Microtubules and biotic interactions


  • Adrienne R. Hardham

    Corresponding author
    • Plant Science Division, Research School of Biology, College of Medicine, Biology and Environment, The Australian National University, Canberra, Australia
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For correspondence (e-mail Adrienne.Hardham@anu.edu.au).


Plant microtubules undergo extensive reorganization in response to symbiotic and pathogenic organisms. During the development of successful symbioses with rhizobia and mycorrhizal fungi, novel microtubule arrays facilitate the progression of infection threads and hyphae, respectively, from the plant surface through epidermal and cortical cells. During viral and nematode infections, plant microtubules appear to be commandeered by the pathogen. Viruses use plant microtubules for intra and intercellular movement, as well as for interhost transmission. Nematodes manipulate spindle and phragmoplast microtubules to enhance mitosis and partial cytokinesis during the development of syncytia and giant cells. Pathogenic bacteria, fungi and oomycetes induce a range of alterations to microtubule arrays and dynamics. In many situations, the pathogen, or the elicitor or effector proteins derived from them, induce depolymerization of plant cortical microtubule arrays. In some cases, microtubule disruption is associated with the plant defence response and resistance. In other cases, microtubule depolymerization increases plant susceptibility to the invading pathogen. The reasons for this apparent inconsistency may depend on a number of factors, in particular on the identity of the organism orchestrating the microtubule changes. Overall, the weight of evidence indicates that microtubules play an important role in both the establishment of functional symbioses and in defence against invading pathogens. Research is beginning to unravel details about the nature of both the chemical and the mechanical signals to which the plant microtubule arrays respond during biotic interactions.


Fifty years ago, Ledbetter and Porter (1963) reported the presence of microtubules in the cortex of interphase plant cells. During the first half of the intervening 50 years, electron microscopy was used to study these microtubule arrays in detail, with the main research goal focused on determining the structure and organization of the arrays and their role in plant cell morphogenesis. The second half of the half-century has seen the introduction of immunofluorescence microscopy and genetic tagging with fluorescent proteins to investigate cortical microtubule organization, dynamics and function in greater depth. These latter investigations have included research on the role of cortical microtubules in the interactions of plant cells with other organisms, both friend and foe. Early immunocytochemical localization of microtubules during attack by fungal pathogens revealed that rapid morphological changes in the plant cytosol included reorganization of the cortical microtubule array (Kobayashi et al., 1992, 1994). Subsequent studies of genetically tagged tubulin or microtubule-associated proteins have added to our understanding of microtubule behaviour following contact with pathogens and symbionts. Microtubule reorganization has now been extensively documented during both resistant and susceptible interactions with a range of organisms.

This review provides a synopsis of our current understanding of the organization and role of plant cortical microtubules during interactions with pathogens and symbionts. The topic has been reviewed previously (Takemoto and Hardham, 2004; Lipka and Panstruga, 2005; Schmidt and Panstruga, 2007), and the emphasis in the present article focuses on information that has become available over the last 5–6 years. When considering the role of plant microtubules in biotic interactions, one factor that should be kept in mind is the identity of the organism that is orchestrating the changes in the plant microtubule arrays. Is it the plant as part of its defence or mutualistic response, or it is the symbiont or pathogen as part of its infection strategy? Could the behaviour of the microtubules be a result of the integration of signals originating in both plant and invader?

Microtubules and Plant–Symbiont Interactions

Responses to rhizobia during nodulation

The plant microtubule cytoskeleton in root hairs and cortical cells responds to contact by symbiotic Rhizobium bacteria, and to the nodulation factors produced by them. There are three main phases in the establishment of a functional root nodule: root hair curling; infection thread growth; and nodule development. The formation of specific microtubule arrays is associated with each of these processes.

Within minutes of bacterial contact or local application of nodulation factors, there is a transient disruption of cortical and endoplasmic microtubule arrays near the tips of root hairs, after which the density of a bundle of microtubules between the nucleus and the root hair tip increases, and the nucleus moves closer to the hair apex (Figure 1a; Timmers et al., 1999; Weerasinghe et al., 2003; Sieberer et al., 2005). As the root hair curls and the infection thread begins to form, a cortical microtubule array assembles next to the plasma membrane surrounding the developing infection thread (Figure 1b). The microtubules are believed to play a role in directing infection thread growth (Sieberer et al., 2005).

Figure 1.

Organization of microtubules (green) during symbiotic interactions with Rhizobium in Medicago sativa (a) and Medicago truncatula (b), and during infection of M. truncatula by Gigaspora gigantea (c–f). Microtubules are visualized by immunolabelling (a, b), and by the expression of GFP-Map4-MBP (c–f). (a) Microtubules in an infected root hair. Microtubules (arrowheads) line the invading infection thread (blue) and form a bundle between the nucleus (red–yellow) and hair tip. (b) Infection thread (blue) growing through cytoplasmic strands containing a dense array of microtubules in root cortical cells; nuclei, red–yellow. (c) Cells in an uninoculated root. (d) Contact of the fungus (red) with the epidermis induces nuclear movement to the contact site and the development of a network of randomly oriented microtubules (arrows). (e) An inverted cone of microtubules (arrow) forms below the contact site (arrowhead). (f) Dense complexes of microtubules (arrows) assemble in bridges of cytoplasm that traverse the epidermal cells prior to invasion by fungal hyphae. Scale bars: (a) 5 μm; (b) 15 μm; (c–f) 20 μm. Images are reproduced with permission from Timmers et al. (1999) (a, b) and Genre et al. (2005) (c–d).

Coincident with infection thread initiation, microtubules in nearby root cortex cells are also reorganized. The normal cortical array disappears, and an anticlinal array is formed within a band of cytoplasm, similar to the pre-penetration apparatus in mycorrhizal infections (see below), bridging the cell between the periclinal cell walls (Timmers et al., 1999). A network of microtubules continues to surround the infection thread as it grows through the cytoplasmic bridge across the cortical cell. After the nodule has developed, the organization of microtubules within the nodule cells is consistent with their playing a role in the stratification of symbiosomes and other organelles within the nodule (Davidson and Newcomb, 2001; Fedorova et al., 2007).

Recent studies employing genetic tagging of tubulin or microtubule-associated proteins with fluorescent proteins have shed light on the dynamics of microtubules in root hairs during the early phase of Rhizobium infection. In Lotus japonicus root hairs, analysis of parameters indicative of microtubule dynamic instability show that microtubules in young root hairs are more dynamic than those in mature root hairs (Vassileva et al., 2005). Inoculation with symbiotic Mesorhizobium loti or purified M. loti nodulation factors leads to a decrease in microtubule dynamics, and to a reduction in microtubule length and axiality. Analysis of microtubule dynamics in Medicago truncatula root hairs also shows that polymerization rates are highest at the tip of growing root hairs, and that treatment with nodulation factors leads to a reduction in microtubule dynamics (Timmers et al., 2007). It may be that stable microtubules are required to maintain the infection thread and guide its progress from the root periphery to the developing nodule.

Responses to mycorrhizal fungi

The establishment of symbiotic associations between plant roots and mycorrhizal fungi occurs in most plants, and is of considerable importance for plant growth (Smith and Smith, 2012). In exchange for sugars from the plant, endomycorrhizal fungi provide plants with minerals, such as phosphate, to which they often have limited access. During endomycorrhizal associations, the fungus degrades localized regions of root cell walls and develops specialized infection structures that, like the intracellular infection hyphae and haustoria of biotrophic fungal pathogens, are always separated from the host cytoplasm by an intact plant plasma membrane.

During the establishment of endomycorrhizal associations, the fungus forms highly branched arbuscules or pelotons within the root cells. The plant–fungal interface consists of the fungal plasma membrane and cell wall surrounded by the perifungal membrane (derived from the host plasma membrane), and has a high surface-to-volume ratio, as is appropriate for a site of intense molecular exchange. The application of immunofluorescence labelling shows that after root cell invasion by fungal hyphae, the organization of plant microtubules is altered substantially and a microtubule array assembles adjacent to the perifungal membrane (Genre and Bonfante, 1997; Uetake et al., 1997; Uetake and Peterson, 1998; Genre and Bonfante, 1998; Matsubara et al., 1999; Blancaflor et al., 2001; Genre and Bonfante, 2002; Armstrong and Peterson, 2002).

One of the many intriguing aspects of the establishment of an arbuscular endomycorrhizal association is that there are a number of similarities in the behaviour of plant cells that are in contact with a mycorrhizal fungus with that in cells that are in contact with a potential pathogenic fungus. Just as rapid cytoplasmic aggregation occurs beneath pathogen appressoria (see below), cytoplasm aggregates beneath the hyphopodia of arbuscular mycorrhizal fungi (Genre et al., 2005) before fungal penetration of the outer wall of the epidermal cell. There follows the development of an anticlinal column of cytoplasm that forms a bridge between the outer and inner periclinal walls of the epidermal cell, similar to the bridge of cytoplasm that precedes infection thread development during rhizobial symbiosis. It is only after the formation of the bridge of cytoplasm – the pre-penetration apparatus – that the mycorrhizal fungal hypha penetrates the outer epidermal wall and grows across the epidermal cell to reach the root cortical cell layer (Genre et al., 2005). The behaviour of microtubules in this process in epidermal and cortical cells has been visualized through the GFP-tagging of Map4-MBD (Figure 1c–f). γ-Tubulin labelling suggests that the microtubules are nucleated at the perifungal membrane, rather than being moved intact from other regions of the plant plasma membrane (Genre and Bonfante, 1999).

Microtubules and Plant–Pathogen Interactions

Microtubule reorganization during virus infections

In contrast to the situation during bacterial or eukaryotic infections, in which plant microtubules play a demonstrated or potential role in plant defence, in the case of virus–plant interactions, changes in the microtubule cytoskeleton generally appear to be associated with the manipulation of the microtubules by the virus to achieve enhanced intracellular or intercellular movement, or even interhost transmission. Although in some situations viral proteins or viruses can move independently of microtubules (Gillespie et al., 2002; Kawakami et al., 2004), since the first reports of the interaction of viral movement proteins with the plant cytoskeleton (Heinlein et al., 1995; McLean et al., 1995) many studies have shown that microtubules may play a role in the movement of viral RNAs and virions within and between plant cells (for reviews, see Takemoto and Hardham, 2004; Harries et al., 2010). For example, two studies of transgenic Nicotiana tabacum (tobacco) plants give evidence for a role of microtubules in the intercellular movement of the tobacco mosaic virus (TMV) movement protein. Tobacco plants that overexpress a tobacco microtubule-associated protein that binds to TMV movement protein (Kragler et al., 2003), or that are resistant to the microtubule assembly inhibitor, ethyl phenyl carbamate (Ouko et al., 2010), display reduced rates of intercellular TMV movement protein transport compared with controls. In the latter case, the tobacco mutant contained lower levels of tyrosinated α-tubulin and lower rates of plus-end growth, as indicated by the rate of movement of the GFP-EB1 marker. Reduced microtubule dynamics in the drug-resistant mutant bestowed increased tolerance to viral infection.

In addition to demonstrations of the interaction of viral movement proteins with microtubules, two recent studies have revealed the apparent manipulation of the plant microtubule cytoskeleton to aid in the formation and movement of cauliflower mosaic virus (CaMV) inclusion bodies (Martinière et al., 2009; Harries et al., 2009). Inclusion bodies are thought to be sites of viral replication, and their association with microtubules has been well documented in animal cells (reviewed by Novoa et al., 2005). Two CaMV-encoded proteins, designated P2 and P3, are transported by microtubules from their site of synthesis to large electron-lucent inclusion bodies (Martinière et al., 2009). Visualization of inclusion bodies via GFP-tagging of a third CaMV inclusion body protein, P6, allowed the demonstration of inclusion body trafficking along actin microfilaments and their association with microtubules when stationary (Harries et al., 2009). Further analysis showed that the P6-containing inclusion bodies increased microtubule resistance to depolymerization by oryzalin (Figure 2a,b). These inclusion bodies are thought to be important for viral uptake into the vector's stylet. This means that the use of plant microtubules by the CaMV is instrumental in interhost transmission of the pathogen.

Figure 2.

Microtubule reorganization and dynamics during viral and bacterial infections. Microtubules are visualized in Nicotiana edwardsonii by the expression of GFP-MBP (a, b), in Vitus rupestris by immunolabelling (c, d) and in Arabidopsis thaliana by the expression of GFP-Map4 (e, f) and GFP-AtEB1 (g, h). (a and b) Expression of GFP-tagged P6 inclusion body protein from cauliflower mosaic virus stabilizes microtubules against depolymerization by oryzalin (50 μm). (c and d) Fragmentation and depolymerization of microtubules induced by treatment with Harpin effector (9 μg ml−1). (e and f) Depolymerization of microtubules induced by HopZ1a effector from Pseudomonas syringae. (g and h) Reduction in the number of growing microtubule ends by Pseudomonas syringae HopZ1a. Scale bars: (a, b) 25 μm; (c, d) 50 μm; (e–h) 25 μm. Images reproduced with permission from Harries et al. (2009) (a, b), Qiao et al. (2010) (c, d) and Lee et al. (2012) (e–h).

Phytopathogenic bacteria

Pathogenic bacteria inject a range of virulence effector proteins into the cytoplasm of their host cells using the type-III secretion system (Cornelis and Van Gijsegem, 2000). In animals, a prime target for these effectors is the actin cytoskeleton (Galan, 2001), but some effectors have been shown to affect the microtubule cytoskeleton (Yoshida et al., 2006). Until recently, there has been little, if any, indication that effectors produced by phytopathogenic bacteria target either actin or microtubule arrays; however, over the last 3 years a series of papers has presented evidence indicating that bacterial elicitors and effectors do affect the plant microtubular cytoskeleton.

One study investigated the bacterial effector, Harpin, a protein produced by the fire blight pathogen Erwinia amylovora and other bacteria (Qiao et al., 2010; Chang et al., 2011). The study compared the response to Harpin treatment in cell suspension cultures derived from two grapevine varieties: Vitis vinifera cv. ‘Pinot Noir’ and Vitis rupestris. Harpin triggers a hypersensitive reaction in V. rupestris but not in V. vinifera: in the pathogen-susceptible variety, V. vinifera, the cortical microtubule arrays are not affected, but in the pathogen-resistant variety, V. rupestris, the cortical arrays are severely disrupted by treatment with 9 μg ml–1 Harpin (Figure 2c,d). The ordered cortical array of parallel microtubules typical of uninfected cells is replaced by a sparse array of short microtubules. A possible influence of microtubule dynamics on the response to Harpin was investigated by using immunoblotting to determine the relative abundance of tyrosinated and detyrosinated α-tubulin. The results show that the V. vinifera cell line has high levels of tyrosinated α-tubulin, indicative of high microtubule turnover, whereas the V. rupestris cell line has low levels of tyrosinated α-tubulin, indicative of low microtubule turnover. These observations suggest that Harpin-induced changes in the microtubule arrays in V. rupestris cells may arise through microtubule fragmentation rather than through the sequestering of tubulin dimers and a consequent interference with normal microtubule treadmilling (Qiao et al., 2010). Although Harpin induces rapid accumulation of stilbene synthase transcripts, treatment with the product of this enzyme, the phytoalexin resveratrol, does not disrupt the microtubule arrays in V. rupestris cells (Chang et al., 2011).

In an extension to this study, Chang and Nick (2012) compared the effects of the Harpin effector with those of the model bacterial elicitor, flagellin. Unlike Harpin, the elicitor-active domain of flagellin, flg22, does not trigger a hypersensitive response in V. rupestris cultured cells. It does, however, cause microtubule disruption. Both Harpin and flg22 induce the expression of defence-related genes and alkalinization of the medium by the Vitis cells, and the extent of the alkalinization is reduced by oryzalin treatment. The authors conclude that microtubules are involved in some way in transducing the signal arising from elicitor or effector recognition, perhaps through the modulation of ion transporters in the plasma membrane (Chang and Nick, 2012).

A second bacterial effector, HopZ1a from Pseudomonas syringae, has been shown to disrupt microtubule arrays in Arabidopsis (Lee et al., 2012). HopZ1a is an acetyltransferase that acetylates itself and tubulin. The acetylation of β-tubulin prevents its incorporation into microtubules, and thus interferes with microtubule polymerization. HopZ1a induces a reduction in the density of cortical microtubules and in the abundance of AtEB1-labelled, growing microtubule ends (Figure 2e–h), with both effects requiring HopZ1a acetyltransferase activity. Downstream effects, again requiring catalytically active HopZ1a, include disruption of secretion and callose deposition, and increased susceptibility of the Arabidopsis plants to infection by P. syringae. The pharmacological destruction of microtubules with oryzalin also increased P. syringae virulence, providing evidence that it is the HopZ1a disruption of the cortical microtubule arrays that is responsible for increased plant susceptibility. This effect was not observed with a P. syringae mutant lacking a functional type-III secretion system, indicating that destruction of the microtubule arrays is just one aspect of the infection mechanism of P. syringae.

Microtubule reorganization during attack by fungi and oomycetes

The first visible sign of a plant reaction to the presence of a potentially pathogenic fungus or oomycete on its surface is the aggregation of cytoplasm beneath the contact site. Both immunocytochemical and GFP-tagging experiments have shown that cytoplasmic aggregation is accompanied by a focusing of actin microfilaments and endoplasmic reticulum, and by an accumulation of Golgi stacks and peroxisomes at the contact site (Kobayashi et al., 1994; Kobayashi and Hakuno, 2003; Takemoto et al., 2003, 2006; Hermanns et al., 2008). These cytoplasmic rearrangements bring about a polarization of secretion that results in the development of a wall apposition (papilla) beneath the adherent microorganism on the outer surface of the wall. Wall appositions constitute a physical and chemical barrier that inhibits pathogen ingress, and they are a key component in basal defence (Collins et al., 2003; Assaad et al., 2004; Hückelhoven, 2007a,b). They form not only in epidermal cells in response to contact with hyphae or appressoria, but also in mesophyll cells in contact with haustorial mother cells.

Pharmacological studies have confirmed that it is the actin cytoskeleton that is responsible for cytoplasmic aggregation and the polarization of secretions; however, the microtubule cytoskeleton also changes in response to attempts by a potential pathogen to invade the plant cell. In contrast to the consistent focused rearrangement of actin microfilaments, a range of alterations to the microtubule cytoskeleton has been observed during the early stages of the plant's response. In the biotrophic interactions between Hordeum vulgare (barley) and Blumeria graminis f. sp. hordei (Bgh; Figure 3a), and Linum usitatissimum (flax) and Melampsora, radial arrays of microtubules have been observed beneath the fungal appressoria (Kobayashi et al., 1992, 1994; Hoefle et al., 2011). In interactions of Phytophthora sojae with Petroselinum crispum (parsley) or Glycine max (soya bean), on the other hand, localized microtubule depolymerization occurs at the contact site (Gross et al., 1993; Cahill et al., 2002). In Arabidopsis inoculated with isolates of Phytophthora sojae or Hyaloperonospora arabidopsidis, which encompass the spectrum of non-host, incompatible and compatible interactions, the cortical cytoplasm beneath the invading fungus rapidly accumulates unstructured but intense GFP-tubulin fluorescence (Figure 3b; Takemoto et al., 2003). In some instances, the microtubule array surrounding the region of diffuse fluorescence displays an overall circumferential alignment (Figure 3b). Both the appearance of the ‘cloud’ of diffuse fluorescence and the apparent change in microtubule alignment in the Arabidopsis epidermal cells could be caused by selective depolymerization of microtubules that are oriented radially relative to the penetration site.

Figure 3.

Microtubule response to infection by fungi, oomycetes and nematodes, or to mechanical stimulation. Microtubules in barley (a) and Arabidopsis (b–h) visualized by the expression of GFP-MAGAP1 (a), GFP-tubulin (b–f) and GFP-Map-MBD (g, h). (a) Microtubule array focused on the contact site (*) with an appressorium of Blumeria graminis f. sp. hordei. The red fluorescence is the result of co-transformation with cytoplasmic/nuclear RFP. Attempted penetration of cotyledon epidermal cells by Hyaloperonospora arabidopsidis (Cala2) (b) or contact with a microneedle (c) induces the development of a cloud of GFP-tubulin and formation of a global circumferential array of microtubules surrounding the infection (*) or contact (arrowhead) site. (d–f) Microtubule depolymerization in leaf epidermal cells 15 min (d), 45 min (e) and 75 min (f) after treatment with VD toxin from Verticillium dahliae. (g, h) Phragmoplast microtubules (arrows) during partial cytokinesis and formation of wall stubs (stained with FM4-64) in developing giant cells in Arabidopsis roots. Scale bars: (a–c, g, h) 10 μm; (d–f) 20 μm. Images reproduced with permission from Hoefle et al. (2011) (a), Hardham et al. (2007) (b), Hardham et al. (2008) (c), Yao et al. (2011) (d-f) and Caillaud et al. (2008) (g, h).

Pharmacological destruction of the actin cytoskeleton inhibits cytoplasmic aggregation, papilla formation, hypersensitive cell death and defence gene activation (Tomiyama et al., 1982; Hazen and Bushnell, 1983; Kobayashi et al., 1997b; Škalamera et al., 1997; Škalamera and Heath, 1998; Takemoto et al., 1999), allowing organisms that are normally not virulent to invade and infect a plant that is normally resistant (Kobayashi et al., 1997b; Yun et al., 2003). Early studies involving treatments with microtubule inhibitors, such as oryzalin, generally showed a less marked effect on papilla formation and the inhibition of penetration (Kobayashi et al., 1997b; Škalamera et al., 1997), although in Triticum spp. (wheat) inoculated with the powdery mildew fungus, Sphaerotheca fuliginea, oryzalin increases penetration and the formation of haustoria (Li et al., 2010). In this latter system and in flax inoculated with Melampsora lini (Kobayashi et al., 1997a), depolymerization of microtubules with oryzalin inhibits hypersensitive cell death and leads to increased plant susceptibility. On the other hand, disruption of microtubules in grapevine cell cultures with oryzalin or taxol induces the expression of defence genes (Qiao et al., 2010).

In addition, a recent study of the barley–Bgh interaction has shown that transient silencing of the gene encoding a barley ROP-binding protein kinase, HvRBK1, leads to the fragmentation of cortical microtubules in the barley epidermal cells and increased susceptibility to penetration by Bgh (Huesmann et al., 2012). ROP proteins (Rho of plants; also called RACs) are small plant-specific GTPases that function in the transduction of signals received at the cell surface during plant development and response to biotic and abiotic stresses (Nibau et al., 2006). They are known to regulate the microtubule cytoskeleton, and are involved in signalling during both susceptibility and resistance to microorganisms (Mucha et al., 2011). Activated barley HvROPs, HvRACB and HvRAC1, interact with HvRBK1 and induce its redistribution from the cytoplasm and nucleoplasm to the cell cortex (Huesmann et al., 2012). The implication of these results is that in this system, perception of Bgh at the cell surface leads to the activation (GTP-binding) of the barley ROPs (HvRACB and HvRAC1), which in turn enhances the activity of the ROP effector, HvRBK1, inducing it to move to the cell cortex where, by an as yet undescribed mechanism, it interacts with the cortical microtubules, facilitating their role in penetration resistance. Activated HvRACB also binds to another barley ROP effector, HvMAGAP1 (microtubule-associated ROP-GTPase activating protein), again inducing its redistribution to the cell cortex (Hoefle et al., 2011). The RFP-tagging of HvMAGAP1 shows that it binds to microtubules. In situations in which penetration of Bgh was successfully inhibited, HvMAGAP1-labelled microtubules formed a dense radial network focused on the contact site beneath the fungal appressorium (Figure 3a). However, in situations in which Bgh was able to penetrate the epidermal cell, there was little change in the organization of the cortical microtubules in the cell, except for the development of a region of diffuse fluorescence around the neck of the haustorium (corresponding to the penetration site; Hoefle et al., 2011).

What underlies the variability in the response of plant microtubules to a potential pathogen? In the barley–Bgh system discussed above, the observed behaviour of the microtubule cytoskeleton suggests differences are associated with the nature of the final outcome of the interaction, i.e. susceptibility or resistance. But which is the cause and which the effect in this situation? Other studies have observed no apparent difference in the response of the cortical microtubule arrays to virulent, avirulent or non-host pathogens (Takemoto et al., 2003). These latter observations have led to the hypothesis that the microtubule cytoskeleton may be responding to the physical pressure exerted by the fungal or oomycete hypha as it begins to penetrate the plant cell surface. This hypothesis was tested by gently contacting, but not puncturing, the epidermal surface of Arabidopsis cotyledons with fine microneedles and following the subcellular changes of GFP-tagged microtubules, microfilaments, endoplasmic reticulum and peroxisomes (Hardham et al., 2008). These experiments revealed rapid alterations in the behaviour and organization of these cell components following microneedle contact. Within 3–5 min, actin, endoplasmic reticulum and peroxisomes begin to accumulate at the contact site. In the GFP-tubulin transgenic plants, a cloud of bright, diffuse tubulin fluorescence, initially surrounded by a microtubule-depleted zone, forms in the cortical cytoplasm at the contact site (Figure 3c). The region of bright diffuse fluorescence follows the needle if it is moved across the cell surface, and disperses if the needle is lifted off the cotyledon surface (Hardham et al., 2008).

Studies of the behaviour of cortical microtubules during plant morphogenesis also indicate that mechanical signals in the form of physical stress can orient cortical microtubules (Hamant et al., 2008). Although there is no evidence that microtubules sense stress forces directly, it is likely that microtubule orientation results from the detection of stress by receptors in either the cell wall or plasma membrane. Possible candidates include mechanosensitive channels in the plasma membrane. Ten MSL genes encoding proteins similar to mechanosensitive channels of small conductance (MscS-like) have been identified in Arabidopsis (Haswell et al., 2008), and GFP-tagging of two proteins, MSL9 and MSL10, shows that they are localized to the plasma membrane in root cells. Genetic deletion studies provide evidence that MSL9 and MSL10 are required for mechanosensitive channel activity in root protoplasts (Haswell et al., 2008).

Studies in which a microneedle is used to apply pressure to leaf epidermal cells indicate that plant cells can detect this physical stress, and the subcellular response that ensues is similar to that occurring during basal defence to potentially pathogenic fungi or oomycetes (Hardham et al., 2008). Two questions that arise from these observations are: (i) what is the exact nature and magnitude of the physical force that induces this response, including changes to the microtubular cytoskeleton, in the plant cell; and (ii) can a full response be induced by a physical signal alone, or is a chemical signal also required?

With regard to the first question, and the parameters of the physical force that is required to induce a response, it has been noted that during infection of Arabidopsis leaves by H. arabidopsidis, cytoplasmic aggregation occurs only in cells that are being or have been invaded by the pathogen (Hermanns et al., 2008). By contrast, in situations in which the plant cell was touched by a hypha growing within the apoplast, no cytoplasmic aggregation was observed. One interpretation of these observations is that there is a threshold value for the physical force that is required to stimulate the subcellular reorganization involved in a defence response (Hermanns et al., 2008). Alternatively, the lack of a response to contact with the lateral wall of the non-invading hypha could be caused by the absence of chemical signals that are associated with an invading hyphal tip. This latter suggestion leads to the second question posed above, namely can a full basal defence response be induced by a physical signal in the absence of an accompanying chemical signal?

The microtubule cytoskeleton does respond to chemical signals from pathogenic fungi or oomycetes. Treatment of tobacco culture cells with cryptogein elictor from Phytophthora cryptogea causes microtubule depolymerization (Binet et al., 2001). Similarly, in a concentration-dependent manner, VD toxin produced by the necrotrophic fungal pathogen, Verticillium dahliae, causes cortical microtubule depolymerization in Arabidopsis suspension culture cells (Figure 3d–f; Yuan et al., 2006; Shi et al., 2009; Yao et al., 2011). Treatment with VD toxin induces the production of NO (nitric oxide) and H2O2 (Shi et al., 2009; Yao et al., 2011). If either NO or H2O2 production is inhibited, microtubule depolymerization is reduced, suggesting that NO and H2O2 act upstream of the microtubule response (Shi et al., 2009). A study of lateral root development in rice has also shown that both mechanical and hormonal signals are required for the induction of epidermal cell death that accompanies normal lateral root emergence (Steffens et al., 2012). A study of parsley suspension culture cells subjected to localized abiotic pressure or elicitor application, or both, investigated structural and biochemical responses in the plant cells to these signals (Gus-Mayer et al., 1998). The results indicated that mechanical stimulation alone induced cytoplasmic reorganization, the production of reactive oxygen species and the expression of some defence-related genes. Elicitor application alone induced production of reactive oxygen species but no morphological changes. The application of both mechanical and chemical stimuli led to a response similar to that obtained after pressure treatment. None of the regimes resulted in papilla formation or hypersensitive cell death, suggesting that other signals were required in order to induce the full basal defence response in this system (Gus-Mayer et al., 1998).

Although cryptogein elicitor and VD toxin effector can induce changes in microtubule organization, investigations of factors that affect cortical microtubule orientation and the direction of plant cell expansion during plant growth and development have shown that microtubules respond to physical rather than chemical signals (Hamant and Traas, 2010). This has been demonstrated in cell ablation experiments, in which a laser is used to destroy selected cells and the behaviour of microtubules in the adjacent cells is analysed. Ablation of cells in the centre of the shoot meristem changes the stress patterns within the central zone, and results in microtubules becoming re-oriented parallel with the main stress direction. By contrast, ablation of cells in boundary domains did not have this effect. The authors interpret these results as indicating that in this situation, microtubule reorientation is not a response to a chemical signal (Hamant et al., 2008).

Another notable observation made in the cell ablation experiments was that a global circumferential arrangement of microtubules develops in epidermal cells surrounding the ablated zone (Hamant et al., 2008; Hamant and Traas, 2010). This reorganization of cortical microtubules is reminiscent of that observed in Arabidopsis epidermal cells surrounding the pathogen infection site, as described above (Figure 3b; Takemoto et al., 2003), and a similar circumferential arrangement of microtubules was observed following mechanical stimulation with a microneedle (Figure 3c) (Hardham et al., 2008).

Plant microtubules in response to nematode attack

There are two main types of sedentary pathogenic nematodes that attack plants: cyst nematodes and root knot nematodes. Following root invasion, cyst nematodes induce many rounds of proliferative divisions within vascular cells to form a syncytium. After invasion, root knot nematodes cause the development of multinucleate giant cells through the degradation of cortical and vascular cell walls, and by the induction of multiple rounds of mitosis and endoploidy (Davis et al., 2004; Gheysen and Mitchum, 2011).

Immunolocalization and GFP-tagging experiments indicate that disruption of normal interphase microtubule arrays in the root cells occurs during both syncytium and giant cell formation (de Almeida Engler et al., 2004, 2005; Klink et al., 2005). There is evidence of microtubule fragmentation and an increase in the level of diffuse tubulin fluorescence; however, transcriptional analyses show an increase in the expression of tubulin and microtubule-associated proteins during nematode infection (de Almeida Engler et al., 2004; Klink et al., 2005; Caillaud et al., 2008; Swiecicka et al., 2009). During the infection of tomato plants by the potato cyst nematode, genes encoding a tubulin-tyrosine ligase family protein, a putative microtubule-binding protein and an AtMAP70-4 homologue are upregulated 1–7 days after root inoculation (Swiecicka et al., 2009). An effector in the Ran-binding protein family that is secreted by the potato cyst nematode may also be involved in microtubule nucleation and/or stabilization of spindle microtubules (Davis et al., 2004). In Arabidopsis roots infected by Meloidogyne incognita, one upregulated transcript encodes the microtubule-associated protein, MAP65-3 (Caillaud et al., 2008).

In contrast to studies of the role of the plant microtubular cytoskeleton in the defence response during plant–fungal or plant–oomycete interactions, the transcription analyses described above, together with pharmacological studies of plant–nematode interactions, point to a role, indeed a requirement, for plant microtubules in giant cell or syncytium formation and successful nematode development. Depolymerization of microtubules in pea roots by colchicine inhibits M. incognita development (Wiggers et al., 2002), and taxol inhibits nematode maturation in Arabidopsis (de Almeida Engler et al., 2004). In Arabidopsis mutants lacking a functional MAP65-3 gene, both the full differentiation of giant cells and the maturation of nematodes are inhibited (Caillaud et al., 2008). Together, the data suggest that changes in the plant microtubular cytoskeleton are associated with the development of mitotic spindles during mitosis and phragmoplasts during a form of partial cytokinesis (Figure 3g,h). These processes appear to be orchestrated by the nematodes as part of their infection strategy, and are required for successful feeding cell and nematode development.

Concluding remarks

Our current understanding of plant microtubules documents their organization (Figure 4) and multiplicity of roles during biotic interactions. During interactions with pathogenic viruses, nematodes, symbiotic bacteria and fungi, plant microtubules play a role in facilitating infection. Clearly, this is to the detriment of the plant's wellbeing in the case of viruses and nematodes, but is to the plant's advantage during the establishment of a symbiotic relationship with rhizobia and mycorrhizal fungi. It seems likely that in the first case, the plant microtubules are being manipulated by the invading pathogens. In the second case, either the symbiont or the plant, or both, could be orchestrating the changes that accompany the development of the symbiosis. In interactions with a range of other pathogenic organisms, changes to the microtubule cytoskeleton are part of the plant's defence response.

Figure 4.

Diagrammatic representations of the organization of plant microtubules (blue lines) during the interaction of plants with symbionts and pathogens. (a) Infection thread formation after inoculation of a root hair with Rhizobium. (a1) In uninoculated hairs, microtubules are axially or helically aligned, and the nucleus (yellow) is positioned about 30–40 μm from the tip of the root hair. (a2) After inoculation with rhizobia (brown spot), a dense array of microtubules assembles and the nucleus moves closer to the tip of the hair. (a3, a4) As the infection thread (brown) grows, the plasma membrane enclosing the thread is lined by microtubules. A bridge of cytoplasm containing parallel microtubules aligned with the advancing infection thread forms in the adjacent cortical cell. (b) Microtubule reorganization during colonization by an arbuscular mycorrhizal fungus. (b1) Before colonization, microtubules are aligned in parallel arrays in root epidermal and cortical cells. (b2) After contact with the fungal hypha, microtubules aggregate and the nucleus moves to the contact site. (b3, b4) Before hyphal penetration into epidermal (b3) or cortical (b4) cells, an inverted funnel of microtubules assembles between the contact site and the opposite cell wall, and the nucleus is moved away from the contact site. (b5) The advancing fungal hypha is surrounded by the plant plasma membrane lined by microtubules. (b6) As the arbuscule develops, microtubules line the perifungal membrane that encloses the highly branched arbuscule. (c) Inclusion bodies containing the cauliflower mosaic virus P6 protein (green spots) associate with and stabilize microtubules in leaf epidermal cells. (d) Microtubule arrays in an untreated (upper) cell are depolymerized by the HopZ1a effector from Pseudomonas syringae (lower cell). (e) During attempted penetration by some fungal pathogens (e.g. Erysiphe), microtubules in epidermal cells change from a parallel array (upper cell) to become focused on the fungal contact site (lower cell). (f) During attempted epidermal penetration by some oomycete pathogens (e.g. Hyaloperonospora arabidopsidis), microtubules often depolymerize close to the contact site (lower cells), sometimes changing from a parallel array (upper cells) to an overall circumferential array that may cross cell borders around the contact site (lower cells). (g) During infection by root knot nematodes, cortical microtubule arrays are disrupted but spindle and phragmoplast microtubules are abundant in giant cells in the root. Adapted from Takemoto and Hardham (2004).

In analysing microtubule redeployment during interactions with fungi or oomycetes, questions to be addressed include: (i) why is the cortical microtubule array disrupted prior to fungal invasion; and (ii) what is the function of the microtubules that assemble at the perifungal membrane? With regard to the first question, studies to date indicate that prior to fungal invasion, the exact nature of alterations to the cortical microtubule arrays at the contact site depends on the identity of the interacting partners; however, the response is generally one of microtubule depolymerization. The simplest explanation is that either partial or complete microtubule depolymerization makes way for the establishment of a new array at the perifungal membrane.

The area of the plant plasma membrane that surrounds ‘intracellular’ fungal or oomycete cells is specialized for its function in intensive molecular exchange. Immunocytochemical studies show the presence of proteins in the extrahaustorial membrane that do not occur elsewhere in the plasma membrane (Mackie et al., 1991). Genetic tagging of membrane proteins confirms that this is also the case in the peri-arbuscular membrane (Pumplin and Harrison, 2009). Perhaps microtubules are involved in the maintenance of perifungal membrane-specific molecules within the perifungal membrane domain.

In plants, actin microfilaments often facilitate targeted transport of secretory vesicles, and are thus likely to be responsible for the establishment of areas of the plasma membrane with locally distinct molecular composition. However, cortical microtubules are known to control the distribution and/or movement of plasma membrane proteins in plants, in particular that of cellulose synthase enzyme complexes during plant cell wall formation (Bringmann et al., 2012). In a number of situations, wall formation is spatially localized, a distribution reflected by a similar localization of cortical microtubules. Recent studies of localized cell wall formation in epidermal pavement cells and xylem elements have demonstrated intricate interactions between activated ROP (Rho-related GTPases of plants) proteins, cortical microtubules and microtubule-associated proteins to achieve localized wall deposition (Fu et al., 2005; Oda et al., 2010; Oda and Fukuda, 2012a,b; Yang and Lavagi, 2012). On the one hand, proteins in localized domains in the plasma membrane may stabilize or de-stabilize adjacent cortical microtubules, but on the other hand cortical microtubules can control the distribution of localized plasma membrane ROP proteins (Yang and Lavagi, 2012; Oda and Fukuda, 2012b). Perhaps the highly specialized perifungal and extrahaustorial membrane domains are maintained by the cortical microtubule arrays that line them.


I would like to thank Drs ACJ Timmers, A Genre, PA Harries, F Qiao, AHY Lee, C Hoefle, L-L Yao, and M-C Caillaud for allowing me to reproduce their images and S Wragg for assistance in preparation of Figure 4. The author declares no conflict of interest.