The Roles of Actin in Tip Growth of Fungi

INTERNATIONALREVIEW OFCYTOLOGY, VOL. 123

The Roles of Actin in Tip Growth of Fungi I. B. HEATH Department of Biology, York University, North York, Ontario, Canada M3J l P 3

I. Introduction The process of tip growth is the hallmark of the fungal kingdom. Certainly the process occurs in other walled cells, such as pollen tubes and root hairs, but in the other kingdoms it is restricted to a few specialized cell types, whereas among the fungi it is the mode of growth that generates the dominant vegetative structure, the hypha. It is shown in all hypha-producing species and among the yeasts, the fission yeasts show a process that seems to differ only in being of determinate duration. Even the process of budding, which involves localized cell wall synthesis and expansion, can be viewed as a form of tip growth constrained to cease when a sphere, as opposed to a tube, is formed. A similar argument applies to blastospore production (Kendrick, 1985) such that many types of fungal spores can also be viewed as resulting from constrained short-term tip growth. It may be argued that the diversity of form of budding and blastosporogenesis makes it impossible to define tip growth, if these are indeed included as examples of the process. However, the essential features of hyphal tip growth are localized synthesis and extensibility of the cell wall at the extreme tip and a suitable gradient of rigidification of the wall to produce the characteristic tube known as a hypha. It seems likely that only very minor perturbations of these features will generate a bud or a spore. Indeed the morphology of many hyphal tips perturbed in various ways is not that dissimilar from, albeit often simpler than, the shape of buds and spores. Consequently it seems that tip growth may well explain not only the formation of hyphae but also buds and spores. Clearly it is a vital feature of fungal biology, yet it is one that remains shrouded in controversy and obscurity after more than a century of research. Until the last decade it is not unfair to say that most concepts of tip growth centered almost exclusively on the properties of the cell wall. However, in 1982 Picton and Steer introduced the important notion that actin may also play an important role in tip growth. Since then substantial data have accumulated to support aspects of their concept. It is these data that will be the focus of this chapter. I shall make occasional reference to supporting data or ideas derived from nonfungal tip-growing cells but concentrate mostly on hyphal tips and budding of yeast cells. While blastospore production may well be a form of modified tip growth as just argued, we currently have no data to support a role for actin in the process; consequently it will not be considered further. I shall first discuss the 95

Copyright Q 1990 by Academic Press,Inc. All rights of reproduction in any form reserved.

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multiple features of tip growth, then examine the organization of actin in the growth zones. Next will follow a synthesis of these data to provide models for the possible roles of actin in the diverse processes. Concluding the article is an analysis of possible regulatory mechanisms. For more extensive and complementary analyses of various aspects of tip growth covering all tip-growing cell types, the book edited by Heath (1990) should be consulted. For convenience, I shall adopt the use of generic names only, when referring to organisms that have been studied because there are no examples where species-specific results are known. 11. Characteristicsof Tip Growth

The fundamental basis of tip growth is that it results from the balance between highly localized cell extensibility and osmotically generated turgor pressure. We shall see that extensibility is likely to be regulated by many factors, and equally, turgor pressure is the result of multiple inputs such as membrane-located transport channels, membrane and cell wall permeability, and intracellular and extracellular solute concentrations-all of which are in turn regulated by many systems. This multiplicity of contributory factors generates three important considerations. First, to develop a consistently shaped hypha and other structures, there must be remarkably complex interrelated control systems. Second, because the critical processes are so localized, it is almost impossible to analyze them with bulk biochemical techniques; cytological analysis with other manipulations is essential to fully explain the process. Finally, the system is likely to be very labile and easily disrupted by investigative techniques. The latter point influences many of the further aspects of this work and needs further expansion. A. LABILITY OF TIPGROWTH

The ease with which tip growth can be disrupted is well known to all who have observed living hyphae. Slight perturbations will cause hyphae to stop growing and induce changes in internal organization within seconds. For example, Robertson (1958) showed that osmotic insult could induce substantial changes within 10 seconds and that cessation of growth, tip swelling, and resumption of normal growth could all occur within 40 seconds. Such lability is not surprising when one considers that, with hyphal growth rates reaching 50 pm/minute (Griffin, 1981), 10 seconds are equivalent to =8 pm of normal growth! The important message from this lability is that great care must be exercised to ensure that tips that are examined by whatever means are in fact grow-

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ing, rather than showing organization characteristic of a recently stopped tip. This point was extensively considered by Bracker (1971) in the context of vesicle complement in hyphal tips, but the message is frequently ignored in many current studies. Clearly treatments that involve buffer washes, centrifugations, temperature or osmotic changes, detergent treatments and so on, on live cells are almost certain to cause growth inhibition that may, or may not, be accompanied by major changes in the organization and activities of tip growth-related structures and processes, including actin arrays.

B. REGULATED CELLWALLAND MEMBRANE EXPANSION Irrespective of mechanisms, the gradient of cell surface extensibility at the hyphal tip must be under very fine control in order to generate the typical, highly reproducible form of a hypha or spore. Aspects of this control must include spatial information to generate the correct shape, quantitative control to produce the right growth rate, and quantitative control to form the correct diameter of the cell. These controls are influenced by both genetic and environmental factors because under constant environment each species generates characteristic shapes and sizes, whereas a single species will vary these parameters in different environments. Furthermore, cell extensibility is fully reversible, again in a precisely controlled manner, because subapical inextensible hyphae can produce a regulated array of extensible new tips, that is, branches. Most models of tip growth have focused on the regulation of cell wall properties (Wessels, 1986, 1988, 1990; Bartnicki-Garcia, 1973, 1990; Bartnicki-Garcia and Lippman, 1972), but neither these nor models involving actin (Picton and Steer, 1982) have yet provided a fully adequate explanation of all the needed controls. Furthermore, since cell wall synthesis in the hyphal tip involves exocytosis of vesicles (which will be termed wall vesicles in this review) that contribute both plasmalemma and cell wall polymers [as well as enzymes (Gooday and Gow, 1990), which are irrelevant to the current context], it is evident that the controls must also produce the correct balance between membrane and cell wall synthesis because normally the plasmalemma is tightly and uniformly appressed to the cell wall (Heath et al., 1985; Grove and Bracker, 1970; Roberson and Fuller, 1988; Howard and Aist, 1979). The ease with which excess plasmalemma can be produced (seen as plasmalemmasomes) and incorporated into the cell wall (seen as lomasomes) (Heath and Greenwood, 1970) shows that independence of production of plasmalemma and cell wall can occur, thus reinforcing the concept of independence of control in normal tips. Because there is currently no evidence for endocytosis of excess plasmalemma in hyphal tips (e.g., via clathrincoated vesicles), it seems that control is exercised at the level of production of wall and membrane.

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C. LOCALIZATION OF PLASMALEMMA-ASSOCIATED TRANSPORT PROTEINS In all fungi examined, tip growth is accompanied by the generation of electrical currents. The direction of these currents and possibly the ions involved in their generation vary in different species, and it is likely that they are not causally related to the tip growth process itself (Gow, 1989; Harold et al., 1985; Harold and Caldwell, 1990). However, their generation strongly implies the differential distribution of membrane transport proteins (Schreurs and Harold, 1988). There is no direct evidence for such distributions in hyphae, but nonrandom distribution of intramembranous particles in other tip-growing cells is seen (Volkmann, 1984). A mechanism for setting up and maintaining the likely nonrandom distribution of transport proteins in hyphal tips is a feature that should be included in any model for tip growth.

D. VESICLE TRANSPORT A universal feature of hyphal tips is the production of wall vesicles by subapical Golgi bodies (or their equivalents) and the transport of these vesicles to a high concentration at the tip where they are exocytosed (Grove, 1978; Gooday, 1983). A mechanism for transporting these vesicles is an essential feature of tip growth but it is not a simple point-to-point mechanism. For example, the producing Golgi bodies extend over a considerable distance (Heath and Kaminskyj, 1989), and exocytosis as indicated by incorporation of cell wall polymers can also extend for tens to hundreds of micrometers subapically (Gooday, 1971; Fevre and Rougier, 1982; Barmicki-Garcia and Lippman, 1969). Furthermore, subapical branch initiation entails the initial accumulation of wall vesicles in a new area and the subsequent bidirectional transport of vesicles to two potentially competing tips (the original tip and the branch tip). There is no evidence to show whether the branch-forming vesicles are derived from the same population of Golgi bodies as those producing the main tip vesicles or a second population. However, either a single “source” exporting bidirectionally or two “sources” exporting unidirectionally clearly need complex vesicle transport systems, which must be a major feature of tip growth.

E. EXOCYTOSIS Exocytosis is the ultimate fate of the wall vesicles, and with fusion rates calculated to range from 1340 to 33,300 vesicles per minute (Grove et al., 1970), it is clearly a major and highly dynamic aspect of tip growth. Two features of the process deserve special emphasis: regulation of the site of exocytosis and regulation of precocious intervesicle fusions. As mentioned previously (Section 11, D), some wall vesicles appear to fuse with the subapical plasmalemma whereas

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most are transported past this region to fuse at the tip. A full description of tip growth must explain how the site of fusion with the plasmalemma is regulated, including the precise localization of a high rate of fusion in a formerly low fusion zone during branch production. Perhaps even more remarkable is the control, which ensures that during transit, wall vesicles do not appear to show major levels of intervesicle fusion, yet at the site of exocytosis the wall vesicle membrane becomes part of the plasmalemma and thus fusion-competent with the same incoming wall vesicles with which it was fusion-incompetent during the immediately preceding transport period.

F. ORGANELLE MOVEMENTS AND POSITIONING Two types of evidence show that hyphae contain one or more systems for moving and positioning all organelles (McKerracher and Heath, 1987). Direct observations of living cells show that structures such as wall vesicles, mitochondria, nuclei, vacuoles, and assorted unidentified vesicles move independently of one another and of the advancing tip. These movements may be rapid, erratic, discontinuous, and bidirectional-in which case they can be described as saltatory-or they may be slower and more even in pace and direction. The latter movements are characteristic of nuclei and mitochondria. The second type of evidence comes from diverse observations of nonrandom organelle distributions in hyphae. These range from well-established longitudinal gradients of organelles (Zalokar, 1959; Girbardt, 1969; Grove and Bracker, 1970; Heath and Kaminskyj, 1989) to radial patterns (Heath and Kaminskyj, 1989) and include the phenomenon of nuclei being maintained in some consistent position relative to the growing tip (Robinow, 1963; McKerracher and Heath, 1985; Herr and Heath, 1982; Heath, 1982). Similarly vacuolation is controlled to well-defined regions of hyphae. It is unclear whether these diverse phenomena can all be explained in the basis of differential regulation of a single system or if multiple systems operate. What is clear is that hyphae contain both mechanochemical force-generating systems that can act on all, or almost all, intracellular structures, and spatial monitoring systems that can be used to regulate the force generators to ensure the correct positions of the organelles.

G. CYTOPLASMIC MIGRATION The dominant, probably universal, pattern of hyphal growth involves the bulk of the cytoplasm and its contained organelles moving forward with the advancing tip and leaving behind a subapical region containing mostly a large central vacuole (actually typically many closely appressed vacuoles) surrounded by a very thin layer of cytoplasm and a few organelles. This behavior is perhaps most

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elegantly seen in Basidiobolus (Robinow, 1963) but occurs in most species. It led Reinhardt (1 892) to introduce the concept that fungi are tube-dwelling amebas whose cytoplasm migrates through the tube it synthesizes, a concept later reconsidered by Isaac (1964). This concept of migration is complex because the bulk of the cytoplasm does not move relative to the hyphal tip but it does move relative to the lateral walls. Consequently the plasmalemma and that peripheral layer of cytoplasm that comes to line the subapical regions probably do not move relative to the cell wall: certainly they do not need to. However, the bulk of the internal cytoplasm and organelles does move and therefore there is most likely some form of shear zone in the peripheral cytoplasm. The generation of this cytoplasmic migration evidently requires a force-generating system, probably located at least in part in the peripheral cytoplasm, and is clearly an important part of the tip growth system. We have seen that tip growth involves many separable components that must be closely coordinated in order to generate a hypha. All of these components function in close proximity and involve one or more mechanochemical forcegenerating or resisting systems. At present in fungi we only have evidence for the existence of two cytoskeletal elements that could form the base of mechanochemical force-handling systems, microtubules (MT) and actin filaments. It is the organization and possible activities of the latter in the aforementioned processes that will form the focus for the rest of this chapter. However, in a number of contexts it seems likely that actin and MT interact with each other; consequently both systems will be discussed to some extent.

In. Organizationof Actin A. GENERAL FEATURES OF ACTIN Actin is one of the most ubiquitous and widely studied proteins found in living cells; consequently there is an extensive body of information on its properties, most of which is beyond the scope of this review. Excellent extensive reviews can be found in Korn (1982). Taylor and Condeelis (1979), Staiger and Schliwa (1987), and Pollard and Cooper (1986). However, some important general features, which can be explored further by reference to the reviews just listed, need to be summarized as follows. 1. Actin occurs as a globular 45-kDa subunit (G actin) that reversibly polymerizes into a 7-nm-thick long filament (F actin) composed of two helically entwined linear chains. The degree of polymerization and stability of the polymer are regulated by many factors including ionic composition of the medium and

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diverse actin-binding proteins (ABP). While actin filaments can exceed several micrometers in length, much shorter oligomers can occur and are undoubtedly functionally important. 2. F actin is polarized, both with respect to its preferred direction of polymerization and its interaction with proteins such as myosin. The polarity is visualized by binding proteolytic subfragments of myosin-ither heavy meromyosin (HMM) or the S 1 subfragment of HMM-to the filaments, in which case an arrowhead pattern is seen. Polymerization occurs preferentially on the barbed end and myosin moves toward the same end. 3. F actin can occur as separate filaments, as variously large parallel bundles, and as complex irregular gels that can undergo reversible ATP-dependent contraction. Actin-binding proteins play an important role in regulating and maintaining the formation of these diverse arrays. 4. While it is common to talk of actin as a single protein, many organisms contain more than one actin gene and produce diverse isoforms that are biochemically distinct. These isoforms can show temporal, intracellular, cellular, and tissue specificity in their distribution. 5. Both F and G actin can bind directly, or indirectly via ABP, to diverse cellular structures such as other elements of the cytoskeleton, most cellular membranes, liposomes, chromatin, ribosomes, and even specific enzymes such as those of the glycolytic pathway. By synthesis from the foregoing characteristics, and by direct observation (Heuser and Kirschner, 1980), one can deduce a plausible general image of the disposition of actin in a cell. The entire cytoplasm would be permeated by a variable concentration of a diffuse three-dimensional network of F actin that is reinforced by bundles of actin and connected to the plasmalemma and probably other cellular constituents. It can perform work by contraction of the network by mutual sliding of filaments or by forming a substrate for sliding interactions with other molecules such as myosin. This assemblage and its properties would be regulated by ABP, diverse ions and nucleotide triphosphates such as ATP and GTP. It is our progress toward verifying and refining our knowledge of this hypothetical assemblage in the apex of tip-growing hyphae that we shall now consider. However, an important aspect of this consideration, one too frequently ignored, is some analysis of the technical merits of the data base. B. METHODS AND TECHNICAL LIMITATIONS There are basically five complementary techniques, data from all of which need to be integrated in order to provide a full understanding of tip growth. Each has its advantages and limitations as follows.

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Biochemical methods can show the presence and diversity of actin and its isoforms, identify factors influencing its properties (Nanomura et al., 1975), and find functionally important ABP (Liu and Bretscher, 1989; Drubin et al., 1988). What they cannot do directly is identify tip growth-specific features or provide spatial information because of the very local scale of the process. However, they provide a very important adjunct to cytological techniques because once ABP or regulatory ions such as Ca2+ are identified, it is possible to generate antibodies for immunocytochemistry (Drubin el al., 1988) or select relevant probes for cytochemistry (Jackson and Heath, 1989). Genetic approaches can identify the processes that depend on actin (Novick and Botstein, 1985) and also describe genes whose products interact with actin, such as ABP (Adams and Botstein, 1989; Adams et al., 1989). The limitation of the genetic approach is differentiating between primary versus indirect secondary roles. For example, the demonstration of secretory vesicle accumulation in actin-defective mutants of Saccharomyces (Novick and Botstein, 1985) could indicate a role for actin in vesicle transport, exocytosis, vesicle synthesis, cell expansion (assuming a feedback between cell expansion and exocytosis), or even regulation of cellular nucleotide triphosphate or ion levels. Furthermore, understanding the role of the changed gene demands careful phenotype analysis (Oakley, 1985), a process that is often difficult to carry out with the same vigor that was applied to the initial genetic analysis. Inhibitor studies are potentially very helpful because they can be applied reversibly and their effects monitored either biochemically or cytologically, but in general they suffer a similar problem to that found with genetic analysis-that of differentiating between primary versus secondary effects. Furthermore, with respect to actin there is a shortage of good probes. The cytochalasins are known to have diverse nonactin targets (Seagull and Thomas, 1976; Poste, 1973; Treves ef al., 1987), and there is reason to believe that stable, functional actin arrays can resist cytochalasin disruption (Forer et al., 1972, and references therein; Seagull and Heath, 1980). The phallotoxins (Faulstich et al., 1988) seem to be very specific for F actin, but again there is evidence that stable, functional actin arrays can exist in the presence of phallotoxins (Faulstich et al., 1988; Jackson and Heath, 1990a) and that some intracellular actin filaments are, for unknown reasons, inaccessible to the phallotoxins (Wilson er al., 1987; Tang et al., 1989). Microinjection of antibodies (Mabuchi and Okuno, 1977) or chemically modified cytoskeletal proteins such as N-ethyl malemide-treated HMM (Meeusen and Cande, 1979) are potentially very useful specific inhibitors (but see Forer, 1985, for a critique of their use), but their applications to fungal cells have not been explored and again there is the problem of primary versus secondary effects. Light microscopy (LM) offers the benefits of direct spatial information at a resolution capable of examining tip-specific processes in living cells but suffers

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a number of limitations. Actin arrays, at least in fungal cells, are only detectable with the aid of fluorescent labels. In fungi, only fluorochrome-labeled phallotoxins and fluorescent antibody-based techniques have been used, although in principle microinjected fluorescently labeled actin or myosin (Sanger et af., 1989) or similarly labeled ABP can also be used. These techniques suffer three major limitations: resolution, sensitivity, and artifactual morphological changes. Resolution is light-limited to ~ 0 . pm, 2 which is good relative to the dimensions of the tip growth zone but poor relative to high-density populations of 0.01-pmdiameter wall vesicles and 0.007-pm-diameter actin filaments. Sensitivity is more difficult to analyze because the minimum number of actin filaments detectable in a cellular environment is unknown. Furthermore, while a clearly organized actin pattern is easily described and analyzed, a diffuse actin network permeating the cytoplasm would give an image that might well be disregarded as “background,” yet such a network would clearly be functionally very important. Consequently analyses based on detected actin patterns may deal with only part of the story, and that part is subjected to the problems of artifact, which are not easily handled in a rigorous manner. Studies based on fixed or permeabilized cells are clearly prone to artifactual changes in the labile actin arrays during prefixation handling, and actin is known to be very sensitive to different fixative buffers (Heath, 1987) and mild detergents (Heath, 1988). However, the demonstration of rhodamine-phalloidin-stained actin in growing, living hyphae (Jackson and Heath, 1990a) shows a way around fixation artifacts. A further, often neglected aspect of the artifact problem is intercellular variability. A perusal of most reports shows considerable variability, but the staining patterns observed are seldom describable in quantitative terms; consequently it is difficult to deal rigorously with intercellular variability. Finally, there is the question of specificity. While there is no evidence for false positive results with current labeling techniques, there is the evidence referred to above that at least rhodamine-labeled phalloidin may not stain all cellular actin. Electron microscopy (EM) is arguably the ultimate foreseeable technique for analysis of the roles of actin in tip growth because it alone has the resolution needed to describe the architecture of all relevant molecules and organelles. However, this promise is presently far from realized because even the best preparation techniques, such as transmission EM of thin-sectioned, freeze-substituted cells fail to reveal fully the actin arrays known to be present by LM (Heath and Kaminskyj, 1989), and other promising techniques (Clarke et al., 1975; Heuser and Kirschner, 1980) involve treatments likely to cause significant losses or rearrangements. From the foregoing comments, it is clear that analysis of the roles of actin in tip growth suffers from many technical limitations, many of which are often not fully acknowledged. Bearing these points in mind, I shall now discuss the currently available data on the organization of actin in hyphal tips.

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c. DIVERSITY AND hlWG"TETAT10N OF PATERNS The unifying feature in all reports of actin in hyphal tips or buds is that it is most abundant in, but not exclusive to, the growing tip or bud. However, in this region there are two dominant patterns of organization. The most common pattern is an aggregation of peripheral spots or plaques. These plaques are most concentrated, and probably exclusively located, on or very close to the plasmalemma. Comparison of face views and optical sections reveals that they do appear to be discoidal (although at this level of resolution the difference is not always obvious), with diameters typically in the 0.3- to 0.5-pm range. This pattern is seen in basidiomycetes such as Schizophyllum (Runeberg et al., 1986), Amanita, Heterobasidion,Paxillus, and Suillus (Salo er al., 1989), and Uromyces (Hoch and Staples, 1983a; Fig. l), ascomycetes such as Saccharomyces (Adams and Pringle, 1984; Kilmartin and Adams, 1984; Novick and Botstein, 1985; Hasek er al., 1987), Schizosacchuromyces (Marks and Hyams, 1985; Marks et af., 1986; Kanbe er af., 1989), Candida (Anderson and Soll, 1986), and zygomycetes such as Neozygites (Butt and Heath, 1988; Fig. 2), Conidiobolus (Fig. 3), and Entomophaga (Fig. 3 . I In contrast to the plaques, oomycetes such as Saprolegnia (Heath, 1987, 1988; Jackson and Heath, 1989, 1990a,b, Figs. 4,8, 9). Pythium (Fig. 7). and Achlya (Fig. 6) show a finely fibrillar apical cap, which is also intimately associated with the plasmalemma. These caps enclose the entire apex and appear to contain most actin at the extreme tip. Subapically they gradually give way to a peripheral array of coarser filaments interspersed with plaques similar to those found at the tips of other fungi (Heath, 1987, 1988; Jackson and Heath, 1989, Figs. 4, 10-12). A third reported pattern of apical actin is seen in the ascomycete Gyromitra. where a diffuse fluorescence permeating the entire tip cytoplasm is seen (Salo et a/., 1989).Plaques and filaments are apparently absent in Gyromitra.While not strictly relevant to the tip growth question, it is notable that the dichotomy between the apical caps of the oomycetes and the plaques of most other fungi is also found in the subapical arrays where the strictly peripheral fibrils and plaques of the oomycetes compare with fewer coarser predominantly central fibrils and peripheral plaques in the other fungi. These dichotomies may relate to the apparent fact that the oomycetes are distantly related to the other fungi (Gunderson et al., 1987). However, the important point is that their tip growth process does not appear to differ significantly from that of the other fungi; consequently one must conclude that tip growth can occur in the presence of both apical caps and plaques. In order to understand how the apical actin arrays are involved in tip growth, it is necessary to understand how they relate to the other structures characteristic of the growing tip. Because actin has not been well preserved in ultrastructural studies 'All figures are of formaldehyde-fixed and rhodamine-phalloidin-stainedcells as per Heath (1987). unless specified to the contrary.

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(Section IIIB) a direct comparison between the LM localization of actin and the EM-based description of apical organelles and structures is not possible. However, there are two current views on the likely interpretation of the actin plaques. Hoch and Staples (1983a) originally noted that they resembled focal contacts or adhesion plaques characteristicof many animal cells (Burridge et ul., 1987).However, partly because the plaque lacked attached actin fibrils extending into the cytoplasm, a characteristic of focal contacts, they favored the alternative interpretation that they corresponded to small vesicular structures termed filasomes (Howard, 1981). Filasomes do have a fibrillar coat in some ultrastructural preparations (Howard, 1981), and Hoch and Staples (1983a) noted that their general distribution in thin sections of Uromyces germ tubes (Hoch and Staples, 1983b)correlated with that of the actin plaques. The function of filasomes is unknown. In contrast, Adams and Pringle (1984) did find actin filaments attached to some plaques in Saccharomyces and favored the analogy with focal contacts. The recent observations of Kanbe et a1 (1989), showing microfilament-associated granules attached to the plasmalemma in the plaque-bearing region of Schizosaccharomyces may also be evidence in support of the equivalence of plaques and focal contacts. Actin is apparently associated with the plasmalemma in a number of fungi (Howard, 1981; Allen et ul., 1974), and Heath (1987) has argued that inducible plaques at the tip of Saprolegniu hyphae are associated with the plasmalemma. Equally, the absence of filasomes (Heath et al., 1985; Heath and Kaminskyj, 1989) and actin plaques in normal hyphal tips of Suprolegnia is a correlation consistent with the argument that in other fungi filasomes and apical actin plaques may be the same structures. However, the morphologically similar subapical plaques in Saprolegnia (Heath, 1987, 1988) are evidently not filasomes. At present the detailed quantitative work or correlative LM and EM needed to prove that actin plaques are coincident with filasomes is lacking for any fungus. Consequently, one can only conclude that apical actin arrays may be associated with either filasomes, when present, or the plasmalemma, or both. However, the ease with which plaquelike aggregates of actin can be induced in tips normally lacking them (Heath, 1987, 1988) is a clear reminder that all plaques may represent preparation-induced artifacts, especially when there is extensive prefixation handling of cells (Adams and Pringle, 1984; Anderson and Soll, 1986). The observation that the normally abundant subapical plaques are rare in living hyphae of Saprolegniu (Jackson and Heath, 1990a; Figs. 8,9) reinforces this point. Nevertheless, even if some or all plaques prove to be fixation-induced artifacts, they are an interesting and potentially valuable artifact that tells us something about the lability and linkages of actin in the cell. The foregoing comments summarize the well-documented arrays of actin associated with tip growth in fungi, but I shall now review the limited evidence suggesting that they may not represent the total actin complement of hyphal tips. This evidence comes from observations of atypical or minority data that hint at alternative arrays and indications of the inadequacies of current technologies.

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Examples of the first type include (a) published pictures suggestive of a diffuse, possibly plasmalemma-associated, pattern of rhodamine-phalloidin staining in some hyphal tips from populations showing typical plaques (Figs. 3 and 7 of Hoch and Staples, 1985; Fig. 16 of Hoch and Staples, 1983a; Fig. 3c of Runeberg et al., 1986; Fig. 5 of Marks and Hyams, 1985); (b) areas of artifactually induced, apparently contracted cytoplasm showing a high concentration of diffuse rhodamine-phalloidin staining (McKerracher and Heath, 1987; Fig. 13); (c) light but uniform levels of rhodaminephalloidin staining in artifactually produced plasmalemma strands (Fig. 14); (d) the inducibility of perinuclear actin shells in hyphae normally lacking them (Figs. 15, 16), a phenomenon that suggests there may normally be low and undetected populations of actin associated with the nuclear envelope and that unknown triggers cause the rest of the cytoplasmic actin to be drawn to the nuclear envelope; (e) reports of actinlike filaments associated with the Spitzenkorper of some species (Howard, 1981; Roberson and Fuller, 1988) (there are currently no rhodamine-phalloidin studies showing comparable arrays in any fungi); (f) the apparently atypical report showing diffuse apical actin in Gyromitru (Salo et al., 1989). This fungus may differ from other eufungi or may simply be showing the normal array that is artifactually transformed to plaques in related species. All of these data suggest that the dominantly reported arrays may not indicate the whole story for actin organization. Examples of the possible inadequacies of current localization techniques include the following: (a) In some cell types (not fungi), rhodamine-phalloidin seems incapable of staining all actin populations (Wilson et al., 1987; Tang et al., 1989). (b) Living cells stained with rhodamine-phalloidin show somewhat different staining patterns relative to fixed cells (Jackson and Heath, 1990a; Figs. 8,9). (c) Increasing “refinement” of technique in some cells reveals increasingly fine and complex (and presumably more complete) actin arrays (Traas e f al., 1987; Pierson, 1988; Sonobe and Shibaoka, 1989; Kakimoto and Shibaoka, 1987). (d) Different buffers in the fixative give different staining patterns (Heath, 1987); consequently further changes in protocol may reveal other patterns yet undetected. (e) Current rhodamine-phalloidin staining protocols give poor-quality ultrastructural images (Fig. 15 of Heath, 1987),suggesting that they are not yet optimal. (f) Both pre- and postfixation detergent treatments (Heath, 1987, 1988; Figs. 17-21) induce changes in actin arrays, thus indicating their lability, even after fixation, and FIG. 1. Tips of three germ tubes of Uromyces vignue, showing apical accumulations of actin plaques. ~ 2 0 0 0I.. B. Heath (unpublished). FIG.2. Cells of Neozygires sp. Plaques of actin occur over the surface of all cells but are most abundant at the growing ends of the cells. These cells contain multiple nuclei, typically four, which are enclosed by an actin shell. The bright equatorial band of actin in one cell is associated with septum formation. Fluorescence image (a) and Nomarski differential interference contrast (DIC) image (b) of the same cells. x725. From Butt and Heath (1988). FIG.3. Hyphal tips of Conidiobolus obscurus viewed with fluorescence (a) and DIC (b), showing the apical accumulation of actin plaques. ~ 7 2 5T.. M. Butt and I. B. Heath (unpublished).

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therefore the potential for artifact. This point is especially relevant in immunocytochemical protocols that involve many postfixation processing steps. Clearly the ideal answer to analyzing the arrangement of actin in cells involves high-resolution and high-sensitivity observations of labeled living cells, but while this has now been achieved to some extent (Jackson and Heath, 1990a), these authors point out a number of remaining shortcomings. The foregoing comments indicate the potential inadequacies in our observations of actin distribution in tip-growing cells. Similar comments apply to the localizations of ABP that selectively colocalize with different actin arrays (Liu and Bretscher, 1989; Drubin er al., 1988). The possibilities of artifactual colocalization due to movement during processing or artifactual differential localizations due to differential losses during processing cannot be discounted, nor can the organization of the arrays themselves be considered reliable in light of the previous comments. Similarly the phenotypic analyses of actin and ABP mutants with current cytological techniques may not be telling the full story because of the potential inadequacies of the techniques discussed here. The foregoing discussion makes it very clear that absolute reliance on current observations of actin distributions in tip-growing cells is very unwise. However, a number of points can be accepted with some confidence: (a) Actin is most concentrated in growing hyphal tips. (b) Actin is most abundant in the vicinity of the plasmalemma, to which at least some of it is likely to be attached. (c) There may well be populations of actin, at low concentration, permeating the cytoplasm and attached to at least some organelles (e.g., filasomes and nuclei). I shall now explore ways in which these observations may help explain how actin is involved in the processes of tip growth.

IV.Roles of Actin in Hyphal Tip Growth A. TIPMORPHOGENESIS Picton and Steer (1982) postulated that tip morphogenesis in pollen tubes might be regulated by cytoplasmic actin in addition to, or instead of, the plastic FIG.4. Hyphal tips of Suprolegniu ferax showing caps of finely filamentous actin and subapical . Heath (1987). arrays of cables and plaques in older regions of hyphae. ~ 2 0 0 0From FIG.5. Hyphal tip and branch of Entomophuga uulicue showing concentrationsof actin plaques T.M. Butt and I. B. Heath (unpublished). at the tips. ~1000. FIG.6. Hyphal tip of Achyla ambisexualis. The DIC image (c) shows it to be a normal-looking tip. In median optical section actin is clearly more concentrated at the apex (b), and in a surface optical section (a) the actin forms a filamentous cap comparable to that seen in Suprolegnia (Fig. 4). ~1000.S. G. W. Kaminskyj (unpublished). FIG.7. Hyphal tip of Pythium uphunidermurum viewed in median (b) and surface (a) optical sections showing the basic similarity of actin patterns relative to the other oomycetes (Figs. 4 and 6). ~ 2 0 0 0R. . J. Howard and I. B. Heath (unpublished).

FIGS.8 and 9. Living and growing hyphae of Suprolegniuferux electroporated with rhodaminephalloidin, showing that the basic organization of the apical actin caps is similar to that seen in fixed . Jackson and Heath (1990a). hyphae (Fig. 4). ~ 1 2 0 0From FIGS.I(L12. Hyphae of Suprolegniuferu-r showing reconstructed sequences of branch formation concomitant with the reorganization of the subapical actin arrays into a cap typical of hyphal tips. ~ 2 0 0 0From . Heath (1987).

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deformability of the apical cell wall. The observed apical actin cap of Saprolegnia and the other oomycetes (Figs. 4 , 6 1 2 ; Heath, 1987) is perhaps the most likely morphology predicted by their hypothesis, although the widespread high accumulation of actin in hyphal tips is also consistent with their model. Numerous observations support the hypothesis that the actin cap in Saprolegnia regulates tip morphogenesis. Its formation at the initiation of branching (Figs. 10-1 2), its consistent presence in growing tips, its disorganization concomitant with cessation of growth (Heath, 1987), the correlation between its length and the rate of tip growth (Jackson and Heath, 1989, 1990b) (faster growing hyphae, which may have longer zones of plastic wall, have correspondingly longer caps), the preference for bursting-induced by osmotic shock-to occur in the cap region where presumably the wall is weakest and the transition from the tapering, expanding region of the tip to the parallel, nonexpanding subapical region being coincident with the base of the cap (Jackson and Heath, 1990b) all support, but do not prove, the hypothesis. Similarly transiently accelerated growth rates induced by cytochalasin (Jackson and Heath, 1990b) and Caz+-induced changes in tip morphology (Jackson and Heath, 1989) support the model. The apical plaques found in other fungi are less easily envisaged as playing a role in tip morphogenesis. However, cytochalasins (Betina et al., 1972; Allen et al., 1980; Grove and Sweigard, 1980; Tucker et al., 1986) and Ca2+ and H+ concentrations (Dow and Rubery, 1975), which are likely to affect actin arrays also influence tip morphology, usually causing swelling, in other fungi. Similarly, there is a correlated appearance of abnormal actin arrays and slow growth rates in “snowflake” mutants of Aspergillus (Allen et al., 1974), and cytochalasin reverses slow growth in other mutants (Allen ef al., 1980). Furthermore, Novick and Botstein (1985) showed that actin mutation led to osmotic sensitivity and that overproduction of an ABP induces abnormal budding (Drubin et al., 1988) in Saccharomyces. All of these observations are consistent with the hypothesis that actin can play a morphogenetic role in fungi other than the oomycetes. However, none of these data are unambiguous for reasons discussed in Section II1,B. Furthermore, experiments on tip morphogenesis must consider the properties of the wall itself. For example Ca*+ may (Virk and Cleland, 1988; Bittisnich and Williamson, 1989), or may not (Rayle, 1989) directly modulate cell wall properties. If actin is important in tip morphogenesis, then it must be tightly linked to the plasmalemma, which is the effective site of turgor pressure application, and possibly also to the cell wall. Gustin et al. (1988) also postulated the need for plasmalemma-linked cytoskeletal elements in their work on mechanosensitive ion channels in Saccharomyces. The microfilament-associated granules on the plasmalemma of Schizosaccharomyces(Kanbe et al., 1989) are evidence for the existence of such linkages in fungal growth zones. There is also evidence in plants for actin-plasmalemma-cell wall linkages (Schindler et al., 1989), and Kropf et

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al. (1988) have indicated the likely existence of such linkages in germinating Fucus zygotes where incipient tip growth needs a cell wall for localization and has an actin cap. Certainly in animal cells there are numerous actin-membrane linking proteins and Ca2+ sensitivity is known (Pollard and Cooper, 1986). The ABP that colocalize with actin plaques in Saccharomyces (Drubin et al., 1988) are candidates for fungal actin-membrane linkers, but there are no data to support this speculation yet. B. LOCALIZATION OF PLASMALEMMA-ASSOCIATED PROTEINS The existence of electrical currents and ion gradients (Harold and Caldwell, 1990; Section II,C) and cell wall synthesis gradients (Girard and Fevre, 1984; Gooday and Gow, 1990) in hyphal tips suggests that ion transport proteins and plasmalemma-based cell wall-synthetic enzymes are nonuniformly distributed on the plasmalemma. Gradients of these plasmalemma-associatedproteins could be generated by localized insertion, diffusion, and degradation, or excision at some other point. Equally, they may be held in a nonuniform pattern by being linked to a skeletal system, and the peripheral, plasmalemma-associated actin arrays are good candidates for such a role. There is no direct evidence to support this hypothesis, but Novick and Botstein (1985) showed that actin mutation altered chitin synthesis location in Saccharomyces (chitin synthase enzymes are almost certainly plasmalemma-located proteins). The widespread actin belts associated with septum formation (Girbardt, 1979) could be responsible for recruiting and localizing chitin-synthetic enzymes, and Butt and Heath (1988) suggested a similar role for a rather different array of actin associated with septum synthesis in a fission yeast. Brawley and Robinson (1985) suggested, with no direct evidence, that actin caps localized Ca2+ channels in germinating Fucus zygotes, and Dictyostelium contains an ABP, fodrin, which is analogous to spectrin, which, in erythrocytes, is an actin-anion channel-binding protein (Bennett and Condeelis, 1988). Clearly the concept of actin determining the position of FIG. 13. A short branch of Saprolegnia ferar fixed with more dilute formaldehyde than usual. Some of the cytoplasm appears to have contracted into the branch from the main hypha (DIC image, b), and this cytoplasm is unusually rich in actin (a). x1124. I. B. Heath (unpublished). FIG. 14. A subapical hypha of Saprolegnia ferar treated for 10 seconds with 0.01% Triton X-100 prior to fixation and staining. Cytoplasm has contracted to the left, leaving strands of apparently little more than membrane (phase image, b), which appears to contain actin (a). ~ 2 0 0 0I.. B. Heath (unpublished). FIGS.15 and 16. Subapical regions of hyphae treated with cytochalasin E prior to fixation and staining. This treatment sometimes, but not always, causes the loss of the normal subapical array of peripheral plaques and fibers of actin (see Figs. 4 and 10) and the formation of perinuclear (e.g., n) shells of actin. In Fig. 15 the hypha appears to be in an intermediate stage with no fibrils but residual plaques and shells, whereas in Fig. 16 all peripheral actin is gone and only shells are present. x1124. I. B. Heath (unpublished).

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plasmalemma proteins enjoys little direct support in fungi, but it is an idea whose time for direct investigation has come.

c.VESICLE TRANSPORT Polarized transport of wall vesicles to the hyphal tip occurs parallel to actin filaments whenever such are in the vicinity of the tip; this is consistent with the filaments forming a track along which the vesicles could be moved by an actin-myosin-based system as reported in other vesicle transport systems (Kohno and Shimmen, 1988; Grolig et al., 1988; Adams and Pollard, 1986). Disruption of actin arrays by actin mutation (Novick and Botstein, 1985) or ABP overproduction (Drubin et al., 1988) in Saccharomyces results in disruption of localized call expansion (due to misdirection of vesicles?), anomalous wall vesicle accumulations and inhibition of invertase secretion (an enzyme likely to be in at least some wall vesicles). Similarly, cytochalasins inhibit cellulase secretion in Achlya (Thomas et al., 1974), and cellulase is likely to be transported in wall vesicles (Nolan and Bal, 1974). Heath and Kaminskyj (1989) showed good concordance between actin arrays and translocating wall vesicle populations in Saprolegniu. In other tip-growing cells the transport of comparable vesicles can be inhibited by cytochalasins (Picton and Steer, 1981; Lancelle and Hepler, 1988; Bartnik and Severs, 1988), and they are associated with myosin (Tang et al., 1989; Heslop-Harrison and Heslop-Harrison, 1989). Fungi also contain myosin (Watts et al., 1985; Drubin et al., 1988). All of the foregoing data strongly suggest that myosin-coated wall vesicles are translocated to the hyphal tip by sliding along actin cables. However, in some fungi there are data that seem to contradict this model. Howard and Aist (1980) reported that disruption of MT disrupted wall vesicle distribution patterns and FIG. 17. Hypha of Suprolegniuferax treated with 0.1% Nonidet P-40for 40 seconds prior to fixation. The apical actin fibrils are fragmented and the subapical plaques are no longer seen. ~2000.I. B. Heath (unpublished). FIG. 18. Hypha of Suprolegniu ferax treated with 0.1%' h e e n 20 for 10 minutes prior to fixation. 124. The normal apical actin cap has been almost entirely replaced by coarse peripheral plaques. XI From Heath (1988). FIG. 19. Hyphal tip of Suprolegniuferax treated with 0.1% Brij 58 for 5 minutes prior to fixation. This mild detergent treatment, which preserves organelle motility, induces changes similar to Tween 20 (Fig. 18). x1124. From Heath (1988). FIG. 20. Subapical hypha of Suprolegniuferar with incipient branch (upper center) treated as in . Heath Fig. 18, showing disruption of normal actin arrays (compare with Figs. 10-12). ~ 1 1 2 4From (1988). FIG.21. Hyphae of Suprolegniu ferax treated with 0.1 ?hsodium dodecyl sulfate ufrer normal fixation but prior to rhodamine-phalloidin staining. Note that the normal actin arrays have been totally displaced by a diffuse staining pattern that specifically permeates all of the cytoplasm but not the vacuoles (lower left region in a). The phase-contrast pictures show that the tips are somewhat extracted but relatively normal in appearance (b and c). ~ 9 3 0I.. B. Heath (unpublished).

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thus implied that wall vesicles were transported by interactions with MT as appears to occur in axons (Sheetz et al., 1986). However, they did observe that growth (and therefore presumably wall vesicle transport) continued in the absence of MT. Herr and Heath (1982), Hoch et al. (1987), and Jacobs el al. (1988) showed similar growth in the absence of MT in other fungi. Since MT appear to interact with actin to determine the properties of the cytoskeleton (McKerracher and Heath, 1986; Heath and Kaminskyj, 1989), it seems likely that Howard and Aist's (1980) data could be attributable to an indirect effect of the MT on an actin-based translocating system, rather than providing evidence for a direct role for MT. Huffaker et al. (1988) also showed that MT are not needed for vesicle transport in Saccharomyces. Clearly, at present the balance of evidence favors an actin-myosin-based wall vesicle transport system, which predicts that the apical actin filaments should be polarized with their "pointed" ends (see Section IKA) away from the hyphal tip. Such a prediction would explain the polarized transport of the vesicles and is testable.

D.EXOCYTOSIS Picton and Steer (1982) summarized the largely circumstantial evidence suggesting that actin is directly involved in effecting exocytosis of wall vesicles in pollen tubes, and Brawley and Robinson (1985) suggested a similar role in Fucus embryo germination. There is little direct evidence to support such a role in fungi. However, clearly the very densely packed filaments of the actin cap in Saprolegnia (Heath, 1987; Jackson and Heath, 1990a) must undergo changes, causal or permissive, to allow exocytosis. The actinlike filaments on the surfaces of some wall vesicles (Hoch and Staples, 1983b; Hoch and Howard, 1980; Howard, 1981; Heath et al., 1985; Roberson and Fuller, 1988) may be involved in exocytosis but could also have more to do with transport. Interestingly, the analogous synaptic vesicles of neurons are attached to actin during transit via a specific protein (synapsin 1). the phosphorylation of which releases the vesicles and thereby permits exocytosis (Hirokawa et al., 1989). Inhibition of secretion by actin mutation or cytochalasins (see Section IV,C) is consistent with a role for actin in exocytosis but, as noted previously, because the secretory pathway involves many steps, including vesicle transport, it is totally obscure whether the actin involvement is at the point of exocytosis.

E. ORGANELLE MOVEMENTSAND POSITIONING We have seen that actin is most likely to be the dominant component in wall vesicle transport (Section IV,C), and 1 argue here that it is also important in the transport and positioning of other organelles. However, contrary to the situation

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with wall vesicles, there is substantial evidence to show that MT are also important in the behavior of the other organelles. This evidence has been reviewed previously (Heath et al., 1982; McKerracher and Heath, 1987) and need only be summarized here. It includes morphological observations of interactions between organelles and MT and disruptions of organelle behavior by anti-MT drugs, MT-selective ultraviolet (UV) microbeams, and tubulin mutations. These data clearly show that MT are involved in organelle motility and positioning, but they do not show how they are involved. Aist and Berns (1981) suggested that the ends of astral MT slide along the plasmalemma to generate mitosis-related nuclear movements in Fusariurn but there are no data to identify the forcegenerating molecules in this or any other fungal system. It is inappropriate to discuss further the MT data here. Instead I shall focus on data that show that MT (with microtubule-associated proteins (MAP) alone are insufficient to account for the relevant movements and that actin (with ABP) is most probably also involved. The best-known MT-based motors utilize mechanochemical translocators such as dynein (Witman, 1989), kinesin (Sheetz, 1989), or vesikin (Sloboda and Gilbert, 1989), which either slide vesicles or organelles along static tracklike MT or move MT relative to a static substrate such as a microscope slide. Clearly the results of these interactions will depend on the balance of forces and introduce the need to consider the anchoring of MT and the resistance encountered by the organelles. These considerations are especially important in the context of the behavior of large organelles such as nuclei and mitochondria in hyphae, which often have diameters only a little larger than the organelles. Clearly, moving a large nucleus through cytoplasm demands that the force-generating system must be firmly anchored or it, not the organelles, will move. If the force generator is attached to a long MT, it is conceptually simple to envisage anchorage of the MT to structures or cytoplasm remote from the moving organelle, which in turn could be in cytoplasm of transiently reduced resistance (e.g., solated gel), thus producing a fixed track and motile organelle. Alternatively, since MT can propel themselves in vitro, then their static attachment to the organelle and their interaction with remote “rigid” cytoplasm could also move the organelle. These concepts are difficult to envisage when the MT extend little or not at all beyond the organelle, because then one has the paradoxical situation of attempting to generate force against cytoplasm that is sufficiently “rigid” to remain static against applied tension yet sufficiently deformable to permit passage of the organelle. At present there are no data on fungi that reveal sites of MT anchorage or describe local differences in cytoplasmic consistency. However, there are data that suggest that MT associated with organelle behavior do not act in either a tracklike manner or as self-motile structures pulling (or pushing) against a static matrix. For example, in Basidiobolus, perinuclear MT are predominantly short and not attached to the

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nuclear envelope (McKerracher and Heath, 1985). Likewise, in Uromyces infection structures the nuclear envelope-associated MT do not appear to extend for long distances (Heath and Heath, 1978). although in germ tubes immunofluorescence data suggest that they may be longer (Hoch and Staples, 1985). Nucleus-associated MT in Suprolegniu seem to be short (Heath and Kaminskyj, 1989) and the postmitotic migrating nuclei of Pleurotus apparently do not utilize the astral MT as force generators (Kaminskyj ef al., 1989). Similarly, mitochondria-associated MT are predominantly short and extend little beyond the organelles in both Uromyces (Heath and Heath, 1978) and Suprolegniu (Heath and Kaminskyj, 1989). Furthermore, Oakley and Rinehart (1985) showed that cells containing defective MT were incapable of sustaining normal nuclear movements but showed normal mitochondria1 positioning. Collectively these data show that, while MT are indeed involved in organelle movements and positioning, they do not seem to function in either a tracklike or towing mode. Most likely they serve as intermediaries with other cytoskeletal systems and there is some evidence to suggest that actin is part of this other system. Heath et ul. (1982), Heath and Heath (1978), and Hoch and Staples (1983b) have shown actinlike filaments associated with organelle-associated MT in Uromyces, and Hoch and Staples (1985) showed both concordance and interdependence of actin and MT arrays in these cells. In a somewhat analogous situation, Uyeda and Furuya (1989) showed actin-microtubule interactions in the flagellar roots of Physurum cells. In a somewhat different, but complementary vein, the perinuclear arrays of actin found in Suprolegnia (Figs. 15, 16), Schizosuccharornyces (Marks ef ul., 1986). and Neozygifes (Butt and Heath, 1988) could be evidence that the nuclear envelope is connected to an extensive cytoplasmic actin array that, under some circumstances, tends to collapse or contract to a concentrated aggregate around the nuclei. Evidently such an array would have some effect on nuclear positioning or movement. In this context, it is important to mention that most studies have not shown pennuclear, or mitochondria-associatedactin but negative results are not compelling. For example Fath and Lasek (1988) have shown actin arrays that are not detected by rhodamine-phalloidin staining in axons. Consequently the absence of evidence for actin arrays suitable for a role in organelle motility cannot be accepted at face value. Certainly the evidence for a role for actin in fungal organelle movements is not compelling, but the data supporting the involvement of actin and myosin in organelle movements in algae (Menzel and Elsner-Menzel, 1989a,b) and pollen tubes (Heslop-Harrison and Heslop-Harrison, 1989; Kohno and Shimmen, 1988; Lancelle and Hepler, 1988; Tang er ul., 1989) clearly show that the possibility is very real. F. CYTOPLASMIC MIGRATION

In Section KG, we saw that hyphal cytoplasm can be considered to show ameboid movement; consequently, it is likely that cytoplasmic migration in-

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volves a similar mechanism to ameboid movement, and this almost certainly is based on actin and myosin (Taylor and Condeelis, 1979). Direct evidence for this suggestion is sparse in fungi. The dominant detected fibrillar actin arrays in hyphae are parallel to the postulated direction of movement and contraction (Section III,C), but it is more likely that a more diffuse population of actin and myosin permeating the entire cytoplasm is important in the process. There is evidence for such in Gyromitrium (Salo et al., 1989), and apparently contracted cytoplasm rich in actin is seen in Saprolegnia (McKerracher and Heath, 1987; Fig. 13). McKerracher and Heath (1986) showed CaZ+-sensitiveinducible cytoplasmic contraction in Basidiobolus, as might be expected for an actin-based system. However, Tucker et al. (1986) provided contrary evidence by showing that cytochalasin E destroyed actin filaments yet cytoplasmic migration was unaffected. Their cytochalasin treatment left a diffuse cytoplasmic rhodaminephalloidin stain pattern, which could be interpreted as the migration-associated actin that was unaffected by the cytochalasin. As discussed previously (Section III,B), considerable uncertainty exists in interpreting cytological data. Perhaps the most compelling argument for a role of actin in cytoplasmic migration comes by comparison with other cell types, where actin is almost universally involved (Taylor and Condeelis, 1979). An especially relevant example is woundinduced cytoplasmic contraction in some algae, where actin and myosin seem to be involved (La Claire, 1989, and references therein). If actin is indeed involved in cytoplasmic migration and contraction, one might expect some form of apex-located actin-plasmalemma attachments, especially since inducible contractions seem to be undirectional toward the tip (McKerracher and Heath, 1986). The microfilament-associated granules on the plasmalemma of Schizosaccharomyces (Kanbe et al., 1989) are obvious excellent candidates for the hypothesized attachments. Adams and Pringle (1984) speculated that the apical plaques in yeast buds served such a role, and the demonstration of ABP in these plaques (Adams et al., 1989; Drubin et al., 1988) supports their speculations. The apical lomasomes induced in Saprolegnia by Heath (1987) also indicate a strong attachment between actin and the apical plasmalemma. Allen ef al. (1974) showed that inducible actin filament bundles could be apparently attached to the plasmalemma in Neurospora. In mammalian cells the focal contacts involved in actin-based cytoplasmic migration involve specific proteins that not only link actin to the plasmalemma but also link the plasmalemma to the cell substrate (Bumdge ef al., 1987). There is no evidence for such proteins in fungi, but they are found in plant cells (Schindler et al., 1989) and can be inferred to exist in the tip-growing Fucus 'germling, since Kropf ef al. (1988) showed that both actin and a cell wall are needed for fixation of the axis of polarity during germination. While all of the foregoing suggest that actin and myosin may indeed be involved in cytoplasmic migration in hyphae, it is important to note that other systems may be involved. For example, Sepsenwol et al. (1989) showed that

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ameboid motility of Ascaris sperm is dependent on a family of low molecular weight nonactin-nonmyosin proteins. Clearly, extrapolating across wide phylogenetic distances is hazardous.

V. Regulation of Actin Arrays The disposition, properties, and functioning of actin arrays are known to be influenced by three general intracellular features: ion composition and concentration (Kom, 1982), ABP (Pollard and Cooper, 1986), and other cytoskeletal elements. In the fungi there is no evidence for the existence of intermediate filaments, so that latter category essentially means MT as far as we know at present. 1 shall now briefly review the evidence for the ways in which each of these features may be functioning in fungal tip growth. A. IONICREGULATION We have seen that actin is intimately associated with tip growth and we know that in many cell types Ca2+ influences the structure of actin gels (Condeelis et al., 1984; Pollard, 1981; Yin et al., 1980) and the interaction between actin and myosin (Williamson, 1984; Kohama and Shimmen, 1985; Pies and WohlfarthBottermann, 1986). Consequently, it is likely that any Ca2+effects on tip growth are mediated via the actin system and therefore data implicating Ca2+ in tip growth can be construed as evidence for Ca2+ regulation of tip growth-related actin. However, this line of evidence is very circumstantial because Ca2+ are likely to influence many other aspects of tip growth, including cell wall extensibility (Togawa and Bonner, 1957; Dow and Rubery, 1975; Virk and Cleland, 1988; Bittisnich and Williamson, 1989; Rayle, 1989). Ca2+ affects hyphal growth rates (McGill and Gow, 1987; Takeuchi et al., 1988; Schmid and Harold, 1988; Jackson and Heath, 1989), branching patterns (Reissig and Kinney, 1983; Harold and Harold, 1986; Schmid and Harold, 1988). and tip morphology (Schmid and Harold, 1988; Jackson and Heath, 1989). Hyphae establish tip-high gradients of membrane-associated Ca2+(Reiss and Herth, 1979; Schmid and Harold, 1988; Jackson and Heath, 1989), and other tip-growing cells show a similar gradient of free cytoplasmic Ca2+ (Brownlee and Wood, 1986), although the latter has yet to be demonstrated in any hyphae. Similar sorts of data show similar effects of Ca2+ in other tip-growing cells (Steer and Steer, 1989; Steer, 1990; Herth et al., 1990), suggesting the universality of Ca2+ involvement in tip growth. However, only in the cases of cytoplasmic contraction of Basidiobolus (McKerracher and Heath, 1986) and the apical actin arrays of Saprolegnia (Jackson and Heath, 1989) is there evidence for a direct effect of Ca2+ on tip growth-related actin arrays. Even in these, and all of

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the other studies, it is not possible to prove that the primary target of Ca2+ is regulation of actin. Because fungi contain calmodulin (Davis and Thorner, 1986) and antagonists of calmodulin also influence growth rate (Brownlee, 1984; Ortega-Perez and Turian, 1987), and because calmodulin mediates Ca2+ regulation of many intracellular processes (Klee et al., 1980), it is very difficult to ascribe Ca2+ effects directly to the actin system alone. In addition to the possible roles of Ca2+in regulation of actin, H+ can also influence actin assemblages (Wang et al., 1989). There is evidence for a tip-high pH gradient in hyphal tips (Turian, 1978, 1981; McGillivray and Gow, 1987), but its role in tip growth is unclear. pH effects on growth rates are very difficult to analyze (Griffin, 1981). However, the availability of sensitive intracellular pH indicators makes it possible to investigate the role of pH in actin regulation and tip growth in a more meaningful way. Before leaving the topic of ionic regulation of actin, it is worth pointing out that the properties of actin, like many other cellular features, can be influenced by physiologically realistic concentrations of ions such as Mg*+, K+, Na+, and POi(Korn, 1982). Furthermore, the properties of ABP are also influenced by diverse ions; consequently the unraveling the possible regulatory behavior of ions on tip growth and actin arrays is very complex and has barely even begun. B. ACTIN-BINDING PROTEINS Mammalian and ameboid cells are known to contain diverse ABP that interact with actin and regulate its polymerization, gel properties, and associations with mechanochemical effectors such as myosin (Korn, 1982; Pollard and Cooper, 1986). The first-described ABP in fungi was myosin, now known from both Saccharomyces (Watts et al., 1985, 1987; Drubin et al., 1988) and probably Neurospora (Van Tuinen el al., 1986). Since the interaction between actin and myosin in muscle cells is regulated by tropomyosin, it is not surprising that this ABP has now been found in Saccharomyces and shown to influence the organization of actin arrays in this organism (Liu and Bretscher, 1989). The array of known ABP in Saccharomyces has been extended by the discovery of six genes (SAC genes) encoding ABP (Novick et al., 1989; Adams and Botstein, 1989; Adams et al., 1989). At least four of these gene products influence the organization of actin arrays, and one of them encodes the same 67-kDa protein previously isolated by actin affinity chromatography by Drubin et al. (1988). Drubin et al. (1988) also isolated an 85-kDa ABP, but it is not yet clear if this corresponds to one of the six SAC genes. The 67-and 85-kDa ABP and 'tropomyosin all colocalize with actin in the cell, but they show different patterns with the 67-kDa protein on both cables and plaques, tropomyosin on cables only, and 85-kDa on plaques only (Drubin et al., 1988; Liu and Bretscher, 1989), as might be expected for molecules involved in determining

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the organization and functioning of the actin arrays. Not surprisingly, all of the yeast ABP analyzed in detail alter various aspects of cell growth and morphology, but at present-partly for reasons discussed in Section IVB-these alterations have not helped describe the precise functions of either actin or the ABP. Nevertheless, the identification and analysis of ABP by both biochemical and genetic techniques is an important advance in our understanding of intracellular actin arrays. C. MICROTUBULES It has been known for a long time that MAP can mediate functionally significant interactions between F actin and MT (Pollard et al., 1984). Consequently, it is entirely likely that these two cytoskeletal elements should have some influence on each other in fungal hyphae. However, direct evidence for this is sparse. Actinlike filaments are known to be associated with MT in Uromyces germ tubes (Heath and Heath, 1978; Heath et al., 1982; Hoch and Staples, 1983b), but only the work of Hoch and Staples (1985) has addressed the question of the possib!e role of the MT in determining the organization of the actin arrays. They found that disruption of the cytoplasmic MT generally had rather little effect on the actin arrays but did find that apical and subapical actin filament arrays were frequently undetected after loss of MT. These results suggest that MT play some role in actin organization but show that other factors are probably more important in Uromyces germ tubes. The possible role of MT in other hyphal tips is unexplored; however, given that MT generally tend to be rather few in number in the periphery of hyphal tips where most actin is organized (Heath and Kaminskyj, 1989), it seems likely that their role in regulating actin arrays is not extensive or pivotal.

VI. Future Directions It is clear from the preceding discussion that actin is likely to be vitally involved in all aspects of tip growth of fungal hyphae. However, the data for many of the suggested functions are underwhelming and are typically available only for a very limited number of species. In order to produce a coherent and general analysis of the way actin contributes to tip growth, many more careful and detailed studies integrating the advantages of molecular, genetic, and cytological techniques are needed from a selected range of diverse species. Work using ultrastructural immunocytochemistry to localize precisely hypothetically interesting ABP such as those responsible for modulating actin gel properties and contractility, and actin-membrane interactions seem potentially particularly rewarding. Equally, high-resolution analysis of ion fluctuations and gradients

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and the identification of intercellular ion distribution effectors and storage systems are now technically feasible and cannot fail to be most rewarding. Perhaps the most important aspect that must be considered in such future work is a greater effort to ensure that the data are demonstrably artifact-free and apply to hyphal tips known with a high degree of certainty to have been actively undergoing tip growth at the time of analysis. These are not trivial requirements in a system so labile as a growing hyphal tip. However, the range of technologies currently available and the conceptual advances emanating from diverse areas of cell biology suggest that now, perhaps more than at any time in the past, we are likely to make very significant progress in understanding all of the beautifully orchestrated interacting processes that are collectively manifest as a growing hyphal tip.

ACKNOWLEDGMENTS It is a pleasure to acknowledge the stimulating discussions with Lisa McKerracher, Sandra Jackson, and Susan Kaminskyj, all of whom have contributed substantially to the development of ideas expressed in this paper. The Natural Sciences and Engineering Research Council of Canada have provided continuing support, which has been crucial to the development of the ideas and the execution of some of the unpublished work described here. Benita Rozario did an excellent job of typing the manuscript.

REFERENCES Adams, A. E. M., and Botstein, D. (1989). Generics 121,675-683. Adams, A. E. M., and Pringle, J. R. (1984). J. Cell Biol. 98,934-945. Adams, A. E. M., Botstein, D., and Drubin, D. G. (1989). Science 243,231-233. Adams, R. J., and Pollard, T. D. (1986). Nurure (London) 322,754-756. Aist, J. R.,and Berns, M. W. (1981).J . CellBiol. 91,446-458. Allen, E. D., Lowry, R. J., and Sussman, A. S. (1974). J. Lllrrusfrucr.Res. 48,455464. Allen, E. D., Aiuto, R., and Sussman, A. S. (1980). Proroplusm 102,63-75. Anderson, J. M., and Soll, D. R. (1986). J . Gen. Microbiol. 132,2035-2047. Bartnicki-Garcia, S . (1973). In “Microbial Differentiation” (J. M. Ashworth and J. E. Smith, eds.), pp. 245-267. Cambridge Univ. Press, London and New York. Bartnicki-Garcia, S. (1990). In “Tip Growth” (I. B. Heath, ed.), pp. 21 1-232. Academic Press, San Diego California (in press). Bartnicki-Garcia, S., and Lippman, E. (1969). Science 165,302-304. Bartnicki-Garcia, S., and Lippman, E. (1972). J. Gen. Microbiol. 73,487-500. Barmik, E., and Severs, A. (1988). Plunru 176, 1-9. Bennett, H., and Condeelis, J. (1988). Cell Motil. Cyroskel. 11,303-317. Betina, V., Micekova, D., and Nemec, P. (1972). J . Gen Microbiol. 71,343-349. Bittisnich, D. J., and Williamson, R. E. (1989). Plantu 178,96102. Bracker, C. E., (1971). Proroplusrnu 72,381-397. Brawley, S . H., and Robinson, K. R. (1985). J . CellBiol. 100, 1173-1184.

124

1. B. HEATH

Brownlee, C. (1984). Newt Phyrol. 97, 15-24, Brownlee, C., and Wood, J. W. (1986).Narure (London) 320,624-626. Burridge, K.. Molony, L., and Kelly, T. (1987). J. Cell Sci., Suppl. 8, 21 1-229. Butt, T. M., and Heath, I. B. (1988). Eur. J. Cell Biol. 46,499-505. Clarke, M., Schatten, G., Mazia, D., and Spudich, J. A. (1975). Pror. Natl. Acad. S r i . U.S.A. 72, 1758-1762. Condeelis. J., Vahey, M.. Carboni, J. M., DeMey, J., and Ogihara, S. (1984). J . Cell Biol. 99, I19~-126~. Davis, T. N.. and Thorner, J. (1986). In “Yeast Cell Biology” (J. Hicks, ed.), pp. 477-503. Alan, R. Liss, New York. Dow. J. M.. and Rubery, P. H. (1975). J. Gen. Microbiol. 91,425428. Drubin, D. G.. Miller, K. G., and Botstein, D. (1988). J . Cell Biol. 107, 2551-2561. Fath. K. R., and Lasek, R. J. (1988). J. Cell Biol. 107,613421. Faulstich, H.. Zobeley, S., Rinnerthaler. G.. and Small, J. V. (1988). J. Musrle Res. Cell Moril. 9, 370-383. Fhvre, M., and Rougier, M. (1982). Arch. Microbiol. 131, 212-215. Forer. A. (1985). Can. J. Biochem. Cell Biol. 63,585-598. Forer, A., Emmersen. J., and Behnke, 0. (1972). Science 175,77L776. Girard, V., and Fkvre, M.(1984). J . Gen. Microbiol. 130, 1557-1562. Girbardt, M. (1969). Protoplasma 67,413-441. Girbardt, M. (1979). Exp. Mycol. 3,215-228. Gooday. G. W. (1971). J. Gen. Microbiol. 67, 125-133. Gooday, G. W. (1983). In “Fungal Differentiation” (J. E. Smith, ed.), pp. 315-356. Dekker, New York. Gooday. G. W., and Cow, N. A. B. (1990). In”Tip Growth (1. B. Heath, ed.), pp. 31-58. Academic Press, San Diego, California. Cow, N. A. R. (1989). Biol. Bull. (Woods Hole, Mass.) 176(S), 31-35. Griffin. D. H. (1981). “Fungal Physiology.” Wiley, New York. Grolig, F.. Williamson, R. E.. Parke. J., Miller, C., and Anderton, 8 . H. (1988). Eur. J . Cell B i d . 47, 22-3 I . Grove, S. N. (1978). In “The Filamentous Fungi” (J. E. Smith and D. R. Berry, eds.), Vol. 3. pp. 28-50. Wiley. New York. Grove, S. N.. and Bracker, C. E. (1970). J. Bacteriol. 104,989-1009. Grove, S . N., and Sweigard, J. A. (1980). Exp. Mycol. 4,239-250. Grove, S. N.. Bracker, C. E., and Morr.5, D. J. (1970). Am. 1.Bot. 57,245-266. Gunderson, J. H., Elwood. H.. Ingold. A., Kindle, K., and Sogin, M. L. (1987). Proc. Natl Acad. Sci. U.S.A. 84,5823-5827. Gustin, M. C., Zhou. X.-L., Martinac, B., and Kung. C. (1988). Scienre 242,762-76s. Harold. F. M., and Caldwell, J. H. (1990). In “Tip Growth’’ (I. B. Heath. ed.), pp. 59-90. Academic Press, San Diego, California. Harold. F. M., Kropf. D. L., and Caldwell, J. H. (1985). Exp. Mycol. 9, 183-186. Harold, R. L., and Harold. F. M. ( 1986). J. Gen. Microbiol. 132,2 13-2 19. Hasek, J., Rupes, I., Svoboda, J., and Streiblova, E. (1987). J. Gen. Microbiol. 133,3355-3363. Heath, I. B. (1982). In Microtubules in Microorganisms” (P. Capuccinelli and N. R. Morris, eds.), pp. 275-3 1 I . Dekker, New York. Heath, 1. B. (1987). Eur. J . Cell Biol. 44, 10-16. Heath, 1. B. (1988). J. Cell Sci. 91,4147. Heath, 1. B., ed. (1990). “Tip Growth.” Academic Press, San Diego, California. Heath, I. B., and Greenwood, A. D. (1970). J. Gen. Microbiol. 62, 129-137. Heath, I. B.. and Heath, M. C. (1978). Cyrobiologie 16,393411.

ROLES OF ACTIN IN TIP GROWTH OF FUNGI

125

Heath, I. B., and Kaminskyj, S. G. W. (1989).J . Cell Sci. 93,41-52. Heath, I. B., Heath, M. C., and Herr, F. (1982). Symp. SOC.Exp. Biol. 35,563-588. Heath, I. B., Rethoret, K., Arsenault, A. L., and Ottensmeyer, F. P. (1985). Proroplusmu 128,81-93. Herr, F.B., and Heath, M. C. (1982). Exp. Mycol. 6,15-24. Herth, W., Reiss, H.-D., and Hartmann E. (1990). In “Tip Growth” (I. B. Heath, ed.), pp. 91-118. Academic Press, San Diego, California. Heslop-Harrison, J., and Heslop-Harrison, Y. (1989). J. Cell Sci. 94, 315-325. Heuser, J. E., and Kirschner, M. W. (1980). J. Cell Biol. 86,212-233. Hirokawa, N.. Sobue, K., Kanda, K., Harada, A., and Yorifuji, H. (1989). J . Cell Biol. 108,111-126. Hoch, H. C., and Howard, R. J. (1980) Protoplusma 103,281-297. Hoch, H. C., and Staples, R. C. (1983a) EUKJ. Cell Biol. 32,52-58. Hoch, H. C., and Staples, R. C. (1983b). Mycologiu 75,795-824. Hoch, H. C., and Staples, R. C. (1985). Proroplusmu 124, 112-122. Hoch, H. C., Tucker, B. E., and Staples, R. C. (1987).Eur. J . Cell Biol. 45,209-218. Howard, R. J. (1981).J. Cell Sci. 48, 89-103. Howard, R. J., and Aist, J. R. (1979). J . Wlfrastrucr.Res. 66,224-234. Howard, R. J., and Aist, J. R. (1980). J. Cell Biol. 87, 55-64. Huffaker, T.C., Thomas, J. H., and Botstein, D. (1988). J. Cell Biol. 106, 1997-2010. Isaac, P. K. (1964). Can. J. Bor. 42,787-792. Jackson, S. L., and Heath, I. B. (1989). Exp. Mycol. 13, 1-12. Jackson, S. L., and Heath, I. B. (1990a). Proroplusmu 154,66-70. Jackson, S . L., and Heath, I. B. (1990b) Protoplasma (in press). Jacobs, C. W., Adams, A. E. M., Szaniszlo, P. J., and Pringle, J. R. (1988). J. Cell Biol. 107, 1409-1426. Kakimoto, T., and Shibaoka, H. (1987). Planr Cell Physiol. 28, 1581-1585. Kaminskyj, S. G. W., Yoon, K. S., and Heath, I. B. (1989). J. Cell Sci. 94,663-674. Kanbe, T., Kobayashi, I., and Tanaka, K. (1989). J. Cell Sci. 94,647656. Kendrick, B. (1985). “The Fifth Kingdom.” Mycologue Publ., Waterloo. Kilmartin, J. V., and Adams, A. E. M. (1984). J Cell Biol. 98,922-933. Klee, C. B., Crouch, T.H., and Richman, P. G. (1980). Annu. Rev. Biochem. 49,489-515. Kohama, K., and Shimmen, T. (1985). Proroplusmu 129,88-91. Kohno, T., and Shimmen, T. (1988). J . Cell Biol. 106, 1539-1543. Korn, E. D. (1982). Physiol. Rev. 62,672-737. Kropf, D. L., Kloareg, B., and Quatrano, R. S. (1988). Science 239, 187-190. La Claire, J. W., II (1989). Plantu 177,47-57. Lancelle, S. A., and Hepler, P. K. (1988). Proroplusmu, Suppl. 2,65-75. Liu, H., and Bretscher, A. (1989). Cell (Cambridge, Mass.) 57,233-242. Mabuchi, I., and Okuno, M. (1977). J. Cell Biol. 74,251-263. Marks, J., and Hyams, J. S. (1985). Eur. J . Cell Biol. 39, 27-32. Marks, J., Hagan, I. M., and Hyams, J. S. (1986). J . Cell Sci., Suppl. 5,229-241. McGillivray, A. M., and Gow,N. A. R. (1987). J. Gen. Microbiol. 133,2875-2881. McKerracher, L. J., and Heath, I. B. (1985).Profoplasma 125, 162-172. McKerracher, L. J., and Heath, I. B. (1986).Cell Moril. Cyroskel. 6 , 3 5 4 7 . McKerracher, L. J., and Heath, I. B. (1987). Exp. Mycol. 11,79-100. Meeusen, R. L., and Cande, W. 2. (1979). J . Cell Biol. 82,5765. Menzel, D., and Elsner-Menzel, C. (1989a). Bor. Acfu 102, 164-171. Menzel, D., and Elmer-Menzel, C. (1989b). Protoplasma 150, 1-8. Nanomura, Y., Katayama, E., and Ebashi, S. (1975). J. Biochem. (Tokyo) 78, 1101-1 104. Nolan, R. A., and Bal, A. K. (1974). J. Bucreriol. 117, 840-843. Novick, P.,and Botstein, D. (1985). Cell (Cambridge, Muss.) 40, 405416.

126

1. B. HEATH

Novick, P., Osmond, B. C., and Botstein. D. (1989). Generics 121, 659-674. Oakley, B. R. (1985). Con. J. Biochem. Cell Biol. 63,479488. Oakley, B. R., and Rinehan, J. E. (1985).J. Cell Biol. 101,2392-2397. mega-Perez, R.. and Turian, G. (1987). Cyrobios 49, 137-145. Picton, J. M., and Steer, M. W. (1981).J. Cell Sri. 49,261-272. Picton, J. M., and Steer, M. W. (1982).J Theor. Biol. 98, 15-20. Pierson, E. S. (1988). Sex. Plant Reprod. 1 , 8 3 4 7 . Pies, N. J., and Wohlfarth-Bottermann. K.-E.(1986). Eur. J. Cell Biol. 40, 139-149. Pollard, T. D. (1981).J . Biol. Chem. 256, 7666-7670.. Pollard, T. D., and Cooper,J. A. (1986). Annu. Rev. Biochem. 55,987-1035. Pollard, T. D., Selden, S. C., and Maupin. P. (1984). J . Cell Biol. 99.33s-37s. Poste, G. (1973). Methods Cell Biol. 7, 21 1-249. Rayle, D. L. (1989). Planru 178,92-95. Reinhardt, M. 0. (1892). Juhrb. Wiss.Bor. 23,479-566. Reiss, H.-D., and Herth, W. (1979). Pluntu 146,615421. Reissig, J. C., and Kinney, S. G. (1983). J. Bacreriol. 154, 1397-1402. Roberson, R. W., and Fuller, M.S. (1988). Proroplusmu 146, 143-149. Robertson, N. F. (1958). Ann. Bot. (London) [N.S.] 22, 159-173. Robinow, C. F. (1963).J. Cell Biol. 17, 123-152. Runeberg. P., Raudaskoski, M., and Virtanen, I. (1986). Eur. J. Cell Biol. 41,25-32. Salo, V., Niini, S. S., Virtanen, I., and Raudaskoski, M. (1989). J. Cell Sci. 94, 11-24. Sanger. J. M.,Mittal, B., Dome, J. S., and Sanger, J. W. (1989). CellMotil. Cyroskel. 14,201-219. Schindler. M., Meiners. S., and Cheresh, D. A. (1989). J . Cell Biol. 108, 1955-1965. Schmid, J.. and Harold, F. M.(1988). J. Gen. Mirrobiol. 134, 2623-2631. Schreurs, W. J. A., and Harofd, F. M. (1988). Proc. Nurl Acad. Sci. U.S.A. 85, 1543-1538. Seagull, R. W., and Heath, I. B. (1980). Protoplusmu 103, 231-240. Seagull, R. W., and Thomas, D. des S. (1976). 2. Pfanzenphysiol. 80,92-94. Sepsenwol. S.. Ris, H., and Roberts, T. M. (1989). J . Cell Biol. 108,5546. Sheetz, M. P. (1989). I n “Cell Movement. Kinesin Dynein and Microtubule Dynamics” (F. D. Warner and J. R. Mchtosh, eds.), Vol. 2, pp. 277-286. Alan R. Liss, New York. Sheetz, M. P.. Vale, R., Schnapp, B.. Schroer, T.,and Reese, T. (1986). J. Cell Sci., Suppl. 5, 181-1 88. Sloboda, R. D., and Gilbert, S. P. (1989). I n “Cell Movement. Kinesin. Dynein and Microtubule Dynamics” (F. D. Warner and J. R. Mclntosh eds.), Vol. 2, pp. 223-232. Alan R. Liss, New York. Sonobe, S.. and Shibaoka, H. (1989). Protoplusmu 148,80436. Staiger, C . J., and Schliwa, M. (1987). Proroplusmu 41, 1-12. Steer, M. W. (1990). I n “Tip Growth” (1. B. Heath, ed.), pp. 119-145. Academic Press, San Diego, California. Steer, M. W., and Steer, J. M. (1989). New Phyrol. 111, 323-358. Takeuchi, Y., Schmid, J., Caldwell, J. H., and Harold, F. M. (1988). J . Membr. Biol. 101, 3341. Tang, X.,Lancelle, S. A., and Hepler, P. K. (1989). Cell. Moril.Cyroskel. 12, 216224. Taylor, D. L., and Condeelis, J. S . (1979). Inr. Rev. Cyrol. 56, 57-144. Thomas, D. des S., Lutzac, M., and Manavathu, E. (1974). Nature (London) 249, 140-142. Togawa. T., and Bonner, J. (1957). Plunr Physiol. 32,207-212. Traas. J. A., Doonan, 1. H., Rawkins, D. J., Shaw. P.J., Watts, J., and Lloyd, C. W. (1987). J . Cell B i d . 105,387-395. Treves, S., Di Virgilio, F., Vaselli, G. M..and Pozzan, T. ( 1987). Exp. Cell Res. 168, 285-298. Tucker, B. E., Hoch, H. C., and Staples, R. C. (1986). Proroplusmu 135,88-101. Turian, G. (1978). Experienriu 34, 1277-1279.

ROLES OF ACTIN IN TIP GROWTH OF FUNGI

127

Turian, G. (1981). Bot. Helv.91, 161-168. Uyeda, T.Q. P., and Furuya, M. (1989). J. Cell B i d . 108, 1727-1735. Van Tuinen, D., Ortega-Perez, R., and Turian, G. (1986). Bor. Helv. 96,299-302. Virk, S . S., and Cleland, R. E. (1988). Planfa 1 7 6 , 6 M 7 . Volkmann, D. (1984). Planfa 162, 392-304. Wang, F.,Sampogna, R. V., and Ware, B. R. (1989). Biophys. J . 55,293-298. Watts, F. Z., Miller, D. M., and Orr, E. (1985). Nature (Londonj 316, 83-85. Watts, F. Z., Shiels, G., and On; E. (1987). EMBO J. 6,3499-3505. Wessels, J. G. H. (1986). Int. Rev. Cytol. 104, 37-79. Wessels, J. G. H. (1988). Acta Bot. Neerl. 37, 3-16. Wessels, J. G. H. (1990). In “Tip Growth” (I. B. Heath, ed.), pp. 1-29. Academic Press, San Diego, California. Williamson, R. E. (1984). Plant, Cell Environ. 7,431-440. Wilson, P., Fuller, E., and Forer, A. (1987). Biochern. Cell B i d . 65,376-385. Witman, G. B. (1989). In “Cell Movement. The Dynein ATPases” (F. D. Warner and J. R. Mclntosh, eds.), Vol. 1, pp. 25-36. Alan R. Liss, New York. Yin, H. L., Zoner, K. S., and Stossel, T. P. (1980). J. Biol. Chern. 225,9494-9500. Zalokar, M. (1959). Am. J . Bof. 46,602-610.