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Apical growth

The Microbial World:

Fungal tip growth and hyphal tropisms

Produced by Jim Deacon
Institute of Cell and Molecular Biology, The University of Edinburgh

Fungal tip growth and hyphal tropisms

Fungal hyphae extend continuously at their extreme tips, where enzymes are released into the environment and where new wall materials are synthesised. The rate of tip extension can be extremely rapid - up to 40 micrometres per minute. It is supported by the continuous movement of materials into the tip from older regions of the hyphae. So, in effect, a fungal hypha is a continuously moving mass of protoplasm in a continuously extending tube.

This unique mode of growth - apical growth - is the hallmark of fungi, and it accounts for much of their environmental and economic significance.

Apical growth enables fungi to extend into fresh zones of substrate.

This is especially important for growth on complex, insoluble polymers such as cellulose, because the enzymes released from hyphae have a limited rate of diffusion. These enzymes create zones of substrate erosion around the hyphae, but the tips can grow continuously out of these nutrient-exhausted zones. For this reason, fungi are the major decomposers of organic matter (see wood decay fungi) and also are efficient in capturing mineral nutrients in mycorrhizal associations. In contrast to hyphae, the fungi that typically grow as yeasts seldom have polymer-degrading enzymes because yeasts cannot extend continuously into new zones of substrate. Instead, they form buds for dispersal in water films - an ideal adaptation for utilising simple, soluble nutrients.

Apical growth gives penetrating power. Hyphal tips can penetrate plant cell walls and insect cuticle, making fungi important as plant and insect pathogens, and as the major degraders of physically hard materials such as wood.

In the sections below, we consider:

  • the mechanisms of hyphal tip growth,
  • the ways in which hyphae orientate their growth (hyphal tropism) to find nutrients, to parasitise their hosts, to find their partners or to disperse their spores.

Mechanism of apical growth

The ultrastructural organisation of fungal hyphae provides an important clue to the mechanism of apical growth (Figures A-C). The extreme tip of a growing hypha has very few organelles; instead, it contains a body termed the Spitzenkörper (apical body). This consists of a cluster of small, membrane-bound vesicles embedded in a meshwork of actin microfilaments.

Figures A-C. Ultrastructure of the hyphae of Sclerotium rolfsii, prepared for electron microscopy by freeze substitution. From photographs supplied by Dr Robby Roberson (see Roberson & Fuller, 1988).

Fig. A. Young region of a hypha, showing progressive changes in ultrastructural organisation behind the hyphal apex. The apex contains a Spitzenkörper (S). Behind this is a zone rich in mitochondria (M, the dark tubular structures), then a zone containing tubular vacuoles (light coloured) and nuclei (N).

Fig. B. Part of a mature region of a hypha (the apex, not shown, is towards the right of the image) showing mitochondria (M), vacuoles (Va), Golgi bodies (G, seen as dark, ring-like structures) and longitudinally running microtubules (MT).

Fig. C. Close-up of the Spitzenkörper - an accumulation of small, membrane-bound vesicles of different sizes and contents, surrounding a central, vesicle-free core. The hyphal plasma membrane is seen as a thin dark line immediately to the right of the Spitzenkörper; two thin wall layers are seen outside the plasma membrane.

The importance of the Spitzenkörper was recognised long before the advent of electron microscopy because it can be seen by phase-contrast microscopy of hyphal tips. It is always present in growing tips, disappears when growth stops, reappears when growth restarts, and its position within the apex changes when hyphae change direction.

Although much remains to be learned about the mechanism of apical growth, we can summarise the process in a simple model, shown in the diagram below.

Simplified model of apical growth of fungal hyphae. G = Golgi body; V = vesicle; M = microtubule. From Deacon (1997).

  • The apical vesicles that make up the Spitzenkörper are thought to be produced from Golgi bodies and then transported to the tip by elements of the cytoskeleton - perhaps the microtubules, actin microfilaments and motor proteins like myosin.

  • The vesicles fuse with the plasma membrane at the tip, and release their contents. These contents almost certainly differ in the different types of vesicle, but are thought to include:

    • enzymes involved in wall synthesis,

    • enzymes involved in wall lysis,

    • enzyme activators,

    • some preformed wall polymers such as mannoproteins, although most wall polymers are synthesised in situ at the tip.

  • The wall is thin and thought to be structurally weak at the extreme tip, enabling new wall materials to be inserted. So the structural integrity of the hyphal tip might depend on the "actin cap" - a meshwork of actin microfilaments. The wall is strengthened progressively behind the apex by cross-linking of wall polymers.

The diagram below shows the process of wall growth in slightly more detail. For example, it is known that the main enzymes that synthesise fungal walls are chitin synthase and glucan synthetase. These enzymes are likely to be delivered to the tip in membrane-bound vesicles or vesicle-like bodies, and they are known to be inserted into the plasma membrane as integral membrane proteins.

Both of these enzymes need to be activated - glucan synthetase by guanosine triphosphate (GTP) and chitin synthase by a protease which probably arrives in another type of vesicle. They receive wall substrates from the cytosol at their inner face, and they extrude the synthesised wall polymers (chitin or glucan chains) at their outer face so that these polymers enter the wall zone.

Simplified model of wall growth at the hyphal apex. From Deacon (1997).

Clearly, the process of tip growth is extremely complex, with many coordinated components. Any change in the balance of these components could alter the shape of the tip or the direction of growth. An example of this is the
deformation of tip growth shown in The Fungal Web. In the next section we see how fungi change their direction of growth in response to environmental signals.

Further reading


JW Deacon (1997) Modern Mycology. Third edn. Blackwell Science, Oxford.

NAR Gow & GM Gadd (1994) The Growing Fungus. Chapman & Hall, London.

Research paper:

RW Roberson & MF Fuller (1988) Ultrastructural aspects of the hyphal tip of Sclerotium rolfsii preserved by freeze substitution. Protoplasma 146, 143-149.


There are many excellent reviews of fungal tip growth, including:

GW Gooday (1995) The dynamics of hyphal growth. Mycological Research 99, 385-394.

SL Jackson & IB Heath (1993) Roles of calcium ions in hyphal tip growth. Microbiological Reviews 57, 367-382.

S Bartnicki-Garcia et al. (1995) Determinants of fungal cell wall morphology: the vesicle-supply center. Canadian Journal of Botany 73, S372-378.


Hyphal tropisms

A tropism is an orientation response of a hypha to an external stimulus. An example is shown in Figure D, where encysted zoospores of Pythium aphanidermatum (Oomycota) were allowed to germinate in water, with a nutrient-rich agar block positioned on one side of the cyst clusters. Zoospore cysts of the oomycota have a pre-determined point of germination, so the young hyphae emerged in all directions but then they reorientated and grew towards the nutrient-rich agar (to the right in the images below).

This type of tropism to organic nutrients is found in several members of the fungus-like group oomycota, including the "water moulds" such as Saprolegnia and Achlya species, some of which parasitise freshwater fish. However, tropism to organic nutrients does not seem to occur in the true (chitin-walled) fungi. Instead, these have other forms of tropism, discussed below.

Figure D. Young hyphae (germ tubes) growing from zoospore cysts of Pythium aphanidermatum, seen at different magnifications in the two images. The cysts germinated from a predetermined (fixed) point but the hyphae then grew towards a nutrient-rich agar block (malt extract and peptone) to the right-hand side (not shown). The cells were stained with the fluorescent brightener, Calcofluor, and photographed with a fluorescence microscope, using near-UV illumination.

Spore tropisms

Unlike the zoospore cysts mentioned above, the spores of most fungi do not have a fixed point of germination. Instead they can germinate from almost any point, which can be influenced by external factors. A spectacular example of this is shown in Figures E-G, where spores of Idriella bolleyi (deuteromycota) were sprayed onto the roots of young, aseptic wheat seedlings growing on a thin film of water agar.

The spores on the surface of living root hairs were almost always seen to germinate away from the root hair, and the germ tubes continued to grow away (see the two spores labelled "s" in Fig. E). In contrast, spores on the surface of dead root hairs germinated towards them, and the germ tubes coiled round the root hairs (Figure F) and penetrated them (Fig. G, arrowhead).

Figures E, F. Spores (s) of Idriella bolleyi germinating away from living cereal root hairs (E) but towards dead root hairs (F). Images taken from videotapes (Allan et al., 1992).

The factors that cause the differential tropism of I. bolleyi to living and dead root cells are still unknown. But this phenomenon seems to be ecologically relevant, because other fungi (Geotrichum candidum, Fusarium oxysporum, Gliocladium roseum) did not show the same behaviour (Allan et al., 1992). Idriella is a specialised weak parasite of cereal and grass roots. It colonises the naturally senescing root cortical cells before these can be invaded by common soil saprotrophs, but it does not cause significant damage to living root cells. In this respect Idriella resembles Phialophora graminicola (see
Biology and control of take-all) and, like Phialophora, it can act as a biological control agent, reducing or preventing infection by several pathogens of cereal roots or stem bases (the take-all fungus, eyespot fungus and Fusarium culmorum which causes cereal foot rot).

Presumably, the tropic responses of the spores enable Idriella rapidly to colonise senescing root cells and exploit their nutrients. Idriella also produces a further batch of spores after it has colonised the dying root cells (Figure G), and these spores might be carried down the roots in percolating water to provide general protection of the root zone. Idriella is found commonly on cereal and grass roots in field conditions, where it can be recognised by its production of characteristic groups of darkly pigmented cells in the dying tissues (Figures H, I).

Figure G. Internal colonisation (black arrowhead) of a dead wheat root hair from spores (s) of Idriella bolleyi. The fungus has already started to produce further spores (white arrowheads) 24-36 hours after colonising the dead root cell.

Figures H, I. Characteristic groups of darkly pigmented resting cells of Idriella bolleyi in the dead surface tissues of a wheat seed and young wheat roots.

Spore tropisms of Verticillium biguttatum

Verticillium biguttatum is a mycoparasite, specialised to invade the hyphae of other fungi, which it exploits as a nutrient source. But it is not aggressive like some other mycoparasites (see Pythium oligandrum). Instead it has a restricted host range and it starts its parasitic phase as a biotroph, feeding from the living hyphae of its hosts in much the same way as the biotrophic plant pathogens. As shown in Figure J, the germinating spores of V. biguttatum show a pronounced tropism towards the host hyphae, leading to penetration of the host and production of club-shaped haustoria (nutrient-absorbing structures) within the living host hypha. Having established this feeding relationship, Verticillium grows over the host colony and produces many sporing structures. It kills the older parasitised cells but produces new haustoria at the advancing edge of the infection.

Figure J. Infection of a hypha of Rhizoctonia solani from germinating spores of Verticillium biguttatum. The germ-tubes showed pronounced tropism, growing in spirals towards the host, then they penetrated the Rhizoctonia hypha and produced club-like haustoria (arrowheads). The parasitised host hypha remained alive, with normal protoplasmic streaming. Image taken from a videotaped interaction on a thin film of water agar (van den Boogert & Deacon, 1994).

Figure K. The strain of Rhizoctonia solani that causes black scurf of potatoes, growing on sterile filter paper in laboratory culture. Most of the fungal growth is inconspicuous, but after the filter paper was colonised the nutrients in the hyphae were mobilised to sites where the fungus produced large sclerotia - the survival structures that cause the black scurf symptoms on potato tubers.

V. biguttatum
has attracted interest because its main host is the important plant pathogen Rhizoctonia solani - a fungus that causes seedling diseases of many crops and that also causes "black scurf" of potato tubers. This name refers to the black crust-like sclerotia (resting structures) of R. solani commonly seen on the surface of potato tubers, reducing their market value. They also are produced in laboratory culture (Figure K).

Even a localised infection by V. biguttatum can reduce the production of sclerotia on colonies of Rhizoctonia (Figure L). Presumably this is caused by the continuous withdrawal of nutrients from the Rhizoctonia hyphal network. This raises the possibility that Verticillium might be used as a biocontrol agent of black scurf in commercial conditions. But there are two major problems to overcome. First, Verticillium can be inoculated onto the "seed tubers" but does not spread efficiently to the daughter tubers which are produced later in the season. Second, Verticillium requires relatively high temperatures (minimum about 13-15oC) whereas Rhizoctonia can grow at much lower temperatures and therefore becomes established early in the season, before Verticillium can take effect.

Figure L. Three plates of cellulose agar inoculated with R. solani at two positions (labelled).

Top plate: inoculated with the same strain (3R41) at both positions. Hyphae of the two colonies fused where they met [see Figure R, below] and formed a single colony over the whole plate. The fungus then produced clusters of sclerotia (arrowheads) on both sides of the plate.

Bottom left plate: as in the top plate, but spores of Verticillium were streaked on one of the colonies. The mycoparasite suppressed the production of sclerotia over the whole agar plate.

Bottom right plate: inoculated with two different strains of R. solani, and spores of Verticillium were streaked on one side of the plate. The two Rhizoctonia strains are mutually incompatible - their hyphae fuse at the zone of contact but the fused cells die (seen as a dark crescent-shaped line on the agar plate). Verticillium has suppressed the production of sclerotia on the inoculated colony (3R09), not on the other colony (3R41). Verticillium seems to suppress the production of sclerotia by withdrawing nutrients from the Rhizoctonia network, so a break in this network has restricted the effect to one side of the plate [From van den Boogert & Deacon, 1994].

Tropism of rust germ tubes

The rust fungi are major plant pathogens that establish infections by producing haustoria in the host cells (see Biotrophic Plant Pathogens). As a prelude to this, these fungi often penetrate a leaf through the stomatal openings, and they use contact sensing to locate these sites. A classic example of this is seen in the rust and powdery mildew fungi of cereals. For example, when uredospores of Puccinia graminis germinate on a cereal leaf the germ tubes grow perpendicular to the rows of leaf cells. The same behaviour is seen on inert replicas of cereal leaves (Figure M), showing that the fungus responds to surface topography and not to chemical signals. This behaviour is thought to maximise the chances of locating a stoma, because the stomata occur in lines (marked "s" in Figure M) on cereal leaves and their positions vary in the different lines.

Figure M. Scanning electron micrograph of germ tubes growing from uredospores (u) of Puccinia graminis on an inert replica of a wheat leaf. Growth of the germ tubes (seen as dark, narrow lines) is orientated at right angles to the pattern of ridges and grooves of the leaf surface cells. Lines of stomata (s) occur at intervals across the leaf surface replica.

Figure N. Scanning electron micrograph of germ tube tips of Puccinia graminis growing perpendicular to precisely spaced ridges and grooves of a polystyrene replica of a microfabricated silicon wafer. The germ tubes are about 4 micrometres diameter; the lower region of their tips is flattened against the replica, presumably enabling them to sense the topography.

[Images supplied by Nick Read; see Read et al., 1992]

The rust fungi of dicotyledonous plants do not show this behaviour, because the leaf surface cells and stomata of dicotyledons are not arranged in rows. However, almost all the rust fungi show another type of topographical sensing - the uredospore germ tubes recognise stomata and respond to them by producing a pre-penetration swelling termed an appressorium.

A remarkable insight into this behaviour was achieved by Harvey Hoch & Richard Staples who used the techniques of the microelectronics industry to produce silicon wafers with precisely defined patterns of ridges and grooves. The wafers were used as templates to produce polystyrene replicas which were then inoculated with uredospores. Working initially with the bean rust Uromyces appendiculatus, it was found that germ tubes produced appressoria when they encountered a single ridge (or groove) of about 0.5 micrometre height, but showed almost no response to heights above 1.0 micrometre. Other rust fungi responded to different ridge heights (see Allen et al., 1991), and these differences are thought to reflect adaptations to different host plants, for which the elevation of the stomatal guard cells may provide the signal for production of an appressorium.

The germ tubes of cereal rust fungi behave differently from the rest. They orientate at right angles to a series of widely spaced ridges and grooves (Figure N) which probably simulate the natural lines of leaf surface cells. They do not form appressoria in response to single ridges; instead, they require a series of very closely spaced ridges, which perhaps signals to them that they have located a stoma.

How do external signals cause growth changes?

This question is relevant to all the examples of tropism discussed so far, but the evidence is best for topographical signalling, where at least four factors have been identified.

1. Close adhesion is required for sensing of topography, and this is achieved by germ tube mucilage. Consistent with this, both adhesion and contact sensing can be abolished by treating the germ tubes with proteolytic enzymes which destroy the proteinaceous component of the adhesive.

2. The position of the Spitzenkörper seems to be important, because germ tubes growing on topographical surfaces have a "nose-down" appearance (Figure N) and the Spitzenkörper is found to be located close to the surface.

3. Electron micrographs and the use of fluorescent dyes show that there is a particularly high density of cytoskeletal elements such as microtubules in the region of a germ tube closest to a surface.

4. By using the "patch clamp" technique on isolated portions of fungal plasma membranes it has been shown that the membrane contains stretch-activated calcium channels.

Calcium is well known as an important signalling ion in eukaryotic cells - it can interact directly with cytoskeletal components and also can act as a second messenger, transducing signals received at the cell surface and leading to changes in gene expression (see Jackson & Heath, 1993).

So, it is suggested that a change in topography might cause a localised stress on the fungal cell membrane, allowing the uptake of calcium ions which then could act either directly on the cytoskeleton to change the orientation of growth or indirectly to alter gene expression leading to differentiation.

Presumably the same underlying mechanisms, but mediated by receptors in the fungal membrane, could explain tropisms to chemical factors.


The spore-bearing structures of several fungi are induced to develop by light (or near-UV). In addition, some spore-bearing structures show a phototropic response, bending towards a light source to facilitate dispersal. This is often found in the dung fungi (coprophilous fungi) which need to disperse their spores onto the surrounding vegetation so that they will be ingested by animals.

Figures O, P. Asexual spore-bearing structures of Pilobolus (zygomycota) growing from a dung pellet. The ring of flavonoid pigment which mediates the phototropic response is faintly visible just below the vesicle in Fig. O. The vesicle acts as a lens to focus light on this pigment - an effect seen in Fig. P where the large black sporangium is pointing towards the camera, with the vesicle behind it.

Pilobolus (Figures O, P) is a classic example of this because it has a mechanism for shooting the whole sporangium clear of a dung pellet. Its spore-bearing structures consist of an erect hypha (sporangiophore) which is swellen into a vesicle below the sporangium. At maturity the vesicle wall ruptures and the vesicle sap squirts the sporangium for a distance of several centimetres. Just below the vesicle is a ring of flavonoid pigment. The vesicle acts as a lens, focusing light on this pigment and causing the sporangiophore to bend so that the sporangium is shot towards the light source.

A similar phototropic response is found in the long sporangiophores of Phycomyces, another genus of coprophilous fungi (Figure Q). In this case the sporangiophore also acts as a lens but there is no vesicle because the spores are released passively.

These phototropic responses differ in one important way from the tropic responses mentioned earlier, because the bending response is achieved by differential wall extension behind the growing tip, probably caused by a combination of localised wall softening (involving wall lytic enzymes) and localised wall growth or stretching. Enzymes with flavin-containing prosthetic groups are quite common in fungi, so a reduction of the flavin pigment by exposure to light could easily change the activity of a wall-associated enzyme.

Figure Q. Sporangiophores of the dung fungus Phycomyces (zygomycota). The left hand image shows stages in elongation of the sporangiophore to a height of 5 cm or more after the tip has swollen to form the sporangium. This occurs by extension of the wall beneath the sporangium. The right hand image shows phototropic bending of the sporangiophore over a period of 15 minutes. [Images based on photographs in W. Shropshire, 1963; Physiological Reviews 43, 38-67. Supplied by Michael Carlile]

Sexual tropisms

The mating reactions of many fungi involve tropic responses to bring two compatible mating types together. For example, the production of sexual spores by the zygomycota (see The Fungal Web) is achieved by the fusion of aerial branches, which grow towards one another under the influence of volatile hormones termed trisporic acids.

Similarly, many basidiomycota undergo fusion (anastomosis) between the normal vegetative hyphae of compatible strains as a prelude to sexual development. These fusions involve highly precise tropic responses, similar to those shown in Figure R for hyphal fusions of Rhizoctonia solani. The fact that these tropic responses (and hyphal fusions) do not occur between different species indicates that the signal molecules must be quite specific, but they have not been identified yet (see Gooday & Adams, 1993).

Figure R. Stages in anastomosis between hyphae of two compatible strains of Rhizoctonia solani. The times shown are minutes after the start of video recording. Several reorientations of the apex of the lower hypha (arrowhead) are seen to occur during this sequence, and fusion is preceded by a directional regrowth of the upper hypha (seen in the 10 minute frame) so that the fusion occurs tip-to-tip. More than 30 minutes elapsed between hyphal contact (frame 3) and complete fusion of the hyphal tips (frame 4). During this time the walls of the hyphal tips dissolved to allow cytoplasmic continuity.

Further reading

Reviews and research papers:

P van den Boogert & JW Deacon (1994) Biotrophic mycoparasitism by Verticillium biguttatum on Rhizoctonia solani. European Journal of Plant Pathology 100, 137-156.

ND Read, LJ Kellock, H Knight & AJ Trewavas (1992) Contact sensing during infection by fungal pathogens. pp. 137-172 in Perspectives in Plant Cell Recognition (eds JA Callow & JR Green). Cambridge University Press.

GW Gooday & DJ Adams (1993) Sex hormones and fungi. Advances in Microbial Physiology 34, 69-145.

EA Allen et al. (1991) Appressorium formation in response to topographical signals in 27 rust species. Phytopathology 81, 323-331.

RH Allan, CJ Thorpe & JW Deacon (1992) Differential tropism to living and dead cereal root hairs by the biocontrol fungus Idriella bolleyi. Physiological and Molecular Plant Pathology 41, 217-226.

SL Jackson & IB Heath (1993) Roles of calcium ions in hyphal tip growth. Microbiological Reviews 57, 367-382.




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