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Concerted Efforts To Develop Handles For Plant Parasitic Nematode Control
Phytosfere'99 - Highlights in European Plant Biotechnology Gert E. de Vries and Karin Metzlaff (Editors). 9 Elsevier Science B.V. All rights reserved.
Concerted Efforts To Develop Handles For Plant Parasitic Nematode Control
Abstract Plant-parasitic nematodes - especially root knot and cyst nematodes - are economically important pests in numerous crops. Chemical soil sterilisation and the use of other unselective pesticides to control plant parasitic nematodes are still a common practice in many European countries and at present no realistic alternatives are available. For the identification of handles to control root knot and cyst nematodes we need to know how they interact with their host. Two main types of endoparasitic nematodes can be distinguished: root-knot and cyst nematodes, both inducing a feeding site in the plant root, but in a different way. To get insight into the molecular mechanisms behind this complex interaction, several strategies to analyse plant gene expression in response to nematode infection have been followed. Random in vivo gus fusions have been particularly successful in identifying plant promoter sequences that are highly activated in nematode feeding sites, with very little expression elsewhere in the plant, but the isolation of the corresponding genes is often not straightforward. Many highly transcribed plant genes have been identified in the feeding sites, but few have been characterised in such detail as to know how important they are for a successful infection. The available data are nevertheless providing interesting tools for novel strategies to engineer nematode resistance into crops. Concomitantly, the signals coming from the nematode that are triggering this plant response or that are important in other steps of the infection process are being characterised. This study has revealed that plant-parasitic nematodes produce many different enzymes to enable them to infect the plant root and to protect themselves against the plant defence response.
Introduction Sedentary plant-parasitic nematodes are important pests of agricultural crops. Traditional management of plant-parasitic nematodes is relying on three basic strategies: crop rotation, agrochemicals, and the use of resistant plants. Crop rotation is sometimes economically disadvantageous and is often not effective, for example in the case of the root-knot nematode Meloidogyne incognita that has a very broad host range. Chemical control is expensive, effective nematicides are difficult to contain and have a broad toxic spectrum, including animals and humans. Host resistance is the most environmentally and economically sound method [ 1]. Unfortunately, natural resistance to plant parasitic nematodes is not available for all cul-
Godelieve Gheysen, Departement Plantengenetica, Universiteit Gent, Belgium
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Challenges of the Environment tivated crop species. Furthermore, a very important limiting factor to the general usefulness of natural resistance genes is the specific nature of their protection, which is often restricted to one nematode species or even certain pathotypes [2]. An alternative strategy that tries to combine effectiveness, broad applicability and durability with environmental friendliness is the engineering of nematode resistance into plants. In the last few years several excellent reviews have been written on this research development, and the reader is referred to them for more general aspects [3-4]. Here, we will summarise the breakthroughs of the last years, mainly in the light of the co-ordinated efforts done by the 12 laboratories involved in the ARENA project of the European Union.
Morphological and histological changes Sedentary nematodes establish and maintain an intimate relationship with their host. After invading the plant root, they migrate to the vascular cylinder in search for a cell that can serve as an initial feeding cell. In response to repeated stimulation by the parasite, this cell develops into either a syncytium (for cyst nematodes, such as Globodera and Heterodera), or several cells are stimulated to form a system of giant cells (as is the case for root-knot nematodes). The combination of light and electron microscopy studies of fixed samples and time-lapse video microscopy on living material has provided detailed insight in the establishment and structure of the nematode feeding site [5-6, reviewed in 3]. The syncytium is formed by breakdown of plant cell wails and subsequent fusion of neighbouring cells. A giant cell is formed as the result of repeated nuclear divisions without cytokinesis. Although these feeding sites differ from each other in ontogenesis and structure, their function is the same: to supply the nematode with sufficient nutrients for growth and reproduction. This common function is reflected in a final analogous ultrastructure. Syncytial and giant cells are hypertrophied and multinucleated cells with a dense granular cytoplasm, an increase in rough endoplasmic reticulum and in the number of mitochondria. Recently the formation of cell wall openings occurring during syncytium differentiation in A. thaliana roots was studied in more detail [7]. The first openings are formed by gradual widening of existing plasmodesmata followed by the fusion of protoplasts from two adjacent cells. At later stages, wall openings form exclusively in cell walls without the involvement of plasmodesmata. The sequence of events leading to these openings is: accumulation of ER membranes accompanying initial wall lesions, expansion of lesions until the middle lamella, and finally dissolution of the middle lamella [7]. Functional plasmodesmata were never found in the outer wall of mature syncytia, although they are present in the walls of the adjacent sieve elements. These plasmodesmata were always closed from the syncytium side by wall material. It can be concluded that the formation of a syncytium is accompanied with many changes in cell wall biology: callose-like deposition around the stylet, cell wall ingrowths adjacent to the xylem, widening or closing of plasmodesmata depending on their location in the syncytium, and de novo formation of wall openings. It will be interesting to see which enzymes are involved in these processes and how they are temporally and spatially controlled. Another aspect of nematode feeding site formation that is being unravelled is the activation of the cell cycle by the nematode stimulus. Sedentary endoparasitic nematodes induce multinucleate feeding cells in the roots of their host plants. These cells undergo multiple rounds of 160
Plant parasitic nematode control shortened cell cycles leading to genome amplification and hypertrophy of the cytoplasm. This aspect of the infection process has been thoroughly reviewed by [8] and more recently by [9]. It is now generally believed that giant cells develop by repeated mitosis without cytokinesis [5]. Jones and Payne [10] showed that cell plate vesicles initially lined up between the two daughter nuclei but then dispersed, resulting in the abortion of the new cell plate formation. No mitosis has been seen in syncytia [11 ], although Piegat and Wilski [ 12] reported an initial mitotic stimulation during syncytium induction by Globodera rostochiensis in infected potato roots. However, it is not clear from the data shown whether this mitosis is really inside the developing syncytium. Indeed, stimulation of cell division has been observed in parenchyma cells surrounding the syncytium and which will be incorporated [13-14]. The enlargement of nuclei indicates that DNA multiplication is taking place within the syncytial tissue during and after the incorporation of new cells through cell wall dissolution [ 11]. Studies with cell cycle inhibiting drugs indicate that mitosis is essential for giant cell development and for normal syncytium expansion [ 14].
Nematode secretions Stylet secretions that originate in the esophageal gland cells of plant parasitic nematodes are believed to play a major role in pathogenesis. Secretions from the dorsal gland cells have always been considered critical for pathogenesis but till recently a role for the subventral glands in the infection process has been questioned. Subventral gland secretions are released into the pharyngeal lumen behind the pump chamber and were thought only to be able to pass posteriorly into the intestine. However, immunolocalisation of HG-ENG-2 demonstrated the presence of this protein in the nematode's subventral glands (where it is produced), as well as in the root cortical tissue after secretion from the stylet [ 15]. This HG-ENG-2 protein belongs to a family of f~-l,4-endoglucanases first identified in secretions of cyst nematodes by [16], and later also in root knot nematodes [ 17]. These enzymes presumably facilitate the migration of the nematode through plant roots by partial cell wall degradation. The corresponding genes represent the first cellulase genes ever cloned from an animal and show highest homology (35-40%) with 13-1,4-endoglucanases from bacteria [16]. Other lytic enzymes identified in the nematode secretions include pectate lyase and proteases (Herman Popeijus and John Jones, unpublished results). Other proteins that are present in nematode secretions (although not necessarily from the esophageal glands) appear to be related to the active protection of the nematode against the plants defense response. GP-SEC2 functions as a fatty acid binding protein capable of binding e.g. linolenic and linoleic acids, that are important precursors in plant defence signalling pathways. In addition, superoxide dismutase and thioredoxin peroxidase secreted by the nematode are involved with inactivation of superoxide radicals and hydrogen peroxide respectively. How important these proteins or the above mentioned lytic enzymes are for nematode pathogenesis, can only be decided if inactivation or absence of these proteins results in a lower percentage of successful nematode infections. Inhibition of the proteins could be achieved by the expression of plantibodies [ 18]. Inactivation of the genes is another theoretical possibility, but transformation of plant-parasitic nematodes is still virtually impossible [19]. A strategy that could be useful is RNA interference, a method capable of efficiently eliminating specific mRNAs in C. elegans [20]. 161
Challenges of the Environment The most intriguing function of the esophageal secretions, their role in formation of the feeding site, is still a black box. Unfortunately, in contrast to comparative studies of nematode development and the genes involved, analysis of the model nematode C. elegans does not help much in respect to pathogenesis-involved genes. However, with the current expertise and molecular technology, at least some clues about possible signalling molecules in the secretions of plant-parasitic nematodes should be identified in the near future.
Screening Screening for nematode-responsive plant regulatory sequences was done by by T-DNA tagging. Plant genes that are up-regulated after nematode infection have first been identified by differential screening of cDNA libraries [21-23] or from subtraction cDNA libraries [24] and later by differential display [25, Vercauteren et al., accepted by MPMI]. To identify promoters that are early and specifically induced in the nematode feeding sites, the most appropriate method is undoubtedly the analysis of promoter-reporter fusions (for example with 13-glucuronidase [gus]). Plant-gus fusions can be made at random in vivo and then screened for specific expression patterns. This "promoter trapping" or "tagging" approach is based on the random integration of a promoterless gus gene after transformation into a plant species, often A. thaliana [26]. When inserted downstream from a plant promoter that is inducible by nematode infection, a higher GUS activity is expected to be seen at the infection site (Fig. 1). The elegance of this method lies in the ability to directly visualise induced expression in nematode feeding sites at various stages of the interaction, while analysing the specificity of expression using uninfected parts of the same plant and control plants. Furthermore, promising plant lines can easily be tested for gus activation upon infection by different types of nematodes. This "tagging" approach has been performed in several labs of the ARENA project and has yielded many interesting promoters [27-30]. The hope was that the homozygous progeny of these T-DNA tagged lines could be knock-outs for the tagged gene,
Figure 1: GUS activity in a Att0025-R/1 syncytium 6 days post inoculation with H e t e r o d e r a s c h a c h t i i (from: Barthels et al. 1997, Plant Cell 9:2119-2134)
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Plant parasitic nematode control
and would therefore yield information on the role of this plant gene, in general or possibly even during nematode infection. Initially none of the lines analysed had an obvious phenotype [28], strengthening the belief that tagging often yields cryptic promoters [31 ]. Although these cryptic promoters are not less useful for expressing a transgene in the nematode feeding sites, from a more fundamental point of view it would be informative to find the endogenous genes that are specifically activated after nematode infection. However, more recently one of the tagged lines was found to have a T-DNA insertion in the rpe gene (D-ribulose-5-phosphate-3-epimerase) resulting in a mutant phenotype. Homozygous rpe plants have a germination deficient phenotype that can be rescued in dwarf plants on sucrose-supplemented medium [30]. Also, upon further study, several other T-DNA insertions appeared to be linked to a "tagged" gene, although in unexpected configuration, such as inside the promoter instead of downstream, and even several kb away from the gene (line Att0001 (M.Karimi), line Att0025 (S. Ohl), line Att0884 (P.Puzio); unpublished observations).
Transcriptional control of nematode-inducible genes In the light of applications, the most interesting A. thaliana promoters are also introduced into crop species to check their activity after nematode infection. The Art0001 (ARM 1) promoter, as well as some other A. thaliana promoters, are clearly up-regulated after nematode infection in oilseed rape and sugarbeet [32, G. Gheysen unpublished], but not in potato, whereas the rpe promoter is activated in potato nematode feeding sites [B. Favery, unpublished]. Besides possible differences in cell-specific or temporal gene regulation between plant species, another reason could be that in potato, the syncytium is formed in cortex cells [33, 34], while in A. thaliana procambium cells are normally used [13]. Maybe the apparently contradictory results on CaMV35S-promoter activity inside syncytia could also be explained by the different plant species used. Goddijn et al. [27] and Urwin et al. [35] describe down-regulation of the 35S promoter in syncytia induced by H. schachtii in A. thaliana, and this was also observed in sugarbeet (G. Gheysen, unpublished), but not in potato syncytia induced by G. rostochiensis [observed up to 13 dpi, 36]. Now that several nematode-induced promoters are available, it could be speculated that common regulatory sequences should be obvious by sequence comparison. Such comparison is however not that straightforward because these promoters have different temporal and spatial expression patterns besides their common expression in nematode feeding sites. Furthermore, different promoter elements or in different arrays may control expression in feeding sites in different promoters. Therefore, deletion analysis combined with genomic footprinting is being done to identify discrete regions that are involved in responsiveness to nematode infection and to distinguish them from other regions in the promoters that dictate their expression in other plant tissues. In a detailed analysis of the Lemmi9 promoter from tomato, a putative nematode responsive element has been identified which is being tested for functionality by fusion to a minimal promoter and mutational studies [37].
Engineering novel nematode resistance genes The cloning, modification and transformation of natural resistance genes is a promising new field for engineering nematode resistance in crop plants. However, to increase the choice of 163
Challenges of the Environment available strategies, and especially to have resistance constructs that broaden the spectrum of target nematodes, synthetic resistance genes are undoubtedly useful alternatives [38]. Broad-spectrum resistance should be feasible if the strategy focuses on common characteristics in the infection process of different nematode species such as the activation of the cell cycle
[8].
Table 1. Genes expressed in feeding sites
N e m a t o d e sensitivity of the rescued rpe m u t a n t s
M. incognita
H. schachtii
G a l l s / plant a
Cysts / plant a
Eggs / cyst b
Wild type
12.9 _+ 3,1
8.2 + 1.6
301 _+ 16
+/rpe
13.0+3,2
7.4+2.4
271 + 2 1
rpe / rpe
0.1 (small)
3.8 + 1.3
320 + 25
Heterozygous +/rpe and the homozygous rpe/rpe mutant plants were
As described above, several tested, in vitro, in the same plates, on medium supplemented with sucrose 2% and kanamycin. Arabidopsis wild type WS were grown in "plant" promoters have been identhe same conditions except kanamycin. tified that are up (or down-) regua The number of galls and cysts per plant was determined four weeks lated upon nematode infection. after inoculation. Values represent the means + SE obtained on 25 plants, each inoculated with one hundred surface-sterilized J2 of M. Depending on the strategy, a proincognita or H. schachtii. moter can be chosen (or engib The number of eggs per cyst was counted 2 months after infection. (Favery et al. 1998, EMBO J. 17 : 6799-6811) neered) to ensure the desired spatial and temporal expression pattern for expressing a protein that is inhibiting one of the steps in pathogenesis. For sedentary endoparasites, on which this text concentrates, two main approaches can be envisaged: either the nematode is directly attacked by expressing a nematoxic protein (reviewed in [39]), or it is indirectly affected by destroying the feeding cells or inhibiting their functioning. The basic concept here is that genes necessary in the compatible interaction are engineered and used against the nematodes. Blocking the expression of plant genes that are normally up-regulated and important for the induction or maintenance of the feeding site is feasible by the antisense strategy [40] and a promoter which is highly induced but not necessarily uniquely expressed inside the feeding cells would be good for this task. Even a slight decrease in efficiency of food supply might be sufficient to significantly reduce nematode multiplication. Genes known to be expressed at high levels in feeding sites and thus probably needed to induce or maintain these structures are cdc2a [41], hmgl [4] and rpe [30, Table 1]. Therefore, transgenic plants are being made with antisense constructs of cdc2a [42], hmgl [43] and rpe [30] fused to a nematode feeding site specific promoter.
Conclusions The techniques to identify differentially expressed genes are constantly being improved. New opportunities are currently being provided by micro-array technologies and cDNA-AFLP transcript profiling. This will allow to complete the collection of genes that are up- or downregulated in the nematode feeding site as well as in the infecting nematode. The challenge will be to elucidate the role of these genes in the infection process. Because of the obligate biotrophic nature of the parasitic lifestyle of sedentary endoparasitic nematodes and the difficulty to obtain sufficient material for analysis, knowledge on the mo164
Plant parasitic nematode control lecular interaction between these nematodes and their host plants has long been very limited. Recent cloning of nematode genes encoding secreted proteins and a detailed analysis of the plant response by a joint effort of European laboratories is now quickly changing the state of the art, with promising perspectives for engineering nematode resistance into crop plants.
Authors of this contribution Godelieve Gheysen 1,2, Pierre Abad 3, Teresa Bleve 4, Vivian Blok 5, Carmen Fenoll 6, John A. Gatehouse 7, Florian Grundler 8, Keith Lindsey 9, Stephan Ohl 1~ Kristina Sagen 11, Robert Shields 12, and Johannes Helder 13 (all authors are partner in Basis and Development of Molecular Approaches to Nematode Resistance (ARENA, 1996-1999, EC grant BIO4-CT96-0318) 1Laboratorium voor Genetica, Departement Plantengenetica, Vlaams Interuniversitair Instituut voor B iotechnologie (VIB), and 2Vakgroep Plantaardige Productie, Faculteit Landbouwkundige en Toegepaste Biologische Wetenschappen, Universiteit Gent, K.L. Ledeganckstraat 35, B-9000 Gent, Belgium; 3 Laboratoire de Biologie des Invert6br6s, Institut National de la Recherche Agronomique, B.E 2078, F-06606 Antibes, France; 4 Istituto di Nematologia Agraria Applicata ai Vegetali, 1-70126 Bari, Italy; 5 SCRI Nematology Department,DD2 5DA Invergowrie, Dundee, United Kingdom; 6 Departamento de Biologfa, Universidad Aut6noma de Madrid, Cantoblanco, E-28049 Madrid, Spain and Facultad de Ciencias del Medio Ambiente, Universidad de Castilla-La Mancha, E-45071 Toledo, Spain; 7 Plant Insect Group, Department of Biological Sciences, University of Durham, Durham DH1 3LE, United Kingdom; 8 Institut ftir Phytopathologie, Christian-Albrechts- Universit/~t, D-24118 Kiel, Germany; 9 Plant Molecular Biology Group, Department of Biological Sciences, University of Durham, Durham DH1 3LE, United Kingdom; 10Zeneca MOGEN International N.V., NL-2333 CB Leiden, The Netherlands; 11IACR-Rothamsted, Entomology and Nematology Department, AL52JQ Harpenden, United Kingdom; 11 Plant Breeding International Cambridge Ltd., Marls Lane, Trumpington, Cambridge CB2 2LQ, United Kingdom; 12 Wageningen University, Laboratory of Nematology, Binnenhaven 10, 6709 PD Wageningen, The Netherlands
Acknowledgement This review was supported by the EU grants BIO4-CT960318 and FAIR-CT96-1714.
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Concerted Efforts To Develop Handles For Plant Parasitic Nematode Control
Abstract Plant-parasitic nematodes - especially root knot and cyst nematodes - are economically important pests in numerous crops. Chemical soil sterilisation and the use of other unselective pesticides to control plant parasitic nematodes are still a common practice in many European countries and at present no realistic alternatives are available. For the identification of handles to control root knot and cyst nematodes we need to know how they interact with their host. Two main types of endoparasitic nematodes can be distinguished: root-knot and cyst nematodes, both inducing a feeding site in the plant root, but in a different way. To get insight into the molecular mechanisms behind this complex interaction, several strategies to analyse plant gene expression in response to nematode infection have been followed. Random in vivo gus fusions have been particularly successful in identifying plant promoter sequences that are highly activated in nematode feeding sites, with very little expression elsewhere in the plant, but the isolation of the corresponding genes is often not straightforward. Many highly transcribed plant genes have been identified in the feeding sites, but few have been characterised in such detail as to know how important they are for a successful infection. The available data are nevertheless providing interesting tools for novel strategies to engineer nematode resistance into crops. Concomitantly, the signals coming from the nematode that are triggering this plant response or that are important in other steps of the infection process are being characterised. This study has revealed that plant-parasitic nematodes produce many different enzymes to enable them to infect the plant root and to protect themselves against the plant defence response.
Introduction Sedentary plant-parasitic nematodes are important pests of agricultural crops. Traditional management of plant-parasitic nematodes is relying on three basic strategies: crop rotation, agrochemicals, and the use of resistant plants. Crop rotation is sometimes economically disadvantageous and is often not effective, for example in the case of the root-knot nematode Meloidogyne incognita that has a very broad host range. Chemical control is expensive, effective nematicides are difficult to contain and have a broad toxic spectrum, including animals and humans. Host resistance is the most environmentally and economically sound method [ 1]. Unfortunately, natural resistance to plant parasitic nematodes is not available for all cul-
Godelieve Gheysen, Departement Plantengenetica, Universiteit Gent, Belgium
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Challenges of the Environment tivated crop species. Furthermore, a very important limiting factor to the general usefulness of natural resistance genes is the specific nature of their protection, which is often restricted to one nematode species or even certain pathotypes [2]. An alternative strategy that tries to combine effectiveness, broad applicability and durability with environmental friendliness is the engineering of nematode resistance into plants. In the last few years several excellent reviews have been written on this research development, and the reader is referred to them for more general aspects [3-4]. Here, we will summarise the breakthroughs of the last years, mainly in the light of the co-ordinated efforts done by the 12 laboratories involved in the ARENA project of the European Union.
Morphological and histological changes Sedentary nematodes establish and maintain an intimate relationship with their host. After invading the plant root, they migrate to the vascular cylinder in search for a cell that can serve as an initial feeding cell. In response to repeated stimulation by the parasite, this cell develops into either a syncytium (for cyst nematodes, such as Globodera and Heterodera), or several cells are stimulated to form a system of giant cells (as is the case for root-knot nematodes). The combination of light and electron microscopy studies of fixed samples and time-lapse video microscopy on living material has provided detailed insight in the establishment and structure of the nematode feeding site [5-6, reviewed in 3]. The syncytium is formed by breakdown of plant cell wails and subsequent fusion of neighbouring cells. A giant cell is formed as the result of repeated nuclear divisions without cytokinesis. Although these feeding sites differ from each other in ontogenesis and structure, their function is the same: to supply the nematode with sufficient nutrients for growth and reproduction. This common function is reflected in a final analogous ultrastructure. Syncytial and giant cells are hypertrophied and multinucleated cells with a dense granular cytoplasm, an increase in rough endoplasmic reticulum and in the number of mitochondria. Recently the formation of cell wall openings occurring during syncytium differentiation in A. thaliana roots was studied in more detail [7]. The first openings are formed by gradual widening of existing plasmodesmata followed by the fusion of protoplasts from two adjacent cells. At later stages, wall openings form exclusively in cell walls without the involvement of plasmodesmata. The sequence of events leading to these openings is: accumulation of ER membranes accompanying initial wall lesions, expansion of lesions until the middle lamella, and finally dissolution of the middle lamella [7]. Functional plasmodesmata were never found in the outer wall of mature syncytia, although they are present in the walls of the adjacent sieve elements. These plasmodesmata were always closed from the syncytium side by wall material. It can be concluded that the formation of a syncytium is accompanied with many changes in cell wall biology: callose-like deposition around the stylet, cell wall ingrowths adjacent to the xylem, widening or closing of plasmodesmata depending on their location in the syncytium, and de novo formation of wall openings. It will be interesting to see which enzymes are involved in these processes and how they are temporally and spatially controlled. Another aspect of nematode feeding site formation that is being unravelled is the activation of the cell cycle by the nematode stimulus. Sedentary endoparasitic nematodes induce multinucleate feeding cells in the roots of their host plants. These cells undergo multiple rounds of 160
Plant parasitic nematode control shortened cell cycles leading to genome amplification and hypertrophy of the cytoplasm. This aspect of the infection process has been thoroughly reviewed by [8] and more recently by [9]. It is now generally believed that giant cells develop by repeated mitosis without cytokinesis [5]. Jones and Payne [10] showed that cell plate vesicles initially lined up between the two daughter nuclei but then dispersed, resulting in the abortion of the new cell plate formation. No mitosis has been seen in syncytia [11 ], although Piegat and Wilski [ 12] reported an initial mitotic stimulation during syncytium induction by Globodera rostochiensis in infected potato roots. However, it is not clear from the data shown whether this mitosis is really inside the developing syncytium. Indeed, stimulation of cell division has been observed in parenchyma cells surrounding the syncytium and which will be incorporated [13-14]. The enlargement of nuclei indicates that DNA multiplication is taking place within the syncytial tissue during and after the incorporation of new cells through cell wall dissolution [ 11]. Studies with cell cycle inhibiting drugs indicate that mitosis is essential for giant cell development and for normal syncytium expansion [ 14].
Nematode secretions Stylet secretions that originate in the esophageal gland cells of plant parasitic nematodes are believed to play a major role in pathogenesis. Secretions from the dorsal gland cells have always been considered critical for pathogenesis but till recently a role for the subventral glands in the infection process has been questioned. Subventral gland secretions are released into the pharyngeal lumen behind the pump chamber and were thought only to be able to pass posteriorly into the intestine. However, immunolocalisation of HG-ENG-2 demonstrated the presence of this protein in the nematode's subventral glands (where it is produced), as well as in the root cortical tissue after secretion from the stylet [ 15]. This HG-ENG-2 protein belongs to a family of f~-l,4-endoglucanases first identified in secretions of cyst nematodes by [16], and later also in root knot nematodes [ 17]. These enzymes presumably facilitate the migration of the nematode through plant roots by partial cell wall degradation. The corresponding genes represent the first cellulase genes ever cloned from an animal and show highest homology (35-40%) with 13-1,4-endoglucanases from bacteria [16]. Other lytic enzymes identified in the nematode secretions include pectate lyase and proteases (Herman Popeijus and John Jones, unpublished results). Other proteins that are present in nematode secretions (although not necessarily from the esophageal glands) appear to be related to the active protection of the nematode against the plants defense response. GP-SEC2 functions as a fatty acid binding protein capable of binding e.g. linolenic and linoleic acids, that are important precursors in plant defence signalling pathways. In addition, superoxide dismutase and thioredoxin peroxidase secreted by the nematode are involved with inactivation of superoxide radicals and hydrogen peroxide respectively. How important these proteins or the above mentioned lytic enzymes are for nematode pathogenesis, can only be decided if inactivation or absence of these proteins results in a lower percentage of successful nematode infections. Inhibition of the proteins could be achieved by the expression of plantibodies [ 18]. Inactivation of the genes is another theoretical possibility, but transformation of plant-parasitic nematodes is still virtually impossible [19]. A strategy that could be useful is RNA interference, a method capable of efficiently eliminating specific mRNAs in C. elegans [20]. 161
Challenges of the Environment The most intriguing function of the esophageal secretions, their role in formation of the feeding site, is still a black box. Unfortunately, in contrast to comparative studies of nematode development and the genes involved, analysis of the model nematode C. elegans does not help much in respect to pathogenesis-involved genes. However, with the current expertise and molecular technology, at least some clues about possible signalling molecules in the secretions of plant-parasitic nematodes should be identified in the near future.
Screening Screening for nematode-responsive plant regulatory sequences was done by by T-DNA tagging. Plant genes that are up-regulated after nematode infection have first been identified by differential screening of cDNA libraries [21-23] or from subtraction cDNA libraries [24] and later by differential display [25, Vercauteren et al., accepted by MPMI]. To identify promoters that are early and specifically induced in the nematode feeding sites, the most appropriate method is undoubtedly the analysis of promoter-reporter fusions (for example with 13-glucuronidase [gus]). Plant-gus fusions can be made at random in vivo and then screened for specific expression patterns. This "promoter trapping" or "tagging" approach is based on the random integration of a promoterless gus gene after transformation into a plant species, often A. thaliana [26]. When inserted downstream from a plant promoter that is inducible by nematode infection, a higher GUS activity is expected to be seen at the infection site (Fig. 1). The elegance of this method lies in the ability to directly visualise induced expression in nematode feeding sites at various stages of the interaction, while analysing the specificity of expression using uninfected parts of the same plant and control plants. Furthermore, promising plant lines can easily be tested for gus activation upon infection by different types of nematodes. This "tagging" approach has been performed in several labs of the ARENA project and has yielded many interesting promoters [27-30]. The hope was that the homozygous progeny of these T-DNA tagged lines could be knock-outs for the tagged gene,
Figure 1: GUS activity in a Att0025-R/1 syncytium 6 days post inoculation with H e t e r o d e r a s c h a c h t i i (from: Barthels et al. 1997, Plant Cell 9:2119-2134)
,f f
~:~~F::~~ /'
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Plant parasitic nematode control
and would therefore yield information on the role of this plant gene, in general or possibly even during nematode infection. Initially none of the lines analysed had an obvious phenotype [28], strengthening the belief that tagging often yields cryptic promoters [31 ]. Although these cryptic promoters are not less useful for expressing a transgene in the nematode feeding sites, from a more fundamental point of view it would be informative to find the endogenous genes that are specifically activated after nematode infection. However, more recently one of the tagged lines was found to have a T-DNA insertion in the rpe gene (D-ribulose-5-phosphate-3-epimerase) resulting in a mutant phenotype. Homozygous rpe plants have a germination deficient phenotype that can be rescued in dwarf plants on sucrose-supplemented medium [30]. Also, upon further study, several other T-DNA insertions appeared to be linked to a "tagged" gene, although in unexpected configuration, such as inside the promoter instead of downstream, and even several kb away from the gene (line Att0001 (M.Karimi), line Att0025 (S. Ohl), line Att0884 (P.Puzio); unpublished observations).
Transcriptional control of nematode-inducible genes In the light of applications, the most interesting A. thaliana promoters are also introduced into crop species to check their activity after nematode infection. The Art0001 (ARM 1) promoter, as well as some other A. thaliana promoters, are clearly up-regulated after nematode infection in oilseed rape and sugarbeet [32, G. Gheysen unpublished], but not in potato, whereas the rpe promoter is activated in potato nematode feeding sites [B. Favery, unpublished]. Besides possible differences in cell-specific or temporal gene regulation between plant species, another reason could be that in potato, the syncytium is formed in cortex cells [33, 34], while in A. thaliana procambium cells are normally used [13]. Maybe the apparently contradictory results on CaMV35S-promoter activity inside syncytia could also be explained by the different plant species used. Goddijn et al. [27] and Urwin et al. [35] describe down-regulation of the 35S promoter in syncytia induced by H. schachtii in A. thaliana, and this was also observed in sugarbeet (G. Gheysen, unpublished), but not in potato syncytia induced by G. rostochiensis [observed up to 13 dpi, 36]. Now that several nematode-induced promoters are available, it could be speculated that common regulatory sequences should be obvious by sequence comparison. Such comparison is however not that straightforward because these promoters have different temporal and spatial expression patterns besides their common expression in nematode feeding sites. Furthermore, different promoter elements or in different arrays may control expression in feeding sites in different promoters. Therefore, deletion analysis combined with genomic footprinting is being done to identify discrete regions that are involved in responsiveness to nematode infection and to distinguish them from other regions in the promoters that dictate their expression in other plant tissues. In a detailed analysis of the Lemmi9 promoter from tomato, a putative nematode responsive element has been identified which is being tested for functionality by fusion to a minimal promoter and mutational studies [37].
Engineering novel nematode resistance genes The cloning, modification and transformation of natural resistance genes is a promising new field for engineering nematode resistance in crop plants. However, to increase the choice of 163
Challenges of the Environment available strategies, and especially to have resistance constructs that broaden the spectrum of target nematodes, synthetic resistance genes are undoubtedly useful alternatives [38]. Broad-spectrum resistance should be feasible if the strategy focuses on common characteristics in the infection process of different nematode species such as the activation of the cell cycle
[8].
Table 1. Genes expressed in feeding sites
N e m a t o d e sensitivity of the rescued rpe m u t a n t s
M. incognita
H. schachtii
G a l l s / plant a
Cysts / plant a
Eggs / cyst b
Wild type
12.9 _+ 3,1
8.2 + 1.6
301 _+ 16
+/rpe
13.0+3,2
7.4+2.4
271 + 2 1
rpe / rpe
0.1 (small)
3.8 + 1.3
320 + 25
Heterozygous +/rpe and the homozygous rpe/rpe mutant plants were
As described above, several tested, in vitro, in the same plates, on medium supplemented with sucrose 2% and kanamycin. Arabidopsis wild type WS were grown in "plant" promoters have been identhe same conditions except kanamycin. tified that are up (or down-) regua The number of galls and cysts per plant was determined four weeks lated upon nematode infection. after inoculation. Values represent the means + SE obtained on 25 plants, each inoculated with one hundred surface-sterilized J2 of M. Depending on the strategy, a proincognita or H. schachtii. moter can be chosen (or engib The number of eggs per cyst was counted 2 months after infection. (Favery et al. 1998, EMBO J. 17 : 6799-6811) neered) to ensure the desired spatial and temporal expression pattern for expressing a protein that is inhibiting one of the steps in pathogenesis. For sedentary endoparasites, on which this text concentrates, two main approaches can be envisaged: either the nematode is directly attacked by expressing a nematoxic protein (reviewed in [39]), or it is indirectly affected by destroying the feeding cells or inhibiting their functioning. The basic concept here is that genes necessary in the compatible interaction are engineered and used against the nematodes. Blocking the expression of plant genes that are normally up-regulated and important for the induction or maintenance of the feeding site is feasible by the antisense strategy [40] and a promoter which is highly induced but not necessarily uniquely expressed inside the feeding cells would be good for this task. Even a slight decrease in efficiency of food supply might be sufficient to significantly reduce nematode multiplication. Genes known to be expressed at high levels in feeding sites and thus probably needed to induce or maintain these structures are cdc2a [41], hmgl [4] and rpe [30, Table 1]. Therefore, transgenic plants are being made with antisense constructs of cdc2a [42], hmgl [43] and rpe [30] fused to a nematode feeding site specific promoter.
Conclusions The techniques to identify differentially expressed genes are constantly being improved. New opportunities are currently being provided by micro-array technologies and cDNA-AFLP transcript profiling. This will allow to complete the collection of genes that are up- or downregulated in the nematode feeding site as well as in the infecting nematode. The challenge will be to elucidate the role of these genes in the infection process. Because of the obligate biotrophic nature of the parasitic lifestyle of sedentary endoparasitic nematodes and the difficulty to obtain sufficient material for analysis, knowledge on the mo164
Plant parasitic nematode control lecular interaction between these nematodes and their host plants has long been very limited. Recent cloning of nematode genes encoding secreted proteins and a detailed analysis of the plant response by a joint effort of European laboratories is now quickly changing the state of the art, with promising perspectives for engineering nematode resistance into crop plants.
Authors of this contribution Godelieve Gheysen 1,2, Pierre Abad 3, Teresa Bleve 4, Vivian Blok 5, Carmen Fenoll 6, John A. Gatehouse 7, Florian Grundler 8, Keith Lindsey 9, Stephan Ohl 1~ Kristina Sagen 11, Robert Shields 12, and Johannes Helder 13 (all authors are partner in Basis and Development of Molecular Approaches to Nematode Resistance (ARENA, 1996-1999, EC grant BIO4-CT96-0318) 1Laboratorium voor Genetica, Departement Plantengenetica, Vlaams Interuniversitair Instituut voor B iotechnologie (VIB), and 2Vakgroep Plantaardige Productie, Faculteit Landbouwkundige en Toegepaste Biologische Wetenschappen, Universiteit Gent, K.L. Ledeganckstraat 35, B-9000 Gent, Belgium; 3 Laboratoire de Biologie des Invert6br6s, Institut National de la Recherche Agronomique, B.E 2078, F-06606 Antibes, France; 4 Istituto di Nematologia Agraria Applicata ai Vegetali, 1-70126 Bari, Italy; 5 SCRI Nematology Department,DD2 5DA Invergowrie, Dundee, United Kingdom; 6 Departamento de Biologfa, Universidad Aut6noma de Madrid, Cantoblanco, E-28049 Madrid, Spain and Facultad de Ciencias del Medio Ambiente, Universidad de Castilla-La Mancha, E-45071 Toledo, Spain; 7 Plant Insect Group, Department of Biological Sciences, University of Durham, Durham DH1 3LE, United Kingdom; 8 Institut ftir Phytopathologie, Christian-Albrechts- Universit/~t, D-24118 Kiel, Germany; 9 Plant Molecular Biology Group, Department of Biological Sciences, University of Durham, Durham DH1 3LE, United Kingdom; 10Zeneca MOGEN International N.V., NL-2333 CB Leiden, The Netherlands; 11IACR-Rothamsted, Entomology and Nematology Department, AL52JQ Harpenden, United Kingdom; 11 Plant Breeding International Cambridge Ltd., Marls Lane, Trumpington, Cambridge CB2 2LQ, United Kingdom; 12 Wageningen University, Laboratory of Nematology, Binnenhaven 10, 6709 PD Wageningen, The Netherlands
Acknowledgement This review was supported by the EU grants BIO4-CT960318 and FAIR-CT96-1714.
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