Journal of Experimental Botany, Vol. 65, No. 7, pp. 1789–1798, 2014 doi:10.1093/jxb/ert415  Advance Access publication 14 December, 2013

Review paper

Vascularization and nutrient delivery at root-knot nematode feeding sites in host roots Derek G. Bartlem1,2, Michael G. K. Jones2 and Ulrich Z. Hammes3,* 1 

Research Faculty of Agriculture, Hokkaido University, Sapporo 060-8589, Japan School of Veterinary and Life Sciences, WA State Agricultural Biotechnology Centre, Murdoch University, Perth WA 6150, Australia 3  Cell Biology and Plant Biochemistry, Regensburg University, Universitaetsstrasse 31, D-93053 Regensburg, Germany 2 

* To whom correspondence should be addressed. E-mail: [email protected]

Abstract Plants are constantly challenged by pathogens and pests, which can have a profound impact on the yield and quality of produce in agricultural systems. The vascular system of higher plants is critical for growth and for their ability to counteract changing external conditions, serving as a distribution network for water, nutrients, and photosynthates from the source organs to regions where they are in demand. Unfortunately, these features also make it an attractive target for pathogens and pests that demand access to a reliable supply of host resources. The vascular tissue of plants therefore often plays a central role in pathogen and parasite interactions. One of the more striking rearrangements of the host vascular system occurs during root-knot nematode infestation of plant roots. These sedentary endoparasites induce permanent feeding sites that are comprised of ‘giant cells’ and are subject to extensive changes in vascularization, resulting in the giant cells being encaged within a network of de novo formed xylem and phloem cells. Despite being considered critical to the function of the feeding site, the mechanisms underlying this vascularization have received surprisingly little attention when compared with the amount of research on giant cell development and function. An overview of the current knowledge on vascularization of root-knot nematode feeding sites is provided here and recent advances in our understanding of the transport mechanisms involved in nutrient delivery to these parasite-induced sinks are described. Key words:  Amino acid transporters, feeding cells, gall, nutrient transport, phloem, plant disease, plant parasite, root-knot nematodes, vascular tissue remodelling, xylem.

Introduction During the establishment of a long-term interaction with a plant host, pathogenic and symbiotic microorganisms can induce a change in plant morphology that results in the formation of unique organs. In the case of rhizobia that form a symbiotic association with legume roots, nodules are induced that encapsulate the nitrogen-fixing bacteroids. Vascularization of these nodules facilitates the collection and redistribution of assimilated nitrogen from the nodule to other plant tissues, in addition to the reverse supply of carbon and amino acids from the host to the bacteroid (Pate et  al., 1969; Prell et  al., 2009; Oldroyd et  al., 2011). Crown

galls induced on plant stems by the pathogenic bacterium Agrobacterium tumefaciens also develop a complex network of vascular tissue that is expected to help provide the pathogen with assimilates from its host (Aloni et al., 1995). In both of these plant–microbe interactions, the de novo formation of the vascular network in the new lateral organs is associated with significant changes in phytohormone levels that appear to be required for the successful interaction (Ullrich and Aloni, 2000; Grunewald et al., 2009). This induction of lateral growth and subsequent vascularization is not limited to plant–microbe interactions. Parasitic animals, in particular

© The Author 2013. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved. For permissions, please email: [email protected]

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Received 17 September 2013; Revised 2 November 2013; Accepted 11 November 2013

1790 | Bartlem et al. Both the root-knot and cyst nematode feeding cells can be viewed as strong, induced terminal sink tissues that require direct access to the plant vascular tissue (Jones, 1981). Vascularization of mature feeding sites is much more profound around root knot nematode giant cells compared with that around syncytia and they are also sourced by different types of phloem (Hoth et  al., 2008). It has generally been thought that both root-knot and cyst nematode feeding sites are symplasmically isolated and that loading into the feeding cells is mediated by transporters (Jones, 1981; Böckenhoff and Grundler, 1994; Grundler et al., 1998). Whilst this was confirmed to hold true for root-knot nematodes, the earlier conclusions did not seem to hold for cyst-nematode syncytia (Hoth et  al., 2005). Taking into account the significant differences in infestation biology between root-knot and cyst nematodes, it is clear that these two types of nematode parasitism are best considered as separate parasitic systems. Here, we specifically focus on root-knot nematode infestation because of the extensive remodelling of the host vascular tissue that occurs at infestation sites, taking into consideration the influence of hormones associated with phloem development and the transport processes that support the supply of plant assimilates to the nematode-induced feeding site.

Root knot galls and vascularization of feeding sites The root-knot nematode infestation cycle can be separated into three distinct stages, invasion, induction, and nutrient acquisition (Fig.  1). After hatching from eggs in the soil, second-stage juveniles penetrate host root cells near the tip, and follow an intercellular migratory path in the cell wall compartment to the zone containing differentiating vascular cells (Wyss et al., 1992; Goto et al., 2013). It is here that they select host target cells and initiate reprogramming events that result in the conversion of these target cells into giant cells. The beginning of this induction stage also marks the point of commitment for the young parasite as it then becomes sedentary and abandons its migratory lifestyle for one dedicated to growth and reproduction. Root-knot nematodes are obligate parasites that depend completely on the supply of nutrients from these induced giant cells, which are accessed using a

Table 1.  Specific characteristics of cyst and root-knot nematode infestation biology Characteristic

Root-knot nematodes

Cyst nematodes

Host specificity: Invasion style: Target cells in host: Feeding cells: Surrounding tissue: Vascularization: Reproduction:

Broad, most vascular plants Intercellular migration around root tip to target site Differentiating vascular cells in zone of elongation Giant cells, induced from distinct initial cells Forms root-knot gall Extensive networks of de novo xylem and phloem Parthenogenetic, rarely sexual. Eggs secreted at root surface

Narrow, defined set of species Destructive piercing through cells direct to target site Cells adjacent to differentiated vascular cells in maturation zone Syncytium, formed by fusion of neighbouring cells Minimal morphological change Primarily original vascular tissue Sexual. Eggs remain within female cyst body until hatching

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the parasitic worms known as root-knot nematodes, also follow a similar strategy in their interaction with plants (Hoth et al., 2008; Jones and Goto, 2011). Plant parasitic nematodes are devastating pests and cause considerable yield losses in many crop plants (Sasser and Freckman, 1987). A large part of this damage is inflicted by the sedentary endoparasitic nematodes, which include two major groups commonly known as cyst (Heterodera spp, Globodera spp) and root-knot nematodes (Meloidogyne spp). These two groups of nematodes induce highly specialized feeding sites inside the roots of their host that, once established, support the growth and development of the parasite throughout their longer term interaction. Because they both share a sedentary lifestyle and induce feeding cells intimately associated with plant vascular tissue, biological insights on host responses to cyst and root-knot nematodes are often integrated and discussed as a generalized view of plant nematode parasitism. However, it is now apparent that the cyst and root-knot nematodes are phylogenetically much more diverse than previously thought (Scholl and Bird, 2005) and there are several important differences in the infestation biology between these two groups (Table  1). Root-knot nematodes can infest a broad range of vascular plants and migrate intercellularly through the root tips to reach the precursor vascular cells in the elongation zone. By contrast, cyst nematodes have a high degree of host specificity and migrate directly to regions of established vascular tissue by destructively piercing and migrating through successive roots cells. The feeding sites of root-knot nematodes consist of multinucleate giant cells that are each induced from a distinct initial precursor cell (Jones and Payne, 1978; Jones, 1981; Jones and Goto, 2011), whereas the feeding sites of cyst nematodes are composed of multinucleate syncytia that are formed by the fusion of neighbouring parenchyma and associated cells (Jones and Northcote, 1972b; Jones and Dropkin, 1976; Sobczak and Golinowski, 2011). Roots infested with root-knot nematodes form gall-like organs that result from changes in the biology of cells surrounding the feeding site, in contrast to the less drastic changes in root morphology at cyst nematode feeding sites. It is likely that root-knot nematodes also differ in the way they interact with host-defence-signalling pathways compared with cyst nematodes and microbial pathogens (Goto et al., 2013).

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needle-like stylet mouth extension that is found in all families of plant parasitic nematode (Hussey and Mims, 1991; Berg et al., 2009). As the giant cells begin to enlarge, the surrounding cells also divide rapidly causing swelling of the root and discontinuity of the vascular tissue. The sedentary nematode also moults a further three times as it initiates its developmental programme towards reproductive adulthood once the nutrient acquisition stage has started. During the nutrient acquisition stage, the growing nematode, giant cells, and surrounding tissue all contribute to the formation of the typical root knot symptom, which, depending on the specific characteristics of the interaction, usually resemble large tumour-like galls that are readily observed by the naked eye. A  single root knot gall can contain multiple nematode infestations and feeding sites, although each nematode induces its own distinct group of giant cells within the gall (Fig. 2). There are usually 4–10 giant cells at a single feeding site that are characterized by multiple, often endopolyploid, nuclei, a dense cytoplasm lacking a large central vacuole, a high density of organelles, and cell wall ingrowths in regions adjacent to surrounding vascular tissue, all indicative of an extremely active metabolism (Jones and Northcote, 1972a; Jones and Payne, 1978; Jones, 1981; Wiggers et  al., 1990; Starr, 1993). The morphological characteristics of these feeding sites indicate that metabolites are delivered efficiently by the plant vasculature. In addition to the induction of giant cells (reviewed in Jones, 1981; Berg et al., 2009; Gheysen and Mitchum, 2011), one of the most striking features of root-knot galls is the de

novo formation of an extensive network of xylem and phloem vascular tissues that encage the cluster of giant cells (Fig. 3). This vascularization is a process that can, in many ways, be compared to angiogenesis (Ullrich and Aloni, 2000). The tight association with xylem and phloem elements surrounding the giant cells was recognized early in classical electron micrographic studies (Jones and Northcote, 1972a; Jones and Gunning, 1976; Jones, 1981), indicative of the need for them to be supplied with water, nutrients and photosynthates from the host. However, our understanding of root-knot nematode infestation biology has been limited to some extent by a lack of genetic mutants in which this vascularization is perturbed. As a result, the mechanisms regulating the vascularization of feeding sites still need to be determined experimentally. Nevertheless, advances are being made in elucidating the function of the giant cell–vascular tissue interface through a combination of advanced molecular and cell biology approaches (Fester et al., 2008; Hoth et al., 2008; Absmanner et al., 2013).

Xylem Xylem elements are readily identifiable in histological sections of feeding sites by their often thicker cell walls and lignified secondary wall depositions (Fig. 2). Depending on which region of the three-dimensional giant cell complex is examined, an extensive network of xylem cells can be found either completely surrounding the giant cells or localized to distinct

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Fig. 1.  Schematic representation of the root-knot nematode infestation life-cycle. Root-knot nematode (N) second stage juveniles penetrate a host root and migrate to the target cells within the endodermis (En) where they induce giant cells (GC). These giant cells, which are multinucleate transfer-like cells, become the feeding cells that support the sedentary nematode for the remainder of its lifecycle. Division of surrounding cells during the induction stage probably disrupt vascular continuity, although the exact identity of the proliferating cells and specific nature of this vascular disruption remain to be experimentally verified. Vascularization of the feeding site during nutrient acquisition and restoration of vascular continuity results in an elaborate network of xylem (Xy, pink) and phloem (Ph, green) in the root galls. Once feeding commences, the nematode moults a further three times inside the root and reaches a spherical shape as an adult, after which the females lay approximately 500 eggs on the root surface in a gelatinous egg mass.

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Fig. 3.  Vascularization of nematode feeding sites. Schematic of the vascular tissue surrounding root-knot nematode-induced giant cells at the nutrient acquisition stage. The same colour scheme for the xylem (Xy), phloem (Ph), nematode (N), and giant cells (GC) is used as for Fig. 1.

regions along the giant cell border (Jones and Northcote, 1972a). This network is thought to be driven by an attempt by the plant to maintain vascular continuity following the disruption that probably occurs during giant cell induction and

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Fig. 2.  Root-knot nematode feeding sites inside host roots. (A) Giant cells (GC, one representative labelled) induced in roots of Lotus japonicus by Meloidogyne hapla root-knot nematode (N) at 2 weeks post-inoculation. Xylem elements can be identified by the thick cells walls, differential toluidine blue staining, and banding patterns of lignified cell wall thickenings (four representative cells marked with black arrowheads). Scale bar=50 μm. (Reproduced from Goto DB, Miyazawa H, Mar JC, Sato M. 2013. Not to be suppressed? Rethinking the host response at a root parasite interface. Plant Science 213, 9–17, with permission from Elsevier.) (B) Giant cells (GC, one representative labelled) induced in roots of Impatiens balsamina by the root-knot nematode Meloidogyne incognita (not visible in this section). The image shown is a scanning electron microscopy picture from an embedded and thick-sectioned gall, in which the cell contents have been etched away (partly, or fully in the case of xylem) using the same method described in Jones and Dropkin (1976). Xylem elements can be identified by the thicker cells walls and lack of cell contents (one representative cluster marked with yellow asterisks). Scale bar=approximately 25 μm.

is probably a result of de novo differentiation (Jones and Goto, 2011). An elegant method has been developed to monitor the formation of the root knot xylem networks in infested roots of Arabidopsis thaliana, which also enables parallel observation of the nematode and giant cell sinks (Fester et al., 2008). This study suggested that, at least in Arabidopsis, there are individual non-connected xylem cells around the giant cells during the juvenile feeding stages, which then develop into the characteristic network ‘cage’ once the adult nematodes begin to produce eggs (Fester et al., 2008). The delayed development of the xylem network observed in Arabidopsis may, however, be a consequence of the spatial constraints of the small Arabidopsis roots as connected xylem cells around the feeding site were observed at earlier root knot stages on the Impatiens balsamina host (Jones and Dropkin, 1976). A highly connected xylem network encasing a large portion of the giant cell complex was also apparent in Lotus japonicus root galls when the nematodes were still at a juvenile stage (Fig. 2A). One clear feature of the xylem cells that differentiate in galls around giant cells is that they are similar to wound-type xylem elements. They are typically asymmetrical and not elongated in the manner of normal xylem cells and they form irregular interconnected networks (Sinnott and Bloch, 1944, 1945; Jones and Dropkin, 1976; Aloni and Plotkin, 1985; Jones and Goto, 2011) (Fig. 4). There are also some parallels with the vascularization that occurs in rhizobium symbiosis and in galls induced by A.  tumefaciens, such as the de novo formation of both phloem and xylem that supply the structures with assimilates (Pate et al., 1969; Aloni et al., 1995). In all these cases, the mechanisms that underlie and control formation and differentiation of such xylem cells is not known, except that it is a product of parenchyma cell differentiation and probably includes some vascular cambial activity to enable extension of the xylem to interface with pre-existing vascular tissues. Root-knot nematode-induced giant cells also develop extensive wall ingrowths on walls that are adjacent to neighbouring vascular cells (Fig. 5). These wall ingrowths which, in different plant species, may be branched or flange-like

Vascularization at nematode feeding sites  |  1793

Fig. 4.  Wound-type xylem at root-knot nematode-induced feeding sites. Scanning electron micrographs of xylem vessels around feeding sites induced by Meloidogyne incognita rootknot nematode in roots of Impatiens balsamina. The cytoplasmic contents were removed by digestion prior to imaging. (A) Irregular, interconnected network of xylem cells (Xy, marked with yellow pseudocolour). The cavity where the nematode was present is pseudocoloured red and marked with N. Modified from Jones and Dropkin (1976). Scale bar=approximately 25 μm. (B) Wound-type xylem elements (Xy, only one representative labelled) bordering a giant cell (GC). Lignified secondary wall deposition patterns typical of wound-like elements are present in the xylem cells. Modified from Jones and Dropkin (1976). Scale bar=approximately 5 μm.

cells may, therefore, represent a strategy for the most efficient and direct access to nutrient requirements. Redirecting the root xylem to the giant cells would allow direct acquisition of essential amino acids and other solutes that are normally assimilated in the roots and transported to transpiring tissues for redistribution in the phloem (Urquhart and Joy, 1982; Dodd, 2005). Under this model, the phloem would also be subject to similar vascularization activity to provide enhanced access to the photosynthates and organic carbon molecules generated in the aerial tissues. Vascularization of the giant cells with respect to phloem and the physiology of the subsequent transport processes is discussed in more detail below.

structures, are like microvilli in the intestine and the plasma membrane of giant cells follows faithfully around them. Extensive wall ingrowths also form on both sides of the walls between neighbouring giant cells. Whereas in external giant cell walls pit fields are lacking, in walls between giant cells extensive pit fields with many plasmodesmatal connections are present in addition to the extensive wall ingrowths. The indication is that wall ingrowths, which are typical of transfer cells that occur naturally in various locations in plants, signal sites where there is substantial movement of water and solutes between the apoplast and symplasm of cells (Jones, 1981; Offler et  al., 2002). The vascularization that envelops giant

Phloem In contrast to the early clarity available for the distribution and differentiation of xylem elements, it was a much more complex case for the phloem associated with developing giant cells (Hoth et al., 2008). Although it is possible that a few plasmodesmata may be present between giant cells and some surrounding living cells (Jones and Dropkin, 1976), GFP expressed in companion cells under the control of the AtSUC2 promoter does not enter these cells. In addition, small fluorescent dyes do not enter giant cells in a time frame that would be consistent with symplasmic transport

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Fig. 5.  The giant cell-xylem interface. Electron micrograph of a cell wall interface between a xylem vessel (Xy) and a giant cell (GC) in Lotus japonicus at 3 weeks after inoculation with the root-knot nematode Meloidogyne hapla. Secondary wall depositions in the xylem vessel are marked (black arrowheads, three marked as representative). The presence of extensive wall ingrowths in the giant cells (white arrowheads, three marked as representative) is indicative of substantial water and solute movement across this interface. Scale bar=2 μm.

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Fig. 6.  Phloem vascularization of root-knot nematode feeding sites in Arabidopsis. Cross-section through an Arabidopsis rootknot gall 3 weeks after inoculation with Meloidogyne incognita root-knot nematode (N). Giant cells (asterisks) are encaged in sieve elements (red), which are decorated with a sieve elementspecific marker antibody (Khan et al., 2007). Nuclei are stained with DAPI. Note the enlarged nuclei visible in one of the giant cells. Nucleated sieve elements are indicated with a white arrow, xylem vessels are indicated with the white arrowheads. Scale bar=50 μm.

around giant cells are highly interconnected by plasmodesmata, thereby forming an extensive symplasmic region that appears to serve as an unloading domain. The consequences, in terms of nutrient unloading into the apoplast as a prerequisite for transporter-mediated uptake into the symplasmically isolated giant cells, are discussed in the following section. The observation that the phloem around giant cells does not have companion cells raises the question about the underlying principles that govern this differential vascularization. Vascular proliferation in plants can be regulated by a secreted CLE peptide, which binds to a plasma membrane-localized receptor and controls cell division rates and differentiation events via a WUSCHEL-like transcription factor (Miyashima et al., 2013). It is plausible that root-knot nematodes could secrete a CLE-like peptide into giant cells, although the only current candidate for such a peptide was suggested to interact with a transcription factor indicating that its action would be exerted in a different fashion (Huang et al., 2006). Furthermore, when this peptide was expressed in plant roots, there was no observed effect on the vascular tissue. In a recent study, the role of the two phytohormones, auxin and cytokinin, was investigated in the phloem throughout feeding site development (Absmanner et al., 2013). Two well-established markers, DR5 and TCS promoter activities, were used to visualize the transcriptional response to auxin or cytokinin, respectively (Ulmasov et  al., 1999; Müller and Sheen, 2008). Immunohistochemistry was also applied to co-localize hormone-responsive cells and sieve elements. The first striking observation was that a constitutive auxin response occurs in companion cells in fully differentiated, uninfested wild-type roots (Absmanner et al., 2013). This was somewhat surprising because the auxin response has previously been associated with the xylem cell lineage in the root tip (Bishopp et al., 2011a, b). This finding also shows that the developmental processes that occur in the phloem in mature sections of the root have been poorly investigated. In rootknot nematode infested roots it was demonstrated that cells around the giant cell exhibited an auxin response prior to their differentiation into sieve elements. All cells that differentiated into sieve elements in the later stages of development had undergone an auxin response, but these sieve elements did not respond to cytokinin. These findings are consistent with a study that showed the Arabidopsis response regulator (ARR) 5, a negative regulator of the cytokinin response, was strongly induced around giant cells (Lohar et al., 2004). Interestingly, a response to either of the two hormones within the mature feeding cells themselves was not detected. These results suggest that the ratio of auxin to cytokinin governs the developmental processes associated with phloem differentiation at root-knot nematode feeding sites. The above work also highlighted the difficulties in identifying phloem cells from histological sections and the phloem was only identified in the above-described study because a sieve element identity marker for Arabidopsis had become available. Without such an identity marker, it is not always possible to distinguish which cells are the phloem, which cells are sieve elements, or which cells are just small parenchyma cells. In early electron microscopy studies the presence

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(Dorhout et al., 1993). Based on these data, it was suggested very early that giant cells are relatively symplasmically isolated and nutrient and assimilate loading occurs by transporters (Jones, 1981). When the phloem cells in root-knot galls and particularly in the vicinity of giant cells were examined in detail using a combination of fluorescence microscopy and immunohistochemistry, it became clear that the phloem in this case is strikingly different from that typically observed in other plant tissues, including the regenerative phloem that forms after wounding or even in other types of plant–biotic interactions (Behnke and Schulz, 1980; Jones, 1981; Schulz, 1986; Hoth et al., 2008; Absmanner et al., 2013). Extensive formation of phloem is induced around giant cells. In some regions, only phloem cells and very few other cell types are found directly adjacent to giant cells (Fig.  6). The interesting feature is that the phloem in which giant cells become virtually embedded does not contain typical companion cells, and exists exclusively of sieve elements that are frequently nucleated and able to perform transcriptional responses (Hoth et  al., 2008; Absmanner et  al., 2013). It is not clear why the companion cells are absent, one explanation is that the companion cells may have been consumed in the vascularization process. Alternatively, they may simply stop expressing the SUC2 gene that defines their identity, which can be reconciled in a physiological sense as this would impair re-uptake of sucrose from the apoplast into the phloem and direct sucrose flow to the giant cells. The sieve elements

Vascularization at nematode feeding sites  |  1795 of ingrowths on regions of the giant cell wall adjacent to small parenchyma cells in addition to more obvious sieve elements was noted, suggesting these too could have been sieve elements (Jones and Northcote, 1972a). It is likely that an updated reassessment of cells surrounding giant cells in other plant systems will reveal a greater presence of phloem than previously thought, although such studies are difficult to achieve because of the requirement for suitable identity markers compatible with the host species being studied.

Physiology and transport processes

Fig. 7.  Apoplastic transport of assimilates into giant cells. Schematic representation of the interface between a sieve element (SE) and a giant cells (GC). The presence of amino acid exporters (UmamiT) (Ladwig et al., 2012) and sucrose exporters, SWEETs (Chen et al., 2012) are only postulated as experimental evidence is not yet available. In Arabidopsis, the proton-coupled amino acid importers AAP6 and CAT6, and the sucrose transporter (SUC1) have been localized to giant cells (Hammes et al., 2005, 2006). The proton gradient necessary to drive uptake would be provided by plasma membrane ATPases, such as PMA4 that has been identified in Solanum lycopercicon (tomato; Bird and Wilson, 1994).

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Giant cells are strong metabolic sinks and it has long been recognized that they are characterized by cell wall ingrowths that increase their plasma membrane surface area (Jones and Northcote, 1972a, b; Jones et  al., 1975; Jones, 1981; Golinowski et al., 1996; Hussey and Grundler, 1998; Hammes et al., 2005). This feature is typically associated with transfer cells that can be found in numerous tissues and are associated with enhanced transport activities (Offler et al., 2002). The wall ingrowths are normally much more extensive where giant cells contact xylem cells compared with where they contact phloem sieve elements, and the amplification of the area of the plasma membrane at sites of extensive wall ingrowth formation has been estimated at up to 15 times (Jones and Dropkin, 1976). Interestingly, it appears that when wall ingrowths do develop opposite the phloem in giant cells they are only of the ‘xylem type’. In most plant species, xylem transfer cells have the same type of ingrowth morphology as phloem transfer cells and it is difficult to distinguish them based on ingrowth morphology alone. However, in the case of Helianthemum, the morphology of xylem and phloem wall ingrowths differ and where the same giant cell contacts both xylem and sieve elements the giant cell ingrowths that form only have the xylem ingrowth morphology (Jones and Gunning, 1976). The mechanisms that govern the formation of the transfer cell-like wall modifications remain elusive. Nevertheless, it seems clear that they serve to increase the cell surface and support fluxes of water and/or nutrients into the root-knot nematode feeding sites. Since nutrient uptake by giant cells from the phloem is not symplasmic, it is dependent on transport across the membranes (Hoth et  al., 2008). Two membranes need to be crossed in these processes, first the sieve element plasma membrane and, subsequently, the giant cell plasma membrane. The molecular players in these processes remain largely unknown but some progress has been made. In order to identify transporters that play a role in providing nutrients to giant cells, a microarray study was performed (Hammes et al., 2005). Entire roots containing galls at 1, 2, and 4 weeks after infestation with root-knot nematodes were examined and compared with non-infested control roots of the same age. Using a stringent statistical analysis it was found that 50 genes encoding transporters are regulated in response to infestation (Hammes et al., 2005). As originally suggested by Jones et al. (1975), the transporters implicated in the uptake of assimilates were all shown to work as proton coupled symporters, which require a proton gradient driven by the

activity of H+-ATPases (Fig. 7). Elevated expression of such a gene has been found at root-knot nematode infestation sites in tomato roots (Bird and Wilson, 1994), further supporting the concept that metabolite fluxes by transporter-mediated processes are important in plant-root knot nematode interactions. Interestingly, the majority of genes that were found to be induced in the root-knot nematode–Arabidopsis microarray study were specifically up-regulated in the galls that contained giant cells and surrounding tissue including the vasculature. A considerable number of these transporters act to transport amino acids or peptides and there was specific up-regulation of some of these in giant cells (Hammes et al., 2005, 2006). In many plant–pathogen interactions there is also good evidence that amino acids play a crucial role in the interaction, and changes in amino acid levels and distribution have also been described in the interaction between maize and Ustilago maydis in addition to plant–nematode interactions (Krauthausen and Wyss, 1982; Hedin and Creech, 1998; Horst et  al., 2009). Root-knot nematodes are heterotrophic organisms and so this need to take up essential amino acids in their diet is not unexpected.

1796 | Bartlem et al. the bacterial pathogen Xanthomonas oryzae, which secretes a transcriptional activator-like protein that directly binds to the OsSWEET11 promoter to induce expression of the transporter. The pathogen induced SWEETs were confirmed to mediate transport of monosaccharides along the concentration gradient, consistent with a role in sugar efflux from the cytosol to the apoplast where they could then be utilized by the pathogen (Chen et al., 2010). It is plausible that SWEET proteins are also involved in the efflux of sucrose from the phloem at root-knot nematode-induced giant cells, although this is yet to be experimentally confirmed.

Concluding remarks Our understanding of long- and short-distance transport processes in plant–nematode interactions is still fragmentary and far from being fully understood. However, it is clear that infestation of plant roots with endoparasitic nematodes such as root-knot nematodes makes such plants more prone to water stress and they also show symptoms of nutrient deficiency and premature senescence. With impending climate change, less availability of irrigation water for crops, and rising fertilizer prices, it is important that a better understanding can be gained of how nematode infestation impairs root function and reduces yields and product quality. What proportion is from the disruption of vascular continuity and efficient functioning and what proportion is from diverting nutrients to the growing nematodes? How do these plant pests control plant cell differentiation? It is clear that a better understanding of nematode–plant interactions is needed and, with such understanding, the knowledge gained can be used to reduce losses caused by these important plant pests and improve nematode control.

Funding Work in the authors’ laboratories is supported by Grantsin-Aid from the Japanese Ministry of Education, Culture, Sports, Science, and Technology (#24687005, #24113501, and #25114503 to DGB), a Research Collaboration Award from the National Institute for Basic Biology, Japan (#13–312 to DGB), Murdoch University and Australian Research Council Grants (MGKJ), a Sir Walter Murdoch Distinguished Collaborator Award from Murdoch University, Australia (DGB and MGKJ), and by the Deutsche Forschungsgemeinschaft (DFG HA 3468/4-1 and through the SFB924 to UZH).

Acknowledgements We are grateful to Toshiaki Ito (Electron Microscope Laboratory, Faculty of Agriculture, Hokkaido University) for assistance with the high-resolution analysis of root-knot nematode feeding sites in Lotus japonicus, to Shuhei Hayashi for images of giant cells in Lotus japonicus, and to Yuka Naraki (Space-Time Inc., Japan) for artwork.

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One of the transporters up-regulated in Arabidopsis thaliana giant cells is cationic amino acid transporter 6 (AtCAT6) (Hammes et al., 2006). This transporter is a member of the amino acid–polyamine–choline (APC) transporter family of the amino acid transporter superfamily (ATF) found in the Arabidopsis genome (Wipf et  al., 2002). In addition to the expression in giant cells, the gene is also expressed in endogenous sink tissues such as the endosperm and seeds. Using Xenopus laevis oocyte expression and the two electrode voltage clamp (TEVC) technique, it was shown that AtCAT6 mediates electrogenic, proton-driven transport of many amino acids with a pronounced preference for large, bulky, and hydrophobic side chains that includes seven out of the eight essential amino acids (Hammes et  al., 2006). Interestingly, atcat6 knock-out mutants fail to grow on glutamine as the sole nitrogen source, indicating a crucial role for AtCAT6 in the transport of this particular amino acid. Root-knot nematode development was not affected in the atcat6 mutant, supporting the idea that several transporters with possibly overlapping substrate specificity operate in giant cells (Hammes et al., 2006). Nevertheless, amino acid transporters have been confirmed to play a role in the interaction between Arabidopsis and root-knot nematodes. The relative proportion of nematode juveniles that matured into females was reduced when feeding on knock-out plants of the amino acid permease genes AtAAP3 and AtAAP6 compared with that observed on wild-type controls (Marella et al., 2013). This effect could have been caused by an altered amino acid distribution in general and not directly linked to the feeding site because aap6 mutants also display an altered amino acid composition of the phloem sap (Hunt et  al., 2010). The molecular nature of the transporters that facilitate amino acid release from the phloem into the apoplast around giant cells is currently unknown and the recently identified bi-directional amino acid facilitators (now annotated as UmamiTs) are promising candidates in this context (Ladwig et al., 2012). Despite their obvious importance in the transport of sugars and particularly the major osmolyte sucrose, the role of sugar transporters in the interaction with root-knot nematodes is less well understood. In earlier studies, electrophysiological examination of the responses of giant cells to added sugars showed a transient depolarization on the addition of a range of sugars, suggesting the involvement of H+ cotransporters (Jones et al., 1975; Jones, 1981). More recently, in root-knot nematode infested Arabidopsis roots a significant induction of AtSUC1 in root knots was observed but its role in the plant–nematode interaction remains to be studied in detail. The molecular identity of the transporters that would mediate efflux of sugar from the phloem around the giant cells is also unknown. To date there is only one group of transporters, the SWEETs, described to be directly regulated by pathogens to provide sugar for their nutrition or the release of sucrose into the apoplast (Chen et al., 2010, 2012). Although not annotated as such at the time, up-regulation of SWEET transcripts was also found in the microarray analysis mentioned above (Hammes et  al., 2005). It was shown that the OsSWEET1 transporter gene in rice is directly induced by

Vascularization at nematode feeding sites  |  1797

References Absmanner B, Stadler R, Hammes UZ. 2013. Phloem development in nematode-induced feeding sites: The implications of auxin and cytokinin. Frontiers in Plant Science 4, 241. Aloni R, Plotkin T. 1985. Wound-induced and naturally occurring regenerative differentiation of xylem in Zea mays L. Planta 163, 126–132. Aloni R, Pradel KS, Ullrich CI. 1995. The three-dimensional structure of vascular tissues in Agrobacterium tumefaciens-induced crown galls and in the host stems of Ricinus communis L. Planta 196, 597–605. Behnke H-D, Schulz A. 1980. Fine structure, pattern of division, and course of wound phloem in Coleus blumei. Planta 150, 357–365.

Goto DB, Miyazawa H, Mar JC, Sato M. 2013. Not to be suppressed? Rethinking the host response at a root parasite interface. Plant Science 213, 9–17. Grundler FMW, Sobczak M, Golinowski W. 1998. Formation of cell wall openings in root cells of Arabidopsis thaliana following infection by the plant-parasitic nematode Heterodera schachtii. European Journal of Plant Pathology 104, 545–551. Grunewald W, van Noorden G, Van Isterdael G, Beeckman T, Gheysen G, Mathesius U. 2009. Manipulation of auxin transport in plant roots during Rhizobium symbiosis and nematode parasitism. The Plant Cell 21, 2553–2562. Hammes UZ, Nielsen E, Honaas LA, Taylor CG, Schachtman DP. 2006. AtCAT6, a sink-tissue-localized transporter for essential amino acids in Arabidopsis. The Plant Journal 48, 414–426.

Bird DM, Wilson MA. 1994. DNA sequence and expression analysis of root-knot nematode-elicited giant cell transcripts. Molecular Plant– Microbe Interactions 7, 419–424.

Hammes UZ, Schachtman DP, Berg RH, Nielsen E, Koch W, McIntyre LM, Taylor CG. 2005. Nematode-induced changes of transporter gene expression in Arabidopsis roots. Molecular Plant– Microbe Interactions 18, 1247–1257.

Bishopp A, Help H, El-Showk S, Weijers D, Scheres B, Friml J, Benkova E, Mahonen AP, Helariutta Y. 2011a. A mutually inhibitory interaction between auxin and cytokinin specifies vascular pattern in roots. Current Biology 21, 917–926.

Hedin PA, Creech RG. 1998. Altered amino acid metabolism in rootknot nematode inoculated cotton plants. Journal of Agricultural and Food Chemistry 46, 4413–4415.

Bishopp A, Lehesranta S, Vaten A, Help H, El-Showk S, Scheres B, Helariutta K, Mahonen AP, Sakakibara H, Helariutta Y. 2011b. Phloem-transported cytokinin regulates polar auxin transport and maintains vascular pattern in the root meristem. Current Biology 21, 927–932. Böckenhoff A, Grundler FMW. 1994. Studies on the nutrient uptake by the beet cyst nematode Heterodera schachtii by in situ microinjection of fluorescent probes into the feeding structures in Arabidopsis thaliana. Parasitology 109, 249–255. Chen LQ, Hou BH, Lalonde S, et al. 2010. Sugar transporters for intercellular exchange and nutrition of pathogens. Nature 468, 527–532. Chen LQ, Qu XQ, Hou BH, Sosso D, Osorio S, Fernie AR, Frommer WB. 2012. Sucrose efflux mediated by SWEET proteins as a key step for phloem transport. Science 335, 207–211. Dodd IC. 2005. Root-to-shoot signalling: assessing the roles of ‘up’ in the up and down world of long-distance signalling in planta. Plant and Soil 274, 251–270. Dorhout R, Gommers FJ, Kollöffel C. 1993. Phloem transport of carboxyfluorescin through tomato roots infected with Meloidogyne incognita. Physiological and Molecular Plant Pathology 43, 1–10. Fester T, Berg RH, Taylor CG. 2008. An easy method using glutaraldehyde-introduced fluorescence for the microscopic analysis of plant biotrophic interactions. Journal of Microscopy (Oxford) 231, 342–348. Gheysen G, Mitchum MG. 2011. How nematodes manipulate plant development pathways for infection. Current Opinion in Plant Biology 14, 415–421. Golinowski W, Grundler FMW, Sobczak M. 1996. Changes in the structure of Arabidopsis thaliana during female development of the

Horst RJ, Doehlemann G, Wahl R, Hofmann J, Schmiedl A, Kahmann R, Kamper J, Sonnewald U, Voll LM. 2009. Ustilago maydis infection strongly alters organic nitrogen allocation in maize and stimulates productivity of systemic source leaves. Plant Physiology 152, 293–308. Hoth S, Schneidereit A, Lauterbach C, Scholz-Starke J, Sauer N. 2005. Nematode infection triggers the de novo formation of unloading phloem that allows macromolecular trafficking of green fluorescent protein into syncytia. Plant Physiology 138, 383–392. Hoth S, Stadler R, Sauer N, Hammes UZ. 2008. Differential vascularization of nematode-induced feeding sites. Proceedings of the National Academy of Sciences, USA 105, 12617–12622. Huang G, Dong R, Allen R, Davis EL, Baum TJ, Hussey RS. 2006. A root-knot nematode secretory peptide functions as a ligand for a plant transcription factor. Molecular Plant–Microbe Interactions 19, 463–470. Hunt E, Gattolin S, Newbury HJ, Bale JS, Tseng HM, Barrett DA, Pritchard J. 2010. A mutation in amino acid permease AAP6 reduces the amino acid content of the Arabidopsis sieve elements but leaves aphid herbivores unaffected. Journal of Experimental Botany 61, 55–64. Hussey RS, Grundler FM. 1998. Nematode parasitism of plants. In: Perry RN, Wright DJ, eds. Physiology and biochemistry of free-living and plant-parasitic nematodes . Wallingford: CAB International Press, 213–243. Hussey RS, Mims CW. 1991. Ultrastructure of feeding tubes formed in giant-cells induced in plants by the root-knot nematode Meloidogyne incognita. Protoplasma 162, 99–107. Jones MGK. 1981. The development and function of plant cells modified by endoparasitic nematodes. In: Zuckerman BM, Rohde RA, eds. Plant parasitic nematodes , Vol. 3. New York: Academic Press, 255–279.

Downloaded from http://jxb.oxfordjournals.org/ at Simon Fraser University on June 8, 2015

Berg RH, Fester T, Taylor CG. 2009. Development of the root-knot nematode feeding cell. In: Berg RH, Taylor CG, eds. Cell biology of plant nematode parasitism . Berlin, Heidelberg: Springer, 115–152.

plant-parasitic nematode Heterodera schachtii. Protoplasma 194, 103–116.

1798 | Bartlem et al. Jones MGK, Dropkin VH. 1976. Scanning electron microscopy of nematode-induced giant transfer cells. Cytobios 15, 149–161. Jones MGK, Goto DB. 2011. Root-knot nematodes and giant cells. In: Jones J, Gheysen G, Fenoll C, eds. Genomics and molecular genetics of plant–nematode interactions . Dordrecht: Springer, 83–100. Jones MGK, Gunning BES. 1976. Transfer cells and nematode induced giant cells in Helianthemum. Protoplasma 87, 273–279. Jones MGK, Northcote DH. 1972a. Multinucleate transfer cells induced in Coleus roots by the root-knot nematode, Meloidogyne arenaria. Protoplasma 75, 381–395.

Oldroyd GE, Murray JD, Poole PS, Downie JA. 2011. The rules of engagement in the legume–rhizobial symbiosis. Annual Review of Genetics 45, 119–144. Pate JS, Gunning BES, Briarty LG. 1969. Ultrastructure and functioning of the transport system of the leguminous root nodule. Planta 85, 11–34. Prell J, White JP, Bourdes A, Bunnewell S, Bongaerts RJ, Poole PS. 2009. Legumes regulate Rhizobium bacteroid development and persistence by the supply of branched-chain amino acids. Proceedings of the National Academy of Sciences, USA 106, 12477–12482.

Jones MGK, Novacky A, Dropkin VH. 1975. Transmembrane potentials of parenchyma cells and nematode-induced transfer cells. Protoplasma 85, 15–37.

Scholl EH, Bird DM. 2005. Resolving tylenchid evolutionary relationships through multiple gene analysis derived from EST data. Molecular Phylogenetics and Evolution 36, 536–545.

Jones MGK, Payne HL. 1978. Early stages of nematode-induced giant-cell formation in roots of Impatiens balsamina. Journal of Nematology 10, 70–84.

Schulz A. 1986. Wound phloem in transition to bundle phloem in primary roots of Pisum sativum L. Protoplasma 130, 12–26.

Khan JA, Wang Q, Sjolund RD, Schulz A, Thompson GA. (2007). An early nodulin-like protein accumulates in the sieve element plasma membrane of Arabidopsis. Plant Physiology 143, 1576–1589. Krauthausen H-J, Wyss U. 1982. Influence of the cyst nematode Heterodera schachtii on relative changes in the pattern of free amino acids at feeding sites. Physiological Plant Pathology 21, 425–436. Ladwig F, Stahl M, Ludewig U, Hirner AA, Hammes UZ, Stadler R, Harter K, Koch W. 2012. Siliques are red1 from Arabidopsis acts as a bidirectional amino acid transporter that is crucial for the amino acid homeostasis of siliques. Plant Physiology 158, 1643–1655. Lohar DP, Schaff JE, Laskey JG, Kieber JJ, Bilyeu KD, Bird DM. 2004. Cytokinins play opposite roles in lateral root formation, and nematode and rhizobial symbioses. The Plant Journal 38, 203–214. Marella HH, Nielsen E, Schachtman DP, Taylor CG. 2013. The amino acid permeases AAP3 and AAP6 are involved in root-knot nematode parasitism of Arabidopsis. Molecular Plant–Microbe Interactions 26, 44–54.

Sinnott EW, Bloch R. 1944. Visible expression of cytoplasmic pattern in the differentiation of xylem strands. Proceedings of the National Academy of Sciences, USA 30, 388–392. Sinnott EW, Bloch R. 1945. The cytoplasmic basis of intercellular patterns in vascular differentiation. American Journal of Botany 32, 151–156. Sobczak M, Golinowski W. 2011. Cyst nematodes and syncytia. In: Jones J, Gheysen G, Fenoll C, eds. Genomics and molecular genetics of plant–nematode interactions . Dordrecht: Springer, 61–82. Starr JL. 1993. Dynamics of the nuclear complement of giant cells induced by Meloidogyne incognita. Journal of Nematology 25, 416–421. Ullrich CI, Aloni R. 2000. Vascularization is a general requirement for growth of plant and animal tumours. Journal of Experimental Botany 51, 1951–1960. Ulmasov T, Hagen G, Guilfoyle TJ. 1999. Dimerization and DNA binding of auxin response factors. The Plant Journal 19, 309–319. Urquhart AA, Joy KW. 1982. Transport, metabolism, and redistribution of xylem-borne amino acids in developing pea shoots. Plant Physiology 69, 1226–1232.

Miyashima S, Sebastian J, Lee JY, Helariutta Y. 2013. Stem cell function during plant vascular development. EMBO Journal 32, 178–193.

Wiggers RJ, Starr JL, Price HJ. 1990. DNA content and variation in chromosome number in plant cells affected by Meloidogyne incognita and M. arenaria. Phytopathology 80, 1391–1395.

Müller B, Sheen J. 2008. Cytokinin and auxin interaction in root stem-cell specification during early embryogenesis. Nature 453, 1094–1097.

Wipf D, Ludewig U, Tegeder M, Rentsch D, Koch W, Frommer WB. 2002. Conservation of amino acid transporters in fungi, plants and animals. Trends in Biochemical Sciences 27, 139–147.

Offler CE, McCurdy DW, Patrick JW, Talbot MJ. 2002. Transfer cells: cells specialized for a special purpose. Annual Review of Plant Biology 54, 431–454.

Wyss U, Grundler FMW, Munch A. 1992. The parasitic behaviour of second-stage juveniles of Meloidogyne incognita in roots of Arabidopsis thaliana. Nematologica 38, 98–111.

Downloaded from http://jxb.oxfordjournals.org/ at Simon Fraser University on June 8, 2015

Jones MGK, Northcote DH. 1972b. Nematode-induced syncytium: a multinucleate transfer cell. Journal of Cell Science 10, 789–809.

Sasser JN, Freckman DW. 1987. A world perspective on nematology: the role of the society. In: Dickson DW, Veech J, eds. Vistas in nematology . Hyattsville, MD: Society of Nematologists Inc., 7–14.