The germ tubes of these fungi exhibit contact sensing and orientation in relation to the topography of the epithelial substratum thigmotropism. Initially, hyphae grow perpendicularly across the plane of axis of the epithelial cells — a strategy that is thought to aid the location of stoma, which in monocotyledons are often positioned in staggered rows. When the germ tube encounters a guard cell of a specific lip height, an appressorium is induced thigmodifferentiation [ 23 ].
Elegant experiments with chemically inert plastic replica surfaces demonstrated that these events are mediated entirely by the topography of the plant surface and not by any chemical gradients [ 23—25 ]. In dicotylenous plants, the stoma occur within a mosaic of cortical cells on the plant surface and certain fungi follow the interstices between cells in order to locate them.
In Uromyces , stretch-activated mechanosensitive channels have been studied [ 26 ] that may act as transducers of topographical information for thigmotropic growth see below.
Tropic growth of fungi within plant tissues. Courtesy of C Voisey. Courtesy of P Tudzynski, with permission from Blackwell. Tropic growth of hyphae of fungal pathogens has been reported for mating interactions for those few species that have recognised sexual cycles Figure 2 [ 16,17 ].
Hyphal aerotropism has also been reported for C. The role of tropic hyphal growth in the penetration of mammalian tissue has been debated [ 28 ] but recent work has shown that C. This lack of penetration also correlated with reduced virulence in an in vivo model of systemic infection, suggesting that normal regulation of tip orientation is required for penetrative growth.
Thigmotropic responses of C. It has been suggested that some dermatophytes are negatively phototropic [ 33 ]. Trichophyton metagrophytes hyphae penetrate through the stratum corneum to locate and colonise hair follicles [ 34 ], but the sensing mechanism by which this targeting is achieved is not known.
The tropic responses of C. Hyphae are oriented randomly in controls g but are cathodally orientated in an applied electric field h direction of the cathode is shown by arrows. If the field polarity is reversed after the establishment of cathodal growth, tips re-orient towards the new cathode position i.
Hyphae sense obstacles such as ridges in the substratum and re-orient tip growth k and l. Sinusoidal and helical growth of hyphae of C. Hyphae of C. The galvanotropic, thigmotropic and sinusoidal orientation responses of C. Whilst the response to environmental cues is likely to be fungus-specific, tip re-orientation may be achieved by the modulation of the conserved machinery that sustains polarised hyphal growth. Recent studies in C. Hyphae normally grow in reasonably straight trajectories; however, reports have shown circumstances in which hyphae meander or exhibit sinusoidal growth.
For example, hyphae of the human pathogen C. Meandering growth in mutants with affected microtubule organisation and polarised actin assembly have also been described see below. These studies suggest that the mechanism of tip orientation is distinguishable from the mechanism that determines apical extension and cell polarity, and they point to important roles for calcium signalling, Ras-type GTPase signalling modules, microtubule plus end organisation and for sterol-rich lipid rafts in regulating hyphal organisation.
Germ tube growth of C. In this fungus a variety of orientation responses have been shown to be dependent on calcium influx and homeostasis. The site of polarity in Saccharomyces cerevisiae is ploidy dependent and is determined by positional markers encoded by BUD genes that position buds at sites either adjacent to or opposite the site of bud scars Figure 3 [ 38 ].
Hypha orientation is, therefore, open to the influence of external cues Figure 3. Directionality of growth is determined by at least three signalling pathways in C. The site of emergence can be influenced by the application of an external electric field [ 42 ], which, in C. Fungal tips are increasingly insensitive to small obstacles in the substrate as the angle of approach steepens [ 47,48 ].
It has been suggested that touch-sensing may not operate efficiently at the extreme apex of the apical dome due to its semi-fluid nature [ 48 ]. However, thigmotropic hyphae are tightly pressed to the substratum and typically have a nose-down morphology [ 48,49 ]. Since mechanosensitive signals must ultimately be translated into changes in the site of vesicle fusion in order to influence thigmotropic hypha orientation, it seems most likely that the tip orientation machinery must also reside within the apical dome.
These signalling proteins ultimately serve to localise the recruitment of actin, and therefore, the vectorial secretion of vesicles to points of growth. Ultimately, it seems likely that there may be core elements that translate signals concerned with vegetative hyphal growth, bud site selection and the growth of mating projections of C. Tea proteins are also involved in the recruitment of formin proteins, such as SepA, that catalyse the polarisation of the actin cytoskeleton at the apex.
TeaC requires microtubules, but not KipA, for its localisation at hyphal tips and at septa [ 53 ]. Mutants in TeaC also exhibited zig-zag growth and were affected in septation. Therefore, the TeaA and TeaC proteins couple the microtubule and actin-based vesicle delivery systems involved in hypha orientation. However, under these conditions zig-zag growth did not occur, suggesting that disruption of the apical sterol-rich domain had multiple effects on hyphal polarity.
The control of directionality in hyphae, therefore, implicates both microtubules in delivering vesicles and localising parts of the actin-dependent vesicle docking system and localised calcium ion uptake and signalling via Ras-type GTPase modules that form part of the polarisome complex within a sterol-rich apical domain Figure 4. Components of the hyphal apex involved in orientation of the growth axis. Directionality involves calcium signalling through the Mid1—Cch1 channel complex, GTP—GDP cycling of the Ras-GTPase, Rsr1, delivery of specific cargo by kinesin motor proteins and proteins that tether microtubule plus ends to complexes in the apical sterol-rich domain.
Vesicles finally are delivered to the membrane in the apex via actinomyosin. The spatial localisation of the KipA kinesin and Tea cell-end proteins in A. The ability to orient the axis of growth of hyphae is a vital aspect of the physiology of fungal cells. Recent work has demonstrated that the molecular machinery that regulates hypha orientation can be distinguished from that inducing polarised growth since mutants and growth conditions can be generated, in which the trajectory of fungal hyphae is influenced without blocking the ability of hyphae to undergo apical growth.
This orientation apparatus apparently involves calcium signalling, GTPase signalling modules and protein complexes that orchestrate sites of actin recruitment and microtubule-tethering at the hyphal apex. These components orchestrate the growth, morphogenesis and various lifestyles of filamentous fungi.
Papers of particular interest, published within the period of review, have been highlighted as:. National Center for Biotechnology Information , U. Sponsored Document from. Current Opinion in Microbiology.
Curr Opin Microbiol. Author information Copyright and License information Disclaimer. Alexandra Brand: ku. This article has been cited by other articles in PMC. Abstract Hypha orientation is an essential aspect of polarised growth and the morphogenesis, spatial ecology and pathogenesis of fungi.
Introduction Most fungi are sessile filamentous organisms that grow by extending the tips of hyphae to form an expanding mycelial network. Tropisms in plant pathogens and saprophytes Tropic alignment of hyphae plays both general and specific roles in the growth of mycelial fungi.
Open in a separate window. Figure 1. Tropisms in human pathogens Tropic growth of hyphae of fungal pathogens has been reported for mating interactions for those few species that have recognised sexual cycles Figure 2 [ 16,17 ]. Figure 2. Molecular mechanisms Whilst the response to environmental cues is likely to be fungus-specific, tip re-orientation may be achieved by the modulation of the conserved machinery that sustains polarised hyphal growth.
Candida albicans Germ tube growth of C. Figure 3. Figure 4. Conclusions The ability to orient the axis of growth of hyphae is a vital aspect of the physiology of fungal cells. References 1. Gow N. Fungal morphogenesis and host invasion. Virag A. Schaller, and B. McKenzie, U. Koser, L. Lewis et al. Mennink-Kersten, J. Donnelly, and P. Brand, A.
Vacharaksa, C. Bendel et al. Harriott, E. Lilly, T. Rodriguez, P. Fidel, and M. Wilson, K. Haedicke, F. Dalle, and B. Mowat, C. Williams, B. Jones, S. McChlery, and G. S—S, Baillie and J. View at: Google Scholar M. Al-Fattani and L. Paramonova, B. Krom, H. Busscher, and P. Ghannoum, R. Jurevic, P. Mukherjee et al. Jarosz, D. Deng, H. Crielaard, and B. Bamford, A.
D'Mello, A. Nobbs, L. Dutton, M. Vickerman, and H. Han, R. Cannon, and S. Hornby, E. Jensen, A. Lisec et al. Chen, M. Fujita, Q. Feng, J. Clardy, and G. Dalle, B. L'Ollivier et al. Lorenz, J. Bender, and G. Phan, C. Myers, Y. Fu et al. Phan, P. Belanger, and S. Hoyer and J. Wasylnka, A. Hissen, A. Wan, and M. S27—S30, Lopes Bezerra and S. Feldmesser, S. Tucker, and A. Nicola, E. Robertson, P. Albuquerque, L. View at: Google Scholar N. Gow and G.
Luo, A. Ibrahim, B. Spellberg, C. Nobile, A. Mitchell, and Y. Staab, S. Bradway, P. Fidel, and P. View at: Google Scholar U. Zeidler, T. Lettner, C. Lassnig et al. Banerjee, D. Thompson, A. Lazzell et al. Zheng, Y. Wang, and Y. Brown Jr. Giusani, X. Chen, and C. View at: Google Scholar C.
Nobile and A. Dwivedi, A. Thompson, Z. Xie et al. Money and F. Aguilar-Uscanga and J. Persat, N. Noirey, J. Diana et al. Staib, C. Zaugg, B. Mignon et al. Mora-Montes, M. Netea, G. Ferwerda et al. Hohl, H. Van Epps, A. Rivera et al.
Aimanianda, J. Bayry, S. Bozza et al. Moyes, M. Runglall, C. Murciano et al. Moyes, C. Murciano, M. Runglall, A. Kohli, A. Islam, and J. In press. Gersuk, D. Underhill, L. Zhu, and K. View at: Google Scholar S. Cheng, F. Lenardon et al. Padilla, G. Langsley, and D. Hise, J. Tomalka, S. Ganesan et al. Ouyang, J. Kolls, and Y. Gow, T. Perera, J. Sherwood-Higham et al. Hoch, R. Staples, B. Whitehead, J. Comeau, and E. Kradin and E. Okuda, M. Ito, and Y.
View at: Google Scholar T. Kanbe and K. Watts, A. Davies, and N. Brand, S. Shanks, V. Duncan, M. Yang, K. Mackenzie, and N. Chen, A. Brand, E. Morrison et al. Lockhart, R. Zhao, K. Daniels, and D. Aoki, S. Ito-Kuwa, K. Colot, H. A high-throughput gene knockout procedure for Neurospora reveals functions for multiple transcription factors.
Phenotype of a mechanosensitive channel mutant, mid-1 , in a filamentous fungus, Neurospora crassa. Cell 7 , — Ptk2 contributes to osmoadaptation in the filamentous fungus Neurospora crassa.
Kaminskyj, S. The relation between turgor and tip growth in Saprolegnia ferax : turgor is necessary, but not sufficient to explain apical extension rates. Harold, R. Growth and morphogenesis in Saprolegnia ferax : is turgor required? Walker, S. Invasive hyphal growth: an F-actin depleted zone is associated with invasive hyphae of the oomycetes Achlya bisexualis and Phytophthora cinnamomi.
Emerson, S. Slime, a plasmodial variant of Neurospora crassa. Genetica 34 , — Perkins, D. Chromosomal loci of Neurospora crassa. Leal-Morales, C. Alterations in the biosynthesis of chitin and glucan in the slime mutant of Neurospora crassa. Heath, I. Mechanisms of hyphal tip growth: tube dwelling amebae revisited. An alternative to a simple version of turgor-driven growth, emphasizing the role of the protoplast within the cell wall.
Mass flow and pressure-driven hyphal extension in Neurospora crassa. The direct verification of mass flow in hyphal networks, and details of the magnitude of pressure gradients that are required to cause mass flow in a low-Reynolds-number environment. Robertson, N. Some observations on the water-relations of the hyphae of Neurospora crassa. Measurement of hyphal turgor.
Measurement of turgor of the oomycete S. The methods would be the same for fungi. This article gives a good introduction to the thermodynamics of water potentials, experimental methodologies and experimental pitfalls. Ng, S. A tether for Woronin body inheritance is associated with evolutionary variation in organelle positioning. PLoS Genet. Plamann, M. Cytoplasmic streaming in Neurospora : disperse the plug to increase the flow?
Heaton, K. Growth-induced mass flows in fungal networks. B , — Sugden, K. Model of hyphal tip growth involving microtubule-based transport. E Stat. Soft Matter Phys. Boudaoud, A. Growth of walled cells: from shells to vesicles. Freitag, M. GFP as a tool to analyze the organization, dynamics and function of nuclei and microtubules in Neurospora crassa. Nelson, G. Calcium measurement in living filamentous fungi expressing codon-optimized aequorin.
Hickey, P. Live-cell imaging of filamentous fungi using vital fluorescent dyes. Methods Microbiol. Virag, A. A mutation in the Neurospora crassa actin gene results in multiple defects in tip growth and branching. Lee, I. Null mutants of the Neurospora actin-related protein 1 pointed-end complex show distinct phenotypes. Cell 12 , — The effects of ropy-1 mutation on cytoplasmic organization and intracellular motility in mature hyphae of Neurospora crassa.
Mourino-Perez, R. Microtubule dynamics and organization during hyphal growth and branching in Neurospora crassa.
Ramos-Garcia, S. Cytoplasmic bulk flow propels nuclei in mature hyphae of Neurospora crassa. Cell 8 , — Roca, M. Nuclear dynamics, mitosis, and the cytoskeleton during the early stages of colony initiation in Neurospora crassa.
Cell 9 , — Tucker, E. Cytoplasmic streaming does not drive intercellular passage in staminal hairs of Setcreasea purpurea. Suei, S. An F-actin-depleted zone is present at the hyphal tip of invasive hyphae of Neurospora crassa.
Visualization of F-actin localization and dynamics with live cell markers in Neurospora crassa. Harris, S. Cell 4 , — Gow, N. Transhyphal electrical currents in fungi. McGillviray, A. The transhyphal electrical current of Neurospora crassa is carried principally by protons. Takeuchi, Y. Transcellular ion currents and extension of Neurospors crassa hyphae. Applied electrical fields polarize the growth of mycelial fungi. Lever, M. Burstaller, W. Transport of small ions and molecules through the plasma membrane of filamentous fungi.
Live imaging of the secretory pathway in hyphae of Neurospora crassa. Slayman, C. Measurement of membrane potentials in Neurospora. Potapova, T. Transcellular ionic currents studied by intracellular potential recordings in Neurospora crassa hyphae. Transfer of energy from proximal to apical cells. FEBS Lett. Zalokar, M. Growth and differentiation of Neurospora crassa. Brody, J. Biotechnology at low Reynolds number.
Microfluidics at low Reynolds number is explored in detail from a physical veiwpoint, pertinent to the nature of mass flow in hyphae.
Berg, H. Physics of chemoreception. Short, M. Flows driven by flagella of multicellular organisms enhance long-range molecular transport. USA , — Goldstein, R. Microfluidics of cytoplasmic streaming and its implications for intracellular transport. Van de Meent, J. Analysis of the dynamic and steady-state responses of growth rate and turgor pressure to changes in cell parameters.
A detailed analysis of the nature of the interplay between turgor and cell expansion during cell growth. This analysis is pertinent to any walled cells that rely on turgor for growth.
Bok, J. Structure and function analysis of the calcium-related gene spray in Neurospora crassa. Sanders, D. Acta , 67—76 Schloemer, R. Nitrate transport system in Neuropora crassa. Blatt, M. Versaw, W. Repressible cation—phosphate symporter in Neurospora crassa. USA 92 , — Depolarization of the plasma membrane of Neurospora during active transport of glucose: evidence for a proton-dependent cotransport system.
USA 71 , — Download references. The author thanks past and present members of the Lew laboratory, where research is funded by the Natural Sciences and Engineering Research Council of Canada. You can also search for this author in PubMed Google Scholar.
0コメント