and cytoplasmic meiotic competence. Dev. Biol. 226, 167–179 (2000).
Dong, J. W. et al. Growth differentiation factor-9 is required during early ovarian folliculogenesis. Nature 383, 531–535 (1996).
Carabatsos, M. J., Elvin, J., Matzuk, M. M. & Albertini, D. F. Characterization of oocyte and follicle development in growth differentiation factor-9-deficient mice. Dev. Biol. 204, 373–384 (1998).
Combelles, C. M. H., Carabatsos, M. J., Kumar, T. R., Matzuk, M. M. & Albertini, D. F. Hormonal control of somatic cell oocyte interactions during ovarian follicle development. Mol. Reprod. Devel. 69, 347–355 (2004). Gene deletion studies and rescue strategies show that FSH directly regulates the formation and stability of TZPs, which are the primary cellular device used to integrate metabolism of the oocyte and its surrounding somatic cells.
Johnson, M. T., Freeman, E. A., Gardner, D. K. & Hunt, P. A. Oxidative metabolism of pyruvate is required for meiotic maturation of murine oocytes in vivo. Biol. Reprod. 77, 2–8 (2007).
Sugiura, K., Pendola, F. L. & Eppig, J. J. Oocyte control of metabolic cooperativity between oocytes and companion granulosa cells: energy metabolism. Dev. Biol. 279, 20–30 (2005).
Su, Y. Q. et al. Oocyte regulation of metabolic cooperativity between mouse cumulus cells and oocytes: BMP15 and GDF9 control cholesterol biosynthesis in cumulus cells. Development 135, 111–121 (2008). Landmark paper identifying the role of oocyte-derived paracrine factors in the regulation of sterol metabolism within mammalian oocytes.
Norris, R. P. et al. Cyclic GMP from the surrounding somatic cells regulates cyclic AMP and meiosis in the mouse oocyte. Development 136, 1869–1878 (2009). Shows that cGMP is the principle cumulus cell modulator of oocyte meiotic arrest and that passage of this signal through gap junctions assures maintenance of oocyte protein kinase A activity and hence cell cycle arrest.
Norris, R. P. et al. Luteinizing hormone causes MAP kinase-dependent phosphorylation and closure of connexin 43 gap junctions in mouse ovarian follicles: one of two paths to meiotic resumption. Development 135, 3229–3238 (2008).
Longo, F. J. & Chen, D. Y. Development of cortical polarity in mouse eggs: involvement of the meiotic apparatus. Dev. Biol. 107, 382–394 (1985). First demonstration that meiotic chromatin is implicated in the induction of oocyte cortical polarity. Demonstrates a crucial role for actin but not microtubules in the movement and maintenance of meiotic chromatin at the subcortical location.
Van Blerkom, J. Microtubule mediation of cytoplasmic and nuclear maturation during the early stages of resumed meiosis in cultured mouse oocytes. Proc. Natl Acad. Sci. USA 88, 5031–5035 (1991).
Albertini, D. F. Cytoplasmic reorganization during the resumption of meiosis in cultured preovulatory rat oocytes. Dev. Biol. 120, 121–131 (1987).
Maro, B., Howlett, S. K. & Webb, M. Non-spindle microtubule organizing centers in metaphase II-arrested mouse oocytes. J. Cell Biol. 101, 1665–1672 (1985).
Messinger, S. M. & Albertini, D. F. Centrosome and microtubule dynamics during meiotic progression in the mouse oocyte. J. Cell Sci. 100, 289–298 (1991).
Hassold, T. & Hunt, P. To err (meiotically) is human: the genesis of human aneuploidy. Nature Rev. Genet. 2, 280–291 (2001).
Albertini, D. F. et al. Meiotic spindle dynamics in human oocytes following slow-cooling cryopreservation. Hum. Reprod. 24, 2114–2123 (2009).
Verlhac, M. H., Lefebvre, C., Guillaud, P., Rassinier, P. & Maro, B. Asymmetric division in mouse oocytes: with or without Mos. Curr. Biol. 10, 1303–1306 (2000).
Combelles, C. M. & Albertini, D. F. Microtubule patterning during meiotic maturation in mouse oocytes is determined by cell cycle-specific sorting and redistribution of γ-tubulin. Dev. Biol. 239, 281–294 (2001).
Jones, K. T. & Lane, S. I. Chromosomal, metabolic, environmental, and hormonal origins of aneuploidy in mammalian oocytes. Exp. Cell Res. 318, 1394–1399 (2012).
Kitajima, T. S., Ohsugi, M. & Ellenberg, J. Complete kinetochore tracking reveals error-prone homologous chromosome biorientation in mammalian oocytes. Cell 146, 568–581 (2011).
Schatten, G. The centrosome and its mode of inheritance: the reduction of the centrosome during gametogenesis and its restoration during fertilization. Dev. Biol. 165, 299–335 (1994).
Gueth-Hallonet, C. et al. γ-tubulin is present in acentriolar MTOCs during early mouse development. J. Cell Sci. 103, 157–166 (1993).
Ozil, J. P. & Huneau, D. Activation of rabbit oocytes: the impact of the Ca2+ signal regime on development. Development 128, 917–928 (2001).
Szollosi, D. & Ozil, J. P. De novo formation of centrioles in parthenogenetically activated, diploidized rabbit embryos. Biol. Cell 72, 61–66 (1991).
Verlhac, M. H., de Pennart, H., Maro, B., Cobb, M. H. & Clarke, H. J. MAP kinase becomes stably activated at metaphase and is associated with microtubule-organizing centers during meiotic maturation of mouse oocytes. Dev. Biol. 158, 330–340 (1993).
Verlhac, M. H. et al. Mos is required for MAP kinase activation and is involved in microtubule organization during meiotic maturation in the mouse. Development 122, 815–822 (1996).
Barrett, S. L. & Albertini, D. F. Allocation of γ-tubulin between oocyte cortex and meiotic spindle influences asymmetric cytokinesis in the mouse oocyte. Biol. Reprod. 76, 949–957 (2007).
Schuh, M. & Ellenberg, J. Self-organization of MTOCs replaces centrosome function during acentrosomal spindle assembly in live mouse oocytes. Cell 130, 484–498 (2007). Elucidates the long-suspected driving force for acentriolar spindle assembly in mouse oocytes using live-cell imaging of MTOCs and tubulin to reconstruct the events involved in meiotic spindle assembly.
Courtois, A., Schuh, M., Ellenberg, J. & Hiiragi, T. The transition from meiotic to mitotic spindle assembly is gradual during early mammalian development. J. Cell Biol. 198, 357–370 (2012). Shows that the transition of a barrel-shaped meiotic spindle to that of a pointed mitotic spindle in blastomeres of the mouse embryo occurs through the gradual reduction of polar MTOCs at successive stages of development.
Siller, K. H. & Doe, C. Q. Spindle orientation during asymmetric cell division. Nature Cell Biol. 11, 365–374 (2009).
Morin, X. & Bellaiche, Y. Mitotic spindle orientation in asymmetric and symmetric cell divisions during animal development. Dev. Cell 21, 102–119 (2011).
Albertini, D. F. & Barrett, S. L. The developmental origins of mammalian oocyte polarity. Semin. Cell Dev. Biol. 15, 599–606 (2004).
Van Blerkom, J. & Bell, H. Regulation of development in the fully grown mouse oocyte: chromosome-mediated temporal and spatial differentiation of the cytoplasm and plasma membrane. J. Embry. Exp. Morph. 93, 213–238 (1986).
Connors, S. A., Kanatsu-Shinohara, M., Schultz, R. M. & Kopf, G. S. Involvement of the cytoskeleton in the movement of cortical granules during oocyte maturation, and cortical granule anchoring in mouse eggs. Dev. Biol. 200, 103–115 (1998).
Maro, B., Johnson, M. H., Webb, M. & Flach, G. Mechanism of polar body formation in the mouse oocyte: an interaction between the chromosomes, the cytoskeleton and the plasma membrane. J. Embryol. Exp. Morphol. 92, 11–32 (1986).
Deng, M. et al. Chromatin-mediated cortical granule redistribution is responsible for the formation of the cortical granule-free domain in mouse eggs. Dev. Biol. 257, 166–176 (2003).
Deng, M., Suraneni, P., Schultz, R. M. & Li, R. The Ran GTPase mediates chromatin signaling to control cortical polarity during polar body extrusion in mouse oocytes. Dev. Cell 12, 301–308 (2007). Shows proximity-dependent induction of oocyte cortical polarity by chromatin through a mechanism that probably involves a RAN·GTP gradient.
Duncan, F. E., Moss, S. B., Schultz, R. M. & Williams, C. J. PAR-3 defines a central subdomain of the cortical actin cap in mouse eggs. Dev. Biol. 280, 38–47 (2005).
Halet, G. & Carroll, J. Rac activity is polarized and regulates meiotic spindle stability and anchoring in mammalian oocytes. Dev. Cell 12, 309–317 (2007).
Deng, M., Williams, C. J. & Schultz, R. M. Role of MAP kinase and myosin light chain kinase in chromosome-induced development of mouse egg polarity. Dev. Biol. 278, 358–366 (2005).
Zheng, Y. G protein control of microtubule assembly. Annu. Rev. Cell Dev. Biol. 20, 867–894 (2004).
Kalab, P. & Heald, R. The RanGTP gradient — a GPS for the mitotic spindle. J. Cell Sci. 121, 1577–1586 (2008).
Caudron, M., Bunt, G., Bastiaens, P. & Karsenti, E. Spatial coordination of spindle assembly by chromosome-mediated signaling gradients. Science 309, 1373–1376 (2005).
Dumont, J. et al. A centriole- and RanGTP-independent spindle assembly pathway in meiosis I of vertebrate oocytes. J. Cell Biol. 176, 295–305 (2007).
Pollard, T. D. Regulation of actin filament assembly by Arp2/3 complex and formins. Annu. Rev. Biophys. Biomol. Struct. 36, 451–477 (2007).
Sun, S. C. et al. Arp2/3 complex regulates asymmetric division and cytokinesis in mouse oocytes. PloS ONE 6, e18392 (2011).
Yi, K. et al. Dynamic maintenance of asymmetric meiotic spindle position through Arp2/3-complex-driven cytoplasmic streaming in mouse oocytes. Nature Cell Biol. 13, 1252–1258 (2011). Documents the presence of cytoplasmic streaming in meiosis II mouse oocytes, its dependence on the ARP2/3 complex and its role in providing the dynamic force that maintains subcortical spindle positioning.
Bielak-Zmijewska, A., Kolano, A., Szczepanska, K., Maleszewski, M. & Borsuk, E. Cdc42 protein acts upstream of IQGAP1 and regulates cytokinesis in mouse oocytes and embryos. Dev. Biol. 322, 21–32 (2008).
Sun, S. C. et al. WAVE2 regulates meiotic spindle stability, peripheral positioning and polar body emission in mouse oocytes. Cell Cycle 10, 1853–1860 (2011).
Zheng, P., Baibakov, B., Wang, X. H. & Dean, J. PtdIns(3,4,5)P3 is constitutively synthesized and required for spindle translocation during meiosis in mouse oocytes. J Cell Sci. 21 Dec 2012 (doi:10.1242/jcs.118042).
Na, J. & Zernicka-Goetz, M. Asymmetric positioning and organization of the meiotic spindle of mouse oocytes requires CDC42 function. Cur.Biol. 16, 1249–1254 (2006).
Luo, J., McGinnis, L. K. & Kinsey, W. H. Fyn kinase activity is required for normal organization and functional polarity of the mouse oocyte cortex. Mol. Reprod. Dev. 76, 819–831 (2009).
Kubiak, J. Z. & Ciemerych, M. A. Cell cycle regulation in early mouse embryos. Novartis Found. Symp. 237, 79–89 (2001).
Leader, B. et al. Formin-2, polyploidy, hypofertility and positioning of the meiotic spindle in mouse oocytes. Nature Cell Biol. 4, 921–928 (2002).
Dumont, J. et al. Formin-2 is required for spindle migration and for the late steps of cytokinesis in mouse oocytes. Dev. Biol. 301, 254–265 (2007).
Schuh, M. & Ellenberg, J. A new model for asymmetric spindle positioning in mouse oocytes. Cur. Biol. 18, 1986–1992 (2008). Uses a live F-actin probe to observe actin dynamics during meiosis I spindle migration and proposes a model in which a pulling force is produced through myosin II acting on an F-actin network to connect the spindle pole with the cortex.
Azoury, J., Lee, K. W., Georget, V., Hikal, P. & Verlhac, M. H. Symmetry breaking in mouse oocytes requires transient F-actin meshwork destabilization. Development 138, 2903–2908 (2011).
Li, H., Guo, F., Rubinstein, B. & Li, R. Actin-driven chromosomal motility leads to symmetry breaking in mammalian meiotic oocytes. Nature Cell Biol. 10, 1301–1308 (2008).
Yi, K., Rubinstein, B., Unruh, J. R., Slaughter, B. D. & Li, R. Sequential actin-based pushing forces drive meiosis I chromosome migration and symmetry breaking in oocytes. J. Cell Biol. (in the press). Uses trajectory analysis to show that meiosis I chromosome migration is biphasic and driven by pushing forces that are generated first from FMN2-mediated actin assembly at the spindle periphery and then by ARP2/3-orchestrated cytoplasmic streaming.
Azoury, J. et al. Spindle positioning in mouse oocytes relies on a dynamic meshwork of actin filaments. Cur. Biol. 18, 1514–1519 (2008).
Cooper, J. A. Effects of cytochalasin and phalloidin on actin. J. Cell Biol. 105, 1473–1478 (1987).
Riedl, J. et al. Lifeact: a versatile marker to visualize F-actin. Nature Methods 5, 605–607 (2008).
Burkel, B. M., von Dassow, G. & Bement, W. M. Versatile fluorescent probes for actin filaments based on the actin-binding domain of utrophin. Cell. Motil. Cytoskeleton 64, 822–832 (2007).
Gutzeit, H. O. & Koppa, R. Time-lapse film analysis of cytoplasmic streaming during late oogenesis of Drosophila. J. Embryol. Exp. Morphol. 67, 101–111 (1982).
Wolke, U., Jezuit, E. A. & Priess, J. R. Actin-dependent cytoplasmic streaming in C. elegans oogenesis. Development 134, 2227–2236 (2007).
Kamiya, N. Physical and chemical basis of cytoplasmic streaming. Ann. Rev. Plant Physiol. 32, 205–236 (1981).
Ajduk, A. et al. Rhythmic actomyosin-driven contractions induced by sperm entry predict mammalian embryo viability. Nature Commun. 2, 417 (2011). Reports oscillatory cytoplasmic flow that is regulated by Ca2+ and actomyosin in fertilized mouse eggs and presents the intriguing link between flow parameters and the developmental potential of the embryo.
Deng, M. & Li, R. Sperm chromatin-induced ectopic polar body extrusion in mouse eggs after ICSI and delayed egg activation. PloS ONE 4, e7171 (2009).
Pinot, M. et al. Confinement induces actin flow in a meiotic cytoplasm. Proc. Natl Acad. Sci. USA 109, 11705–11710 (2012).
Nothnagel, E. A. & Webb, W. W. Hydrodynamic models of viscous coupling between motile myosin and endoplasm in Characean algae. J. Cell Biol. 94, 444–454 (1982).
Kachar, B. & Reese, T. S. The mechanism of cytoplasmic streaming in characean algal cells: sliding of endoplasmic reticulum along actin filaments. J. Cell Biol. 106, 1545–1552 (1988).
Swann, K. et al. Phospholipase C-ζ-induced Ca2+ oscillations cause coincident cytoplasmic movements in human oocytes that failed to fertilize after intracytoplasmic sperm injection. Fertil. Steril. 97, 742–747 (2012).
Theurkauf, W. E., Alberts, B. M., Jan, Y. N. & Jongens, T. A. A central role for microtubules in the differentiation of Drosophila oocytes. Development 118, 1169–1180 (1993).
Theurkauf, W. E., Smiley, S., Wong, M. L. & Alberts, B. M. Reorganization of the cytoskeleton during Drosophila oogenesis: implications for axis specification and intercellular transport. Development 115, 923–936 (1992).
Palacios, I. M. & St Johnston, D. Kinesin light chain-independent function of the kinesin heavy chain in cytoplasmic streaming and posterior localisation in the Drosophila oocyte. Development 129, 5473–5485 (2002).
Serbus, L. R., Cha, B. J., Theurkauf, W. E. & Saxton, W. M. Dynein and the actin cytoskeleton control kinesin-driven cytoplasmic streaming in Drosophila oocytes. Development 132, 3743–3752 (2005).
Ganguly, S., Williams, L. S., Palacios, I. M. & Goldstein, R. E. Cytoplasmic streaming in Drosophila oocytes varies with kinesin activity and correlates with the microtubule cytoskeleton architecture. Proc. Natl Acad. Sci. USA 109, 15109–15114 (2012).
Forrest, K. M. & Gavis, E. R. Live imaging of endogenous RNA reveals a diffusion and entrapment mechanism for nanos mRNA localization in Drosophila. Cur. Biol. 13, 1159–1168 (2003).
Lantz, V. A., Clemens, S. E. & Miller, K. G. The actin cytoskeleton is required for maintenance of posterior pole plasm components in the Drosophila embryo. Mech. Dev. 85, 111–122 (1999).
Nishida, H. Specification of developmental fates in ascidian embryos: molecular approach to maternal determinants and signaling molecules. Int. Rev. Cytol. 217, 227–276 (2002).
Sardet, C., Paix, A., Prodon, F., Dru, P. & Chenevert, J. From oocyte to 16-cell stage: cytoplasmic and cortical reorganizations that pattern the ascidian embryo. Dev. Dyn. 236, 1716–1731 (2007).
Zernicka-Goetz, M. & Huang, S. Stochasticity versus determinism in development: a false dichotomy? Nature Rev. Genet. 11, 743–744 (2010).
Hiiragi, T., Louvet-Vallee, S., Solter, D. & Maro, B. Embryology: does prepatterning occur in the mouse egg? Nature 442, E3–E4 (2006).
Van Blerkom, J. Mitochondria in human oogenesis and preimplantation embryogenesis: engines of metabolism, ionic regulation and developmental competence. Reproduction 128, 269–280 (2004).
Poulton, J. et al. Transmission of mitochondrial DNA diseases and ways to prevent them. PLoS Genet. 6, e1001066 (2010).
Van Blerkom, J. & Runner, M. N. Mitochondrial reorganization during resumption of arrested meiosis in the mouse oocyte. Am. J. Anat. 171, 335–355 (1984).
Tokura, T., Noda, Y., Goto, Y. & Mori, T. Sequential observation of mitochondrial distribution in mouse oocytes and embryos. J. Assist Reprod. Genet. 10, 417–426 (1993).
Sun, Q. Y. et al. Translocation of active mitochondria during pig oocyte maturation, fertilization and early embryo development in vitro. Reproduction 122, 155–163 (2001).
Van Blerkom, J., Davis, P., Mathwig, V. & Alexander, S. Domains of high-polarized and low-polarized mitochondria may occur in mouse and human oocytes and early embryos. Hum. Reprod. 17, 393–406 (2002).
Van Blerkom, J., Davis, P. & Alexander, S. Differential mitochondrial distribution in human pronuclear embryos leads to disproportionate inheritance between blastomeres: relationship to microtubular organization, ATP content and competence. Hum. Reprod. 15, 2621–2633 (2000).
Herr, J. C. et al. Distribution of RNA binding protein MOEP19 in the oocyte cortex and early embryo indicates pre-patterning related to blastomere polarity and trophectoderm specification. Dev. Biol. 314, 300–316 (2008).
Ohsugi, M., Zheng, P., Baibakov, B., Li, L. & Dean, J. Maternally derived FILIA–MATER complex localizes asymmetrically in cleavage-stage mouse embryos. Development 135, 259–269 (2008).
Li, L., Baibakov, B. & Dean, J. A subcortical maternal complex essential for preimplantation mouse embryogenesis. Dev. Cell 15, 416–425 (2008). Presents the first biochemical and functional characterization of a protein complex in the oocyte cortex that directly links oogenesis to embryogenesis in mammals.
Zheng, P. & Dean, J. Role of Filia, a maternal effect gene, in maintaining euploidy during cleavage-stage mouse embryogenesis. Proc. Natl Acad. Sci. USA 106, 7473–7478 (2009).
Yurttas, P. et al. Role for PADI6 and the cytoplasmic lattices in ribosomal storage in oocytes and translational control in the early mouse embryo. Development 135, 2627–2636 (2008).
Tashiro, F. et al. Maternal-effect gene Ces5/Ooep/Moep19/Floped is essential for oocyte cytoplasmic lattice formation and embryonic development at the maternal-zygotic stage transition. Genes Cells 15, 813–828 (2010).
Morency, E., Anguish, L. & Coonrod, S. Subcellular localization of cytoplasmic lattice-associated proteins is dependent upon fixation and processing procedures. PloS ONE 6, e17226 (2011).
Flemr, M., Ma, J., Schultz, R. M. & Svoboda, P. P-body loss is concomitant with formation of a messenger RNA storage domain in mouse oocytes. Biol. Reprod. 82, 1008–1017 (2010).
Acknowledgements
This work is supported in part by a grant from the US National Institutes of Health (NIH) (grant P01 GM 066311) (to R.L.) and an ESHE fund (to D.F.A.). The authors apologize to the researchers whose work could not be cited owing to space limitations, especially those who have developed many transgenic lines that have revealed so much about mouse oocyte biology.
Author information
Authors and Affiliations
Ethics declarations
Competing interests
The authors declare no competing financial interests.
Related links
Glossary
- Polar body
-
The daughter cell which is much smaller than the oocyte that results from each of the two meiotic cell divisions. The first polar body mostly degenerates within hours of formation, whereas the second polar body persists intact through the early cleavage stages of embryonic development.
- Gap junctions
-
Intercellular channel structures formed by connexin proteins that connect neighbouring cells to allow passage of nutrients, ions and signalling molecules.
- Adherens junctions
-
Protein complexes that contain cadherin and catenin proteins. They are formed between neighbouring cells in the tissue and serve not only to maintain cell–cell adhesion but also to regulate intracellular signalling and cytoskeletal organization.
- Maternal effect
-
Refers to when the phenotype of an organism reflects the genotype of the mother (rather than its own genotype). This is often due to the mother supplying gene products (mRNA and/or proteins) to the embryo.
- Germinal vesicle
-
The large nucleus of the primary oocyte before meiosis is completed.
- Polyspermy
-
Referring to one egg being fertilized by multiple sperms.
- Microtubule-organizing centres
-
(MTOCs). Major sites of microtubule nucleation and anchoring. MTOCs in mammalian meiotic spindles lack centrioles and are inherited by the embryos until embryos are able to assemble centriole-containing centrosomes.
- Bivalents
-
Paired homologous chromosomes in meiosis I.
- Reductional chromosome segregation
-
During meiosis I, pairs of homologous chromosomes segregate to opposite sides of the cell division plane, resulting in daughter cells with half of the chromosome number of the immature oocyte or somatic cells.
- Equational chromosome segregation
-
During meiosis II, sister chromatids separate and segregate to the two daughter cells, thus maintaining the same chromosome number as the post-meiosis I oocyte.
- Centrosomes
-
Organelles that function as the main microtubule organizing centres (MTOCs) in animal cells. They comprise two orthogonally arranged centrioles surrounded by an amorphous protein mass termed the pericentriolar material (PCM). Centrosomes nucleate the mitotic spindle and regulate cell cycle progression.
- Parthenogenetic embryos
-
Embryos obtained by asexual reproduction, whereby embryo growth and development occur without fertilization.
- Partitioning defective 3
-
(PAR3). A member of the conserved family of PAR proteins, which polarize cells during animal development.
- Actin-related protein 2/3
-
(ARP2/3). A highly conserved seven-subunit protein complex that contains two actin-related proteins, ARP2 and ARP3, and nucleates an actin filament at a ∼70° angle from the side of an existing actin filament.
- Formin
-
Formins are an evolutionarily conserved family of actin nucleators. Their nucleation activity is accomplished by a dimer of the formin homology 2 (FH2) domain that then tracks along the elongating actin barbed end.
Rights and permissions
About this article
Cite this article
Li, R., Albertini, D. The road to maturation: somatic cell interaction and self-organization of the mammalian oocyte. Nat Rev Mol Cell Biol 14, 141–152 (2013). https://doi.org/10.1038/nrm3531
Published:
Issue Date:
DOI: https://doi.org/10.1038/nrm3531
This article is cited by
-
Distinct characteristics of the DNA damage response in mammalian oocytes
Experimental & Molecular Medicine (2024)
-
Identification of transcriptome characteristics of granulosa cells and the possible role of UBE2C in the pathogenesis of premature ovarian insufficiency
Journal of Ovarian Research (2023)
-
Integrated bioinformatics analysis elucidates granulosa cell whole-transcriptome landscape of PCOS in China
Journal of Ovarian Research (2023)
-
Novel variants in TUBB8 gene cause multiple phenotypic abnormalities in human oocytes and early embryos
Journal of Ovarian Research (2023)
-
Simulated microgravity reduces quality of ovarian follicles and oocytes by disrupting communications of follicle cells
npj Microgravity (2023)