The machinery of mitochondrial fusion, division, and distribution, and emerging connections to apoptosis

Mitochondrion 4 (2004) 285–308 www.elsevier.com/locate/mito

Invited review

The machinery of mitochondrial fusion, division, and distribution, and emerging connections to apoptosis Karen G. Hales* Department of Biology, Davidson College, Davidson, NC 28036-7118, USA Received 9 March 2004; received in revised form 19 May 2004; accepted 25 May 2004

Abstract Mitochondrial undergo regulated fusion and division in many organisms and cell types, and each event is mediated by a different complex of proteins each containing at least one large GTPase. The mitochondrial fusion and division molecular machinery is in large part conserved; recent studies show a functional connection between some of these proteins and the apoptotic cascade. Mitochondria also undergo directed movement in cells, and the gene products that attach and propel mitochondria along cytoskeletal elements (actin filaments in some organisms, microtubules in others) are becoming gradually elucidated. q 2004 Elsevier B.V. and Mitochondria Research Society. All rights reserved. Keywords: Fission; Fusion; Distribution; Morphogenesis; Apoptosis

1. Introduction 1.1. Overview Mitochondria are dynamic organelles which undergo fusion, division, and directed movement in many cell types. Tight regulation of mitochondrial morphogenesis during growth and differentiation may enable more efficient generation and distribution of ATP as well as more accurate partitioning of mitochondria during cell division. Research in yeast and other model systems has begun to elucidate the molecular machinery underlying the fusion, division, * Tel.: C1-704-894-2324; fax: C1-704-894-2512. E-mail address: [email protected]

and distribution of mitochondria. In recent years, a human genetic disorder has been attributed to mutations in a gene involved in mitochondrial fusion, and more recent results suggest roles for mitochondrial fission and fusion mediators in apoptosis. In this review, I summarize our knowledge to date of the mechanisms of mitochondrial fusion, fission, and distribution, as well as the relationship of mitochondrial morphogenesis to programmed cell death. 1.2. Contexts for and adaptive advantages of regulated mitochondrial fusion and division In many single celled eukaryotes, mitochondria fuse into a branched reticulum under certain growth conditions and at particular points in the cell cycle or

1567-7249/$ - see front matter q 2004 Elsevier B.V. and Mitochondria Research Society. All rights reserved. doi:10.1016/j.mito.2004.05.007

286

K.G. Hales / Mitochondrion 4 (2004) 285–308

life cycle. The existence of a mitochondrial reticulum in the budding yeast Saccharomyces cerevisiae was initially demonstrated through transmission electron microscopy and reconstruction of mitochondrial shape from serial thin sections of cells (Hoffmann and Avers, 1973). Mitochondrial shape in wild type S. cerevisiae cells is always in flux, with overall form dictated by an equilibrium between fusion and fission events (Nunnari et al., 1997; Sesaki and Jensen, 1999). When haploid cells mate, mitochondria from each parent tend to fuse together in the zygote (Nunnari et al., 1997). The balance is tipped toward fusion also during meiosis and sporulation of diploids (Miyakawa et al., 1994). In interphase S. cerevisiae cells, mitochondrial reticula tend to predominate during anaerobic respiration and under aerobic conditions when excess glucose is present; fragmentation predominates with limited glucose present, and also during mitosis under any condition (Stevens, 1981; Visser et al., 1995). Similar cell cycle patterns of mitochondrial fusion and division are seen in the alga Chlamydomonas reinhardtii (Ehara et al., 1995), some strains of the slime mold Physarum polycephalum (Kawano et al., 1993), and other single celled eukaryotes (reviewed in Bereiter-Hahn and Voth (1994) and Kawano et al. (1995)). During spermatogenesis in many non-mammalian higher organisms, mitochondria undergo fusion on a large scale. In Drosophila melanogaster and other insects, all the mitochondria in a post-meiotic spermatid aggregate beside the nucleus and fuse into exactly two giant mitochondrial derivatives, which interweave topologically to form a complex spherical structure called the Nebenkern (Tates, 1971; Fuller, 1993). The two mitochondrial derivatives within the Nebenkern subsequently untangle from each other (perhaps requiring mitochondrial fission events) and elongate beside the growing flagellar axoneme (Fuller, 1993). Mitochondrial fusion occurs during spermatogenesis in some nematodes (Kruger, 1991), mollusks (Hodgson and Bernard, 1986), and plants (Renzaglia and Duckett, 1989). Mitochondria in mammalian sperm form an end-to-end helical array around the sperm midpiece without fusing; however, this array is characterized by ‘stud-like bridging elements’ between adjacent mitochondria (Olson and Winfrey, 1992) that appear to allow aqueous

connection between mitochondrial matrices (Zorov et al., 1990). Mitochondrial fusion and division has been documented in other mammalian tissues. In perinatal rat liver, mitochondrial reticula divide and later reform during early postnatal development (Smith, 1931; Aprille, 1986). In rat diaphragm muscle, mitochondria exist as separate columns parallel to the muscle fibers before birth; later, branches form and connect the columns into complete reticula by the end of two months (Bakeeva et al., 1981). Mitochondrial fusion and division occur in a variety of cultured mammalian cell lines (Legros et al., 2002; Ishihara et al., 2003; Mattenberger et al., 2003), suggesting that perhaps (as in yeast) an equilibrium of fusion and division may determine mitochondrial form in many cell types. Genetic evidence (described later) is beginning to elucidate the roles of mitochondrial morphogenesis in mammalian model systems. Mitochondrial fusion may provide an adaptive advantage in three ways: by enabling ATP production in oxygen-poor regions of cells, by allowing coordinated calcium buffering across the mitochondrial network, and by minimizing the effects of mitochondrial mutations through complementation between fused organelles. The formation and maintenance of a mitochondrial reticulum is thought to allow coordinated and efficient ATP production throughout a cell via lateral transmission of membrane potential (Skulachev, 2001). The potential across the inner membrane generated during electron transport can, in theory, be transmitted over the entire mitochondrion, since the cytosol and mitochondrial matrix are good conductors separated by a membrane of low conductance. This membrane potential is a convertible form of energy currency whose putative transmission would occur faster than ATP diffusion (Bereiter-Hahn and Voth, 1994). Indeed, local photodamage in one region of a mitochondrial network causes almost instant depolarization of the entire reticulum (Skulachev, 2001), consistent with the idea that mitochondrial networks are intracellular proton cables that send energy in the form of a membrane potential deep into the interior of a cell, allowing ATP production in regions where nutrients or oxygen may be limited. Skulachev (1990) extended his model by proposing that mitochondrial reticular membranes could also facilitate lateral transport of oxygen (which is more

K.G. Hales / Mitochondrion 4 (2004) 285–308

soluble in lipids than in water) and fatty acids. It has indeed been shown that gradients of ATP, oxygen, and pH can exist across a cell and that mitochondria in different regions of a cell can show different membrane potentials (reviewed in Bereiter-Hahn and Voth (1994)). Recent results suggest that mitochondrial reticula may also facilitate calcium buffering upon release of calcium from the endoplasmic reticulum or outside the cell. When mitochondria in cultured mammalian cells were induced to fragment by overexpression of the fission protein hFis1, calcium uptake (as measured via fluorescence of a mitochondrial-specific calcium sensor) occurred more slowly and heterogeneously in the fragmented mitochondria as opposed to in the reticulum of control transfected cells (Frieden et al., 2004). In S. cerevisiae, mitochondrial fusion enables complementation between respiratory deficient strains with distinct mitochondrial mutations (Clark-Walker and Miklos, 1975). Inter-mitochondrial complementation in multi-cellular organisms may help minimize the effects of pathogenic mitochondrial mutations that can arise as cells age. Mitochondrial complementation has been documented in human cell culture experiments (Enriquez et al., 2000; Ono et al., 2001), though the biological relevance of this phenomenon is questioned by the vastly different complementation efficiencies noted in these two reports. Ono et al. (2001) electrofused HeLa (cervical carcinoma) cells and 143B (osteosarcoma) cells that each carried different mitochondrial tRNA mutations, using a system that allowed selection of cell hybrids. All isolated hybrids were respiratory-competent and showed normal translation levels, indicating that complementation had occurred efficiently. In contrast, Enriquez et al. (2000) used polyethylene glycol (PEG) to fuse two 143B cell lines, one carrying a mitochondrial tRNA mutation and one carrying a mutation in an NADH dehydrogenase subunit gene. Of the estimated number of cell hybrids formed, fewer than 1% showed respiratory capability and thus complementation. The contradictory complementation results could result from differing activity of the mitochondrial fusion machinery in the cell lines used, though later assays show similar fusion efficiency and seem to eliminate this as a possible explanation (Legros et al., 2002). Nevertheless, experiments in transgenic mice support an in vivo role for

287

mitochondrial fusion in protecting against the effects of pathogenic mitochondrial mutations: mice whose cells contained both wild type and partially deleted mitochondrial genomes showed consistently similar respiratory levels across different tissues carrying wildly varying ratios of the mitochondrial genotypes (Nakada et al., 2001). Furthermore, direct examination of mitochondrial DNA (mtDNA) diffusion in fused cultured cells indicates that mtDNA nucleoids spread fully within fused mitochondria by 12–14 h after polykaryon formation, suggesting that the inefficient functional complementation described above may result from the slower rate at which respiratory complexes are assembled in newly formed heteroplasmic mitochondria (Legros et al., 2004). The conservation of the capacity for mitochondrial division probably stems in part from the need to segregate distinct organelles during cell division, and also in part from the need to balance mitochondrial fusion during different growth stages. Furthermore, mitochondrial division is an event of apoptosis, and recent results (described in a later section) suggest that the division machinery may play a role in the propagation of the apoptotic cascade. The capability of mitochondria to divide may have been conserved in part because of the benefit conferred by the killing of unnecessary, diseased, or neoplastic cells. 1.3. Contexts for and adaptive advantages of regulated mitochondrial distribution In most eukaryotic cells, accurate mitochondrial partitioning during cytokinesis ensures viability of both daughter cells; mechanisms for mitochondrial distribution during mitosis and meiosis have therefore been conserved. During budding in S. cerevisiae, a subset of the mother cell’s mitochondria are transported to the growing daughter cell, while the remainder are anchored in place at the distal end of the mother cell (Yang et al., 1999; Boldogh et al., 2001b). In the fission yeast Schizosaccharomyces pombe, mitochondria appear to segregate evenly at mitosis via association with spindle poles (Yaffe et al., 2003). In other cells that undergo equatorial cytokinesis, mitochondria are often positioned evenly along spindle microtubules, ensuring that each daughter cell inherits a complement (Fuller, 1993; Pereira et al., 1997).

288

K.G. Hales / Mitochondrion 4 (2004) 285–308

In many specialized cells, mitochondria reside in close apposition to energy-consuming subcellular structures, such as between myofibrils in muscle cells (Kirkwood et al., 1986) or near axonemes in ciliated and flagellated cells (e.g. sperm; Fuller, 1993). Cells that have a central requirement for ion transport across the plasma membrane, either for restoration of an electrochemical potential (as in nerve cells) or for inducing osmotic water movement (as in some epithelial cells), show abundant mitochondria at the membrane regions where ion exchange occurs (reviewed in Tyler (1992) and Bereiter-Hahn and Voth (1994)). A well-studied example of regulated mitochondrial transport is in axons, where mitochondria can remain stationary or move back and forth between the cell body and synaptic terminal (Hollenbeck, 1996). Such movement enables ATP generation near the various energy-requiring proteins of the presynaptic membrane and may also buffer Ca2C concentrations (Werth and Thayer, 1994). Regulated placement of mitochondria appears to allow sufficient ATP to be delivered quickly where needed, consistent with Skulachev’s (2001) model above that ATP diffuses slowly. Genetically controlled mitochondrial distribution therefore may have been conserved because of the energy efficiency conferred upon specialized cell types.

2. Machinery of mitochondrial fusion Five gene families that promote mitochondrial fusion have been identified. Two include large, conserved mitochondrial membrane-associated GTPases: Drosophila Fzo and homologs, and S. cerevisiae Mgm1p and homologs. A third component of the mitochondrial fusion machinery (Ugo1p) is also mitochondrial membrane-associated and seems to be unique to single-celled eukaryotes. These three fusion mediators appear to interact physically in vivo. Two other S. cerevisiae proteins affect mitochondrial fusion through regulation of the large GTPases: Mdm30p affects Fzo1p levels in yeast, and Rbd1p/Pcp1p/Ugo2p (conserved in many eukaryotes) cleaves Mgm1p. These five protein families are described below. A seemingly unrelated mitochondrial fusogen from the slime mold P. polycephalum (not elaborated below) is known to be encoded on

a mitochondrial plasmid; continued examination of a candidate gene whose product localizes to the mitochondrial outer membrane may elucidate mitochondrial fusion mechanisms in that system (Takano et al., 2002). 2.1. The Fzo/mitofusin family and the Mdm30p F-box protein Members of the Fzo1p family have been characterized in flies, yeast, and mammals; database searches indicate the existence of nematode homologs. All family members include a GTPase domain near the amino terminus, two closely spaced transmembrane domains near the carboxy terminus, and predicted coiled-coil regions near both ends (Hales and Fuller, 1997; Santel and Fuller, 2001; Hwa et al., 2002). Drosophila Fzo, the founding member of this GTPase family, is required for mitochondrial fusion during spermatogenesis (and thus required for fertility), and is detectable on spermatid mitochondria only around the time of mitochondrial fusion (Hales and Fuller, 1997). While the fzo gene is transcribed only in the male germ line, a second Drosophila homolog, dmfn, is expressed more broadly (Hwa et al., 2002) but has not been characterized genetically. S. cerevisiae FZO1 was identified by homology to Drosophila fzo. Yeast strains deleted for FZO1 contain fragmented mitochondria and eventually lose their mtDNA as a secondary effect (Hermann et al., 1998; Rapaport et al., 1998). Using a mitochondrial fusion assay (Nunnari et al., 1997) in which parental haploid yeast strains were labeled with different mitochondrially targeted fluorophores, Hermann et al. (1998) demonstrated with a temperature sensitive allele that Fzo1p is required for mitochondrial fusion during mating. Protease protection assays indicated that the Fzo1p amino terminal GTPase domain faces the cytosol (Hermann et al., 1998; Rapaport et al., 1998); cofractionation experiments variously showed Fzo1p to be associated with the mitochondrial outer membrane (Rapaport et al., 1998) or with both the outer and inner membranes (Hermann et al., 1998). Fritz et al. (2001) performed further protease protection assays and found that the carboxy terminus of Fzo1p, like the amino terminus, faces the cytosol, suggesting that both transmembrane domains

K.G. Hales / Mitochondrion 4 (2004) 285–308

span the mitochondrial outer membrane with a short linker region in the intermembrane space. This linker region associates peripherally with the inner membrane, since inclusion of a protein denaturant in cofractionation experiments eliminated the association of Fzo1p with the inner membrane (Fritz et al., 2001). Genetic manipulation to change the residues between the transmembrane domains showed that peripheral association of Fzo1p with the mitochondrial inner membrane is crucial for mitochondrial fusion (Fritz et al., 2001). A role for Fzo family members in mitochondrial fusion is conserved in mammals. The human genome includes two Fzo homologs, also known as mitofusins (Mfn1 and Mfn2) (Santel and Fuller, 2001), which show distinct but overlapping expression patterns in brain, heart, muscle, and other tissues (Eura et al., 2003; Santel et al., 2003). Overexpression of Mfn2 in cultured cells leads to perinuclear clustering of mitochondria, apparently mediated by the coiled coil domains (Santel and Fuller, 2001; Rojo et al., 2002; Eura et al., 2003). In some transfected cells (perhaps those with stochastically lower Mfn2 expression levels), or if mitochondrial division is concurrently inhibited, perinuclear clustering does not occur, and abnormally interconnected mitochondrial networks are revealed (Santel and Fuller, 2001; Rojo et al., 2002). Overexpression of Mfn1 increases mitochondrial interconnectivity in cultured cells (Legros et al., 2002; Eura et al., 2003; Santel et al., 2003) and leads to protrusion of outer membrane tubules from the network, which disappear upon simultaneous Mfn2 expression (Eura et al., 2003). The induction of extended mitochondrial networks by overexpressed mitofusins is eliminated when the GTPase domain is altered (Santel and Fuller, 2001; Santel et al., 2003), consistent with results in flies and yeast (Hales and Fuller, 1997; Hermann et al., 1998). Overexpression of these GTPase-altered mitofusins seems to have a dominant negative effect on mitochondrial fusion (Eura et al., 2003; Santel et al., 2003). Alteration of the GTPase region does not affect mitochondrial targeting of Fzo family members (Hales and Fuller, 1997; Hermann et al., 1998; Santel and Fuller, 2001; Santel et al., 2003). Targeting of mitofusins to mitochondria depends instead upon residues near the two adjacent transmembrane domains (Fritz et al., 2001; Santel and Fuller, 2001; Rojo et al., 2002);

289

the resulting membrane topology of the human mitofusins matches that deduced in yeast (Fritz et al., 2001; Rojo et al., 2002). Two research groups used RNA interference (RNAi) to examine mitofusin-mediated mitochondrial fusion in hybrids of human parental cell lines each marked with different mitochondrial fluorophores. Eura et al. (2003) saw different phenotypes in hybrids of Mfn1-repressed cells as compared with Mfn2repressed cells. When Mfn1 was depleted, mitochondria from the two parental cell lines intermingled but did not fuse; however, when Mfn2 was depleted, the two parental mitochondria populations did not seem to even to intermingle, let alone fuse. Ishihara et al. (2003) found that if one mitofusin was eliminated in either parental cell line, fusion failed, while elimination of both mitofusins in one parental cell line (but retention of both mitofusins in the other) did not hinder mitochondrial fusion. The authors conclude that both mitofusins must be present in cis for fusion to occur. This conclusion is somewhat at odds with results from knockout mouse-derived cell lines cell lines (Chen et al., 2003); see below. Three research groups studied the role of membrane potential in mitochondrial fusion assays between differentially marked parental cell lines; complete fusion was recorded only if the mitochondrial inner membrane potential was intact (Legros et al., 2002; Ishihara et al., 2003; Mattenberger et al., 2003). The membrane potential per se, and not its effect on ATP levels, enables mitochondrial fusion, since depletion of ATP by other means did not inhibit fusion (Legros et al., 2002). Mitochondria regained fusion capability upon restoration of the membrane potential, even in the absence of new protein synthesis or if cytoskeletal elements were destabilized (Legros et al., 2002; Ishihara et al., 2003; Mattenberger et al., 2003). It is not clear what effect membrane potential has, if any, on the mitochondrial fusion machinery. Experiments in mammalian model systems are beginning to elucidate the developmental requirements for mitofusins. Chen et al. (2003) made knockout mice lacking either Mfn1 and Mfn2 and then analyzed phenotypes at both the organismal and cellular level. Homozygous mutant mice of either strain die as mid-gestation embryos; Mfn1 mutant embryos are small and deformed but survive

290

K.G. Hales / Mitochondrion 4 (2004) 285–308

slightly longer than Mfn2 embryos, which are morphologically normal though also small. Curiously, Mfn2 (but not Mfn1) mutant placentae show defective giant cell layers, suggesting that mitochondrial fusion mediated by Mfn2 may be unusually important for the viability and function of these polyploid cells. In fibroblast cell lines derived from homozygous knockout mice, Mfn1deficient mitochondria are small, fragmented, and spherical, and they do not appear to move along cytoskeletal tracks. Mfn2-deficient mitochondria are also spherical and fragmented, though slightly larger, and occasionally show what appears to be directed movement. Immunoprecipitation experiments from wild type and knockout cell lines indicate that Mfn1 and Mfn2 can form both homoand heterotypic complexes (Chen et al., 2003), consistent with similar studies of rat mitofusins (Eura et al., 2003). Mitochondrial fusion is severely, but not completely, inhibited in both Mfn1 and Mfn2 knockout cell lines. Exogenous Mfn1 can restore mitochondrial morphology and fusion in the Mfn2 cell line, while exogenous Mfn2 can only partially rescue the Mfn1 phenotype (Chen et al., 2003); these results seem to contradict the RNAi experiments (Ishihara et al., 2003) that show a requirement for both mitofusins in cis; however, as proposed by Chen et al. (2003), perhaps the relative importance of the homotypic and heterotypic mitofusin complexes varies among different cell types, with heterotypic complexes being crucial in the HeLa cells utilized for the RNAi experiments (Ishihara et al., 2003). A study of Mfn2 function in developing rat muscle tissue supports the idea that mitochondrial fusion is important for efficient metabolism. Bach et al. (2003) showed that Mfn2 is upregulated during the formation of mitochondrial reticula in developing muscle cells but is abnormally reduced in a strain of rats genetically predisposed to obesity. Overexpression of mitofusins in cultured myotubes led to perinuclear clustering of mitochondria, consistent with results in other cultured cells, while depletion of mitofusins using antisense techniques let to mitochondrial fragmentation (Bach et al., 2003). In these mitofusin-depleted myotubes, glucose oxidation and oxygen consumption were significantly reduced, raising the possibility that similar metabolic changes in obese

individuals may result at least in part from altered regulation of mitofusins. Mitofusins may be regulated at least in part at the level of protein stability. S. cerevisiae MDM30 encodes an F-box protein which may be part of the ubiquitin/26S proteasome system; cells lacking Mdm30p show elevated Fzo1p, perhaps as a result of hampered degradation, though there is no evidence to date for ubiquitination of Fzo1p in wild type cells (Fritz et al., 2003). Mitochondria in mdm30 mutants display mitochondrial aggregation, consistent with effects of Fzo1p overexpression, and at high temperatures the fzo1-like mtDNA loss is recapitulated in mdm1 mutants (Fritz et al., 2003). The precise relationship between Fzo1p, Mdm30p, and the ubiquitin proteasome system remains to be determined. Since no MDM30 homologs in other organisms are apparent in genome databases, it also remains to be determined whether other Fzo family members are regulated at the level of protein stability. Such regulation is plausible in the case of Drosophila Fzo, which quickly becomes undetectable on spermatid mitochondria (perhaps due to regulated degradation) following the mitochondrial fusion events that form the Nebenkern (Hales and Fuller, 1997). 2.2. UGO1p and its role in mitochondrial fusion Ugo1p was identified as a component of the fusion machinery in S. cerevisiae through a genetic screen based on genetic interactions between FZO1 and DNM1, the latter of which encodes a mitochondrial division protein (described in more detail in Section 3). The mitochondrial fragmentation (and mtDNA loss) seen in fzo1 mutants depends on Dnm1p-mediated division; in other words, dnm1 mutations suppress fzo1 phenotypes (Bleazard et al., 1999; Sesaki and Jensen, 1999). Sesaki and Jensen (2001) identified ugo1 cells in a screen for new mutants that (like fzo1) show mtDNA loss and are suppressible by dnm1. Mating assays between parental strains marked with different mitochondrial fluorophores (Nunnari et al., 1997) and deleted for UGO1 demonstrated that Ugo1p is required for mitochondrial fusion. Ugo1p is a mitochondrial outer membrane protein whose amino terminus faces the cytosol and whose carboxy terminus

K.G. Hales / Mitochondrion 4 (2004) 285–308

resides in the intermembrane space. Coimmunoprecipitation experiments indicate that Ugo1p and Fzo1p physically interact (Sesaki et al., 2003b; Wong et al., 2003). This interaction occurs between the Ugo1p cytoplasmic domain and portions of the two cytoplasmic domains of Fzo1p, and is stronger in the absence of the Fzo1p GTPase domain; perhaps Fzo1p GTPase activity modulates physical interaction with Ugo1p (Sesaki and Jensen, 2004). A version of Fzo1p that lacks carboxy terminal residues required for Ugo1p binding (but that is properly localized to the outer membrane) is unable to promote mitochondrial fusion (Sesaki and Jensen, 2004). Ugo1p binds via its intermembrane space domain to Mgm1p (described below), thus connecting the two GTPases involved in mitochondrial fusion in yeast (Sesaki and Jensen, 2004). No UGO1 homologs besides in fission yeast are represented in sequence databases; it remains to be seen whether other proteins connect Fzo1p and Mgm1p homologs in other organisms. 2.3. The Mgm1p family of dynamin-related proteins and their cleavage by Rbd1p/Pcp1p/Ugo2p rhomboid proteases Mgm1p is a conserved member of the dynamin family of large GTPases, other branches of which are involved in membrane remodeling processes such as the pinching off of endocytic vesicles (Danino and Hinshaw, 2001). In S. cerevisiae, the mgm1 deletion phenotype resembles the fzo1 and ugo1 phenotypes in that (1) mitochondria become fragmented, sometimes aggregating near the nucleus; (2) mtDNA is lost; (3) mitochondrial networks are restored if Dnm1pmediated mitochondrial division is compromised; (4) mitochondrial fusion cannot occur during mating assays, even if dnm1 is mutated (Sesaki et al., 2003b; Wong et al., 2003). Mitochondrial inner membrane structure is aberrant in mgm1 mutants, and this phenotype is rescued if DNM1 is mutated, even though fusion remains defective, indicating that the inner membrane morphology is not the proximate effector of fusion capability (Sesaki et al., 2003b). Mutations in S. cerevisiae mgm1 and its S. pombe homolog msp1 were originally identified by the mtDNA loss phenotype (Jones and Fangman, 1992; Guan et al., 1993; Pelloquin et al., 1999). Faulty

291

transport of mitochondria to a growing bud was also reported in mgm1 mutants (Shepard and Yaffe, 1999), and is likely a secondary effect of perinuclear aggregation. Mutations affecting the Mgm1p GTPase domain severely compromise or eliminate the ability of Mgm1p to promote mitochondrial fusion (Shepard and Yaffe, 1999; Wong et al., 2000, 2003; Sesaki et al., 2003b). In some cases, overexpression of mgm1 alleles mutated in the GTPase domain lead to a dominant negative effect (Shepard and Yaffe, 1999; Wong et al., 2003), consistent with the possibility that Mgm1p (like other dynamins) may form homotypic assemblies. Accordingly, mutations in the Mgm1p putative assembly domain inhibit Mgm1p-mediated mitochondrial fusion in vivo (Wong et al., 2003). Further evidence for Mgm1p self-assembly is that certain pairs of mgm1 alleles show intragenic complementation (Wong et al., 2003). Coimmunoprecipitation experiments indicate that Mgm1p physically interacts with Fzo1p and Ugo1p (Sesaki et al., 2003b; Wong et al., 2003). Ugo1p has distinctbinding sites for Fzo1p and Mgm1p and is required for establishing a complex thereof (Sesaki and Jensen, 2004); it is not clear whether Fzo1p and Mgm1p directly bind to each other once the initial complex is established. The human homolog of MGM1, called OPA1, is mutated in people with autosomal dominant optic atrophy (ADOA) (Alexander et al., 2000; Delettre et al., 2000). While the gene is expressed in many cell types, patients show symptoms only in the eye, becoming visually impaired due to degeneration of retinal ganglion cells. Studies of mouse OPA1 confirm wide expression across many cell types and subcellular localization of the gene product to mitochondria (Misaka et al., 2002; Griparic et al., 2004). It is likely that a mitochondrial morphogenesis defect in ADOA patients leads to moderate respiratory deficiency, to which retinal ganglion cells are known to be exquisitely sensitive. In cells from ADOA patients as well as in cultured wild type cells with OPA1 depleted by siRNA transfection, mitochondria are fragmented and sometimes clustered (Delettre et al., 2000; Griparic et al., 2004), consistent with observations in yeast. Overexpression of GTPase-defective OPA1 in mammalian cells causes similar effects (Misaka et al., 2002; Griparic et al.,

292

K.G. Hales / Mitochondrion 4 (2004) 285–308

2004). Curiously, overexpression of wild type OPA1 in human cells also leads to fragmented mitochondria (Misaka et al., 2002; Griparic et al., 2004), suggesting that perhaps the proper balance of cleavage products (see below) or of OPA1 with other fusion mediators is important for normal fusogenic activity. Griparic et al. (2004) examined mitochondrial morphology during intermediate stages of siRNAmediated OPA1 depletion in mammalian cells, observing swelling and stretching and then constriction of mitochondria before fragmentation. Transmission electron microscopy revealed changes in cristae morphology (such as reorientation and condensation) during mitochondrial fragmentation (Griparic et al., 2004); however, these changes may be a secondary effect from excessive mitochondrial division, since in mgm1 yeast strains (as noted above), abnormal cristae morphology is suppressed in the absence of Dnm1p-mediated division (Sesaki et al., 2003b). Mgm1p family members exists in two isoforms (90- and 100-kDa in budding yeast) which differ at the amino terminus (Herlan et al., 2003). The existence of a single functional start codon in MGM1 suggests that protease cleavage converts the large form to the small form (Shepard and Yaffe, 1999). The larger isoform contains two amino terminal hydrophobic regions, cleavage within the second of which leads to formation of the smaller isoform (Herlan et al., 2003). Originally, the subcellular localization of Mgm1p family members was reported, variously, as the mitochondrial outer membrane (Shepard and Yaffe, 1999), the matrix side of the mitochondrial inner membrane (Pelloquin et al., 1999), and the intermembrane space peripherally associated with the inner membrane (Wong et al., 2000). The current emerging consensus places both Mgm1p isoforms in the intermembrane space, as shown by protease protection assays (Olichon et al., 2002; Herlan et al., 2003; Sesaki et al., 2003b; Griparic et al., 2004) and measurement of susceptibility to differently targeted versions of a tobacco etch virus protease (Wong et al., 2003). Mgm1p remains insoluble during fractionation, indicating membrane association (Herlan et al., 2003). Regarding the nature and location of Mgm1p membrane association, immunoelectron microscopy indicates an association with the mitochondrial inner membrane (Wong et al., 2000; Olichon et al., 2002).

Treatment of mitochondrial fractions with high salt or carbonate solubilizes the short Mgm1p isoform (Herlan et al., 2003), indicating a peripheral membrane association. The large isoform stays membraneassociated upon salt treatment and only in some cases is released by carbonate treatment, suggesting an unusually tight peripheral membrane association (Shepard and Yaffe, 1999; Olichon et al., 2002; Herlan et al., 2003; Griparic et al., 2004). Cleavage of the large Mgm1p isoform by the Rbd1p/Pcp1p/Ugo2p rhomboid protease is necessary for normal mitochondrial morphology and retention of mtDNA (Herlan et al., 2003; McQuibban et al., 2003; Sesaki et al., 2003a). Rbd1p/Pcp1p/Ugo2p localizes to the mitochondrial inner membrane; cells lacking this protease contain only the large Mgm1p isoform and show fragmented mitochondria and mtDNA loss identical to mgm1 cells. The short Mgm1p isoform thus plays an important role in mitochondrial morphology and fusion. However, mitochondria lacking Rbd1p/Pcp1p/Ugo2p and thus containing only the large Mgm1p isoform retain at least some residual fusion activity in mating assays (Sesaki et al., 2003a), unlike mgm1 mutants which lack all mitochondrial fusion capability. The activity of both Mgm1p isoforms seems to be important for wild type function, since a gene encoding just the short isoform cannot rescue an mgm1 deletion mutant (which lacks both isoforms) but does partially rescue a rbd1/pcp1/ugo2 mutant, which lacks just the short isoform (Herlan et al., 2003). It has been postulated that a heterooligomeric complex containing the uncleaved and cleaved Mgm1p isoforms may mediate mitochondrial fusion under wild type conditions (Herlan et al., 2003), with the balance of short and long isoforms playing a crucial role. McQuibban et al. (2003) report that the varying extent of S. cerevisiae mitochondrial fusion under different growth conditions might be controlled in part by Rbd1p/Pcp1p levels, which are higher during logarithmic growth than during stationary phase. In contrast, a recent report implicates respiratory capacity as the primary determinant of alternative production of the two Mgm1p isoforms. Herlan et al. (2004) showed that the ability of the Mgm1p hydrophobic regions to insert in the mitochondrial inner membrane directly influences the proportion of large and small isoforms generated. In the model

K.G. Hales / Mitochondrion 4 (2004) 285–308

postulated, Mgm1p is targeted across the inner membrane, with the first hydrophobic region acting as a temporary stop-transfer sequence. If conditions are right (e.g. sufficient ATP for the import machinery to function efficiently), the protein is imported further, so that the second hydrophobic region reaches the inner membrane and is cleaved by Rbd1p/Pcp1p/ Ugo2p to generate the small Mgm1p isoform. Otherwise, Mgm1p stays as the long isoform. Indeed, reduced ATP synthesis shifted the balance of Mgm1p isoforms such that more of the large form was generated (Herlan et al., 2004). A similar effect was observed upon genetic downregulation of the inner membrane import machinery or upon increasing the hydrophobicity of the stop-transfer sequence (Herlan et al., 2004). Perhaps this level of Mgm1p regulation provides an adaptive advantage to the organism by causing respiratory deficient mitochondria to produce an imbalance of Mgm1p isoforms, leading to fusion incompetence and subsequent separation of these mitochondria from the healthy mitochondrial population (Herlan et al., 2004).

293

mitochondrial fusion mechanisms are among the next issues for the community to address.

3. Machinery of mitochondrial division As with mitochondrial fusion, several members of the mitochondrial division machinery have been identified. Mitochondrial outer membrane-associated proteins that promote fission include (in yeast) Dnm1p, Fis1p, and Mdv1p, the first two of which are conserved in many organisms. The Rab32 small GTPase may also play a role. Inner membrane proteins that may act in mitochondrial fission have been identified in two subsets of single-celled eukaryotes: Mdm33p in yeasts and FtsZ proteins in primitive algae. However, these proteins are not conserved outside their respective groups of species. Fission mediators associated with the mitochondrial inner membrane are yet to be identified in higher eukaryotes.

2.4. Possible mechanisms of mitochondrial fusion

3.1. The Dnm1p/Drp1 family and its role in outer mitochondrial membrane fission

The precise molecular choreography by which mitochondrial fusion occurs is unknown. Fzo1p family members pass through the outer membrane twice, and the loop in the intermembrane space appears to interact with Mgm1p family members, which are peripherally associated with the inner membrane. In yeast, Ugo1p binds to Fzo1p and Mgm1p separately, perhaps helping to form the complex. Could the portion of Fzo1p facing the cytosol (along with Ugo1p in yeast) mediate membrane docking (Sesaki et al., 2003b)? It remains to be determined whether the Fzo1p GTPase has a mechanochemical function or regulates assembly of other proteins involved in membrane fusion. Similarly, it is unclear whether the Mgm1p GTPase has a mechanical fusogenic function and/or coordinates assembly of the other members of the fusion complex (Wong et al., 2003). Is Mgm1p GTPase activity modulated by the ratio of small and large isoforms? Do Fzo1p and Mgm1p work together to coordinate fusion of the outer and inner membranes? What plays the connecting role of Ugo1p in metazoans? These and other questions concerning

The yeast dynamin-related protein Dnm1p and its homologs in worms, mammals, and plants play a central role in mitochondrial division. Like other dynamin family members, Dnm1p is a large protein with several domains, including a GTPase domain near the amino terminus and a carboxy-terminal GTPase effector domain (GED) (van der Bliek, 1999). Dnm1p binds and hydrolyzes GTP (Fukushima et al., 2001). S. cerevisiae mutants that lack Dnm1p or that have mutations in the GTPase domain show an aggregated and collapsed mitochondrial network (Otsuga et al., 1998), similar to that seen in cultured human cells transfected with a dominant negative version of the mammalian homolog Drp1 (Smirnova et al., 1998). These collapsed networks consist of abnormally interconnected net-like mitochondrial structures, which can be visualized upon genetic or chemical cytoskeletal perturbation (Bleazard et al., 1999; Smirnova et al., 2001). Functional Dnm1p is required for the mitochondrial fragmentation normally seen in S. cerevisiae fzo1 fusion mutants, confirming the role of Dnm1p as a division mediator and suggesting that division and fusion are usually in

294

K.G. Hales / Mitochondrion 4 (2004) 285–308

dynamic balance (Bleazard et al., 1999; Sesaki and Jensen, 1999). Studies of C. elegans DRP-1 suggest a role for this Dnm1p homolog in mitochondrial outer membrane division only; overexpressed wild type C. elegans DRP-1 causes excess mitochondrial fragmentation, while anti-sense constructs or dominant negative versions with mutations in the GTPase domain inhibit outer but not inner membrane division (Labrousse et al., 1999). Dynamin-related proteins also seem to act in mitochondrial division in plants. In Arabidopsis, disruption of the ADL1C and ADL1E dynamins (or expression of dominant negative versions) leads to unusually elongated and interconnected mitochondria (Jin et al., 2003). Dominant negative versions of the Arabidopsis homolog ADL2b, when introduced into cultured tobacco cells, are associated with increased size and decreased number of mitochondria, consistent with a fission role (Arimura and Tsutsumi, 2002). Subcellular localization experiments indicate that Dnm1p and homologs (while partially cytosolic) are also in the right place at the right time to mediate mitochondrial division. In budding yeast, Dnm1p was shown by immunolocalization to exist in punctate structures on the cytoplasmic side of the mitochondrial outer membrane, often at constriction sites or at the end of a mitochondrial tubule which has presumably recently divided (Bleazard et al., 1999). Tagged versions of Arabidopsis ADL2b, ADL1C, and ADL1E are similarly detectable on mitochondrial tips and at constriction sites (Arimura and Tsutsumi, 2002; Jin et al., 2003). Time lapse experiments in C. elegans and cultured mammalian cells confirm that puncta containing GFP-tagged DRP-1 or Drp1 indeed correspond to sites of mitochondrial division (Labrousse et al., 1999; Smirnova et al., 2001). Various lines of evidence suggest that Dnm1p and homologs, like their more distant dynamin relatives, homo-oligomerize during membrane remodeling. Coimmunoprecipitation and yeast two-hybrid experiments indicate that Dnm1p physically interacts with itself (Shin et al., 1999; Fukushima et al., 2001). Tagged and purified Drp1 can assemble in vitro into ring- or spiral-shaped structures (Smirnova et al., 2001). A version of Drp1 unable to hydrolyze GTP causes extensive tubulation of mitochondria, with Drp1-associated striations surrounding the organelle (Yoon et al., 2001). Does GTP hydrolysis by

Dnm1p/Drp1 family members power the constriction of these rings to cause mitochondrial membrane scission, or does GTP-bound Dnm1p/Drp1 recruit other factors that have a mechanochemical function? That question has not yet been resolved, though the latter model is supported by studies with a version of Dnm1p mutated in the GED. Expression in yeast of this Dnm1p variant, which presumably is impaired in GTP hydrolysis and thus stabilized as Dnm1p-GTP, results in an increased mitochondrial fission rate (Fukushima et al., 2001). Therefore, Dnm1p binding to (but not hydrolysis of) GTP seems to regulate a rate-limiting step in mitochondrial fission, consistent with a possible role in recruiting other factors with mechanochemical membrane severing activity. A recent study implicates the Sumo1 conjugation system as a regulator of DRP1 activity in mammalian cells (Harder et al., 2004). Sumo1, a ubiquitin-related protein, is conjugated to many subcellular substrates to induce not proteolysis but instead modulation of stability or activity (reviewed in Seeler and Dejean (2003)). Harder et al. (2004) used a combination of yeast two hybrid analysis, GST pull-down assays, and Western blotting with inhibition of Sumo1 removal to show that DRP1 is a substrate for Sumo1 conjugation. Sumo1-associated puncta were visualized at sites of mitochondrial fission but also at other non-DRP1associated sites on mitochondria and elsewhere in the cell. Overexpression of wild type (but not nonconjugatable) Sumo1 led to significant stabilization of DRP1 and increased mitochondrial fragmentation, suggesting that this post-translational ‘sumoylation’ of DRP1 may be central to regulation of mitochondrial fission (Harder et al., 2004). The interplay between Dnm1p/DRP1-binding partners (described in Section 3.2) and the Sumo1 conjugation system is yet to be explored. 3.2. Two Dnm1p-binding partners/mitochondrial fission participants: Mdv1p and Fis1p Two gene products have been found to interact with Dnm1p and help mediate mitochondrial fission in yeast; at least one of the two is conserved in mammals. The MDV1 gene (alternately known as FIS2, GAG3, and NET2) was found through a genetic screen for fzo1 suppressors (Mozdy et al., 2000; Tieu and Nunnari, 2000), as well as a genetic screen for

K.G. Hales / Mitochondrion 4 (2004) 285–308

mgm1 suppressors (Fekkes et al., 2000) and a yeast two-hybrid screen for Dnm1p-interactors (Cerveny et al., 2001). A second gene, FIS1 (also known as MDV2), was found simultaneously in the same fzo1 suppressor screens (Mozdy et al., 2000; Tieu and Nunnari, 2000). Mitochondria in single mdv1 and fis1 mutants are abnormally clustered and net-like, similar to those in dnm1 mutants (Fekkes et al., 2000; Mozdy et al., 2000; Tieu and Nunnari, 2000; Cerveny et al., 2001). Mdv1p contains WD40 repeats and predicted coiled coil regions (Fekkes et al., 2000; Tieu and Nunnari, 2000; Cerveny et al., 2001) and is a peripheral mitochondrial outer membrane protein that faces the cytoplasm, as shown by fractionation and protease protection experiments (Fekkes et al., 2000; Cerveny et al., 2001). Fis1p is a relatively small transmembrane protein that spans the outer mitochondrial membrane with the bulk of the protein facing the cytoplasm (Mozdy et al., 2000). The localization and interactions between Dnm1p, Mdv1p, and Fis1p have been examined. Normally, Mdv1p localizes in a Dnm1p-dependent manner to puncta on the mitochondrial surface (Tieu and Nunnari, 2000; Cerveny et al., 2001), while Fis1p is localized diffusely across the mitochondria (Mozdy et al., 2000). If Dnm1p is absent, Mdv1p localizes diffusely to mitochondria in a manner dependent on Fis1p (Mozdy et al., 2000; Tieu and Nunnari, 2000). The presence of Fis1p is required for mitochondrial localization of both Dnm1p and Mdv1p (Fekkes et al., 2000; Mozdy et al., 2000; Tieu and Nunnari, 2000; Cerveny et al., 2001). Yeast two hybrid experiments confirm that Mdv1p physically interacts (via different domains) with both Dnm1p and Fis1p (Tieu et al., 2002; Cerveny and Jensen, 2003). According to a recently proposed model for mitochondrial fission (Shaw and Nunnari, 2002), Dnm1p puncta first form on the mitochondrial surface via interaction with Fis1p, and Mdv1p assembles onto Dnm1p. Mdv1p then physically interacts directly with Fis1p, triggering the fission event. This latter interaction is inferred from studies of a mutant Fis1p that can still recruit Dnm1p (and therefore Mdv1p) puncta but cannot bind directly to Mdv1p; in this fis1 strain, mitochondrial fission is defective (Tieu et al., 2002). Fis1p, like Dnm1p, has homologs in many organisms (James et al., 2003; Yoon et al., 2003; Stojanovski et al., 2004). In contrast, database

295

searches for Mdv1p homologs indicate close relatives only in other single celled eukaryotes, with resemblance of yet uncertain significance to various WD40 proteins in higher organisms. A human homolog of Fis1p shows subcellular localization to mitochondria (dependent on a targeting signal in the hFis1 carboxy terminus) similar to that in yeast (James et al., 2003; Yoon et al., 2003; Stojanovski et al., 2004). Overexpression of hFis1 enhances mitochondrial fragmentation (James et al., 2003; Yoon et al., 2003), while hFis1 depletion by RNAi or antibody injection inhibits fragmentation (Yoon et al., 2003; Stojanovski et al., 2004), consistent with a conserved role for Fis1p family members in mitochondrial fission, and suggesting that in mammalian cells, hFis1 is a ratelimiting factor, unlike in yeast. Mitochondrial fragmentation induced by excess hFis1 depends on Drp1, since dominant negative Drp1 minimizes the mitochondrial fragmentation caused by overexpression of hFis1 (James et al., 2003; Stojanovski et al., 2004). Like in yeast, Drp1 and hFis1 interact physically, as demonstrated by coimmunoprecipitation and other techniques (Yoon et al., 2003). 3.3. Regulation of the mitochondrial outer membrane fission machinery and a possible role for a Rab family protein It remains to be determined how the assembly of the Dnm1p/Mdv1p/Fis1p complex (and the subsequent triggering of mitochondrial fission) is regulated. The enhancement of fission by stabilized Dnm1p-GTP (Fukushima et al., 2001) is consistent with a possible regulatory role of the Dnm1p GTPase cycle in assembling the fission machinery. Recent results hint at the possibility that another type of GTPase may be involved, at least in mammalian cells. The Rab32 small GTPase, when GTP-bound, is associated with mitochondria, and a mutant version unable to hydrolyze GTP causes mitochondrial morphology suggestive of fission defects (Alto et al., 2002). This effect is seen only when the carboxyterminal cysteines enabling mitochondrial association are retained in Rab32 (Alto et al., 2002). It is interesting to speculate that the Rab32 GTPase might help regulate assembly of the Drp1 complex. Rab32 is also known to anchor protein kinase

296

K.G. Hales / Mitochondrion 4 (2004) 285–308

A (PKA) to mitochondria, and it is unclear whether PKA has a role in mitochondrial dynamics. 3.4. Independence of outer and inner membrane fission events; the Mdm33p and FtsZ families and their possible roles in mitochondrial inner membrane fission A recent study in yeast suggests that the Dnm1pcontaining complexes mediate only one of two distinct processes that govern mitochondrial fission. Time lapse fluorescence imaging and three-dimensional reconstruction of mitochondrial shape indicated that the narrowing of mitochondrial tubules occurs independently of the formation of Dnm1p rings (Legesse-Miller et al., 2003). Furthermore, Dnm1p puncta form and sometimes disappear without concurrent mitochondrial fission. Legesse-Miller et al. (2003) speculate that mitochondrial fission occurs only when Dnm1p and Mdv1p happen to assemble at a location where the organelle has already narrowed enough to be fully encircled. At wider points of the organelle, Dnm1p complexes appear only on one side, not completely encircling the mitochondrion (Legesse-Miller et al., 2003). The independence of mitochondrial narrowing and Dnm1p complex formation raises the question of the physical basis of mitochondrial constriction. Do additional components in the mitochondrial inner membrane mediate constriction and perhaps also inner membrane fission, independent of outer membrane events? Indeed, inner membrane fission proceeds in the absence of outer membrane fission when C. elegans DRP-1 is inhibited (Labrousse et al., 1999) and when yeast Fis1p is mutated (Jakobs et al., 2003). In yeast, Mdm33p may be a component of the mitochondrial inner membrane fission machinery. Mdm33p spans the mitochondrial inner membrane and, when overexpressed, triggers excessive septation of this compartment (Messerschmitt et al., 2003). Cells lacking Mdm33p contain aberrantly extended mitochondria, often in ring-shaped or spherical conformations; cells lacking both Mdm33p and Fis1p are similar, suggesting that the net-like conformation seen in fis1 mutants depends on functional Mdm33p (Messerschmitt et al., 2003). It remains to be seen whether Mdm33p interacts with other components and whether its activity is somehow

coordinated with the activity of the mitochondrial outer membrane fission machinery. Mdm33p has homologs only in other yeast species (Messerschmitt et al., 2003). Primitive eukaryotes may depend on FtsZ family members for mitochondrial inner membrane fission while still utilizing Dnm1p family proteins for outer membrane division. FtsZ proteins are widely known as mediators of bacterial cytokinesis, forming a ring on the cytoplasmic side of the membrane at the middle of the dividing cell (reviewed in Margolin (2001)). The discovery of a mitochondrial FtsZ protein in the chromophyte alga Mallomonas splendens (Beech et al., 2000) led to the initial supposition that the FtsZ system of mitochondrial division, a relic of the bacterial origin of mitochondria, was simply replaced by the Dnm1p/Drp1 system during evolution of higher eukaryotes. However, the alga Cyanidioschyzon merolae appears to utilize both an FtsZ homolog and a Dnm1p homolog (called CmDnm1) for mitochondrial division, with the former found on the inner membrane and the latter on the outer membrane (Nishida et al., 2003). The timing of the localization of these two proteins is consistent with a model in which FtsZ participates in the initial constriction of mitochondria while CmDnm1 acts at a later step to effect final membrane severing (Nishida et al., 2003). With FtsZ family members perhaps mediating mitochondrial inner membrane fission in primitive eukaryotes, and with Mdm33p family members perhaps doing the same in yeasts, it remains to be seen what gene products, if any, are fulfilling an analogous role in other eukaryotes. Mechanisms for coordinating the inner and outer membrane fission machinery also are yet to be determined.

4. Apoptosis: possible roles of the mitochondrial fission and fusion machinery Changes in mitochondrial morphology and the associated loss of membrane potential and release of cytochrome c play a central role in apoptosis (Desagher and Martinou, 2000). The appearance of numerous small mitochondria during apoptosis is consistent with possible roles for enhanced mitochondrial fission and/or inhibited mitochondrial fusion during this process. To explore whether the machinery

K.G. Hales / Mitochondrion 4 (2004) 285–308

of normal mitochondrial division is used during apoptosis, Frank et al. (2001) traced the localization of Drp1 in apoptotic cultured mammalian cells. They confirmed that mitochondria do increase in number after apoptotic stimulation, and they noted that Drp1 (normally partially cytosolic with some punctate distribution on mitochondria) redistributed during apoptosis so that a much larger fraction of the protein was detectable in mitochondrial patches (Frank et al., 2001). Furthermore, Drp1-mediated mitochondrial division was shown to have a central role during the progression of apoptotic events. Cells treated with staurosporine normally are triggered to undergo apoptosis, with all the hallmark morphological changes. A dominant negative version of Drp1 in cells treated with staurosporine inhibited not only mitochondrial fragmentation but also mitochondrial membrane depolarization, cytochrome c release, and apoptotic DNA cleavage as detected by a TUNEL assay (Frank et al., 2001). This Drp1-mediated inhibition of apoptosis occurred even when Bax, a pro-apoptotic Bcl-2 family member, was overexpressed. These results suggest that Drp-1-mediated mitochondrial division is a crucial early step of apoptosis. How is mitochondrial division triggered during apoptosis? A study using cultured mammalian cells hints at an intimate physical connection between Bax and two mediators of mitochondrial morphogenesis. Drp1 and Mfn2. Upon staurosporine treatment, Bax migrated to clusters on the mitochondrial surface, colocalizing with the fission mediator Drp1 (Karbowski et al., 2002), and perhaps helping to recruit a larger pool of Drp1 to the mitochondrial surface to enhance fission activity. Bax localized to Drp1 puncta even when dominant negative Drp1 inhibited apoptotic events, suggesting that Drp1 control of apoptosis (Frank et al., 2001) is downstream of Bax redistribution. Another study indicated that Drp1 redistribution to mitochondria occurred after caspase-induced cleavage of the endoplasmic reticulum protein BAP31 and subsequent Ca2C release from the ER and uptake by mitochondria (Breckenridge et al., 2003). The second component of the mitochondrial morphogenesis machinery with which Bax associates is the fusion mediator Mfn2 (Karbowski et al., 2002). Recent results indeed suggest that inhibition of mitochondrial fusion contributes to mitochondrial

297

fragmentation during apoptosis. Karbowski et al. (2004) developed a new system to assay mitochondrial fusion in mammalian cells by transfecting with and then tracking the dilution of a photoactivatable GFP fusion protein targeted to mitochondria. In various mammalian cell types, mitochondrial fusion was inhibited upon chemical induction of apoptosis; this fusion inhibition was accelerated if Bax was overexpressed, and the timing of inhibition coincided with Bax clustering on the mitochondrial membrane (Karbowski et al., 2004). The exact choreography by which Bax association with Mfn2 and Drp1 inhibits mitochondrial fusion and enhances mitochondrial division is unknown. Experiments with two other components of the mitochondrial morphogenesis machinery indicate possible links to apoptosis. In HeLa cells, depletion by RNAi of OPA1, the mammalian homolog of mitochondrial fusion mediator Mgm1p, leads to aberrant inner membrane structure and mitochondrial fragmentation, as well as to events of apoptosis: mitochondrial membrane potential reduction, cytochrome c release, and an apoptotic nuclear morphology (Olichon et al., 2003). Overexpression of the fission mediator hFis1 also causes mitochondrial fragmentation, cytochrome c release, and subsequent events of apoptosis (James et al., 2003). In both the OPA1 depletion and hFis1 overexpression experiments, excess levels of anti-apoptotic Bcl-2 family members inhibited cell death but, curiously, did not inhibit mitochondrial fragmentation (James et al., 2003; Olichon et al., 2003). These results suggest that the level of mitochondrial fragmentation per se does not appear to be a determinant of apoptosis. Rather, apoptosis may be triggered in response to particular levels of protein complexes containing Bax and various mitochondrial fission and fusion mediators. Many experiments lie ahead to dissect the precise roles of fission and fusion mediators in apoptosis.

5. Mechanisms of mitochondrial distribution In a few single-celled eukaryotes, actin filaments are the primary mediators of mitochondrial movement; recent evidence in budding yeast points to the involvement of two primary protein complexes, one to nucleate actin polymerization and propel

298

K.G. Hales / Mitochondrion 4 (2004) 285–308

mitochondrial movement, and one to connect actin cables to mitochondrial membranes and the mtDNA within. In other single-celled eukaryotes as well as in higher eukaryotes, mitochondrial distribution is mediated mainly by microtubules, though in neurons there is evidence for actin-based motility as well. Genetic and biochemical analysis has partially elucidated the basis of microtubule-based mitochondrial motility. Various lines of evidence in different contexts also suggest the involvement of intermediate filaments, fatty acid metabolism, and the ubiquitin pathway in mitochondrial distribution. 5.1. Molecular mediators of actin-based mitochondrial distribution in some fungi Mitochondria travel along actin cables in fungi such as S. cerevisiae and Aspergillus nidulans. Disruption of microtubules has no direct effect on mitochondrial organization in those organisms (Huffaker et al., 1988; Smith et al., 1995; Suelmann and Fischer, 2000). During cell division in S. cerevisiae, a subset of mitochondria move along polarized actin cables into the bud, while other mitochondria colocalize with actin in a ‘retention zone’ in the mother cell (Simon et al., 1995; Yang et al., 1999), ensuring that each cell contains a mitochondrial complement following mitosis. Meiotic S. cerevisiae cells also display rapid actin-associated mitochondrial movement (Smith et al., 1995). Yeast strains with particular mutations in the actin (ACT1) gene show aberrant mitochondrial motility and morphology, even in some cases when actin cables remain visibly present; presumably in the latter, the moiety of the actin molecule to which mitochondria connect is altered (Drubin et al., 1993; Lazzarino et al., 1994; Smith et al., 1995). Other mutations in yeast that affect actin cytoskeleton stability, such as in the MDM20-encoded tropomyosin acetylator, cause mitochondrial inheritance defects (Hermann et al., 1997; Singer and Shaw, 2003). Chemical disruption of actin cables in S. cerevisiae (Boldogh et al., 1998) or A. nidulans (Suelmann and Fischer, 2000) also inhibits mitochondrial movement. Mitochondrial binding to and movement along actin cables in vitro requires low ATP levels; at high ATP levels the interaction is disrupted (Lazzarino et al., 1994; Simon et al., 1995). Myosin seems not to

be involved in mitochondrial movement, as mutations in various myosin genes do not affect mitochondrial distribution both in S. cerevisiae (Simon et al., 1995) and A. nidulans (Suelmann and Fischer, 2000). Instead, mitochondrial motility along actin cables in S. cerevisiae involves the Arp2/3 complex (Boldogh et al., 2001a), which is comprised of two actin-related proteins and five other subunits, and which has previously been shown to nucleate actin polymerization for propulsion of certain pathogenic bacteria within cells (reviewed in Fehrenbacher et al. (2003)). Boldogh et al. (2001a) copurified two members of the Arp2/3 complex with mitochondria and showed that the complex localizes to S. cerevisiae mitochondria in vivo. They showed that a temperature sensitive strain mutated in an Arp2/3 complex subunit, when shifted to the restrictive temperature, was defective in mitochondrial movement though mitochondria still were localized to actin cables and were shaped normally. Also, wild type mitochondria showed actin nucleating activity; chemically limiting mitochondrial polymerization/depolymerization (while maintaining the overall levels of cables and patches) also decreased mitochondrial mobility. All of these observations are consistent with a role for Arp2/3mediated mitochondrial dynamics in mitochondrial distribution in S. cerevisiae (Boldogh et al., 2001a). Curiously, Arp2/3-mediated mitochondrial movement occurs along existing polarized actin cables, unlike the movement of pathogenic bacteria via Arp2/3generated actin comets. Since mitochondria still colocalize with actin cables even when Arp2/3 complex function is disrupted, other proteins must provide a physical connection between the mitochondrial outer membrane and actin filaments. Three S. cerevisiae gene products implicated in this capacity are Mmm1p, Mdm10p, and Mdm12p, each of which is required for mitochondrial movement during budding (Burgess et al., 1994; Sogo and Yaffe, 1994; Berger et al., 1997; Boldogh et al., 1998, 2003). Mmm1p and Mdm10p have been shown to mediate in vivo and in vitro mitochondrial association with actin cables, and reassociation to mitochondria of previously washed-away peripheral membrane protein(s) needed for ATP-sensitive actin association (Boldogh et al., 1998). Mmm1p, Mdm10p, and Mdm12p appear to form a complex, as suggested by

K.G. Hales / Mitochondrion 4 (2004) 285–308

coimmunoprecipitation as well as experiments to test mutual localization dependence (Boldogh et al., 2003). All three proteins were initially thought to be mitochondrial outer membrane proteins (Burgess et al., 1994; Sogo and Yaffe, 1994; Berger et al., 1997), though the recent finding via protease protection assays and other methods that Mmm1p spans both the mitochondrial outer and inner membranes with the small amino terminus in the matrix (Kondo-Okamoto et al., 2003), suggests that the complex may actually reside at contact sites between the two mitochondrial membranes. Indeed, all three proteins colocalize in an actin-independent, punctate manner with each other and with a subset of mtDNA nucleoids; inactivation of any of the three proteins causes (initially) aberrant mitochondrial nucleoid structure and (ultimately) mtDNA to be lost (Aiken Hobbs et al., 2001; Boldogh et al., 2003). Furthermore, mitochondrial inner membrane structure becomes dramatically altered upon Mmm1p inactivation (Aiken Hobbs et al., 2001). The complex containing Mmm1p, Mdm10p, and Mdm12p has therefore been deemed a ‘mitochore’, a structure connecting mtDNA with the actin cytoskeleton for faithful segregation of mitochondrial genetic material during cell division (Aiken Hobbs et al., 2001; Boldogh et al., 2003). A hypomorphic allele mmm1-6 was independently identified in a screen for strains in which mtDNA escapes to the nucleus at a higher than normal rate (Thorsness and Fox, 1993; Hanekamp et al., 2002). Mitochondrial morphology defects and mtDNA loss are observed in this strain only when grown on non-fermentable carbon sources, as opposed to the inevitable mtDNA loss under any condition in mmm1 deletion strains (Hanekamp et al., 2002). Suppressors of mmm1- and mdm10-associated mitochondrial morphology defects have been identified, and cloning of the associated genes should shed more light on the mechanism by which this complex controls mitochondrial movement and mtDNA distribution (Hanekamp et al., 2002). The mitochore, also called the ‘two membrane spanning structure’ (TMS), includes the DNA-binding protein Mgm101, which is required for mtDNA maintenance (Meeusen et al., 1999; Meeusen and Nunnari, 2003). Curiously, in wild type strains chemically triggered to lose mtDNA, Mgm101p and Mmm1p still associate with mitochondria in

299

self-replicating TMS/mitochore foci (Meeusen and Nunnari, 2003), and such mitochondria exhibit faster than normal motility on actin filaments (Boldogh et al., 2003), suggesting a regulatory role (but not an absolute requirement) for mtDNA on mitochore function. BrdU labeling experiments indicated that the TMS/mitochore is associated only with replicating nucleoids (Meeusen and Nunnari, 2003). It is not clear whether Mmm1p (despite spanning both mitochondrial membranes) actually contacts mtDNA, as alteration of the matrix-localized Mmm1p amino terminus does not affect mtDNA maintenance (Kondo-Okamoto et al., 2003). The specific interactions among members of the mitochore complex remain to be determined, as does the exact nature of the contact between the complex and mtDNA and actin. The fact that mitochondrial movement in N. crassa is microtubule-based (see below) but also requires an Mmm1p homolog suggests that additional adaptors may provide cytoskeletal specificity in each species. An Mdm10p homolog in A. nidulans, MdmB, seems to function in mitochondrial morphology, though its even distribution across mitochondria (in contrast to punctate localization of Mdm10p in S. cerevisiae) raises some doubt regarding functional equivalence (Koch et al., 2003). In strains deficient for MdmB, some but not all mitochondria are abnormally collapsed and spherical, and only at low temperatures. It is not yet known whether MdmB has a role in mtDNA nucleoid integrity or acts in a complex. Evidence is emerging for a second complex that governs nucleoid integrity and mitochondrial connections to the cytoskeleton in S. cerevisiae. MMM2 (identified as a genetic interactor with MMM1) encodes a novel mitochondrial outer membrane protein that also colocalizes with a subset of mtDNA nucleoids, though not always with Mmm1passociated nucleoids (Youngman et al., 2004). Mmm2p fractionates as part of a large complex apparently distinct from that containing Mmm1p, Mdm10p, and Mdm12p (Youngman et al., 2004). Cells with MMM2 disrupted show misshapen and collapsed mitochondrial networks, though more varied in shape than in cells disrupted for MMM1. The functional differences between Mmm2p-containing complexes and those containing Mmm1p, Mdm10p, and Mdm12p remain to be determined.

300

K.G. Hales / Mitochondrion 4 (2004) 285–308

5.2. Molecular mediators of microtubule-based mitochondrial distribution in some fungi In contrast to actin-based mitochondrial motility in S. cerevisiae and A. nidulans, mitochondria seem to move primarily associated with microtubules in other single-celled eukaryotes such as Neurospora crassa (Steinberg and Schliwa, 1993) and S. pombe (Yaffe et al., 1996). Chemical disruption of the actin cytoskeleton in N. crassa does not affect mitochondrial distribution (Prokisch et al., 2000). Mitochondria from N. crassa bind to microtubules in vitro in a manner dependent on unknown salt-extractable peripheral proteins of the mitochondrial outer membrane (Fuchs et al., 2002). An MMM1 homolog in N. crassa, when mutated, is associated with abnormally collapsed spherical mitochondria, analogous to the budding yeast mutant phenotype (Prokisch et al., 2000). Despite initial speculation that Mmm1p family members might have evolved to connect mitochondria to different cytoskeletal elements in different organisms, recent results show that N. crassa mitochondria from mmm1 mutant lines can still bind microtubules, indicating that MMM1 is not a crucial connector between mitochondria and microtubules (Fuchs et al., 2002). Instead, perhaps the mmm1 phenotype results from altered regulation of motor proteins already attached to mitochondria. It is not known whether N. crassa MMM1 affects mtDNA nucleoid stability as in S. cerevisiae. In S. pombe, aberrant mitochondrial distribution during mitosis occurs as a result of mutations in genes encoding either alpha or beta tubulin subunits (Yaffe et al., 1996). Observation of wild type S. pombe mitosis has hinted that mitochondrial association with spindle poles may govern equal segregation of mitochondrial material (Yaffe et al., 2003). Time lapse observation of mitochondria and microtubules in wild type S. pombe cells suggests that mitochondria may not move along existing microtubules but rather may change shape in concert with microtubule polymerization and depolymerization (Yaffe et al., 2003). The S. pombe genome includes a homolog of S. cerevisiae MDM10 (Berger et al., 1997), but it is not known whether this homolog functions analogously to connect mitochondria to the cytoskeleton. Recent experiments have identified S. pombe Mmd1p as a novel cytosolic protein required for proper

mitochondrial distribution (Weir and Yaffe, 2004). In mmd1 mutant cells, mitochondria appear to associate with microtubules at the cellular poles but do not form the lateral contacts needed for extension along these cytoskeletal elements (Weir and Yaffe, 2004). Mmd1p interactors and mechanisms of mitochondrial movement in fission yeast remain to be determined. 5.3. Molecular mediators of mitochondrial distribution in higher eukaryotes: primarily microtubule-based but some actin-based mitochondrial mobility For decades, it has been the common view that mitochondria travel on microtubule tracks in many higher eukaryotic cell types, an idea supported by numerous observations of mitochondria/microtubule relative positioning and the effects of microtubule inhibitors on mitochondrial motility (reviewed in Bereiter-Hahn and Voth (1994)). Functional analysis of microtubule motors expressed broadly in mice and Drosophila supports this view. Tanaka et al. (1998) generated mice lacking the kif5B kinesin heavy chain, a ubiquitous component of the kinesin plus enddirected microtubule motor. Cells cultured from the inviable homozygous knockout embryos showed abnormal clustering of mitochondria near the nucleus; the mitochondria in mutant cells were still colocalized with perinuclear microtubules but did not move near the cell periphery. Subsequent disruption of microtubules in the mutant cells resulted in random dispersion of mitochondria throughout the cytoplasm, suggesting that mitochondria in kif5B mutant cells are still bound to microtubules but perhaps (as predicted) do not move toward the plus ends. Fractionation of cells from wild type mice gave results consistent with mitochondrial association of KIF5B (Tanaka et al., 1998). A Drosophila kinesin family member also appears to mediate mitochondrial distribution in many cell types, particularly during mitosis. The KLP67A kinesin heavy chain-related gene is expressed throughout embryonic and larval development in a pattern reflecting mitotic domains (Pereira et al., 1997). KLP67A is detectable by immunolocalization on mitochondria on or near the spindle aster during embryonic mitosis, consistent with a possible role for

K.G. Hales / Mitochondrion 4 (2004) 285–308

KLP67A in mitochondrial positioning for accurate segregation during mitotic cell divisions. Gametogenesis in Drosophila provides additional contexts for studying mitochondrial distribution. During male meiosis in Drosophila, mitochondria are visibly associated with the spindle during meiotic cell divisions, also presumably to ensure accurate segregation (Fuller, 1993). Following meiosis, mitochondria aggregate beside each haploid spermatid nucleus in a manner consistent with minus-enddirected movement along microtubules toward the microtubule organizing center (Fuller, 1993). The molecular basis for this movement is not yet known, though analysis of recessive male sterile mutants with defects in mitochondrial aggregation may elucidate the molecular mechanisms. In Drosophila males homozygous for mutations in no mitochondrial derivative (nmd), mitochondria fail to aggregate in post-meiotic spermatids and instead remain dispersed in the cytoplasm; nmd encodes a predicted mitochondrial outer membrane ATPase (S. Baxley, S. Holmberg, S.T. Burke, A. Aldridge, N. Wolf, M.T. Fuller and K.G. Hales, unpublished data). In male flies homozygous for mutations in the mitoshell gene, mitochondria aggregate around the nucleus instead of beside it (T. Plowshay, M. Wilson, A. Aldridge, and K.G. Hales, unpublished data). Further analysis and identification of the mitoshell gene will elucidate whether the defect is in mitochondrial transport or farther upstream in microtubule organization or another process. Mitochondrial transport during Drosophila oogenesis also may be microtubule based. In the ovary are 16-cell cysts connected by cytoplasmic bridges, and within each cyst one of the cells is destined to become the oocyte. Mitochondria and other cytoplasmic components are transported from the 15 nurse cells into the future oocyte along a structure called a fusome, which is rich in microtubules whose minus ends are oriented near the oocyte (Grieder et al., 2000; Cox and Spradling, 2003). Midway through oogenesis, a subset of mitochondria move to the oocyte and contribute to the Balbiani body, a posterior aggregate of organelles that is postulated to contribute to the germ cells of the individual the oocyte will become (Cox and Spradling, 2003). The molecular mechanisms by which these mitochondria travel along

301

the fusome (and are distinguished from mitochondria left behind in the nurse cells) is unknown. In neurons, mitochondria must travel great distances along axons from the cell body to the synaptic terminal and back. This long-distance anterograde and retrograde mobility seems to occur primarily along microtubules, though some limited actin filamentassociated mobility is retained upon microtubule depolymerization in vertebrate neurons (Morris and Hollenbeck, 1995; Ligon and Steward, 2000). In a few contexts, actin filaments seem to the primary track for mitochondrial transport; for example, in photoreceptors of the locust Schistocerca gregaria, mitochondria move along actin filaments towards microvilli in response to light (Sturmer et al., 1995). The role of microtubules in axonal itochondrial transport is supported by genetic analysis in Drosophila. Mutations in the kinesin heavy chain (Hurd and Saxton, 1996) or light chain (Gindhart et al., 1998), which together make up the microtubule plus enddirected kinesin heterotetramer, or in dynactin complex members (associated with minus-end-directed motor activity) (Martin et al., 1999) lead to ‘organelle jams’ in axons and ultimately to paralysis and larval death. Mitochondria are among the organelles that are not transported properly along axons in the absence of these microtubule motor proteins. What are the adaptors that connect mitochondria to these multipurpose motor proteins? Recent work in Drosophila has identified the milton gene as required for proper transport of mitochondria to nerve terminals (Stowers et al., 2002). The milton mutant was identified in a mosaic screen to identify photoreceptor defects caused by homozygous mutagenized chromosomes in the eye in otherwise heterozygous backgrounds. The only defect in homozygous milton photoreceptors is anterograde mitochondrial transport; other organelle cargo moves properly in the axon. The Milton protein immunolocalizes to mitochondria and coimmunoprecipitates with kinesin heavy chain, though additional tests suggest the kinesin interaction is not direct (Stowers et al., 2002). Milton is homologous to mammalian HAP-1, which binds the huntingtin (Huntington disease) protein. Expressed in a variety of tissues, Milton may be an adaptor between mitochondria and kinesins in many contexts other than neurons. Indeed, preliminary data hint that Milton may mediate mitochondrial elongation along

302

K.G. Hales / Mitochondrion 4 (2004) 285–308

the growing microtubule-based flagellar axoneme in developing Drosophila spermatids (Siegenthaler and Hales, unpublished data). In mammals, Kif1B is a monomeric neuronal kinesin-related microtubule motor that localizes to mitochondria and can transport mitochondrial cargo in vitro (Nangaku et al., 1994). A splice variant of Kif1B lacks the mitochondrial cargo domain and instead transports synaptic vesicles along axons; defects in Kif1B are associated with Charcot-MarieTooth disease type 2A (Zhao et al., 2001), though it is thought that lack of the vesicle carrying variant causes the major symptoms. Thus, little is known about the adaptors that connect mitochondria to microtubule based motors. Even less is known about the regulation of the direction of mitochondrial movement on microtubules (plus end- versus minus end-directed), though two recent hints concern phosphatidylinositol lipids, and a third hint concerns a possible correlation between mitochondrial membrane potential and the directionality of transport. In cultured neurons, transfection of a pleckstrin homology domain that specifically binds phosphatidylinositol (4,5) bisphosphate (PIP2) shifts the balance toward plus-end-directed mitochondrial movement without altering speed or overall level of transport (De Vos et al., 2003). In developing axons responding to nerve growth factor (NGF), mitochondria undergo preferential anterograde (plus enddirected) movement in a manner dependent on NGF-triggered phosphoinositide 3-kinase signaling (Chada and Hollenbeck, 2003). Perhaps different phosphoinositides on the outer mitochondrial membrane regulate activity of motor proteins. A recent study of chicken neurons demonstrated that most of the mitochondria moving in an anterograde direction have high membrane potential, whereas retrogrademoving mitochondria tend to have lower potential (Miller and Sheetz, 2004). This observation is consistent with speculation that the mitochondrial subpopulation traveling along the fusome to the Drosophila oocyte consists of the most highly respiring mitochondria (Cox and Spradling, 2003). In both the neuron and the developing oocyte, selective mitochondrial transport based on membrane potential would provide an adaptive advantage, allowing the healthiest mitochondria to fuel ion transport at the synapse or to populate the germ

cells for the benefit of the next generation. The molecular mechanism by which membrane potential influences directionality of mitochondrial transport is unknown, though it is tempting to speculate that proteins connecting and/or transporting mitochondria along cytoskeletal elements take on potential-dependent conformations (Miller and Sheetz, 2004). 5.4. Intermediate filaments and other factors influencing mitochondrial distribution Various data hint that intermediate filaments may serve to anchor mitochondria in some cell types. Descriptive studies (Toh et al., 1980; Mose-Larsen et al., 1982) have noted association of mitochondria with intermediate filaments. In budding yeast the MDM1 gene encodes an intermediate filament-like protein and is required for mitochondrial inheritance (McConnell and Yaffe, 1992, 1993; Fisk and Yaffe, 1997). In mice engineered to lack the desmin intermediate filament, skeletal muscle mitochondria are mislocalized. In rat neurons overexpressing the mutant neurofilament versions associated with some Charcot-Marie-Tooth disease subtypes, mitochondria do not undergo proper axonal transport. To determine the molecular basis and function for mitochondrial association with various types of intermediate filaments, significant biochemical and genetic analysis will be required. Finally, two phenomena that affect mitochondrial distribution by unknown mechanisms are ubiquitination and fatty acid saturation. S. cerevisiae strains mutated at the ubiquitin ligase gene RSP5 show mitochondrial inheritance defects, and rsp5 mutations suppress mutations in the MDM1 gene, which encodes an intermediate filament-like protein also needed for mitochondrial inheritance (Fisk and Yaffe, 1999). The relevant ubiquitinated substrate is unknown, but it is postulated to be a protein that regulates mitochondrial binding to Mdm1p (Fisk and Yaffe, 1999). Unsaturated fatty acids are required for proper mitochondrial inheritance in budding yeast, as shown by the fact that defects in the Ole1p fatty acid desaturase inhibit proper movement of mitochondria to the bud (Stewart and Yaffe, 1991). A lack of fatty acids in mitochondria could render them less fluid and mobile, less able to undergo fusion or fission, or less able to bind motor

K.G. Hales / Mitochondrion 4 (2004) 285–308

proteins or present adaptor proteins on the mitochondrial surface (Stewart and Yaffe, 1991).

6. Future directions With some of the major protein players in mitochondrial fusion and fission now identified, the focus in those areas now turns to (1) specific molecular mechanisms by which these proteins alter membrane topology and (2) questions of coordination and regulation. Topics for the future include: What are the three dimensional structures of the fusion and fission complexes? In what way does each complex change its shape to bring together or divide mitochondrial membranes? What are the triggers for and inhibitors of those shape changes? Do the dynamin homologs in each complex function as ‘pinchases’ similar to the prototypical dynamin? How are fusion and fission of the outer and inner mitochondrial membranes coordinated? How are tissue-specific variants of the fusion and fission complexes different from each other? How much of a regulatory role do fusion and fission mediators have in apoptosis? In the area of mitochondrial distribution, questions to be addressed include: What are the adaptors connecting mitochondria to motor proteins that traverse cytoskeletal elements? What are the adaptors that connect mitochondria to proteins involved in polymerization and depolymerization of cytoskeletal elements, such as the Arp2/3 complex in S. cerevisiae? How does membrane potential regulate the direction in which mitochondria move along cytoskeletal elements? What other factors regulate directionality of transport? What is the role of intermediate filaments in mitochondrial distribution? Much remains to be discovered in the field of mitochondrial dynamics.

Note added in proof As this article was going to press, it was reported that mutations in human MFN2, a mitochondrial fusion mediator, are associated with Charcot-MarieTooth neuropathy type 2A in some families (Zu¨chner et al., 2004, Nature Genetics 36: 449–451).

303

Acknowledgements K.G.H. is supported by CAREER grant 0133335 from the National Science Foundation and by Davidson College.

References Aiken Hobbs, A.E., Srinivasan, M., McCaffery, J.M., Jensen, R.E., 2001. Mmm1p, a mitochondrial outer membrane protein, is connected to mitochondrial DNA (mtDNA) nucleoids and required for mtDNA stability. J. Cell Biol. 152, 401–410. Alexander, C., Votruba, M., Pesch, U.E., Thiselton, D.L., Mayer, S., Moore, A., et al., 2000. OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome 3q28. Nat. Genet. 26, 211–215. Alto, N.M., Soderling, J., Scott, J.D., 2002. Rab32 is an A-kinase anchoring protein and participates in mitochondrial dynamics. J. Cell Biol. 158, 659–668. Aprille, J.R., 1986. Perinatal development of mitochondria in rat liver, in: Fiskum, G. (Ed.), Mitochondrial Physiology and Pathology. Van Nostrand Reinhold, New York, NY, pp. 66–99. Arimura, S., Tsutsumi, N., 2002. A dynamin-like protein (ADL2b), rather than FtsZ, is involved in Arabidopsis mitochondrial division. Proc. Natl. Acad. Sci. USA 99, 5727–5731. Bach, D., Pich, S., Soriano, F.X., Vega, N., Baumgartner, B., Oriola, J., et al., 2003. Mitofusin-2 determines mitochondrial network architecture and mitochondrial metabolism. A novel regulatory mechanism altered in obesity. J. Biol. Chem. 278, 17190–17197. Bakeeva, L.E., Chentsov, Y.S., Skulachev, V.P., 1981. Ontogenesis of mitochondrial reticulum in rat diaphragm muscle. Eur. J. Cell Biol. 25, 175–181. Beech, P.L., Nheu, T., Schultz, T., Herbert, S., Lithgow, T., Gilson, P.R., McFadden, G.I., 2000. Mitochondrial FtsZ in a chromophyte alga. Science 287, 1276–1279. Bereiter-Hahn, J., Voth, M., 1994. Dynamics of mitochondria in living cells: shape changes, dislocations, fusion, and fission of mitochondria. Microsc. Res. Tech. 27, 198–219. Berger, K.H., Sogo, L.F., Yaffe, M.P., 1997. Mdm12p, a component required for mitochondrial inheritance that is conserved between budding and fission yeast. J. Cell Biol. 136, 545–553. Bleazard, W., McCaffery, J.M., King, E.J., Bale, S., Mozdy, A., Tieu, Q., et al., 1999. The dynamin-related GTPase Dnm1 regulates mitochondrial fission in yeast. Nat. Cell Biol. 1, 298– 304. Boldogh, I., Vojtov, N., Karmon, S., Pon, L.A., 1998. Interaction between mitochondria and the actin cytoskeleton in budding yeast requires two integral mitochondrial outer membrane proteins, Mmm1p and Mdm10p. J. Cell Biol. 141, 1371–1381. Boldogh, I.R., Yang, H.C., Nowakowski, W.D., Karmon, S.L., Hays, L.G., Yates 3rd., J.R., Pon, L.A., 2001a. Arp2/3 complex and actin dynamics are required for actin-based mitochondrial motility in yeast. Proc. Natl. Acad. Sci. USA 98, 3162–3167.

304

K.G. Hales / Mitochondrion 4 (2004) 285–308

Boldogh, I.R., Yang, H.C., Pon, L.A., 2001b. Mitochondrial inheritance in budding yeast. Traffic 2, 368–374. Boldogh, I.R., Nowakowski, D.W., Yang, H.C., Chung, H., Karmon, S., Royes, P., Pon, L.A., 2003. A protein complex containing Mdm10p, Mdm12p, and Mmm1p links mitochondrial membranes and DNA to the cytoskeleton-based segregation machinery. Mol. Biol. Cell 14, 4618–4627. Breckenridge, D.G., Stojanovic, M., Marcellus, R.C., Shore, G.C., 2003. Caspase cleavage product of BAP31 induces mitochondrial fission through endoplasmic reticulum calcium signals, enhancing cytochrome c release to the cytosol. J. Cell Biol. 160, 1115–1127. Burgess, S.M., Delannoy, M., Jensen, R.E., 1994. MMM1 encodes a mitochondrial outer membrane protein essential for establishing and maintaining the structure of yeast mitochondria. J. Cell Biol. 126, 1375–1391. Cerveny, K.L., Jensen, R.E., 2003. The WD-repeats of Net2p interact with Dnm1p and Fis1p to regulate division of mitochondria. Mol. Biol. Cell 14, 4126–4139. Cerveny, K.L., McCaffery, J.M., Jensen, R.E., 2001. Division of mitochondria requires a novel DMN1-interacting protein, Net2p. Mol. Biol. Cell 12, 309–321. Chada, S.R., Hollenbeck, P.J., 2003. Mitochondrial movement and positioning in axons: the role of growth factor signaling. J. Exp. Biol. 206, 1985–1992. Chen, H., Detmer, S.A., Ewald, A.J., Griffin, E.E., Fraser, S.E., Chan, D.C., 2003. Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. J. Cell Biol. 160, 189–200. Clark-Walker, G.D., Miklos, G.L., 1975. Complementation in cytoplasmic petite mutants of yeast to form respiratory competent cells. Proc. Natl. Acad. Sci. USA 72, 372–375. Cox, R.T., Spradling, A.C., 2003. A Balbiani body and the fusome mediate mitochondrial inheritance during Drosophila oogenesis. Development 130, 1579–1590. Danino, D., Hinshaw, J.E., 2001. Dynamin family of mechanoenzymes. Curr. Opin. Cell Biol. 13, 454–460. Delettre, C., Lenaers, G., Griffoin, J.M., Gigarel, N., Lorenzo, C., Belenguer, P., et al., 2000. Nuclear gene OPA1, encoding a mitochondrial dynamin-related protein, is mutated in dominant optic atrophy. Nat. Genet. 26, 207–210. Desagher, S., Martinou, J.C., 2000. Mitochondria as the central control point of apoptosis. Trends Cell Biol. 10, 369–377. De Vos, K., Sable, J., Miller, K.E., Sheetz, M.P., 2003. Expression of phosphatidylinositol (4,5) bisphosphate-specific pleckstrin homology domains alters direction but not the level of axonal transport of mitochondria. Mol. Biol. Cell 14, 3636–3649. Drubin, D.G., Jones, H.D., Wertman, K.F., 1993. Actin structure and function: roles in mitochondrial organization and morphogenesis in budding yeast and identification of the phalloidinbinding site. Mol. Biol. Cell 4, 1277–1294. Ehara, T., Osafune, T., Hase, E., 1995. Behavior of mitochondria in synchronized cells of Chlamydomonas reinhardtii (Chlorophyta). J. Cell Sci. 108 (Pt 2), 499–507. Enriquez, J.A., Cabezas-Herrera, J., Bayona-Bafaluy, M.P., Attardi, G., 2000. Very rare complementation between

mitochondria carrying different mitochondrial DNA mutations points to intrinsic genetic autonomy of the organelles in cultured human cells. J. Biol. Chem. 275, 11207–11215. Eura, Y., Ishihara, N., Yokota, S., Mihara, K., 2003. Two mitofusin proteins, mammalian homologues of FZO, with distinct functions are both required for mitochondrial fusion. J. Biochem. (Tokyo) 134, 333–344. Fehrenbacher, K., Huckaba, T., Yang, H.C., Boldogh, I., Pon, L., 2003. Actin comet tails, endosomes and endosymbionts. J. Exp. Biol. 206, 1977–1984. Fekkes, P., Shepard, K.A., Yaffe, M.P., 2000. Gag3p, an outer membrane protein required for fission of mitochondrial tubules. J. Cell Biol. 151, 333–340. Fisk, H.A., Yaffe, M.P., 1997. Mutational analysis of Mdm1p function in nuclear and mitochondrial inheritance. J. Cell Biol. 138, 485–494. Fisk, H.A., Yaffe, M.P., 1999. A role for ubiquitination in mitochondrial inheritance in Saccharomyces cerevisiae. J. Cell Biol. 145, 1199–1208. Frank, S., Gaume, B., Bergmann-Leitner, E.S., Leitner, W.W., Robert, E.G., Catez, F., et al., 2001. The role of dynamin-related protein 1, a mediator of mitochondrial fission, in apoptosis. Dev. Cell 1, 515–525. Frieden, M., James, D., Castelbou, C., Danckaert, A., Martinou, J.C., Demaurex, N., 2004. Ca(2C) homeostasis during mitochondrial fragmentation and perinuclear clustering induced by hFis1. J. Biol. Chem. 279, 22704–22714. Fritz, S., Rapaport, D., Klanner, E., Neupert, W., Westermann, B., 2001. Connection of the mitochondrial outer and inner membranes by Fzo1 is critical for organellar fusion. J. Cell Biol. 152, 683–692. Fritz, S., Weinbach, N., Westermann, B., 2003. Mdm30 is an F-box protein required for maintenance of fusion-competent mitochondria in yeast. Mol. Biol. Cell 14, 2303–2313. Fuchs, F., Prokisch, H., Neupert, W., Westermann, B., 2002. Interaction of mitochondria with microtubules in the filamentous fungus Neurospora crassa. J. Cell Sci. 115, 1931–1937. Fukushima, N.H., Brisch, E., Keegan, B.R., Bleazard, W., Shaw, J.M., 2001. The GTPase effector domain sequence of the Dnm1p GTPase regulates self-assembly and controls a ratelimiting step in mitochondrial fission. Mol. Biol. Cell 12, 2756– 2766. Fuller, M.T., 1993. Spermatogenesis, in: Bate, M., MartinezArias, A. (Eds.), The Development of Drosophila melanogaster. Cold Spring Harbor, New York, pp. 71–147. Gindhart Jr., J.G., Desai, C.J., Beushausen, S., Zinn, K., Goldstein, L.S., 1998. Kinesin light chains are essential for axonal transport in Drosophila. J. Cell Biol. 141, 443–454. Grieder, N.C., de Cuevas, M., Spradling, A.C., 2000. The fusome organizes the microtubule network during oocyte differentiation in Drosophila. Development 127, 4253–4264. Griparic, L., van der Wel, N.N., Orozco, I.J., Peters, P.J., van der Bliek, A.M., 2004. Loss of the intermembrane space protein Mgm1/OPA1 induces swelling and localized constrictions along the lengths of mitochondria. J. Biol. Chem. 279, 18792–18798.

K.G. Hales / Mitochondrion 4 (2004) 285–308 Guan, K., Farh, L., Marshall, T.K., Deschenes, R.J., 1993. Normal mitochondrial structure and genome maintenance in yeast requires the dynamin-like product of the MGM1 gene. Curr. Genet. 24, 141–148. Hales, K.G., Fuller, M.T., 1997. Developmentally regulated mitochondrial fusion mediated by a conserved, novel, predicted GTPase. Cell 90, 121–129. Hanekamp, T., Thorsness, M.K., Rebbapragada, I., Fisher, E.M., Seebart, C., Darland, M.R., et al., 2002. Maintenance of mitochondrial morphology is linked to maintenance of the mitochondrial genome in Saccharomyces cerevisiae. Genetics 162, 1147–1156. Harder, Z., Zunino, R., McBride, H., 2004. Sumo1 conjugates mitochondrial substrates and participates in mitochondrial fission. Curr. Biol. 14, 340–345. Herlan, M., Vogel, F., Bornhovd, C., Neupert, W., Reichert, A.S., 2003. Processing of Mgm1 by the Rhomboid-type protease Pcp1 is required for maintenance of mitochondrial morphology and of mitochondrial DNA. J. Biol. Chem. 2003;. Herlan, M., Bornhovd, C., Hell, K., Neupert, W., Reichert, A.S., 2004. Alternative topogenesis of Mgm1 and mitochondrial morphology depend on ATP and a functional import motor. J. Cell Biol. 165, 167–173. Hermann, G.J., King, E.J., Shaw, J.M., 1997. The yeast gene, MDM20, is necessary for mitochondrial inheritance and organization of the actin cytoskeleton. J. Cell Biol. 137, 141– 153. Hermann, G.J., Thatcher, J.W., Mills, J.P., Hales, K.G., Fuller, M.T., Nunnari, J., Shaw, J.M., 1998. Mitochondrial fusion in yeast requires the transmembrane GTPase Fzo1p. J. Cell Biol. 143, 359–373. Hodgson, A.N., Bernard, R.T.F., 1986. Ultrastructure of the sperm and spermatogenesis of three species of Mytilidae Mollusca bivalvia. Gamete Res. 15, 123–136. Hoffmann, H.P., Avers, C.J., 1973. Mitochondrion of yeast: ultrastructural evidence for one giant, branched organelle per cell. Science 181, 749–751. Hollenbeck, P.J., 1996. The pattern and mechanism of mitochondrial transport in axons. Front Biosci. 1, d91–d102. Huffaker, T.C., Thomas, J.H., Botstein, D., 1988. Diverse effects of beta-tubulin mutations on microtubule formation and function. J. Cell Biol. 106, 1997–2010. Hurd, D.D., Saxton, W.M., 1996. Kinesin mutations cause motor neuron disease phenotypes by disrupting fast axonal transport in Drosophila. Genetics 144, 1075–1085. Hwa, J.J., Hiller, M.A., Fuller, M.T., Santel, A., 2002. Differential expression of the Drosophila mitofusin genes fuzzy onions (fzo) and dmfn. Mech. Dev. 116, 213–216. Ishihara, N., Jofuku, A., Eura, Y., Mihara, K., 2003. Regulation of mitochondrial morphology by membrane potential, and DRP1dependent division and FZO1-dependent fusion reaction in mammalian cells. Biochem. Biophys. Res. Commun. 301, 891–898. Jakobs, S., Martini, N., Schauss, A.C., Egner, A., Westermann, B., Hell, S.W., 2003. Spatial and temporal dynamics of budding yeast mitochondria lacking the division component Fis1p. J. Cell Sci. 116, 2005–2014.

305

James, D.I., Parone, P.A., Mattenberger, Y., Martinou, J.C., 2003. hFis1, a novel component of the mammalian mitochondrial fission machinery. J. Biol. Chem. 2003;. Jin, J.B., Bae, H., Kim, S.J., Jin, Y.H., Goh, C.H., Kim, D.H., et al., 2003. The Arabidopsis dynamin-like proteins ADL1C and ADL1E play a critical role in mitochondrial morphogenesis. Plant Cell 15, 2357–2369. Jones, B.A., Fangman, W.L., 1992. Mitochondrial DNA maintenance in yeast requires a protein containing a region related to the GTP-binding domain of dynamin. Genes Dev. 6, 380–389. Karbowski, M., Lee, Y.J., Gaume, B., Jeong, S.Y., Frank, S., Nechushtan, A., et al., 2002. Spatial and temporal association of Bax with mitochondrial fission sites, Drp1, and Mfn2 during apoptosis. J. Cell Biol. 159, 931–938. Karbowski, M., Arnoult, D., Chen, H., Chan, D.C., Smith, C.L., Youle, R.J., 2004. Quantitation of mitochondrial dynamics by photolabeling of individual organelles shows that mitochondrial fusion is blocked during the Bax activation phase of apoptosis. J. Cell Biol. 164, 493–499. Kawano, S., Takano, H., Imai, J., Mori, K., Kurioiwa, T., 1993. A genetic system controlling mitochondrial fusion in the slime mould, Physarum polycephalum. Genetics 133, 213–224. Kawano, S., Takano, H., Kuroiwa, T., 1995. Sexuality of mitochondria: fusion, recombination, and plasmids. Int. Rev. Cytol. 161, 49–110. Kirkwood, S.P., Munn, E.A., Brooks, G.A., 1986. Mitochondrial reticulum in limb skeletal muscle. Am. J. Physiol. 251, C395– C402. Koch, K.V., Suelmann, R., Fischer, R., 2003. Deletion of mdmB impairs mitochondrial distribution and morphology in Aspergillus nidulans. Cell Motil. Cytoskeleton 55, 114–124. Kondo-Okamoto, N., Shaw, J.M., Okamoto, K., 2003. Mmm1p spans both the outer and inner mitochondrial membranes and contains distinct domains for targeting and foci formation. J. Biol. Chem. 278, 48997–49005. Kruger, J.C.d., 1991. Ultrastructure of sperm development in the plant-parasitic nematode Xiphinema theresiae. J. Morph. 210, 163–174. Labrousse, A.M., Zappaterra, M.D., Rube, D.A., van der Bliek, A.M., 1999. C. elegans dynamin-related protein DRP-1 controls severing of the mitochondrial outer membrane. Mol. Cell 4, 815–826. Lazzarino, D.A., Boldogh, I., Smith, M.G., Rosand, J., Pon, L.A., 1994. Yeast mitochondria contain ATP-sensitive, reversible actin-binding activity. Mol. Biol. Cell 5, 807–818. Legesse-Miller, A., Massol, R.H., Kirchhausen, T., 2003. Constriction and dnm1p recruitment are distinct processes in mitochondrial fission. Mol. Biol. Cell 14, 1953–1963. Legros, F., Lombes, A., Frachon, P., Rojo, M., 2002. Mitochondrial fusion in human cells is efficient, requires the inner membrane potential, and is mediated by mitofusins. Mol. Biol. Cell 13, 4343–4354. Legros, F., Malka, F., Frachon, P., Lombes, A., Rojo, M., 2004. Organization and dynamics of human mitochondrial DNA. J. Cell Sci. 117, 2653–2662.

306

K.G. Hales / Mitochondrion 4 (2004) 285–308

Ligon, L.A., Steward, O., 2000. Role of microtubules and actin filaments in the movement of mitochondria in the axons and dendrites of cultured hippocampal neurons. J. Comp. Neurol. 427, 351–361. Margolin, W., 2001. Spatial regulation of cytokinesis in bacteria. Curr. Opin. Microbiol. 4, 647–652. Martin, M., Iyadurai, S.J., Gassman, A., Gindhart Jr., J.G., Hays, T.S., Saxton, W.M., 1999. Cytoplasmic dynein, the dynactin complex, and kinesin are interdependent and essential for fast axonal transport. Mol. Biol. Cell 10, 3717–3728. Mattenberger, Y., James, D.I., Martinou, J.C., 2003. Fusion of mitochondria in mammalian cells is dependent on the mitochondrial inner membrane potential and independent of microtubules or actin. Fed. Eur. Biol. Soc. Lett. 538, 53–59. McConnell, S.J., Yaffe, M.P., 1992. Nuclear and mitochondrial inheritance in yeast depends on novel cytoplasmic structures defined by the MDM1 protein. J. Cell Biol. 118, 385–395. McConnell, S.J., Yaffe, M.P., 1993. Intermediate filament formation by a yeast protein essential for organelle inheritance. Science 260, 687–689. McQuibban, G.A., Saurya, S., Freeman, M., 2003. Mitochondrial membrane remodelling regulated by a conserved rhomboid protease. Nature 423, 537–541. Meeusen, S., Nunnari, J., 2003. Evidence for a two membranespanning autonomous mitochondrial DNA replisome. J. Cell Biol. 163, 503–510. Meeusen, S., Tieu, Q., Wong, E., Weiss, E., Schieltz, D., Yates, J.R., Nunnari, J., 1999. Mgm101p is a novel component of the mitochondrial nucleoid that binds DNA and is required for the repair of oxidatively damaged mitochondrial DNA. J. Cell Biol. 145, 291–304. Messerschmitt, M., Jakobs, S., Vogel, F., Fritz, S., Dimmer, K.S., Neupert, W., Westermann, B., 2003. The inner membrane protein Mdm33 controls mitochondrial morphology in yeast. J. Cell Biol. 160, 553–564. Miller, K.E., Sheetz, M.P., 2004. Axonal mitochondrial transport and potential are correlated. J. Cell Sci. 117, 2791–2804. Misaka, T., Miyashita, T., Kubo, Y., 2002. Primary structure of a dynamin-related mouse mitochondrial GTPase and its distribution in brain, subcellular localization, and effect on mitochondrial morphology. J. Biol. Chem. 277, 15834–15842. Miyakawa, I., Higo, K., Osake, F., Sando, N., 1994. Double staining of mitochondria and mitochondrial nucleoids in the living yeast during the life cycle. J. Gen. Appl. Microbiol. 40, 1–14. Morris, R.L., Hollenbeck, P.J., 1995. Axonal transport of mitochondria along microtubules and F-actin in living vertebrate neurons. J. Cell Biol. 131, 1315–1326. Mose-Larsen, P., Bravo, R., Fey, S.J., Small, J.V., Celis, J.E., 1982. Putative association of mitochondria with a subpopulation of intermediate-sized filaments in cultured human skin fibroblasts. Cell 31, 681–692. Mozdy, A.D., McCaffery, J.M., Shaw, J.M., 2000. Dnm1p GTPasemediated mitochondrial fission is a multi-step process requiring the novel integral membrane component Fis1p. J. Cell Biol. 151, 367–380.

Nakada, K., Inoue, K., Ono, T., Isobe, K., Ogura, A., Goto, Y.I., et al., 2001. Inter-mitochondrial complementation: mitochondria-specific system preventing mice from expression of disease phenotypes by mutant mtDNA. Nat. Med. 7, 934–940. Nangaku, M., Sato-Yoshitake, R., Okada, Y., Noda, Y., Takemura, R., Yamazaki, H., Hirokawa, N., 1994. KIF1B, a novel microtubule plus end-directed monomeric motor protein for transport of mitochondria. Cell 79, 1209–1220. Nishida, K., Takahara, M., Miyagishima, S.Y., Kuroiwa, H., Matsuzaki, M., Kuroiwa, T., 2003. Dynamic recruitment of dynamin for final mitochondrial severance in a primitive red alga. Proc. Natl. Acad. Sci. USA 100, 2146–2151. Nunnari, J., Marshall, W.F., Straight, A., Murray, A., Sedat, J.W., Walter, P., 1997. 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. Olichon, A., Emorine, L.J., Descoins, E., Pelloquin, L., Brichese, L., Gas, N., et al., 2002. The human dynaminrelated protein OPA1 is anchored to the mitochondrial inner membrane facing the inter-membrane space. Fed. Eur. Biol. Soc. Lett. 523, 171–176. Olichon, A., Baricault, L., Gas, N., Guillou, E., Valette, A., Belenguer, P., Lenaers, G., 2003. Loss of OPA1 perturbates the mitochondrial inner membrane structure and integrity, leading to cytochrome c release and apoptosis. J. Biol. Chem. 278, 7743–7746. Olson, G.E., Winfrey, V.P., 1992. Structural organization of surface domains of sperm mitochondria. Mol. Reprod. Dev. 33, 89–98. Ono, T., Isobe, K., Nakada, K., Hayashi, J.I., 2001. Human cells are protected from mitochondrial dysfunction by complementation of DNA products in fused mitochondria. Nat. Genet. 28, 272–275. Otsuga, D., Keegan, B.R., Brisch, E., Thatcher, J.W., Hermann, G.J., Bleazard, W., Shaw, J.M., 1998. The dynamin-related GTPase, Dnm1p, controls mitochondrial morphology in yeast. J. Cell Biol. 143, 333–349. Pelloquin, L., Belenguer, P., Menon, Y., Gas, N., Ducommun, B., 1999. Fission yeast Msp1 is a mitochondrial dynamin-related protein. J. Cell Sci. 112 (Pt 22), 4151–4161. Pereira, A.J., Dalby, B., Stewart, R.J., Doxsey, S.J., Goldstein, L.S., 1997. Mitochondrial association of a plus end-directed microtubule motor expressed during mitosis in Drosophila. J. Cell Biol. 136, 1081–1090. Prokisch, H., Neupert, W., Westermann, B., 2000. Role of MMM1 in maintaining mitochondrial morphology in Neurospora crassa. Mol. Biol. Cell 11, 2961–2971. Rapaport, D., Brunner, M., Neupert, W., Westermann, B., 1998. Fzo1p is a mitochondrial outer membrane protein essential for the biogenesis of functional mitochondria in Saccharomyces cerevisiae. J. Biol. Chem. 273, 20150–20155. Renzaglia, K.S., Duckett, J.G., 1989. Ultrastructural studies of spermatogenesis in Antherocerophyta V. Nuclear metamorphosis and the posterior mitochondrion of Notothylas orbicularis and Phaeoceros laevis. Protoplasma 151, 137–150.

K.G. Hales / Mitochondrion 4 (2004) 285–308 Rojo, M., Legros, F., Chateau, D., Lombes, A., 2002. Membrane topology and mitochondrial targeting of mitofusins, ubiquitous mammalian homologs of the transmembrane GTPase Fzo. J. Cell Sci. 115, 1663–1674. Santel, A., Fuller, M.T., 2001. Control of mitochondrial morphology by a human mitofusin. J. Cell Sci. 114, 867–874. Santel, A., Frank, S., Gaume, B., Herrler, M., Youle, R.J., Fuller, M.T., 2003. Mitofusin-1 protein is a generally expressed mediator of mitochondrial fusion in mammalian cells. J. Cell Sci. 116, 2763–2774. Seeler, J.S., Dejean, A., 2003. Nuclear and unclear functions of SUMO. Nat. Rev. Mol. Cell Biol. 4, 690–699. Sesaki, H., Jensen, R.E., 1999. Division versus fusion: Dnm1p and Fzo1p antagonistically regulate mitochondrial shape. J. Cell Biol. 147, 699–706. Sesaki, H., Jensen, R.E., 2001. UGO1 encodes an outer membrane protein required for mitochondrial fusion. J. Cell Biol. 152, 1123–1134. Sesaki, H., Jensen, R.E., 2004. Ugo1p links the Fzo1p and Mgm1p GTPases for mitochondrial fusion. J. Biol. Chem. 2004;. Sesaki, H., Southard, S.M., Hobbs, A.E., Jensen, R.E., 2003a. Cells lacking Pcp1p/Ugo2p, a rhomboid-like protease required for Mgm1p processing, lose mtDNA and mitochondrial structure in a Dnm1p-dependent manner, but remain competent for mitochondrial fusion. Biochem. Biophys. Res. Commun. 308, 276–283. Sesaki, H., Southard, S.M., Yaffe, M.P., Jensen, R.E., 2003b. Mgm1p, a dynamin-related GTPase, is essential for fusion of the mitochondrial outer membrane. Mol. Biol. Cell 14, 2342–2356. Shaw, J.M., Nunnari, J., 2002. Mitochondrial dynamics and division in budding yeast. Trends Cell Biol. 12, 178–184. Shepard, K.A., Yaffe, M.P., 1999. The yeast dynamin-like protein, Mgm1p, functions on the mitochondrial outer membrane to mediate mitochondrial inheritance. J. Cell Biol. 144, 711–720. Shin, H., Takatsu, H., Mukai, H., Munekata, E., Murakami, K., Nakayama, K., 1999. Intermolecular and interdomain interactions of a dynamin-related GTP-binding protein, Dnm1p/Vpsp-like protein. J. Biol. Chem. 274, 2780–2785. Simon, V.R., Swayne, T.C., Pon, L.A., 1995. Actin-dependent mitochondrial motility in mitotic yeast and cell-free systems: identification of a motor activity on the mitochondrial surface. J. Cell Biol. 130, 345–354. Singer, J.M., Shaw, J.M., 2003. Mdm20 protein functions with Nat3 protein to acetylate Tpm1 protein and regulate tropomyosin– actin interactions in budding yeast. Proc. Natl. Acad. Sci. USA 100, 7644–7649. Skulachev, V.P., 1990. Power transmission along biological membranes. J. Membr. Biol. 114, 97–112. Skulachev, V.P., 2001. Mitochondrial filaments and clusters as intracellular power-transmitting cables. Trends Biochem. Sci. 26, 23–29. Smirnova, E., Shurland, D.L., Ryazantsev, S.N., van der Bliek, A.M., 1998. A human dynamin-related protein controls the distribution of mitochondria. J. Cell Biol. 143, 351–358. Smirnova, E., Griparic, L., Shurland, D.L., van der Bliek, A.M., 2001. Dynamin-related protein Drp1 is required for mitochondrial division in mammalian cells. Mol. Biol. Cell 12, 2245– 2256.

307

Smith, D.M., 1931. The ontogenic history of the mitochondria of the hepatic cell of the white rat. J. Morph. Physiol. 52, 485–512. Smith, M.G., Simon, V.R., O’Sullivan, H., Pon, L.A., 1995. Organelle-cytoskeletal interactions: actin mutations inhibit meiosis-dependent mitochondrial rearrangement in the budding yeast Saccharomyces cerevisiae. Mol. Biol. Cell 6, 1381–1396. Sogo, L.F., Yaffe, M.P., 1994. Regulation of mitochondrial morphology and inheritance by Mdm10p, a protein of the mitochondrial outer membrane. J. Cell Biol. 126, 1361–1373. Steinberg, G., Schliwa, M., 1993. Organelle movements in the wild type and wall-less fz;sg;os-1 mutants of Neurospora crassa are mediated by cytoplasmic microtubules. J. Cell Sci. 106 (Pt 2), 555–564. Stevens, B., 1981. Mitochondrial structure, in: Strathern, J.M., Jones, E.W., Broach, J.R. (Eds.), The Molecular Biology of the Yeast Saccharomyces: Life Cycle and Inheritance. Cold Spring Harbor, Cold Spring Harbor, NY, pp. 471–504. Stewart, L.C., Yaffe, M.P., 1991. A role for unsaturated fatty acids in mitochondrial movement and inheritance. J. Cell Biol. 115, 1249–1257. Stojanovski, D., Koutsopoulos, O.S., Okamoto, K., Ryan, M.T., 2004. Levels of human Fis1 at the mitochondrial outer membrane regulate mitochondrial morphology. J. Cell Sci. 117, 1201–1210. Stowers, R.S., Megeath, L.J., Gorska-Andrzejak, J., Meinertzhagen, I.A., Schwarz, T.L., 2002. Axonal transport of mitochondria to synapses depends on milton, a novel Drosophila protein. Neuron 36, 1063–1077. Sturmer, K., Baumann, O., Walz, B., 1995. Actin-dependent lightinduced translocation of mitochondria and ER cisternae in the photoreceptor cells of the locust Schistocerca gregaria. J Cell Sci 108 (Pt 6), 2273–2283. Suelmann, R., Fischer, R., 2000. Mitochondrial movement and morphology depend on an intact actin cytoskeleton in Aspergillus nidulans. Cell Motil. Cytoskeleton 45, 42–50. Takano, H., Kawano, S., Sasaki, N., Kuroiwa, H., Kuroiwa, T., 2002. Characterization of a putative fusogen encoded in a mitochondrial plasmid of Physarum polycephalum. J. Plant Res. 115, 255–261. Tanaka, Y., Kanai, Y., Okada, Y., Nonaka, S., Takeda, S., Harada, A., Hirokawa, N., 1998. Targeted disruption of mouse conventional kinesin heavy chain, kif5B, results in abnormal perinuclear clustering of mitochondria. Cell 93, 1147–1158. Tates, A.D., 1971. Cytodifferentiation during spermatogenesis in Drosophila melanogaster: an electron microscope study. Rijksuniversiteit, Leiden. Thorsness, P.E., Fox, T.D., 1993. Nuclear mutations in Saccharomyces cerevisiae that affect the escape of DNA from mitochondria to the nucleus. Genetics 134, 21–28. Tieu, Q., Nunnari, J., 2000. Mdv1p is a WD repeat protein that interacts with the dynamin-related GTPase, Dnm1p, to trigger mitochondrial division. J. Cell Biol. 151, 353–366. Tieu, Q., Okreglak, V., Naylor, K., Nunnari, J., 2002. The WD repeat protein, Mdv1p, functions as a molecular adaptor by interacting with Dnm1p and Fis1p during mitochondrial fission. J. Cell Biol. 158, 445–452.

308

K.G. Hales / Mitochondrion 4 (2004) 285–308

Toh, B.H., Lolait, S.J., Mathy, J.P., Baum, R., 1980. Association of mitochondria with intermediate filaments and of polyribosomes with cytoplasmic actin. Cell Tissue Res. 211, 163–169. Tyler, D., 1992. The Mitochondrion in Health and Disease. VCH, New York, NY. van der Bliek, A.M., 1999. Functional diversity in the dynamin family. Trends Cell Biol. 9, 96–102. Visser, W., van Spronsen, E.A., Nanninga, N., Pronk, J.T., Gijs Kuenen, J., van Dijken, J.P., 1995. Effects of growth conditions on mitochondrial morphology in Saccharomyces cerevisiae. Antonie Van Leeuwenhoek 67, 243–253. Weir, B.A., Yaffe, M.P., 2004. Mmd1p, a novel, conserved protein essential for normal mitochondrial morphology and distribution in the fission yeast, Schizosaccharomyces pombe. Mol. Biol. Cell 2004;. Werth, J.L., Thayer, S.A., 1994. Mitochondria buffer physiological calcium loads in cultured rat dorsal root ganglion neurons. J. Neurosci. 14, 348–356. Wong, E.D., Wagner, J.A., Gorsich, S.W., McCaffery, J.M., Shaw, J.M., Nunnari, J., 2000. The dynamin-related GTPase, Mgm1p, is an intermembrane space protein required for maintenance of fusion competent mitochondria. J. Cell Biol. 151, 341–352. Wong, E.D., Wagner, J.A., Scott, S.V., Okreglak, V., Holewinske, T.J., Cassidy-Stone, A., Nunnari, J., 2003. The intramitochondrial dynamin-related GTPase, Mgm1p, is a component of a protein complex that mediates mitochondrial fusion. J. Cell Biol. 160, 303–311.

Yaffe, M.P., Harata, D., Verde, F., Eddison, M., Toda, T., Nurse, P., 1996. Microtubules mediate mitochondrial distribution in fission yeast. Proc. Natl. Acad. Sci. USA 93, 11664–11668. Yaffe, M.P., Stuurman, N., Vale, R.D., 2003. Mitochondrial positioning in fission yeast is driven by association with dynamic microtubules and mitotic spindle poles. Proc. Natl. Acad. Sci. USA 100, 11424–11428. Yang, H.C., Palazzo, A., Swayne, T.C., Pon, L.A., 1999. A retention mechanism for distribution of mitochondria during cell division in budding yeast. Curr. Biol. 9, 1111–1114. Yoon, Y., Pitts, K., McNiven, M.A., 2001. Mammalian dynaminlike protein DLP1 tubulates membranes. Mol. Biol. Cell 12, 2894–2905. Yoon, Y., Krueger, E.W., Oswald, B.J., McNiven, M.A., 2003. The mitochondrial protein hFis1 regulates mitochondrial fission in mammalian cells through an interaction with the dynamin-like protein DLP1. Mol. Cell Biol. 23, 5409–5420. Youngman, M.J., Aiken Hobbs, A.E., Burgess, S.M., Srinivasan, M., Jensen, R.E., 2004. Mmm2p, a mitochondrial outer membrane protein required for mitochondrial shape and maintenance of mtDNA nucleoids. J. Cell Biol. 164, 677–688. Zhao, C., Takita, J., Tanaka, Y., Setou, M., Nakagawa, T., Takeda, S., et al., 2001. Charcot-Marie-Tooth disease type 2A caused by mutation in a microtubule motor KIF1Bbeta. Cell 105, 587–597. Zorov, D.B., Skulachev, V.P., Khalangk, V., 1990. Membranous electric cable: 4. Mitochondrial helix of rat spermatozoan. Biologicheskie Membrany 7, 243–249.