1. Gow N.A., Brown A.J., Odds F.C. Fungal morphogenesis and host invasion. Curr Opin Microbiol. 2002;5:366–371.[PubMed]
2•. Virag A., Harris S.D. The Spitzenkörper: a molecular perspective. Mycol Res. 2006;110:4–13.[PubMed]An excellent overview of the process of hyphal tip growth covering the main protein molecular complexes that are required for polarised tip growth.
3. Steinberg G. Hyphal growth: a tale of motors, lipids, and the Spitzenkörper. Eukaryot Cell. 2007;6:351–360.[PMC free article][PubMed]
4. Machesky L.M., Gould K.L. The Arp2/3 complex: a multifunctional actin organizer. Curr Opin Cell Biol. 1999;11:117–121.[PubMed]
5. Lipschutz J., Mostov K. Exocytosis: the many masters of the exocyst. Curr Biol. 2002;1:212–214.[PubMed]
6. Sudbery P., Court H. Polarised growth in fungi. In: Howard R.J., Gow N.A.R., editors. The Mycota VIII. edn 2. Springer-Verlag; 2007. pp. 137–166.
7•. Martin S.W., Konopka J.B. Lipid raft polarization contributes to hyphal growth in Candida albicans. Eukaryot Cell. 2004;3:675–684.[PMC free article][PubMed]Here the authors show the apical distribution of sterol-rich membranes in the germ tubes of C. albicans and that pharmacological disruption of sphingolipid or sterol biosynthesis caused abnormal hyphal development.
8. Pearson C.L., Xu K., Sharpless K.E., Harris S.D. MesA, a novel fungal protein required for the stabilization of polarity axes in Aspergillus nidulans. Mol Biol Cell. 2004;15:3658–3672.[PMC free article][PubMed]
9. Alvarez J.F., Douglas L.M., Konopka J. Sterol-rich plasma membrane domains in fungi. Eukaryot Cell. 2007;6:755–763.[PMC free article][PubMed]
10. Irazoqui J.E., Gladfelter A.S., Lew D.J. Cdc42p, GTP hydrolysis, and the cell's sense of direction. Cell Cycle. 2004;3:861–864.[PubMed]
11. Harris S.D., Read N.D., Roberson R.W., Shaw B., Seiler S., Plamann M., Momany M. Polarisome meets Spitzenkörper: microscopy, genetics and genomics coverage. Eukaryot Cell. 2005;4:225–229.[PMC free article][PubMed]
12. Steinberg G. On the move: endosomes in fungal growth and pathogenicity. Nat Rev Microbiol. 2007;5:309–316.[PubMed]
13•. Fischer R., Zekert N., Takeshita N. Polarized growth in fungi — interplay between the cytoskeleton, positional markers and membrane domains. Mol Microbiol. 2008;68:813–826.[PubMed]This is an excellent review of polarised growth in fungi, with particular emphasis on hyphal growth in Aspergillus.
14. Roca M.G., Artl J., Jeffree C.E., Read N.D. Cell biology of conidial anastomosis tubes in Neurospora crassa. Eukaryot Cell. 2005;4:911–919.[PMC free article][PubMed]
15. Roca M.G., Read N.D., Wheals A.E. The conidial anastomosis tubes in filamentous fungi. FEMS Microbiol Lett. 2005;249:191–198.[PubMed]
16. Gooday G.W., Adams D.J. Sex hormones and fungi. Adv Microb Physiol. 1993;34:69–145.[PubMed]
17. Daniels K.J., Srikantha T., Lockhart S.R., Pujol C., Soll D.R. Opaque cells signal white cells to form biofilms in Candida albicans. EMBO J. 2006;25:2240–2252.[PMC free article][PubMed]
18. Gooday G.W. Chemotaxis and chemotropism in fungi and algae. In: Carlile M.J., editor. Primitive Sensory and Communication Systems. Academic Press; 1975. pp. 155–204.
19. Jansson H.-B., Johansson T., Nordbring-Herts B., Tunlid A., Odham G. Chemotropic growth of germ tubes of Cochliobolus sativus to barley roots or root exudates. Trans Br Mycol Soc. 1988;90:647–650.
20. De Silva L., Youatt J., Gooday G.W., Gow N.A.R. Inwardly directed ionic currents of Allomyces macrogynus and other water moulds indicate sites of proton-driven nutrient transport but are incidental to tip growth. Mycol Res. 1992;96:925–931.
21•. Christensen M.J., Bennett R.J., Ansari H.A., Koga H., Johnson R.D., Bryan G.T., Simpson W.R., Koolaard J.P., Nickless E.M., Voisey C.R. Epichloë endophytes grow by intercalary hyphal extension in elongating grass leaves. Fungal Genet Biol. 2008;45:84–93.[PubMed]In this paper the authors demonstrate that plant epiphytes have to undergo intercalary growth in order to be able to be maintained within the growing meristematic regions of plants. This paper therefore points to a notable departure from the dogma of apical hyphal growth in fungi.
22. Rolke Y., Tudzynski P. The small GTPase Rac and the p21-activated kinase Cla4 in Claviceps purpurea: interaction and impact on polarity, development and pathogenicity. Mol Microbiol. 2008;68:405–423.[PubMed]
23. Allen E.A., Hoch H.C., Stavely J.R., Steadman J.R. Uniformity among races of Uromyces appendiculatus in response to topographical signalling for appressorium formation. Phytopathology. 1991;81:883–887.
24. Collins T.J., Read N.D. Appressorium induction by topographical signals from six cereal rusts. Physiol Mol Plant Pathol. 1997;51:169–179.
25. Gow N.A.R. Nonchemical signals used for host location and invasion by fungal pathogens. Trends Microbiol. 1993;1:45–50.[PubMed]
26. Zhou X.-L., Stumpf M.A., Hoch H.C., Kung C. A mechanosensitive channel in whole cells and membrane patches of the fungus Uromyces. Science. 1991;253:1415–1417.[PubMed]
27. Aoki A., Ito-Kuwa S., Nakamura K., Vidotta V., Takeo K. Oxygen as a possible tropic factor in hyphal growth of Candida albicans. Mycoscience. 1998;39:231–238.
28. Davies J.M., Stacey A.J., Gilligan C.A. Candida albicans hyphal invasion: thigmotropism or chemotropism? FEMS Microbiol Lett. 1999;171:245–249.[PubMed]
29••. Brand A., Vacharaksa A., Bendel C., Norton J., Haynes P., Henry-Stanley M., Wells C., Ross K., Gow N.A.R., Gale C.A. An internal polarity landmark is important for externally induced hyphal behaviors in Candida albicans. Eukaryot Cell. 2008;7:712–720.[PMC free article][PubMed]The authors show evidence for the internal polarity landmark proteins — the Bud1/Rsr1 Ras-GTPase and its GAP, Bud2, are required for the normal thigmotropic and galvanotropic orientation of germ tubes of C. albicans. It is also shown that mutants lacking these proteins are less able to penetrate into oral epithelial cells and kidneys of experimental animals.
30. Sherwood J., Gow N.A.R., Gooday G.W., Gregory D., Marshall D. Contact sensing in Candida albicans: a possible aid to epithelial penetration. J Med Vet Mycol. 1992;30:461–469.[PubMed]
31. Perera T.H.S., Gregory D.W., Marshall D., Gow N.A.R. Contact sensing in hyphae of dermatophytic and saprophytic fungi. J Med Vet Mycol. 1997;35:289–294.[PubMed]
32••. Brand A., Shanks S., Duncan V.M.S., Yang M., Mackenzie K., Gow N.A.R. Hyphal orientation of Candida albicans is regulated by a calcium-dependent mechanism. Curr Biol. 2007;17:347–352.[PMC free article][PubMed]Using a range of mutants in calcium transport systems this article shows that galvanotropism and thigmotropism of C. albicans requires the high and low affinity uptake mechanisms and components of the underlying calcium-signalling pathways.
33. Brasch J., Menz A. UV susceptibility and negative phototropism of dermatophytes. Mycoses. 1995;38:197–203.[PubMed]
34. Hutton R.D., Kerbs S., Yee K. Scanning electron microscopy of experimental Trichophyton mentagrophytes infections in guinea pig skin. Infect Immun. 1978;21:247–253.[PMC free article][PubMed]
35. Sherwood-Higham J., Zhu W.-Y., Devine C.A., Gooday G.W., Gow N.A.R., Gregory D.W. Helical growth of Candida albicans. J Med Vet Mycol. 1995;32:437–445.[PubMed]
36. Brand A., Lee K., Veses V., Gow N.A.R. Calcium homeostasis is required for contact-dependent helical and sinusoidal tip growth in Candida albicans hyphae. Mol Microbiol. 2009;71:1155–1164.[PMC free article][PubMed]
37. Crombie T., Gow N.A.R., Gooday G. Influence of applied electrical fields on yeast and hyphal growth of Candida albicans. J Gen Microbiol. 1990;136:311–317.[PubMed]
38. Casamayor A., Snyder M. Bud-site selection and cell polarity in budding yeast. Curr Opin Microbiol. 2002;5:179–186.[PubMed]
39. Herrero A.B., López M.C., Fernández-Lago L., Domínguez A. Candida albicans and Yarrowia lipolytica as alternative models for analysing pudding patterns and germ tube formation in dimorphic fungi. Microbiology. 1999;145:2727–2737.[PubMed]
40. Chaffin W.L. Site selection for bud and germ tube emergence in Candida albicans. J Gen Microbiol. 1984;130:431–440.
41•. Kumamoto C., Vinces M.D. Alternative Candida albicans lifestyles: growth on surfaces. Annu Rev Microbiol. 2005;59:113–133.[PubMed]Excellent supplementary information looking specifically at the role of mechanosensitive phenomena in fungal physiology, including aspects not covered in the present review on cell wall stress regulation and the electrophysiology of mechanosensitive ion channels.
42. McGillivray A.M., Gow N.A.R. Applied electrical fields polarize the growth of mycelial fungi. J Gen Microbiol. 1986;132:2515–2525.
43. Pu R., Robinson K.R. Cytoplasmic calcium gradients and calmodulin in the early development of the fucoid algs Pelvetia compressa. J Cell Sci. 1998;111:3197–3207.[PubMed]
44. Iwano M., Shiba H., Miwa T., Che F.S., Takayama S., Nagai T., Miyawaki A., Isogai A. Ca2+ dynamics in a pollen grain and papilla cell during pollination of Arabidopsis. Plant Physiol. 2004;136:3562–3571.[PMC free article][PubMed]
45. Lever M., Robertson B., Buchan A.D.B., Gooday G.W., Gow N.A.R. pH and Ca2+ dependent galvanotropism of filamentous fungi: implications and mechanisms. Mycol Res. 1994;98:301–306.
46. Kumamoto C. Molecular mechanisms of mechanosensing and their roles in fungal contact sensing. Nat Rev Microbiol. 2008;6:667–673.[PMC free article][PubMed]
47. Watts H.J., Véry A.-A., Perera T.H.S., Davies J.M., Gow N.A.R. Thigmotropism and stretch-activated channels in the pathogenic fungus Candida albicans. Microbiology. 1998;144:689–695.[PubMed]
48. Bowen A.D., Davidson F.A., Keatch R., Gadd G.M. Induction of contour sensing in Aspergillus niger by stress and its relevance to fungal growth mechanics and hyphal tip structure. Fungal Genet Biol. 2007;44:484–491.[PubMed]
49. Read N.D., Kellock L.K., Knight H., Trewavas A.J. Contact sensing during infection by fungal pathogens. In: Callow J.A., Green J.R., editors. Vol. 48. Cambridge University Press; 1992. pp. 137–172. (Perspectives in Plant Cell Recognition).
50. Hausauer D.L., Gerami-Nejad M., Kistler-Anderson C., Gale C.A. Hyphal guidance and invasive growth in Candida albicans require the Ras-like GTPase Rsr1p and its GTPase-activating protein Bud2p. Eukaryot Cell. 2005;4:1273–1286.[PMC free article][PubMed]
51. Steinberg G. Tracks for traffic: microtubules in the plant pathogen Ustilago maydis. New Phytol. 2007;174:721–733.[PubMed]
52••. Takeshita N., Higashitsuji Y., Konzack S., Fischer R. Apical sterol-rich membranes are essential for localizing cell end markers that determine growth directionality in the filamentous fungus Aspergillus nidulans. Mol Biol Cell. 2008;19:339–351.[PMC free article][PubMed]The authors characterise the roles of two cell-end marker proteins, TeaA and TeaR, and the CENP-E kinesin KipA in hyphal orientation and in microtubule architecture. The Tea mutants have meandering or zig-zag hyphae. KipA is shown to be important for the localisation of TeaA and TeaR, but not for their cytoplasmic transport. In addition they present evidence that the sterol-rich apical membrane domain is vital for the localisation of the Tea proteins and for hyphal polarity.
53. Higashitsuji Y, Herrero S, Takeshita N, Fischer R: The cell end marker protein TeaC determines growth directionality and is involved in cell septation inAspergillus nidulans.Eukaryot Cell 2009, doi:10.1128/EC.00251-08, in press. [PMC free article][PubMed]
54. Wu Q., Sandrock T.M., Turgeon B.G., Yoder O.C., Wirsel S.G., Aist J.R. A fungal kinesin required for organelle motility, hyphal growth, and morphogenesis. Mol Biol Cell. 1998;9:89–101.[PMC free article][PubMed]
55. Konzack S., Rischitor P.E., Enke C., Fischer R. The role of the kinesin motor KipA in microtubule organization and polarised growth of Aspergillus nidulans. Mol Biol Cell. 2005;16:497–506.[PMC free article][PubMed]
56. Côte P., Whiteway M. The role of Candida albicans FAR1 in regulation of pheromone-mediated mating, gene expression and cell cycle arrest. Mol Microbiol. 2008;68:392–404.[PubMed]
Microtubules Are Essential for Rapid Hyphal Growth
Our data demonstrate that the cytoplasmic microtubule array is necessary for rapid hyphal growth of A. nidulans. By observing the microtubules in living cells, we were able to follow the events that occurred in rapidly growing hyphae in response to the addition of the antimicrotubule agent benomyl. For the first few minutes after the addition of benomyl, the cytoplasmic microtubules remained intact, presumably because this period of time was required for benomyl to diffuse through the medium and penetrate into cells. The microtubule array then disassembled rapidly and was gone ≈6 min after the addition of benomyl. Tip growth initially continued at an unreduced rate after microtubule disassembly but slowed down rapidly, reaching a steady, slower rate ∼11 min after benomyl addition. It should be noted, however, that although microtubules are essential for rapid tip growth, they are not absolutely required for polarized growth. Tip growth continued slowly even when microtubules were depolymerized, and it has been shown previously that conidia germinate and polarized tip growth occurs in concentrations of benomyl that completely inhibit microtubule assembly (Oakley and Morris, 1980).
The present and previous data suggest a model for the roles of the microtubule and actin cytoskeletons in tip growth. We suggest that the main function of microtubules in tip growth is to transport vesicles containing cell wall materials to the vicinity of the hyphal tip. This transport system is not essential for tip growth but is certainly rate limiting for the growth of hyphal tip cells. The vesicles transported by the microtubule cytoskeleton are captured by a system involving actin and the myoA myosin, and this system is required for the fusion of these vesicles into the membrane at the growing tip. This acto-myosin system is essential for tip growth under all circumstances. When microtubules are disassembled by benomyl, there is initially enough material in the tip vicinity to allow rapid growth, but these materials become depleted quickly in the absence of microtubules and growth slows dramatically. The residual, slow growth that persists afterward is presumably due to wall materials reaching the tip area by nonmicrotubule-dependent mechanisms such as Brownian motion. In germlings, tip growth is slow (less than twice the growth rate in benomyl-treated tip cells), and, consequently, microtubule-based movement of wall materials is relatively unimportant for tip growth.
Our data are also consistent with the possibility that microtubules play a secondary role in focusing actin at the tip of growing hyphae. This would explain why tips bulge slightly after benomyl addition (Figure 2). It also would explain the larger bulges that are often seen when benomyl is washed out. By our model, when microtubules reassemble, they transport wall materials to the vicinity of the tips. These are captured by the poorly focused actomyosin system and used to synthesize new wall, and this creates a larger bulge. However, the microtubules quickly focus the actomyosin system and normal growth is resumed. It has previously been shown that microtubules are required for the apical localization of actin in Candida albicans germlings (Akashi et al., 1994); Torralba et al., (1998a) have reported, however, that treatment of A. nidulans hyphae with 0.25 μg/ml antimicrotubule agent methyl benzimidazole-2-yl carbamate actually causes actin to move closer to the hyphal tip. However, it should be noted that microtubules were not disassembled completely under these conditions, and, consequently, the role of microtubules in the localization of actin to hyphal tips in A. nidulans remains unresolved.
The role of cytoskeletal elements in tip-growing systems has been studied in a variety of organisms. In all cases, the actin cytoskeleton is absolutely required for tip extension. In growing neurites, microtubules are also essential (reviewed by Gordon-Weeks, 1991; Dent and Gertler, 2003). In plants, the role of microtubules has been evaluated in root hair cell growth and pollen tube extension. The growth rate of root hair cells was not significantly affected by the presence of antimicrotubule agents, but the cells no longer grew straight and started to exhibit waviness and branching (Bibikova et al., 1999; Ketelaar et al., 2003). In angiosperm pollen tubes, microtubules are essential for tip elongation and, in addition, loss of microtubules leads to tip swelling and bifurcation (Anderhag et al., 2000; Justus et al., 2004). In angiosperms, the available data suggest that microtubules are relatively unimportant in pollen tube tip growth (Heslop-Harrison et al., 1988; Ålström et al., 1995), although it would be useful to repeat these experiments with current techniques and reagents. These observations, and our data, suggest that the importance of microtubule-dependent transport varies among tip growth systems. In some systems it is critical but in others it is relatively unimportant. These data also suggest that microtubules often play a role in positioning and organizing the site of tip growth.
A Remarkable Spatial Regulation of Microtubule Disassembly Facilitates Tip Growth during Mitosis
We have confirmed the finding of Riquelme et al. (2003) that hyphal tip growth is continuous during mitosis. Our data also resolve a paradox with the Riquelme et al. (2003) data. If cytoplasmic microtubules are required for rapid tip growth as we have shown, and they disassemble in mitosis as reported previously (Oakley et al., 1990; Jung et al., 2001), the obvious expectation is that tip growth would slow during or shortly after mitosis. We have found, however, that, in hyphal tip cells, microtubules near the cell apex generally do not depolymerize completely during mitosis (although they do depolymerize completely in germlings as previously reported). We hypothesize that the remnant microtubules transport adequate amounts of cell tip precursors to the vicinity of the tip to allow continuous growth. It also should be noted that enough cell wall precursors are normally at the tip to allow growth for some time after microtubule disassembly by benomyl, so the remnant microtubules do not need to be as efficient in supplying cell wall precursors as the interphase microtubule cytoskeleton. They only need to supply enough materials to tide the growing tip over until the interphase microtubule cytoskeleton reassembles at the end of mitosis.
The maintenance of microtubules near the hyphal tip during mitosis constitutes a remarkable spatial regulation of microtubule disassembly. Cytoplasmic microtubules disassemble rapidly throughout most of the cytoplasm but, in the same cell, they remain intact near the hyphal tip. Cytoplasmic microtubules disassemble in many types of cells as they enter mitosis, and the regulation of microtubule assembly and disassembly is generally thought to involve a balance between the activities of proteins that stabilize microtubules (e.g., microtubule-associated proteins) and proteins that promote microtubule disassembly (catastrophe factors). Because relatively little is known about microtubule stabilizers and catastrophe factors in A. nidulans, it is premature to speculate on the mechanism by which disassembly is inhibited near the hyphal tip. Whatever the mechanism, it would seem to require a system for sensing tip proximity (e.g., release of factors by the growing tip), and the tip proximity signal must act directly or indirectly on the factors that regulate microtubule disassembly. Because the regulation occurs over a very short time span, it is more likely to involve protein modification than protein synthesis.
Rapid Tip Growth, Asymmetric Division, and the Coenocytic Growth Form
We have found that hyphal tip cells grow 5.4 times as rapidly as germlings. It is obviously advantageous for tip growth to be rapid because it allows the organism to find new sources of nutrients quickly, but why are hyphal tip cells able to grow so much more rapidly than germlings? We suggest that a part of the answer has to do with the ratio of nuclei to cytoplasmic volume. It is likely that there is an optimal nucleus-to-cytoplasm ratio (Clutterbuck, 1969; Fiddy and Trinci, 1976). Certainly, some variation in the nucleus-to-cytoplasm volume ratio will be tolerated, and this ratio might vary to some extent in different growth conditions. However, the general principle must apply that tip growth cannot significantly outpace nuclear division because the nucleus-to-cytoplasm ratio would decrease and the nuclei would eventually be unable to supply the cytoplasm with mRNAs sufficient to support maximum growth. Because the diameters of hyphae are relatively consistent under a given set of growth conditions, it follows that if there is an optimal nucleus-to-cytoplasm ratio, there must be an optimal hyphal length per nucleus. Fiddy and Trinci (1976) have found that for diploid hyphae such as the ones we have used for our experiments, the average hyphal length per nucleus is 22 ± 7 μm. Thus, binucleate germlings would average 44 μm in length, and such a germling would grow in one cell cycle to be a germling with four nuclei and an average length of 88 μm (i.e., would grow 44 μm in one cell cycle). A tip cell with 10 nuclei would be ∼220 μm in length and in one cell cycle it would become a cell with 20 nuclei and a length of 440 μm. It would, thus, grow 220 μm in one cell cycle (5 times as much as a binucleate germling) with no alteration in the nucleus to cytoplasm ratio.
We suggest that maintaining an appropriate nucleus-to-cytoplasm ratio while maintaining a maximal growth rate is an important reason that A. nidulans cells divide asymmetrically and that A. nidulans has a coenocytic growth form. When A. nidulans spores germinate, three to four rounds of nuclear division occur before the first septum forms (Fiddy and Trinci, 1976). The septum divides the cell asymmetrically creating a tip cell that contains the majority of the nuclei. Additional rounds of asymmetric septation follow successive rounds of nuclear division and these eventually result in multinucleate tip cells. We suggest having many nuclei in the tip cell is necessary for maximal tip growth and that the delay of septation until at least the third nuclear division coupled with asymmetric division at this and subsequent stages are adaptations that allow tip cells to have many nuclei. We also note that A. nidulans must have mechanisms that govern the transition from slow germling growth to rapid growth of hyphal tip cells.