Building the cell: design principles of cellular architecture (original) (raw)

• Marsh, B. J., Mastronarde, D. N., Buttle, K. F., Howell, K. E. & McIntosh, J. R. Organellar relationships in the Golgi region of the pancreatic β cell line, HIT-T15, visualized by high resolution electron tomography. Proc. Natl Acad. Sci. USA 98, 2399–2406 (2001).
  CAS PubMed PubMed Central Google Scholar
• Minton, A. P. How can biochemical reactions within cells differ from those in test tubes? J. Cell Sci. 119, 2863–2869 (2006).
  CAS PubMed Google Scholar
• Karsenti, E. Self-organization in cell biology: a brief history. Nature Rev. Mol. Cell Biol. 9, 255–262 (2008).
  CAS Google Scholar
• West, G. B., Brown, J. H. & Enquist, B. J. in Scaling in Biology (eds J. H. Brown & G. B. West) (Oxford University Press, New York, 2000).
  Google Scholar
• Wheatley, D. N. & Bowser, S. S. Length control of primary cilia: analysis of monociliate and multiciliate PtK1 cells. Biol. Cell 92, 573–582 (2000).
  CAS PubMed Google Scholar
• Tam, L. W., Wilson, N. F. & Lefebvre, P. A. A CDK-related kinase regulates the length and assembly of flagella in Chlamydomonas. J. Cell Biol. 176, 819–829 (2007).
  CAS PubMed PubMed Central Google Scholar
• Yan, M., Rayapuram, N. & Subramani, S. The control of peroxisome number and size during division and proliferation. Curr. Opin. Cell Biol. 17, 376–383 (2005).
  CAS PubMed Google Scholar
• Guo, Y. et al. Functional genomic screen reveals genes involved in lipid-droplet formation and utilization. Nature 453, 657–661 (2008).
  CAS PubMed PubMed Central Google Scholar
• Katsura, I. Determination of bacteriophage λ tail length by a protein ruler. Nature 327, 73–75 (1987). Demonstrates that the length of a biological structure can be determined by the length of a single protein, which apparently acts as a ruler during assembly.
  CAS PubMed Google Scholar
• Shibata, S. et al. FliK regulates flagellar hook length as an internal ruler. Mol. Microbiol. 64, 1404–1415 (2007).
  CAS PubMed Google Scholar
• Journet, L., Agrain, C., Broz, P. & Cornelis, G. R. The needle length of bacterial injectisomes is determined by a molecular ruler. Science 302, 1757–1760 (2003).
  CAS PubMed Google Scholar
• Fowler, V. M., McKeown, C. R. & Fischer, R. S. Nebulin: does it measure up as a ruler? Curr. Biol. 16, R18–R20 (2006).
  CAS PubMed Google Scholar
• Stephens, R. E. Quantal tektin synthesis and ciliary length in sea-urchin embryos. J. Cell Sci. 92, 403–413 (1989).
  CAS PubMed Google Scholar
• Rosenbaum, J. L., Moulder, J. E. & Ringo, D. L. Flagellar elongation and shortening in Chlamydomonas. The use of cycloheximide and colchicine to study the synthesis and assembly of flagellar proteins. J. Cell Biol. 41, 600–619 (1969).
  CAS PubMed PubMed Central Google Scholar
• Lefebvre, P. A. & Rosenbaum, J. L. Regulation of the synthesis and assembly of ciliary and flagellar proteins during regeneration. Ann. Rev. Cell Biol. 2, 517–546 (1986).
  CAS PubMed Google Scholar
• Marshall, W. F., Qin, H., Rodrigo Brenni, M. & Rosenbaum, J. L. Flagellar length control system: testing a simple model based on intraflagellar transport and turnover. Mol. Biol. Cell 16, 270–278 (2005).
  CAS PubMed PubMed Central Google Scholar
• Sriburi, R., Jackowski, S., Mori, K. & Brewer, J. W. XBP1: a link between the unfolded protein response, lipid biosynthesis, and biogenesis of the endoplasmic reticulum. J. Cell Biol. 167, 35–41 (2004).
  CAS PubMed PubMed Central Google Scholar
• Cox, J. S., Chapman, R. E. & Walter, P. The unfolded protein response coordinates the production of endoplasmic reticulum protein and endoplasmic reticulum membrane. Mol. Biol. Cell 8, 1805–1814 (1997).
  CAS PubMed PubMed Central Google Scholar
• Bernales, S., McDonald, K. L. & Walter, P. Autophagy counterbalances endoplasmic reticulum expansion during the unfolded protein response. PLoS Biol. 4, e413 (2006).
  Google Scholar
• Coyne, B. & Rosenbaum, J. L. Flagellar elongation and shortening in Chlamydomonas. II. Re-utilization of flagellar proteins. J. Cell Biol. 47, 777–781 (1970).
  CAS PubMed PubMed Central Google Scholar
• Marshall, W. F. & Rosenbaum, J. L. Intraflagellar transport balances continuous turnover of outer doublet microtubules: implications for flagellar length control. J. Cell Biol. 155, 405–414 (2001). Presents a simple size-control mechanism in which size-dependent assembly counteracts size-independent disassembly, with the two processes balancing only at a unique value for size.
  CAS PubMed PubMed Central Google Scholar
• Rosenbaum, J. L. & Witman, G. B. Intraflagellar transport. Nature Rev. Mol. Cell Biol. 3, 813–825 (2002).
  CAS Google Scholar
• Tyska, M. J. & Mooseker, M. S. MYO1A (brush border myosin I) dynamics in the brush border of LLC-PK1-CL4 cells. Biophys. J. 82, 1869–1883 (2002).
  CAS PubMed PubMed Central Google Scholar
• Rzadzinska, A. K., Schneider, M. E., Davies, C., Riordan, G. P. & Kachar, B. An actin molecular treadmill and myosins maintain stereocilia functional architecture and self-renewal. J. Cell Biol. 164, 887–897 (2004).
  CAS PubMed PubMed Central Google Scholar
• Keener, J. How Salmonella typhimurium measures the length of flagellar filaments. Bull. Math. Biol. 68, 1761–1778 (2006).
  CAS PubMed Google Scholar
• Varga, V. et al. Yeast kinesin-8 depolymerizes microtubules in a length-dependent manner. Nature Cell Biol. 8, 957–962 (2006).
  CAS PubMed Google Scholar
• Burbank, K. S., Mitchison, T. J. & Fisher, D. S. Slide-and-cluster models for spindle assembly. Curr. Biol. 17, 1373–1383 (2007).
  CAS PubMed Google Scholar
• Hennis, A. S. & Birky, C. W. Stochastic partitioning of chloroplasts at cell division in the alga Olisthodiscus, and compensating control of chloroplast replication. J. Cell Sci. 70, 1–15 (1984).
  CAS PubMed Google Scholar
• Marshall, W. F. Stability and robustness of an organelle number control system: modeling and measuring homeostatic regulation of centriole abundance. Biophys. J. 93, 1818–1833 (2007).
  CAS PubMed PubMed Central Google Scholar
• Marshall, W. F., Vucica, Y. & Rosenbaum, J. L. Kinetics and regulation of de novo centriole assembly. Implications for the mechanism of centriole duplication. Curr. Biol. 11, 308–317 (2001).
  CAS PubMed Google Scholar
• La Terra, S. et al. The de novo centriole assembly pathway in HeLa cells: cell cycle progression and centriole assembly/maturation. J. Cell Biol. 168, 713–722 (2005).
  CAS PubMed PubMed Central Google Scholar
• Paulsson, J. & Ehrenberg, M. Noise in a minimal regulatory network: plasmid copy number control. Quart. Rev. Biophys. 34, 1–59 (2001).
  CAS Google Scholar
• Umen, J. G. The elusive sizer. Curr. Opin. Cell Biol. 17, 435–441 (2005).
  CAS PubMed Google Scholar
• Bergeland, T., Widerberg, J., Bakke, O. & Nordeng, T. W. Mitotic partitioning of endosomes and lysosomes. Curr. Biol. 11, 644–651 (2001).
  CAS PubMed Google Scholar
• Ludford, R. J. & Gatenby, J. B. Dictyokinesis in germ cells. Proc. R. Soc. Lond., B, Biol. Sci. 92, 235–244 (1921).
  Google Scholar
• Lucocq, J. M. & Warren, G. Fragmentation and partitioning of the Golgi apparatus during mitosis in HeLA cells. EMBO J. 6, 3239–3246 (1987).
  CAS PubMed PubMed Central Google Scholar
• Warren, G. Membrane partitioning during cell division. Annu. Rev. Biochem. 62, 323–348 (1993).
  CAS PubMed Google Scholar
• Farré, J. C. & Subramani, S. Peroxisome turnover by micropexophagy: an autophagy-related process. Trends Cell Biol. 14, 515–523 (2004).
  PubMed Google Scholar
• Morgan, T. H. Regeneration of proportionate structures in Stentor. Biol. Bull. 2, 311–328 (1901).
  Google Scholar
• Bakowska, J. & Jerka-Dziadosz, M. Ultrastructural aspect of size dependent regulation of surface pattern of complex ciliary organelle in a protozoan ciliate. J. Embryol. Exp. Morph. 59, 355–375 (1980).
  CAS PubMed Google Scholar
• Malawista, S. E. & Van Blaricom, G. Cytoplasts made from human blood polymorphonuclear leukocytes with or without heat: preservation of both motile function and respiratory burst oxidase activity. Proc. Natl Acad. Sci. USA 84, 454–458 (1987).
  CAS PubMed PubMed Central Google Scholar
• Verkhkovsky, A. B., Svitkina, T. M. & Borisy, G. G. Self-polarization and directional motility of cytoplasm. Curr. Biol. 9, 11–20 (1999).
  Google Scholar
• Wedlich-Soldner, R., Altschuler, S., Wu, L. & Li, R. Spontaneous cell polarization through actomyosin-based delivery of the Cdc42 GTPase. Science 299, 1231–1235 (2003). Demonstrates a simple self-organizing polarity system by combining modelling and experimental measurements.
  CAS PubMed Google Scholar
• Xu, J. et al. Divergent signals and cytoskeletal assemblies regulate self-organizing polarity in neutrophils. Cell 114, 201–214 (2003).
  CAS PubMed Google Scholar
• Yam, P. T. et al. Actin–myosin network reorganization breaks symmetry at the cell rear to spontaneously initiate polarized cell motility. J. Cell Biol. 178, 1207–1221 (2007).
  CAS PubMed PubMed Central Google Scholar
• Ozbudak, E. M., Becskei, A. & van Oudenaarden, A. A system of counteracting feedback loops regulates Cdc42p activity during spontaneous cell polarization. Dev. Cell 9, 565–571 (2005).
  CAS PubMed Google Scholar
• Jones, C. et al. Ciliary proteins link basal body polarization to planar cell polarity regulation. Nature Genet. 40, 69–77 (2008).
  CAS PubMed Google Scholar
• Montcouquiol, M. et al. Identification of Vangl2 and Scrb1 as planar polarity genes in mammals. Nature 423, 173–177 (2003).
  CAS PubMed Google Scholar
• Kupfer, A., Dennert, G. & Singer, S. J. Polarization of the Golgi apparatus and the microtubule-organizing center within cloned natural killer cells bound to their targets. Proc. Natl Acad. Sci. USA 80, 7224–7228 (1983).
  CAS PubMed PubMed Central Google Scholar
• Sanchez-Madrid, F. & Serrador, J. M. Mitochondrial redistribution: adding new players to the chemotaxis game. Trends Immunol. 38, 193–196 (2007).
  Google Scholar
• Kupfer, A., Louvard, D. & Singer, S. J. Polarization of the Golgi apparatus and the microtubule-organizing center in cultured fibroblasts at the edge of an experimental wound. Proc. Natl Acad. Sci. USA 79, 2603–2607 (1982).
  CAS PubMed PubMed Central Google Scholar
• Zmuda, J. F. & Rivas, R. J. The Golgi apparatus and the centrosome are localized to the sites of newly emerging axons in cerebellar granule neurons in vitro. Cell. Motil. Cytoskel. 41, 18–38 (1998).
  CAS Google Scholar
• Agutter, P. S. & Wheatley, D. N. Random walks and cell size. Bioessays 22, 1018–1023 (2000).
  CAS PubMed Google Scholar
• Verkman, A. S. Solute and macromolecule diffusion in cellular aqueous compartments. Trends Biochem. Sci. 27, 27–33 (2002).
  CAS PubMed Google Scholar
• Sun, J. & Weinstein, H. Toward realistic modeling of dynamic processes in cell signaling: quantification of macromolecular crowding effects. J. Chem. Phys. 127, 155105 (2007).
  PubMed Google Scholar
• Ridgway, D. et al. Coarse-grained molecular simulation of diffusion and reaction kinetics in a crowded virtual cytoplasm. Biophys. J. 94, 3748–3759 (2008).
  CAS PubMed PubMed Central Google Scholar
• Prodon, F., Sardet, C. & Nishida, H. Cortical and cytoplasmic flows driven by actin microfilaments polarize the cortical ER–mRNA domain along the a–v axis in ascidian oocytes. Dev. Biol. 313, 682–699 (2008).
  CAS PubMed Google Scholar
• Gomes, E. R., Jani, S. & Gundersen, G. G. Nuclear movement regulated by Cdc42, MRCK, myosin, and actin flow establishes MTOC polarization in migrating cells. Cell 121, 451–463 (2005).
  CAS PubMed Google Scholar
• Carvalho, P., Tirnauer, J. S. & Pellman, D. Surfing on microtubule ends. Trends Cell Biol. 13, 229–237 (2003).
  CAS PubMed Google Scholar
• Gundersen, G. G., Gomes, E. R. & Wen, Y. Cortical control of microtubule stability and polarization. Curr. Opin. Cell Biol. 16, 106–112 (2004).
  CAS PubMed Google Scholar
• Wu, X., Xiang, X. & Hammer, J. A. Motor proteins at the microtubule plus-end. Trends Cell Biol. 16, 135–143 (2006).
  PubMed Google Scholar
• Pearson, C. G. & Bloom, K. Dynamic microtubules lead the way for spindle positioning. Nature Rev. Mol. Cell Biol. 5, 481–492 (2004).
  CAS Google Scholar
• Grill, S. W. & Hyman, A. A. Spindle positioning by cortical pulling forces. Dev. Cell 8, 461–465 (2005).
  CAS PubMed Google Scholar
• Watanabe, T., Noritake, J. & Kaibuchi, K. Regulation of microtubules in cell migration. Trends Cell Biol. 15, 76–83 (2005).
  CAS PubMed Google Scholar
• Allan, V. J., Thompson, H. M. & McNiven, M. A. Motoring around the Golgi. Nature Cell Biol. 4, E236–E242 (2002).
  CAS PubMed Google Scholar
• Rios, R. M. & Bornens, M. The Golgi apparatus at the cell centre. Curr. Opin. Cell Biol. 15, 60–66 (2003).
  CAS PubMed Google Scholar
• Barr, F. A. & Egerer, J. Golgi positioning: are we looking at the right MAP? J. Cell Biol. 168, 993–998 (2005).
  CAS PubMed PubMed Central Google Scholar
• Chabin-Brion, K. et al. The Golgi complex is a microtubule-organizing organelle. Mol. Biol. Cell 12, 2047–2060 (2001).
  CAS PubMed PubMed Central Google Scholar
• Efimov, A. et al. Asymmetric CLASP-dependent nucleation of noncentrosomal microtubules at the _trans_-Golgi network. Dev. Cell 12, 917–930 (2007).
  CAS PubMed PubMed Central Google Scholar
• Drabek, K. et al. Role of CLASP2 in microtubule stabilization and the regulation of persistent motility. Curr. Biol. 16, 2259–2264 (2006).
  CAS PubMed Google Scholar
• Feldman, J. L., Geimer, S. & Marshall, W. F. The mother centriole plays an instructive role in defining cell geometry. PLoS Biol. 5, e149 (2007).
  PubMed PubMed Central Google Scholar
• Svetina, S. & Zeks, B. Shape behavior of lipid vesicles as the basis of some cellular processes. Anat. Rec. 268, 215–225 (2002).
  CAS PubMed Google Scholar
• Deuling, H. J. & Helfrich, W. Curvature elasticity of fluid membranes — catalog of vesicle shapes. Journal De Physique 37, 1335–1345 (1976).
  CAS Google Scholar
• Kas, J. & Sackmann, E. Shape transitions and shape stability of giant phospholipid vesicles in pure water induced by area-to-volume changes. Biophys. J. 60, 825–844 (1991).
  CAS PubMed PubMed Central Google Scholar
• Seifert, U. Configurations of fluid membranes and vesicles. Advances in Physics 46, 13–137 (1997).
  CAS Google Scholar
• McMahon, H. T. & Gallop, J. L. Membrane curvature and mechanisms of dynamic cell membrane remodeling. Nature 438, 590–596 (2005).
  CAS PubMed Google Scholar
• Veatch, S. L. & Keller, S. L. Organization in lipid membranes containing cholesterol. Phys. Rev. Lett. 89, 268101 (2002).
  PubMed Google Scholar
• Baumgart, T., Hess, S. T. & Webb, W. W. Imaging coexisting fluid domains in biomembrane models coupling curvature and line tension. Nature 425, 821–824 (2003).
  CAS PubMed Google Scholar
• Dobereiner, H. G., Kas, J., Noppl, D., Sprenger, I. & Sackmann, E. Budding and fission of vesicles. Biophys. J. 65, 1396–1403 (1993).
  CAS PubMed PubMed Central Google Scholar
• Julicher, F. & Lipowsky, R. Domain-induced budding of vesicles. Phys. Rev. Lett. 70, 2964–2967 (1993).
  CAS PubMed Google Scholar
• Roux, A. et al. Role of curvature and phase transition in lipid sorting and fission of membrane tubules. EMBO J. 24, 1537–1545 (2005).
  CAS PubMed PubMed Central Google Scholar
• Zimmerberg, J. & Kozlov, M. M. How proteins produce cellular membrane curvature. Nature Rev. Mol. Cell Biol. 7, 9–19 (2006).
  CAS Google Scholar
• Farsad, K. & De Camilli, P. Mechanisms of membrane deformation. Curr. Opin. Cell Biol. 15, 372–381 (2003).
  CAS PubMed Google Scholar
• Antonny, B. Membrane deformation by protein coats. Curr. Opin. Cell Biol. 18, 386–394 (2006).
  CAS PubMed Google Scholar
• Frost, A. et al. Structural basis of membrane invagination by F-BAR domains. Cell 132, 807–817 (2008). Describes how a class of membrane curvature-inducing proteins operates at a structural level.
  CAS PubMed PubMed Central Google Scholar
• Shnyrova, A. V. et al. Vesicle formation by self-assembly of membrane-bound matrix proteins into a fluidlike budding domain. J. Cell Biol. 179, 627–633 (2007).
  CAS PubMed PubMed Central Google Scholar
• Shibata, Y., Voeltz, G. K. & Rapoport, T. A. Rough sheets and smooth tubules. Cell 126, 435–439 (2006).
  CAS PubMed Google Scholar
• Dabora, S. L. & Sheetz, M. P. The microtubule-dependent formation of a tubulovesicular network with characteristics of the ER from cultured cell extracts. Cell 54, 27–35 (1988).
  CAS PubMed Google Scholar
• Vale, R. D. & Hotani, H. Formation of membrane networks in vitro by kinesin-driven microtubule movement. J. Cell Biol. 107, 2233–2241 (1988).
  CAS PubMed Google Scholar
• Leduc, C. et al. Cooperative extraction of membrane nanotubes by molecular motors. Proc. Natl Acad. Sci. USA 101, 17096–17101 (2004).
  CAS PubMed PubMed Central Google Scholar
• Dreier, L. & Rapoport, T. A. In vitro formation of the endoplasmic reticulum occurs independently of microtubules by a controlled fusion reaction. J. Cell Biol. 148, 883–898 (2000).
  CAS PubMed PubMed Central Google Scholar
• Waterman-Storer, C. M. & Salmon, E. D. Endoplasmic reticulum membrane tubules are distributed by microtubules in living cells using three distinct mechanisms. Curr. Biol. 8, 798–806 (1998).
  CAS PubMed Google Scholar
• Vedrenne, C. & Hauri, H. P. Morphogenesis of the endoplasmic reticulum: beyond active membrane expansion. Traffic 7, 639–646 (2006).
  CAS PubMed Google Scholar
• Voeltz, G. K., Prinz, W. A., Shibata, Y., Rist, J. M. & Rapoport, T. A. A class of membrane proteins shaping the tubular endoplasmic reticulum. Cell 124, 573–586 (2006). Shows how proteins can induce specific changes in membrane shape.
  CAS PubMed Google Scholar
• De Craene, J. O. et al. Rtn1p is involved in structuring the cortical endoplasmic reticulum. Mol. Biol. Cell 17, 3009–3020 (2006).
  CAS PubMed PubMed Central Google Scholar
• Hu, J. et al. Membrane proteins of the endoplasmic reticulum induce high-curvature tubules. Science 319, 1247–1250 (2008).
  CAS PubMed Google Scholar
• Bereiter-Hahn, J. Behavior of mitochondria in the living cell. Int. Rev. Cytol. 122, 1–63 (1990).
  CAS PubMed Google Scholar
• Okamoto, K. & Shaw, J. M. Mitochondrial morphology and dynamics in yeast and multicellular eukaryotes. Annu. Rev. Genet. 39, 503–536 (2005).
  CAS PubMed Google Scholar
• Boldogh, I. R. & Pon, L. A. Mitochondria on the move. Trends Cell Biol. 17, 502–510 (2007).
  CAS PubMed Google Scholar
• Hoppins, S., Lackner, L. & Nunnari, J. The machines that divide and fuse mitochondria. Annu. Rev. Biochem. 76, 751–780 (2007).
  CAS PubMed Google Scholar
• Merz, S., Hammermeister, M., Altmann, K., Durr, M. & Westermann, B. Molecular machinery of mitochondrial dynamics in yeast. Biol. Chem. 388, 917–926 (2007).
  CAS PubMed Google Scholar
• Nunnari, J. et al. Mitochondrial transmission during mating in Saccharomyces cerevisiae is determined by mitochondrial fusion and fission and the intramitochondrial segregation of mitochondrial DNA. Mol. Biol. Cell 8, 1233–1242 (1997).
  CAS PubMed PubMed Central Google Scholar
• Arakaki, N. et al. Dynamics of mitochondria during the cell cycle. Biol. Pharm. Bull. 29, 1962–1965 (2006).
  CAS PubMed Google Scholar
• Taguchi, N., Ishihara, N., Jofuku, A., Oka, T. & Mihara, K. Mitotic phosphorylation of dynamin-related GTPase Drp1 participates in mitochondrial fission. J. Biol. Chem. 282, 11521–11529 (2007).
  CAS PubMed Google Scholar
• Youle, R. J. & Karbowski, M. Mitochondrial fission in apoptosis. Nature Rev. Mol. Cell Biol. 6, 657–663 (2005).
  CAS Google Scholar
• Egner, A., Jakobs, S. & Hell, S. W. Fast 100-nm resolution three-dimensional microscope reveals structural plasticity of mitochondria in live yeast. Proc. Natl Acad. Sci. USA 99, 3370–3375 (2002).
  CAS PubMed PubMed Central Google Scholar
• Keren, K. et al. Mechanism of shape determination in motile cells. Nature 453, 475–480 (2008).
  CAS PubMed PubMed Central Google Scholar
• Solomon, F. Specification of cell morphology by endogenous determinants. J. Cell Biol. 90, 547–553 (1981).
  CAS PubMed Google Scholar
• Bouck, G. B. & Brown, D. L. Microtubule biogenesis and cell shape in Ochromonas: II. The role of nucleating sites in shape development. J. Cell Biol. 56, 360–378 (1987).
  Google Scholar
• Albrecht-Buehler, G. Daughter 3T3 cells. Are they mirror images of each other? J. Cell Biol. 72, 595–603 (1977). Shows images of sister cells that seem to be mirror images of each other, suggesting the propagation of structural determinants to the two daughter cells in opposite directions by the mitotic spindle during division.
  CAS PubMed Google Scholar
• Solomon, F. Detailed neurite morphologies of sister neuroblastoma cells are related. Cell 16, 165–169 (1979).
  CAS PubMed Google Scholar
• Locke, M. Is there somatic inheritance of intracellular patterns? J. Cell Biol. 96, 563–567 (1990).
  Google Scholar
• Tawk, M. et al. A mirror-symmetric cell division that orchestrates neuroepithelial morphogenesis. Nature 446, 797–800 (2007).
  CAS PubMed Google Scholar
• Beisson, J. & Sonneborn, T. M. Cytoplasmic inheritance of the organization of the cell cortex in Paramecium aurelia. Proc. Natl Acad. Sci. USA 53, 275–282 (1965). A classic study showing that when part of the cortex of a ciliate is rearranged, the rearrangement can propagate for multiple cell divisions without any accompanying genetic change.
  CAS PubMed PubMed Central Google Scholar
• Shi, X. B. & Frankel, J. Morphology and development of mirror-image doublets of Stylonychia mytilus. J. Protozool. 37, 1–13 (1990).
  CAS PubMed Google Scholar