Mechanisms of fast and slow axonal transport.

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Annu. Rev. Neurosci. 1991.14:59-92. Downloaded from www.annualreviews.org by Lomonosov Moscow State University on 01/31/14. For personal use only.

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1991

1991. 14:59-92

by Annual Reviews Inc. All rights reserved

MECHANISMS OF FAST AND SLOW AXONAL TRANSPORT Richard B. Vallee

Worcester Foundation for Experimental Biology, Cell Biology Group, Shrewsbury, M assachusetts 01545 G('orge S. Bloom

Department of Cell Biology and Neuroscience, University of Texas Southwestern Medical Center, Dallas, Texas 75235 KEY WORDS:

microtubule, kinesin, dynein, dynamin, tubulin.

INTRODUCTION One of the features of the neuron that most strikingly distinguishes it from other cells is its highly elongated processes. These specializations pose a particular challenge to the normal metabolic machinery of the cell. Because biosynthesis is largely restricted to the region of the cell body and the dendrites, there must be a constant flow of material from these portions of the cell out into the axon. This process is known as "anterograde" (or "orthograde") axonal transport. The return of materials toward the cell body is by "retrograde" axonal transport. Axonal transport is probably mediated by several distinct mechanisms. Evidence for multiple rate classes for anterograde transport has come from an elegant and relatively simple type of experiment. Radiolabeled amino acids are injected into the eye or into PNS ganglia to label neuronal cell bodies. Labeled proteins are then detected in the nerve as a function of time and distance, and analyzed by sodium dodecyl sulfate (SDS) poly acrylamide gel electrophoresis. This analysis has revealed that different proteins travel at different rates within the axon, allowing discrete classes of transported proteins to be defined. Anterograde transport alone exhibits at least five distinct rate classes 59 o I 47-006Xj91 j030 1-0059

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based on observations of the time-course of movement of radio labeled proteins outward from the cell body (see e.g. Willard et al 1 97 4, Hoffman & Lasek 1975; reviewed in Grafstein & Forman 1 980, Brady & Lasek 1 982). "Rapid" anterograde transport is represented by at least two rate classes, both of which involve the movement of membranous material: In mammals, class I polypeptides migrate at rates of '" 1 00-400 mm/day, and class II polypeptides at '" 20-70 mm/day (Grafstein & Forman 1 980). The difference in rate classes could be due to sieving of membranous organelles of different sizes as they pass through the cytoskeletal meshwork of the axoplasm, rather than a differencc in transport mechanism (Allen et al 1 985, Vale et al 1 985c). Rate classes IV and V represent slow axonal transport and are discussed below. Rate class III, whose polypeptides migrate at '" 3-20 mm/day, seems more like slow transport in polypeptide composition, but very little is known about this transport component. Retrograde transport has not been as precisely analyzed as anterograde transport because of the inability to introduce label in a coherent pulse to large numbers of axon termini. Reported rates for retrograde transport rates have varied over a wide range, but have generally been of the same order as the rapid anterograde rates (up to '" 300 mm/day; Grafstein & Forman 1 980). The subcellular components involved in rapid anterograde and retro grade transport are largely distinct. Rapid anterograde transport involves the movement of smooth endoplasmic reticulum, small vesicles, including synaptic vesicles, and plasma membrane components (Grafstein & Forman 1 980). Retrograde transport seems to be an exaggerated manifestation of the normal endocytic process seen in cells in general. Among the more noticeable structures included in retrograde transport are lysosomes and "multivesiculate" bodies, which may be degradative structures but whose function is incompletely understood (Tsukita & Ishikawa 1 980, Smith 1 980). Pinocytic vesicles containing nonspecific labels such as horseradish peroxidase also travel in the retrograde direction, as do ligands such as nerve growth factor, taken up by receptor-mediated endocytosis. Mito chondria appear to travel in both the anterograde and retrograde direc tions, and individual mitochondria may move bidirectionally (Smith 1 972, Forman et aI1977, Allen et aI1 982). Slow axonal transport (rate classes IV and V, also referred to as slow components b and a (SCb and SCa): "'0. 1-4 mm/day), involves the move ment of components of the cytoskeleton and membrane skeleton (see e.g. Hoffman & Lasek 197 5) and associated proteins, including cytoplasmic enzymes of intermediary metabolism (Brady & Lasek 1 9 8 1 ) . The form in which the cytoskeletal proteins are transported is not certain. In metabolic labeling studies, however, coherent peaks of radiolabeled cytoskeletal pro-


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teins are observed to persist over prolonged periods. This suggests that these proteins may be transported in particulate or polymeric form. Slow anterograde transport may serve to replenish worn out structural proteins and to supply such proteins to growing axons. Attempts to monitor slow retrograde transport were made by direct chemical modification of nerves, but labeling of extracellular proteins obscured the behavior of the intra neuronal proteins (Fink & Gainer 1 980). The mechanism of axonal transport has been a long-standing mystery; however, recent advances have led to new insights into how fast transport may occur and some new views on the nature and mechanism of slow transport. After many years of uncertainty, it is now clear that micro tubules play a central role in rapid transport. Three force-producing "motor proteins" capable of interacting with microtubules have also been identified in neural tissue-kinesin, dynein, and dynamin. The properties of kinesin and dynein strongly suggest a role in fast anterograde and retrograde axonal transport, respectively. The function of dynamin is less obvious, though a role in slow transport is suggested by some of its properties (see below). In this chapter we explore current models of rapid and slow transport. In particular, we try to understand axonal transport from the new vantage point afforded by the identification of the motor proteins. In the case of slow transport, we also discuss recent evidence on the biochemical nature of the transported components. This chapter is not intended as a com prehensive review of the physiology of axonal transport, which has been thoroughly reviewed elsewhere (Lasek & Hoffman 1 976, Grafstein & For man 1 980, Brady & Lasek 1 982, Brady & Black 1 986, Hollenbeck 1 989b, Nixon 1 987, 1 990). We refer the reader to more extensive reviews on the properties of the motor proteins (Vale 1987, Porter & Johnson 1 989, Vallee & Shpetner 1 990), as well as shorter discussions of the role of kinesin (Schnapp & Reese 1 986) and cytoplasmic dynein (Vallee et al 1 989a) in anterograde and retrograde transport, respectively.

FAST AXONAL TRANSPORT General Principles

For directed axonal transport to occur, some element of the transport machinery must have a directional polarity. The microtubules fulfill this requirement in that they are themselves polar structures. Within the axon, virtually all of the microtubules have been reported to be oriented with their so-called plus ends toward the axon terminus, and the minus ends toward the cell body (Heidemann et al 1 98 1 , Burton & Paige 1 98 1). [This


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is in contrast to dendritic microtubules, which have been found to have a mixed polarity, with only half oriented with their plus ends away from the cell body (Burton 1 988, Baas et al 1 988).] Present evidence on the structure of intermediate filaments, including neurofilaments-the other major cyto skeletal polymers in the axon-indicates that they are probably nonpolar (Fraser et al 1 986). More direct evidence for the involvement of microtubules in rapid axonal transport has come from a variety oflines of evidence. For example, rapid bidirectional transport occurs in arthropod axons, as well as newly forming vertebrate axons, despite the absence of neurofilaments (e.g. see Fernandez et al 1 97 1 , Samson 1 97 1 , Peters & Vaughn 1 967). In mature mammalian neurons, which do contain neurofi1aments, exposure to the toxin fJ-fJ'-iminodipropionitri1e (IDPN) dissociates microtubules and neuro filaments into discrete domains within the axon; under these conditions, rapidly transported radiolabel is associated only with the microtubules, (Papasozomenos et al 1 9 8 1 1 982, Griffin et al 1 983). Most dramatically, organelles have been demonstrated to move along individual microtubules in vitro (Allen et a1 1 985, Vale et al 1 985a, Schnapp et aI1 985). A substantial body of evidence obtained during the past few years points to kinesin and cytoplasmic dynein as being responsible for generating the forces for the organelle movements that underlie anterograde and retrograde axonal transport. In light of these new developments, the remainder of this section focuses on the properties of these two proteins and evidence regarding their functional role in the cell. ,

Properties of Kinesin

The discovery of kinesin and cytoplasmic dynein emerged from two inde pendent, but related lines of investigation. The identification of kinesin derived from the use of video-enhanced light microscopy to study fast axonal transport in axoplasm extruded from squid giant axons. Such preparations retain many of the structural features of the intact axon and are capable of supporting bidirectional organelle transport for several hours following isolation. Because extruded axoplasm lacks a surrounding plasma membrane, the axon interior is freely accessible to externally applied solutions. By varying the composition of the perfusion buffers, it has been possible to define many of the biochemical requirements and pharmacological properties of fast axonal transport in this system. With this approach, it became clear that ATP is obligatory for fast axonal transport in both directions (Brady et al 1 982, 1 985). The ATP requirement suggested that one or more ATPases capable of interacting with both organelles and microtubules might generate the forces for antero grade and retrograde organelle motility in the axon. A potential route for


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identifying such enzymes was provided by the effect on axoplasm of 5' adenylylimidodiphosphate (AMPPNP), a nonhydrolyzable ATP analog. AMPPNP not only abolished organelle motility in isolated axoplasm, but also appeared to promote the binding of organelles to microtubules (Lasek & Brady 1 98 5). This was reminiscent of the rigor complexes observed between actin and myosin and between ciliary and flagellar dynein and microtubules in the absence of ATP, and suggested that the organelles might be linked to microtubules by motor proteins that were trapped in a rigor state by AMPPNP (Lasek & Brady 1 98 5). Based on this assump tion, purification schemes were devised which made use of AMPPNP to induce the specific binding of the putative motor proteins to microtubules (Brady 1 98 5, Vale et aI 1 98 5a). Taxol-based microtubule purification was used (Vallee 1 982, Vallee & Bloom 1 983) because it obviated the need for nuc1eotides required in traditional reversible assembly purification procedures. Taxol-stabilized micro tubules isolated from chick brain cytosolic extracts in the presence of AMPPNP showed a dramatically increased level of ATPase activity and were enriched in a 1 3 0 kD polypeptide (Brady 1 985). Polypeptides of similar size were identified and purified from both squid optic lobe ( 1 1 0 kD) and calf brain tissue ( 1 20 kD; Vale et al 1 98 5a). Co-purifying with the protein was a novel mechanochemical activity moni tored by a light microscopic assay. The purified protein fractions were adsorbed onto glass coverslips, and micro tubules were then applied in the presence of ATP. Microtubules could be seen to glide along the coverslips, by using video enhanced light microscopy to visualize individual micro tubule polymers (Vale et aI1 98 5a). The movement seen in the presence of the protein led to its name: kinesin (Vale et aI 1 98 5a). Later studies showed that kinesin is an ATPase, and that its activity can be potently stimulated by microtubules (Kuznetsov & Gelfand 1 986, Cohn et a1 1 987, Murofushi et a1 1 988, Wagner et al 1989). Kinesin is now known to be widely dis tributed in neuronal and nonneuronal cells (Scholey et al 1 98 5, Saxton et a1 1 988 , Neighbors et al 1 988 , Murofushi et al 1 988, Pfister et al 1989b, Hollenbeck 1 989a). To determine the direction of force production by kinesin, synthetic microspheres were coated with the protein and applied to microtubules assembled onto isolated centrosomes. The microtubules in these complexes are oriented uniformly with their + ends out (Bergen et al 1980). In the presence of ATP the beads moved away from the centro somes, thereby exhibiting motility in the direction corresponding to anterograde axonal transport in axons (Vale et aI 198 5b). Significant progress has been made toward eludicating the structure of kinesin. The original description 'of purified kinesin provided evidence that


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the protein is formed from two groups of polypeptides in the size ranges of � 1 20 kD and �65 kD (Vale et aI 1 985a). Subsequent studies ruled out the possibility that the smaller polypeptides were proteolytic fragments of the larger species, and revealed that the two sets of polypeptides co-purified through multiple fractionation steps to constant and equimolar stoichiometry (Bloom et a1 1 988, 1 989, Kuznetsov et al 1 988, Wagner et al 1 989). A native molecular mass of � 380,000 was determined for kinesin, leading to the conclusion that the protein is tetrameric, consisting of two copies each of the � 1 20 kD and �6 5 kD polypeptides (Bloom et a1 1 988, Kuznetsov et al 1 988). These have come to be known as the heavy and light chains of kinesin, respectively. Multiple electrophoretic variants exist for both types of subunits (Vale et a1 1 98 5a, Kuznetsov & Gelfand 1 986, Bloom et al 1 988, Murofushi et al 1 988, Saxton et al 1 988), and the hypothesis that these represent distinct isoforms of the kinesin heavy and light chains is supported by western blotting experiments with monoclonal antibodies to each of the two subunit classes (Pfister et al 1989b). The heavy chains have been identified as the ATP-binding subunits on the basis of ATP-photoaffinity labeling (Gilbert & Sloboda 1 986, Penningroth et al 1 987, Bloom et al 1 988), proteolytic modification (Ingold et al 1 988, Kuznetsov et al 1 989), in vitro expression studies (Yang et aI1 988), and primary structural analysis (Yang et aI 1 989). Kinesin purified from bovine brain (Hirokawa et al 1 989), adrenal medulla (Hisanaga et al 1 989), or sea urchin eggs (Scholey et al 1 989) has a highly elongated appearance in the electron microscope (l 80 nm). Located at one end of the protein are a pair of globular heads, each '" 1 0 nm in diameter. At the opposite end is a "fan-shaped" or "feathered" tail � 1 5 nm across at its widest point. Connecting the head and tail domains is a long, fibrous stalk that has a conspicuous flex point near its center. A roughly similar morphology has been reported for porcine brain kinesin, as well (Amos 1 987). Three different anti-heavy chain monoclonal antibodies decorated the head region of the molecule (Hirokawa et al 1 989, Scholey et al 1 989), whereas two anti-light chain antibodies recognized the tail region (Hirokawa et al 1 989). The kinesin heavy chain from Drosophila has been analyzed via molec ular biological methods (Yang et al 1 988, 1 989), and the information obtained has been strikingly consistent with that from electron microscopy and biochemical analysis. A full-length cDNA encoding the kinesin heavy chain was isolated, and the predicted amino acid sequence of the entire polypeptide was determined. The complete heavy chain and a series of truncated versions of the heavy chain were synthesized in vitro and tested for their ability to bind microtubules in the presence of AMPPNP or ATP. Based on these experiments, three distinct domains of the kinesin heavy �

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chain were distinguished. Beginning a t the amino terminus i s an 50 kD, apparently globular, region that contains binding sites for ATP and microtubules. At the carboxyl end of this region is an abrupt transition to a long stretch of sequence predicted to form a nearly uninterrupted coiled coil IX-helix that probably corresponds to the stalk observed by electron microscopy. The one apparent interruption of the IX-helical domain is near the middle and may represent the location of the flex point seen in the stalk by electron microscopy. A small region near the carboxy terminus is predicted to be globular, and probably contributes to the morphology of the tail end of the molecule. The full sequence for the heavy chain of squid kinesin was also reported recently, and homology with the equivalent Drosophila polypeptide was found to be extensive (Kosik et al 1 990).


�

Properties of Cytoplasmic Dynein (MAP 1 C)

Dynein had long been known to be the force-producing ATPase respon sible for sliding between the outer doublet microtubules of ciliary and flagellar axonemes (Gibbons 1 965). The identification of a cytoplasmic form of this enzyme derived from the investigation of the high-molecular weight, microtubule-associated proteins (MAPs) of brain tissue. Microtubules purified from brain by the conventional GTP-dependent reversible assembly method contained two high molecular weight proteins known as MAP 1 and MAP 2 (Murphy & Borisy 1 975, Sloboda et al 1 975). When taxol was used to purify micro tubules from brain white matter, however, three MAP 1 species were seen, termed MAP lA, IB, and 1 C (Bloom et aI1 984). Analysis of the microtubule-binding properties of these proteins revealed that MAP 1 C, like flagellar dynein and kinesin, bound to microtubules in an ATP-sensitive fashion (Paschal et al 1 987). This was the first suggestion that the protein might have mechanochemical properties. MAP l C was purified by dissociation from micro tubules with ATP followed by sucrose density gradient sedimentation (Paschal et al 1 987). Subsequent analysis revealed that, like kinesin, it had an ATPase activity that could be markedly stimulated by microtubules (Paschal et al 1 987, Shpetner et aI 1 988). Direct evidence for force production by MAP 1 C came from the use of a microtubule-gliding assay (Paschal et al 1 987). The direction of force production was determined by using in this assay axonemes isolated from Chlamydomonas reinhardtii, a unicellular, biflagellate alga. Axonemal microtubules have a uniform orientation, and the + ends of Chlamydo monas axonemes can be distinguished from the - ends by the tendency of the + ends to fray during isolation. As for. individual microtubules, the axonemes were found to glide along MAP 1 C-coated coverslips in the


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presence of ATP (Paschal & Vallee 1 987). The axonemes moved exclusively towards their + ends, demonstrating that force production was directed toward the - ends of microtubules. This direction was opposite to that seen for kinesin, and could account for retrograde transport in the axon (Paschal & Vallee 1 987). The identification of MAP 1 C as a bona fide dynein resulted from a combination of structural, biochemical, and enzymatic studies. MAP IC had a sedimentation coefficient of 20 S, comparable to ciliary and flagellar dyneins, and consisted of multiple subunits (Paschal et aI1 987). The MAP 1 C electrophoretic species, now referred to as the heavy chain of the complex (Vallee et a1 1 988), was comparable in mobility to the heavy chains of ciliary and flagellar dyneins. Like the heavy chains of ciliary and flagellar enzymes (Lee-Eiford et aI 1 986), the MAP 1 C heavy chain could be cleaved at a single site by exposure to UV light in the presence of the phosphate analogue and active site inhibitor vanadate. This served to localize the ATPase-active site to the MAP 1 C heavy chain; in addition, its size was determined as 4 1 0 kD, based on the sum of the sizes of the photocleavage fragments (Paschal et a1 1 987, Vallee et aI 1 988). Analysis of purified MAP 1 C by scanning transmission electron microscopy (STEM) revealed that its morphology and molecular mass were indistinguishable from that of two-headed forms of ciliary and flagellar dynein (Vallee et al 1 988). Together with the biochemical and pharmacological analysis of the protein (Paschal & Vallee 1 987, Shpetner et al 1 988), these data identified MAP lC as a cytoplasmic form of dynein, and it has subsequently been referred to simply as dynein, or cytoplasmic dynein (Vallee et al 1 988). What appear to be equivalent proteins have also been found in a variety of nonneuronal tissues and cells from a variety of organisms (Lye et al 1 987, Euteneuer et al 1 988, Neely & Boeckelheide 1 988, Collins & Vallee 1 989, Gilbert & Sloboda 1 989, Schnapp & Reese 1 989, Schroeder et a1 1 989, Koonce & McIntosh 1990). The mass of cytoplasmic dynein determined by STEM was 1.2 x 1 06 (Vallee et aI 1 988), and the relative content of cytoplasmic dynein subunits was determined by quantitative densitometry of SDS polyacrylamide gels of the purified complex (Paschal et al 1 987). Together, these approaches indicated that the native protein is composed of two 4 1 0 kD heavy chains, three 74 kD subunits, and one subunit each of 59, 57, 55, and 53 kD (Vallee et al 1 988). A 1 50 kD subunit was also found in liver and testis dyneins (Collins & Vallee 1 989). In calf brain tissue, this polypeptide is found only in trace amounts in dynein purified by ATP extraction of microtubules. An additional fraction of dynein can be extracted using elevated salt, however, and this fraction shows a high content of the 1 50 kD polypeptide. These data have been interpreted as indicating that the


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1 50 kD species may be a stoichiometric component of a subset of dynein molecules, perhaps defining a functionally distinct dynein species (Holz baur et al 1 989). A 1 73 kD polypeptide was also observed to co-purify with testis dynein (Neely & Boekelheide 1 988). Its relationship to the 1 50 kD species is uncertain. Like muscle myosin and kinesin, cytoplasmic dynein was observed to have two globular "heads" by both STEM (Vallee et a1 1 988) and negative stain electron microscopy (Vallee et al 1 989c). As is the case for ciliary and flagellar dyneins, the remainder of the molecule showed a variety of configurations. Typically, the dynein molecules showed two distinct stalks attached to the heads, which merged into a globular basal structure, though the degree of condensation of the stalks and basal structure varied considerably. Similar images have since been obtained for cytoplasmic dyneins isolated from other sources by using low-angle rotary shadowing or negative stain electron microscopy (Neely & Boekelheide 1 988, Amos 1 989, Gilbert & Sloboda 1989, Schnapp & Reese 1 989, Schroeder et al 1989). Subcellular Localization and Analysis of In Vivo Function

The initial description of kinesin suggested a role in organelle transport along microtubules, though direct evidence for this was lacking (Brady 1 98 5, Vale et al 1 98 5a). Potentially, transport could be accomplished by either of two basic mechanisms. Kinesin could be distributed along the outer surface of microtubules, and organelle motility would occur by passage of the organelle along a series of kinesin molecules. In this scheme, the interaction of kinesin with the microtubule is seen to be stable, while the interaction with the organelle would be transient (Figure l A). This model predicts an immunocytochemical staining pattern for kinesin com parable to the distribution of microtubules in the cell. In an alternative model (Figure IB), the organelle surface would serve as the stable attach ment site for kinesin, which would interact transiently with the microtubule wall during its force-producing cross-bridge cycle. This model predicts an immunocytochemical staining pattern reflecting the distribution of organelles as well as microtubules. The latter pattern is favored by the biochemical analysis of kinesin, which has revealed a reversible force producing interaction with micro tubules. Despite the implication that kinesin should show a more permanent interaction with the surface of organelles, however, it behaves as a soluble cytosolic protein during tissue extraction, with little apparent tendency to co-fractionate with membranes (e.g. Vale et a1 1 985a, Kuznetsov et a1 1 988, Bloom et aI 1 988). To evaluate the subcellular distribution of kinesin more directly, immu nofluorescence microscopy of cultured cells (Pfister et al 1 989b) and iso-


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Figure 1 Alternative mechanisms for organelle movement along microtubules. Kinesin molecules are depicted with permanent associations to the surface of either a microtubule (A) or an organelle (B). In (A) an organelle interacts transiently with a procession of fixed kinesin molecules. The predicted pattern of immunocytochemical staining with antikinesin antibodies would be similar to the distribution of microtubules. In (B) the organelle-bound kinesin molecules interact transiently with the microtubule, and a staining pattcrn com parable to the distribution of organelles, perhaps aligned with microtubules, is predicted. The results of immunocytochemical analysis support model B (Pfister et al 1 989b; Brady et aI 1 990).

lated squid axoplasm (Brady et al 1 990) was performed with a library of monoclonal antikinesin antibodies. Three anti-heavy chain and two anti light chain antibodies were used, and cells were fixed with formaldehyde, methanol, or ethanol/acetic acid. Staining was observed to be restricted to punctate, vesicle-like structures in the cytoplasm of primary neuronal and glial cells, several nonneural cell lines, and squid axoplasm. The immunoreactive structures frequently, though not exclusively, aligned with microtubules, which were not directly stained by the antibodies. These results thus support the model illustrated in Figure 1 B, which indicates that the more permanent attachment site for kinesin is the organelle surface, whereas the transient interaction is with the microtubule. One additional study produced similar results, and provided evidence that tubulovesicular membranes-perhaps those of the endoplasmic reticulum-also had associated kinesin immunoreactivity (Hering & Borisy 1 988). Not all kine sin antibodies have yielded these results, however, and a variety of staining


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patterns have been reported (Neighbors et al 1 988, Hollenbeck 1 989a, Murofushi et al 1 988). Additional information regarding the distribution of kinesin has been obtained by quantitative immunoblotting with a polyclonal antiserum to chick brain kinesin (Hollenbeck 1 989a). In an effort to define the dis tribution of kinesin within different subcellular pools, cultured fibroblasts were subjected to two successive detergent extractions in a microtubule stabilizing buffer. Kinesin that was released from the cells by saponin, which permeabilized the plasma membrane, was considered to represent the soluble cytoplasmic pool. Kinesin that was subsequently solubilized by Triton X-tOO was judged to be organelle-bound. Kinesin that resisted both detergents was considered to be microtubule-associated. The results of this analysis indicated that about one third of the fibroblast kinesin was associated with membranous organelles, approximately two thirds were freely soluble in the cytoplasm. and virtually none was stably bound to microtubules. How can these results be reconciled with the results of the immuno cytochemical analysis? Immunofluorescence microscopy will, in general, reveal the regions of highest antigen density. Thus, the punctuate, organelle like staining pattern observed by this method indicates that a significant fraction ofkinesin is organelle bound, but it does not preclude the existence of a soluble kinesin pool. In fact, a soluble pool might not be detected even if it were larger than the organelle-bound pool, as long as the average concentration of kinesin in the cytoplasm were significantly lower than on organelle surfaces. Nonetheless, the selective extraction experiments must also be inter preted with caution, since the normal binding equilibria between cellular proteins may be unbalanced by this procedure. The detergent extractions must be performed in large excesses of buffer relative to cell volume, which would tend, therefore, to shift the equilibrium between soluble and organelle-bound kinesin to the unbound state. Other variables, such as the composition of the extraction buffer, may also affect the kinesin-membrane interaction. Thus, the evidence supports the existence of organelle-associ ated kinesin, though the absolute fraction in the free and bound states is uncertain. How this equilibrium is regulated and whether it relates to the activity state of kinesin remain interesting questions. Additional support for a role of kinesin in anterograde organelle trans port in vivo has come from an examination of the distribution of the protein (Hirokawa et al 1 990a) after inhibiting transport. Ligated or crushed rodent peripheral nerves were examined by antikinesin immuno fluorescence and immunoelectron microscopy. Intense immunoreactivity was observed at the proximal edges of the lesions, where anterograde


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organelles had accumulated. Little staining was seen at the distal border of the lesions, where retrograde organelles had accumulated. In regions far removed from the lesions, most labeling was found on vesicle-like objects in axons. The identity of the immunoreactive structures as mem brane-bounded organelles was confinned by immunogold electron micros copy. Gold particles were preferentially found in close proximity to organ elle surfaces both on the proximal edges of the lesions and in undamaged areas of the axons. Further evidence for an involvement of kinesin in organelle movement was provided by antibody inhibition experiments (Brady et al 1 990). A monoclonal antikinesin antibody potently interfered with organelle trans port along microtubules in extruded squid axoplasm. Under optimal con ditions, the number of motile organelles was substantially diminished, and the organelles that did move travelled at velocities ",30% the nonnal rate. Inhibition was maximal in the range of 0. 1 -0.6 mg/ml purified antibody, but was less pronounced at either higher or lower antikinesin concen trations. This suggestcd that local immunoprecipitation of kinesin con tributed significantly to the observed inhibition. Curiously, the antibody exerted its effects bidirectionally. This could be due to steric interference with all intracellular movement. Alternatively, it could mean that organelles in the squid giant axon bear both anterograde and retrograde motor proteins on their surface (see Figure 2). A related study in macrophages also revealed inhibition of organelle transport by a polyclonal antikinesin heavy chain (Hollenbeck & Swanson 1 989). Tubular lysosomes in these cells extend from a perinuclear location toward the plasma membrane. Introduction of the antikinesin into macro phages by scrape loading caused a partial retraction of the Iysosomes toward the nucleus. This suggested that lysosomes were drawn outward from the nuclear area by kinesin but owed their perinuclear anchorage to some other factor. Attempts to analyze the interaction of kinesin with organelles by bio chemical means have indicated that the interaction may be complex (Schroer et al 1 988). Squid axoplasmic organelles were extracted with KI to remove extrinsic membrane proteins. Whole axonal cytosol stimulated organelle movement along microtubules, but purified kinesin was inactive. Nonetheless, immunoadsorption of kinesin from the cytosol removed '" 70% of the motility activity. These results supported a role for kinesin in organelle motility and suggested the existence of kinesin-bound factors that were necessary for full activity. The nature of these factors, however, is unknown. Antibodies that react specifically with cytoplasmic dynein have recently become available (Koonce & McIntosh 1 990, Pfarr et al 1 990, Hirokawa et al 1 990b, Steuer et al 1 990), making it possible to begin exploring

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A


Cell Blldy

+---

DVNEIN

+ KINESIN �

*¥**dl'

Axon Terminus

*§

eM

#0

B + KINESIN

DVNEIN Figure 2

Alternative models for the role of kinesin and cytoplasmic dynein in bidirectional axonal transport. (A) Organelles are depicted using either kinesin for anterograde transport or cytoplasmic dynein for retrograde transport. (B) An organelle is depicted with both kinesin and cytoplasmic dynein on its surface; only kinesin is active, allowing for the transport of dynein out to the axon terminus. Molecules are drawn roughly to scale. A substantial portion of the kinesin and dynein molecules must be obscured in electron micrographs of axoplasm, since the two motor proteins are much larger than the bridges observed between organelles and microtubules in the celL The extreme length of the kinesin tail suggests a possible role in ordering kinesin molecules on the organelle surface (Vallee & Shpetner 1990). Adapted from Vallee et al (1989a).

its intracellular distribution. Immunofluorescence microscopy of ligated mouse peripheral nerves revealed punctate organelle-like staining in undis turbed regions (Hirokawa et aI1990b), much like that seen with antikinesin (Pfister et a1 1 989b, Brady et a1 1 990, Hirokawa et aI 1 990a). This suggested that dynein, like kinesin, has a primary association with organelles, rather than microtubules. Near the lesions, however, a marked difference in the


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distribution of dynein and kinesin was noted. Dynein accumulated on both the distal and proximal sides. The distal localization was clearly consistent with a role for cytoplasmic dynein in retrograde organelle transport. A possible explanation for the proximal localization was the presence of dynein on anterograde as well as retrograde organelles. This explanation makes good sense considering a basic problem of retrograde transport: How does the retrograde motor reach the axon terminus to serve in retrograde transport? One possibility is that it "hitches a ride" with an terograde organelles (Figure 2; Vallee et al 1 989a). Presumably it must be inactive in this state, or it would interfere with anterograde transport. Further evidence for the in vivo role of axonal dynein was provided by using a reconstituted system for axoplasmic organelle motility along micro tubules (Schnapp & Reese 1 989). Centrosome-nucleated micro tubules were incubated in the presence of ATP with cytosol and organelles isolated from squid axoplasm . Bidirectional organelle transport was observed along the micro tubules. UV illumination of the cytosol in the presence of vanadate and ATP preferentially inhibited retrograde trans port, presumably by photocleavage and concomitant inactivation of endogenous dynein (Paschal et al 1 987, Lye et al 1 987). Similar results were obtained with lysed mammalian fibroblasts (Schroer et al 1 989). Again, UV irradiation in the presence of vanadate and ATP resulted in the preferential inhibition of retrograde movements. In this study, addition of purified brain dynein was found to restore approximatcly one third of the retrograde organelle motility. The enzymatic, force-producing and pharmacological properties of kinesin and cytoplasmic dynein have been examined in considerable detail. This has provided a basis for comparison with the physiological properties of fast axonal transport. Quantitation of organelle movement in vivo is not standardized, however, thereby making comparisons difficult. Of additional importance, anterograde movements of small vesicles, such as synaptic vesicles, are extremely difficult to see and quantitate. Thus, reported values for anterograde transport often reflect values obtained only for larger organelles. The Mg2+ salts of nucleoside triphosphates have been tested for their ability to be hydrolyzed in vitro by kinesin or cytoplasmic dynein, to promote microtubule gliding by the purified enzymes, and to support organelle transport in the axon. ATP is the preferred substrate for both kinesin (Kuznetsov & Gelfand 1 98 6) and cytoplasmic dynein (Shpetner et al 1 988, Collins & Vallee 1989). The ATPase activities of kinesin (Kuz netsov & Gelfand 1 98 6, Cohn et a1 1 987, Murofushi et a1 1 988, Wagner et a1 1 989) and cytoplasmic dynein (Paschal et al 1987, Shpetner et a1 1 988, Collins & Vallee 1 989) are stimulated several-fold by microtubules, a


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reflection of a mechanochemical cross-bridge cycle in which ATP hydroly sis is coupled to force production. Both kinesin (Kuznetsov & Gelfand 1 986) and cytoplasmic dynein (Shpetner et al 1 988) are capable of hydro lyzing every other conventional nucleoside triphosphate tested, but not all of these nucleotides support motility. ATP, GTP, and ITP all support kinesin-mediated microtubule gliding or bead movement (Porter et al 1987); however, only ATP promotes dyncin-mediated microtubule gliding (Paschal & Vallee 1987). Curiously, the nucleotide specificities for anterograde and retrograde organelle transport did not reflect this difference. UTP, GTP, CTP, and ITP were all found to support organelle motility equally well in both directions in isolated squid axoplasm, although the velocities of transport were only about one third of those observed with ATP, and high levels of nucleotides were required (Leopold et al 1 990). These results appear generally consistent with a role for kinesin in transport but seem to con tradict a role for dynein. Persistent retrograde transport could have been due to low levels of A TP in the other nucleotide preparations or to gen eration of ATP from the other nucleotides as the result of nucleoside diphosphokinase activity. This is unlikely, however, as hexokinase and 2deoxyglucose were included to eliminate possible trace levels of ATP. To prevent further ATP production, moreover, dinitrophenol and diadenosine pentaphosphate were included in the perfusion buffers as inhibitors of oxidative phosphorylation and adenylate kinase, respectively. An alter native explanation for the observed retrograde motility with nucleotides other than ATP is that they can, in fact, support a low level of force production by cytoplasmic dynein, which the microtubule gliding assay is not sufficiently sensitive to detect (Paschal & Vallee 1 987). Rates obtained from in vitro motility assays are in the range seen for organelle movement in axoplasm and for rapid axonal transport in vivo (Grafstein & Forman 1980). In the case of brain cytoplasmic dynein, the rate of microtubule gliding (1.25 ttm/sec; Paschal et al 1 987) was closely consistent with that for retrograde organelle transport in squid axoplasm (Brady et al 1982, 1 985), '" 1 . 3 m/sec. Kinesin-mediated motility in vitro (",0.5 ttm/sec; Vale et a1 1 985a, Porter et a1 1987) is somewhat slower than the rate of anterograde organelle motility ( 2 m/sec; Brady et al 1 982, 1 985). The effect of pharmacological agents has been generally consistent with a role for kinesin and dynein in axonal transport. Vanadate is known as a potent inhibitor of ciliary and flagellar dynein, acting in the micromolar concentration range (Gibbons et al 1 978, Kobayashi et al 1 978). It is somewhat less effective with cytoplasmic dynein, showing a 1 1 pm/sec at room temperature (Shpetner & Vallee 1 989), and microtubule sliding and bead translocation rates for kinesin and cytoplasmic dynein were in the same range (see above). Thus, for these proteins to be involved in slow transport, modulation of the rate of microtubule gliding must be postulated, perhaps by a mechanism as simple as viscous drag through the dense neuronal "cytomatrix." It may be remembered that rates of SCAP movement in contracting microtubule gels were reported as 1 .4 ,urn/min, much closer to the range of 0. 1 5- 1 . 5 ,urn/min for SCa (Weisenberg et al 1 986; and see above). This could reflect the presence of a novel slow motor. However, the slow rates observed for this movement could also result


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from the activity of one of the known motor proteins subjected to the physical constraint of the microtubule gel. The highly elongated neuronal MAPs, which occupy the space between microtubules in the axon (e.g. Hirokawa et a1 1 985) are obvious candidates for modulating intraaxonal motility. In fact, MAP 2 and tau have been found to inhibit microtubule gliding and the microtubule-activated ATPase activity of cytoplasmic dynein (Paschal et aI 1 989). Similar analysis of the effect of MAPs on kinesin has not so far indicated an effect on the mechanochemical activities of this protein (Rodionov et aI 1 990), though steric hindrance of kinesin-mediated microtubule gliding was observed when the MAPs were allowed to absorb to the coverslip (von Massow et al 1 989). Analysis of the effect of MAPs on dynamin has not been per formed, but may be of interest in further understanding its role in axo nal transport.

CONCLUSION The advances in our understanding of axonal transport over the past decade have been dramatic. The role of microtubules in fast transport has become clear, and the discovery of kinesin and cytoplasmic dynein has provided a likely mechanistic basis for microtubule-associated movement. How close are we, in fact, to a complete understanding of rapid transport? The existing pharmacological evidence is consistent with a role for dynein and kinesin in rapid transport. In the case of dynein, the effects of both EHNA and of vanadate-mediated photocleavage have pointed to a specific role in retrograde transport. In the case ofkinesin, the sensitivity of anterograde transport to vanadate and to NEM are somewhat surprising. These results may, in fact, undetscore the lack of specificity of these agents and the difficulty in quantifying anterograde transport. A particularly glaring inconsistency in models involving kinesin and dynein in transport has been the limited biochemical evidence for mem brane association. Recent immunocytochemical evidence has revealed a punctate organelle-like distribution for some fraction of both proteins in the cell. Ultimately, therefore, the difficulty in demonstrating a membrane association in vitro may prove to be more an artifact of biochemical manipulation than a reflection of the normal physiological behavior of these proteins. Thus, although more evidence is certainly needed to establish con clusively a role for kinesin and dynein in rapid transport, such a role is appearing more and more likely. What remains an important question is the regulation of transport. How is dynein transported to the axon ter minus; what is the basis for bidirectional and saltatory movement of individual organelles; and, perhaps of greatest interest, what signals. within


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the organelle membrane are responsible for sorting between anterograde and retrograde pathways? In the case of slow axonal transport, many fascinating experimental observations have contributed to the base of knowledge about this phenomenon, and candidate molecules for microtubule-based transport have been identified. Considerable controversy remains over the nature of this phenomenon, however, especially regarding the identity of the trans port species. This issue may be resolved by the use of new photoprobes to monitor transport. In addition, as in the case of rapid transport, further analysis of the motor proteins should provide important new insight into the mechanism of this phenomenon. NOTE ADDED IN PROOF Recently, a cDNA encoding the 1 00 kD dynamin polypeptide was clone and sequence, which revealed striking homology with a new family of GTP-binding proteins (Obar et aI 1 990). Biochemical evidence has now suggested that GTP, rather than ATP, may be the physiological substrate for dynamin (Shpetner & Vallee 1 990). ACKNOWLEDGMENT

We thank Dr. Scott Brady for critical reading of the manuscript, Wendy Hiller for the preparation of Figure 1 , and Drs. C. Keith, R. Lasek, R. Nixon, P. Hollenbeck, R. Miller, L. Gambetti, and 1. McQuarrie for their helpful comments. Supported by NIH Grant GM2670 1 to R.B.V., and NIH Grant NS23868 and Welch Foundation Grant 1- 1 077 to G.S.B. Literature Cited

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A coupled model of fast axonal transport of organelles and slow axonal transport of tau protein.

Slow axonal transport.

Dynamics of fast axonal transport.

Fast axonal transport in motor nerve regeneration.

Axonal transport: components, mechanisms, and specificity.

Slow axonal transport mechanisms move neurofilaments relentlessly in mouse optic axons.

Assessment of slow axonal transport in lead-exposed rats.

Axotomy accelerates slow component b of axonal transport.

Actin depolymerizing factor is a component of slow axonal transport.

Slow transport of the cytoskeleton after axonal injury.

Local modulation of neurofilament phosphorylation, axonal caliber, and slow axonal transport by myelinating Schwann cells.

Automatic quantification of fast axonal transport in neuronal cell cultures.

Continuation of fast axonal transport in regenerating axons in vitro.

Axonal transport: cargo-specific mechanisms of motility and regulation.

A new method to study fast axonal transport in vivo.

Does nerve impulse activity modulate fast axonal transport?

What tau distribution maximizes fast axonal transport toward the axonal synapse?

Organization and slow axonal transport of cytoskeletal proteins under normal and regenerating conditions.

Slow axonal transport of soluble proteins and calpain in retinal ganglion cells of aged rabbits.

The smooth endoplasmic reticulum: structure and role in the renewal of axonal membrane and synaptic vesicles by fast axonal transport.

Slow axonal transport or proteins; blockade by interruption of contact between cell body and axon.

Axonal transport.

Fast axonal transport of acetylcholinesterase and protein in cold-blocked frog sciatic nerves in vitro.

The direct effects of graded axonal compression on axoplasm and fast axoplasmic transport.