Plant Cell, Vol. 10, 905-916, June 1998, Copyright © 1998, American Society of Plant Physiologists
Srchi13, a Novel Early Nodulin from Sesbania rostrata, Is Related to Acidic Class III Chitinases
Sofie Goormachtiga,
Sam Lievensa,
Willem Van de Veldea,
Marc Van Montagua, and
Marcelle Holstersa
a Laboratorium voor Genetica, Department of Genetics, Flanders Interuniversity Institute for Biotechnology, Universiteit Gent, K.L. Ledeganckstraat 35, B-9000 Gent, Belgium
Correspondence to:
Marcelle Holsters, mahol{at}gengenp.rug.ac.be (E-mail), 32-9-2645349 (fax).
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ABSTRACT |
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On the tropical legume Sesbania rostrata, stem-borne nodules develop after inoculation of adventitious root primordia with the microsymbiont Azorhizobium caulinodans. A cDNA clone, Srchi13, with homology to acidic class III chitinase genes, corresponds to an early nodulin gene with transiently induced expression during nodule ontogeny. Srchi13 transcripts accumulated strongly 2 days after inoculation, decreased from day 7 onward, and disappeared in mature nodules. Induction was dependent on Nod factor–producing bacteria. Srchi13 was expressed around infection pockets, in infection centra, around the developing nodule and its vascular bundles, and in uninfected cells of the central tissue. The specific and transient transcript accumulation together with the lipochitooligosaccharide degradation activity of the recombinant protein hint at a role of Srchi13 in normal nodule ontogeny by limiting the action of Nod factors.
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INTRODUCTION |
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Gram-negative bacteria of the genera Rhizobium, Bradyrhizobium, Azorhizobium, and Sinorhizobium (collectively called rhizo-bia) establish a beneficial interaction with specific leguminous host plants. Compatible rhizobia induce the formation of root nodules, which are new organs that, after invasion and intracellular colonization by the bacteria, become the sites of symbiotic nitrogen fixation. The development of functional nodules is the consequence of specific molecular recognition between the two partners (reviewed in Schultze et al. 1994 
). The communication starts when the bacteria sense plant-produced compounds, most often flavonoids, and activate the expression of their nodulation (nod) genes. These genes are responsible for the production and secretion of chemical return signals, the Nod factors. Nod factors are lipochitooligosaccharides (LCO), which are oligomers of ß-1,4– linked N-acetylglucosamine (GlcNAc) molecules, with an acyl chain replacing the acetyl group of the nonreducing end sugar and with strain-specific decorations at the reducing and the nonreducing ends (reviewed in Mergaert et al. 1997 ).
The Nod factors are specific morphogens that trigger the plant program for nodule development. When applied in purified form to a compatible host, Nod factors may exert nodulation-related effects, such as root hair deformation, preinfection thread formation, nodule primordium formation, and, in some cases, the appearance of nodulelike structures (Heidstra and Bisseling 1996 ; Long 1996 ). Moreover, pure Nod factors directly or indirectly induce the expression of several early nodulin (ENOD) genes, which encode plant proteins that appear at early developmental stages and bring about events connected with cellular signaling (Long 1996 ). Nod factors contribute to the host specificity of nodulation by their strain-specific differences (Schultze et al. 1994 ; Heidstra and Bisseling 1996 ).
Many researchers in the plant–microbe interaction field have called attention to analogies between aspects of pathogenesis and symbiosis and have suggested that rhizobia may be seen as "refined parasites" (Vance 1983 ; Djordjevic et al. 1987 ; Long and Staskawicz 1993 ; Baron and Zambryski 1995 ). Nod factors, with their chitin backbone, resemble the fungal chitin elicitors; they are recognized by the same signaling pathway as chitin elicitors in tomato (Staehelin et al. 1994a ), they provoke elicitor-like effects in alfalfa (Savoure et al. 1997 ), and they can be degraded by chitinases (Heidstra et al. 1994 ; Staehelin et al. 1994b ).
Chitinases hydrolyze oligomers of ß-1,4–linked GlcNAc residues, and some chitinases also have a lysozyme activity (reviewed in Collinge et al. 1993 ). The plant chitinases are separated into different classes (I, II, III, IV, and V) according to primary structural features (Collinge et al. 1993 ; Meins et al. 1994 ). Plant chitinases have been best characterized as tools in the defense response against microbes. The expression of chitinase genes is induced in coordination with other pathogenesis-related proteins in response to the attack of incompatible and compatible pathogens and during nonhost resistance phenomena (Collinge et al. 1993 ). Chitinases may generate elicitors for induction of defense responses, inhibit pathogen growth (Collinge et al. 1993 ), or be involved in plant development. The latter role is illustrated by, for instance, the rescue of a carrot mutant in somatic embryo formation by a class IV chitinase (De Jong et al. 1992 ; Kragh et al. 1996 ), the presence of chitinases in selective parts of flowers (Lotan et al. 1989 ; Neale et al. 1990 ), and the appearance of chitinases during leaf senescence (Hanfrey et al. 1996 ).
The involvement of chitinases in nodulation has also been reported. In the alfalfa–R. meliloti interaction, chitinases are activated as part of a feedback response of the plant to limit the number of infection threads once the first nodule primordia are formed (Vasse et al. 1993 ). Chitinases participate in a defense response in ineffective soybean nodules induced by a mutant B. japonicum strain, and they have been found in the cortex of mature nodules induced by wild-type strains, presumably for protection (Staehelin et al. 1992 ). Finally, based on the observation of an induced Nod factor hydrolytic activity in alfalfa, controlled Nod factor degradation by chitinases or other LCO-degrading enzymes has been suggested to be a feature of regular nodule development (Staehelin et al. 1995 ).
We studied the interaction between the tropical legume Sesbania rostrata and the microsymbiont A. caulinodans ORS571. S. rostrata, endemic to the Sahel region of West Africa, is adapted to growth in waterlogged conditions. All along its stem, the plant carries numerous adventitious root primordia that develop into roots when immersed. After inoculation with A. caulinodans, nodules develop at the adventitious root primordia. The development of these nodules is marked by some interesting characteristics (Tsien et al. 1983 ; Duhoux 1984 ; Goormachtig et al. 1997 ). The primordia pierce through the stem epidermis, creating a circular fissure in which inoculated bacteria proliferate and form densely populated intercellular infection pockets. Opposite the infection pockets, globular nodule primordia are formed. Infection threads (first intercellular and then intracellular) lead the bacteria toward the nodule primordia where invasion occurs. In later stages, different developmental zones are found much the same as those found in indeterminate nodules, with a distal meristem, an infection zone, and a proximal fixation zone being present (Goormachtig et al. 1997 ). For a time, these zones coexist; after a week, however, meristem activity ceases, resulting in round-shaped mature nodules of the determinate type.
By using the differential display approach, Goormachtig et al. 1995 found an ENOD cDNA (Srchi13, previously called didi-13) with homology to class III chitinase genes. In the present article, we analyze the spatiotemporal pattern of Srchi13 transcript accumulation. We also show that a Nod factor–degrading activity is associated with the gene product. Our observations provide evidence for a role of this chitinase in nodule development, presumably by the control of LCO levels and perhaps also by affecting bacterial viability.
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RESULTS |
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Sequence Analysis of Srchi13 cDNA
By using a differential display approach, Goormachtig et al. 1995 identified a partial cDNA sequence that corresponds to a gene with induced transcript accumulation during the early stages of stem nodule development. The fragment didi-13 is homologous to class III chitinase genes. The full-length cDNA sequence, obtained by screening a cDNA library derived from developing stem nodules (see Methods), was named Srchi13.
Figure 1 provides the deduced amino acid sequence from Srchi13 aligned to the primary sequences of other class III chitinases. Database searches (EMBL and SwissProt) revealed a 66 to 70% overall similarity between Srchi13 and other members of class III (Figure 1). The consensus sequence is shown in Figure 1 (Levorson and Chlan 1997 ). Srchi13 shows 66% similarity to this consensus. Two central amino acid motifs, designated regions I and II (Kuranda and Robbins 1991 ), which are highly conserved between class III chitinases of plants and their bacterial homologs (Tsujibo et al. 1993 ; Watanabe et al. 1993 ), are found in Srchi13 (Figure 1). Application of the Genetics Computer Group program (version 7; Genetics Computer Group, Madison, WI) revealed that Srchi13 contains two potential N-glycosylation sites (at positions 19 and 234). Srchi13 encodes an acidic chitinase with a pI value of 5.3. An N-terminal signal peptide was predicted. In comparison to related class III chitinases, Srchi13 has a C-terminal extension of 31 amino acids (Figure 1).
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Figure 1.
Alignment of Srchi13 to Class III Chitinases.
The amino acid sequence of Srchi13 (a; EMBL accession number Z48674) is aligned to those of class III chitinases from para rubber tree (hevamine A) (b; EMBL accession number P23472), grape (c; EMBL accession number P51614), Arabidopsis (d; EMBL accession number A45511), cowpea (e; EMBL accession number S57475), adzuki bean (f; EMBL accession number S36932), and chickpea (g; EMBL accession number P36908). At bottom is the class III consensus sequence (Levorson and Chlan 1997 ). Lowercase letters denote residues of weak consensus (<50%). Residues that are identical between the consensus sequence and Srchi13 are indicated in gray. The arrowhead indicates the possible cleavage sites of the signal peptides. I and II indicate conserved motifs (Kuranda and Robbins 1991 ). Dots indicate gaps introduced to allow better alignment.  , ß,  , and  are places with two consensus residues and correspond to h/n, s/t, k/s, and t/p, respectively.
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DNA gel blot analysis revealed that Srchi13 may belong to a small gene family. As shown in Figure 2, besides a main band, a smaller, less intense DNA fragment appeared after stringent hybridization when the genomic DNA was digested with EcoRI and PstI. The two bands obtained after HindIII digestion resulted from the presence of this restriction site in the Srchi13 probe (see Methods).
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Figure 2.
DNA Gel Blot Analysis of Srchi13.
The enzymes that were used for digestion of genomic DNA are indicated above each lane. The numbers at right indicate the length of the fragments in kilobases.
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Srchi13 Is Differentially Expressed during Nodule Development
The expression of Srchi13 during the development of stem-borne nodules was analyzed by RNA gel blot analysis. The results are shown in Figure 3. RNA was prepared from carefully excised adventitious root primordia before inoculation and 1, 2, 3, 4, 5, 7, and 17 days after inoculation with A. caulinodans and then hybridized with an Srchi13 probe (see Methods). Srchi13 transcripts accumulated 2 days after inoculation (Figure 3). Low Srchi13 transcript accumulation was observed in root primordia 1 day after infection, as indicated by an overexposed RNA gel blot (data not shown). Reverse transcription–polymerase chain reaction (PCR) confirmed that the earliest time point at which Srchi13 transcripts could be detected was 1 day after infection (data not shown). The transcript level slowly increased with time and dropped between 7 and 17 days after infection. In mature nodules (17 days), low expression levels were detected.
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Figure 3.
Temporal Expression Pattern of Srchi13 during Stem Nodule Development.
Srchi13 transcript levels were determined by using RNA gel blot analysis in uninfected root primordia (-) and in developing nodules 1, 2, 3, 4, 5, 7, and 17 days after azorhizobial infection (+1, +2, +3, +4, +5, +7, and +17, respectively). To control for equal loading and blotting, we stained the filter with methylene blue (bottom).
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As shown in Figure 4, a similar expression pattern was obtained during root nodulation on S. rostrata. No expression was detected in uninfected root primordia or in uninoculated roots (Figure 3 and Figure 4). Moreover, no transcripts were found in other parts of the plants, such as leaves, apices, and flowers, or in young seedlings (data not shown). Therefore, Srchi13 can be considered as a true early nodulin gene.
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Figure 4.
Temporal Expression Pattern of Srchi13 during Root Nodule Development.
Srchi13 transcript levels were determined by using RNA gel blot analysis in uninfected roots (-) and in developing nodules 1, 2, 3, 4, 8, 10, and 30 days after azorhizobial infection (+1, +2, +3, +4, +8, +10, and +30, respectively). To control for equal loading and blotting, we stained the filter with methylene blue (bottom).
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Srchi13 Expression Is Specific for Nod Factor–Producing A. caulinodans
Previously, it was shown that Srchi13 transcript accumulation is caused by the presence of A. caulinodans in the inoculum and not by the inoculation procedure itself or by residual traces of the bacterial yeast extract–based growth medium that might have contained elicitors (Goormachtig et al. 1995 ).
Because many chitinases play a role in the general defense response of plants against microbes (Collinge et al. 1993 ), we controlled whether the observed expression patterns of Srchi13 were caused by the mere presence of A. caulinodans or whether there was a correlation with Nod factor production. S. rostrata stems were infected with two A. caulinodans mutants. Each was deficient in some aspect of nodulation. ORS571-V44 carried a Tn5 insertion in the nodA gene and failed to produce Nod factors and to provoke nodulation (Van den Eede et al. 1987 ; Mergaert et al. 1993 ). A second mutant, ORS571-X15, with a Tn5 insertion in a rhamnose biosynthesis locus, had altered surface polysaccharides and was defective in invasion but not in Nod factor production. Infection with the latter mutant resulted in the appearance of bacteria-free pseudonodules (Goethals et al. 1994 ; W. D'Haeze and M. Holsters, unpublished results). RNA was isolated from adventitious root primordia 3 and 6 days after infection with either bacterial mutant. The expression levels of Srchi13 were compared with those obtained after wild-type infection by using RNA gel blot hybridization analysis. The results are shown in Figure 5.
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Figure 5.
Srchi13 Expression upon Inoculation with A. caulinodans Mutants.
Transcript levels were determined by using RNA gel blot analysis in uninfected root primordia (-) and primordia infected with wild-type A. caulinodans ORS571 (WT3 and WT6), the mutant ORS571-X15 (surface polysaccharide mutant defective in invasion; LPS3 and LPS6), and the mutant ORS571-V44 (NODA-, deficient in Nod factor production; NODA-3, NODA-6) 3 and 6 days after infection. We stained the filter with methylene blue to confirm equal loading and blotting (bottom).
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Srchi13 transcripts accumulated after infection with ORS571-X15. Equal amounts of transcripts were detected 3 days after infection with either wild-type or mutant bacteria. However, in the case of ORS571-X15 infection, the expression level dropped markedly at day 6, whereas transcripts still accumulated during wild-type infection. In contrast, no induction of the chitinase gene was observed after the application of strain ORS571-V44. Thus, the observed expression was specific for Nod factor–producing azorhizobia and not a general response of the plants toward the bacterium.
In Situ Localization of Srchi13 Transcripts
To obtain indications for the function of Srchi13 during nodulation, in situ hybridizations were performed on 10-µm sections of developing stem nodules (see Methods). After counterstaining with toluidine blue, photographs were taken using dark-field or bright-field optics. The results are shown in Figure 6.
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Figure 6.
In Situ Localization of Srchi13 Transcripts.
Longitudinal and transverse sections (10 µm) of uninfected root primordia and developing nodules were hybridized with a 35S-labeled antisense RNA probe.
(A) Dark-field microscopy of a longitudinal section of an infected root primordium 1 day after inoculation with A. caulinodans.
(B) Dark-field microscopy of a longitudinal section of an infected root primordium 2 days after bacterial inoculation.
(C) Bright-field microscopy of section shown in (B).
(D) Enlargement of section shown in (C).
(E) Bright-field microscopy of a transverse section of a developing nodule 3 days after bacterial inoculation.
(F) Dark-field microscopy of a longitudinal section of a developing nodule 3 days after bacterial inoculation. Two nodule primordia are seen, one on either side of the root stele.
(G) Dark-field microscopy of a longitudinal section of a 6-day-old developing nodule. Two nodules have developed. An arrow indicates where no nodule parenchyma has developed yet.
Figure 6. (continued).
(H) Bright-field microscopy of section shown in (F).
(I) Bright-field microscopy of section shown in (G).
(J) Magnification of the region indicated by the rectangle in (I).
(K) Dark-field microscopy of a longitudinal section of an 8-day-old nodule.
(L) Dark-field microscopy of a longitudinal section of a 10-day-old nodule.
Signals are seen as white and black dots in dark-field and bright-field micrographs, respectively. c, cortex; f, fixation zone; fi, fissure; i, infection zone; ic, infection center; ip, infection pocket; m, meristem; na, nodule parenchyma; np, nodule primordium; rm, root meristem; rv, root vascular bundle. Bars = 100 µm.
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No Srchi13 transcripts were detected by in situ hybridization in sections of uninfected adventitious root primordia (data not shown). One day after infection, a few cortex cells began to accumulate Srchi13 transcripts (Figure 6A). After 2 days, strong expression was detected in the cortex all around the incipient nodule primordium (Figure 6B and Figure 6C). Magnification clearly showed that the nodule primordium itself did not express Srchi13 (Figure 6D). Also, no signal was detected in the dormant root meristem. Srchi13 transcripts were very prominent around infection pockets (Figure 6D and Figure 6E). Three days after infection, the globular nodule primordia formed open-basket structures around infection centra (regions with many infection threads, connecting outer infection pockets with inner parts of the nodule primordia; Goormachtig et al. 1997 ). Strong expression of Srchi13 was seen in the differentiating nodule parenchyma, in the infection center, and in the cortical cells surrounding the nodule primordia (Figure 6F and Figure 6H). A transverse section clearly shows Srchi13-expressing cells, which surround completely the developing nodule and the invading bacteria (Figure 6E).
Expression became more localized at 4 to 6 days after infection (Figure 6G, Figure 6I, and Figure 6J). At that stage, transient zonation exists, consisting of a distal meristem, an infection zone (prefixation zone), and a zone with invaded cells (young fixation zone; Goormachtig et al. 1997 ). The nodule parenchyma showed high expression of Srchi13 (Figure 6G, Figure 6I, and Figure 6J). Where no parenchyma was differentiated as of yet, for example, distal to the nodule meristem, Srchi13 was expressed in the adjacent cortical cells (Figure 6G, arrow). Srchi13 transcripts were detected in cells of the infection center, in some cells of the infection zone, and in the uninfected cells of the fixation zone (Figure 6G, Figure 6I, and Figure 6J).
When meristem activity ceased, Srchi13 transcripts were found in the nodule parenchyma, in some cells of the infection zone, and in uninfected cells of young fixation zones. The latter point is illustrated in Figure 6K. The uninfected cells of the most proximally located fixation zone did not contain Srchi13 transcripts, whereas those from the distally located cell layers did. Srchi13 transcripts disappeared, first in the uninfected cells of the central tissue and next in the nodule parenchyma in a proximal–distal direction (cf. Figure 6K and Figure 6L). In 20-day-old nodules, Srchi13 could no longer be detected (data not shown).
A Nod Factor–Degrading Activity Is Associated with a Recombinant MBP–Srchi13 Fusion Protein
For alfalfa, it has been shown that Nod factors are degraded by endogenous chitinases (Staehelin et al. 1994b ). To control whether Srchi13 was able to degrade Nod factors of A. caulinodans, the recombinant Srchi13 protein was expressed in Escherichia coli as a fusion with the maltose binding protein (MBP; see Methods). This fusion protein was used in a Nod factor degradation assay with the 14C-labeled peak I (PI) Nod factor fraction (peak I corresponds to Nod factors that carry a C18:1 fatty acid; Mergaert et al. 1993 ) as substrate (see Methods). Figure 7 shows the reaction products analyzed by reversed-phase thin-layer chromatography.
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Figure 7.
Nod Factor Degradation Activity of Srchi13.
Reversed-phase thin-layer chromatography was performed on a 14C-labeled Nod factor fraction (peak I; Mergaert et al. 1993 ) after overnight incubation with the enriched MBP fraction (lane 1), enriched MBP–Srchi13 fraction (lane 2), protein extracts from uninfected root primordia (lane 3), protein extracts from developing nodules 4 days after inoculation (lane 4), and control (lane 5), which represents equal treatment of the Nod factors without incubation with protein extracts. F, solvent front; O, the place of spotting the samples; S, PI Nod factor substrate.
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Figure 7, lanes 1 and 2, represent the reaction products after incubation of the PI Nod factor fraction with the enriched MBP and the MBP–Srchi13 fusion, respectively. No degradation was observed after incubation with MBP (Figure 7, cf. lanes 1 and 5, control). Incubation with the MBP–Srchi13 yielded two spots (Figure 7, cf. lanes 2 and 5), of which the lower, more intensive spot presumably corresponded to trimeric LCOs and the upper to the tetrameric form. This was deduced from comparison to the degradation patterns obtained after incubation with crude extracts from uninfected root primordia or developing nodules. When extracts from uninfected root primordia were used, as shown in Figure 7, lane 3, four different spots were observed most probably corresponding to the pentameric Nod factor substrate and to the tetrameric, trimeric, and dimeric LCO degradation products. This partial degradation is probably due to the action of plant chitinases other than Srchi13, because no Srchi13 transcripts were detected in uninfected root primordia. When extracts from developing nodules were used, as shown in Figure 7, lane 4, dimeric LCO degradation products were obtained. These molecules probably result from the combined action of several induced chitinases, one of which is Srch13.
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DISCUSSION |
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The Early Nodulin Srchi13 Is Related to Acidic Class III Chitinases
Nodule development is accompanied by new plant gene expression, and the genes that are expressed early are thought to play a role in organ formation and bacterial invasion (Franssen et al. 1992 ; Schultze et al. 1994 ). Stem nodulation on S. rostrata offers a good system for the isolation of early nodulin genes because of the predetermined nodulation sites, their accessibility, and their synchronous development. The deduced amino acid sequence of a cDNA Srchi13, with induced transcript levels during early stages of nodule formation (Goormachtig et al. 1995 ), was related to acidic class III chitinases, a family of proteins that are often targeted to the extracellular compartment (Metraux et al. 1988 ; Lawton et al. 1992 ; Nielsen et al. 1993 ). Srchi13 has a putative signal peptide and glycosylation sites and contains consensus regions I and II of the class III chitinases.
RNA gel blot analysis and reverse transcription–PCR revealed that Srchi13 transcripts accumulate in early stages of stem nodulation, before the onset of N2 fixation (which is ~4 or 5 days after infection in this system), and that this expression depends on bacterial LCO production. Srchi13 transcripts were not found in other organs (flowers, stems, and roots). Thus, Srchi13 belongs to the group of the early nodulins. Currently, it is not known whether the gene is activated during wounding or pathogen infection, but it is clear from this study that Srchi13 transcripts transiently accumulate during normal nodule development as a specific response to Nod factor–producing bacteria.
Srchi13 Degrades LCO Molecules
During nodule development, obvious substrates for induced chitinases are the Nod factors. The production of LCO signals by the bacteria probably is counterbalanced by a feedback control, which is a plant-directed degradation mechanism, to avoid signal accumulation. In vetch and alfalfa, Nod factors are degraded rapidly by root chitinases; for alfalfa, a specific, induced Nod factor–hydrolyzing activity has been reported (Heidstra et al. 1994 ; Staehelin et al. 1994b , Staehelin et al. 1995 ). Also during stem nodulation on S. rostrata, Nod factor–degrading activities are induced (Figure 7). Thus, Srchi13 could be involved, directly or preventively, in Nod factor control. Functional analysis revealed that the protein has Nod factor degradation activity, presumably generating trimeric LCOs. This is not surprising because it is typical for class III chitinases to generate trimeric LCO products from LCO molecules that contain chemical modifications at both the reducing and nonreducing ends (M. Schultze, personal communication).
Another putative target for chitinases during nodulation is the bacterial peptidoglycan. Class III chitinases often are associated with lysozyme activity. Nevertheless, such activity as yet cannot be demonstrated (W. Van de Velde and M. Holsters, unpublished results).
A Role Is Proposed for Srchi13 during Nodulation
The expression pattern of Srchi13 is intricate. The first observable expression is 1 day after inoculation in the cortex of the root primordium, at some distance from the bacteria. Because of their physicochemical properties, Nod factors probably do not move over long distances, and therefore gene expression could be caused by diffusible secondary signals (Vijn et al. 1995 ). Two to 6 days after inoculation, Srchi13 expression in cells that delimit the developing nodule compartment could be seen as a precaution against eventual escape of Nod factors and azorhizobia, and at the later stages, Srchi13 could be operational in preventing the parenchyma to take up bacteria. Similarly, expression associated with sporadic cells in the invasion zone (not associated with infection threads) and with uninfected cells in the young fixation zone indicates a role for Srchi13 in rendering cells resistant against uptake of bacteria. Expression of Srchi13 in cells that flank sites of bacterial invasion (around infection pockets and in the infection center) may represent a direct feedback control of Nod factor levels. In the context of these hypotheses, it is important to note that the nod genes of A. caulinodans are expressed during all stages of nodule invasion and inside bacteroids for 20 to 30 days after inoculation (W. D'Haeze, M. Gao, R. De Rycke, G. Engler, M. Van Montagu, and M. Holsters, manuscript in preparation).
Rhizobia use a form of chitin signaling to trigger plant responses for nodule formation. Therefore, it comes as no surprise that a plant chitinase is mobilized in feedback control for signal inactivation. Tools have been borrowed seemingly from defense reactions, but here they are integrated into a developmental pathway. They are used in a constructive way to build a protected enclosure. Indeed, excessive amounts of mitogenic Nod factors could disturb plant growth or lead to elicitation of defense responses that might disturb the interaction (Schmidt et al. 1993 ; Savoure et al. 1997 ).
Chitinases in Beneficial and Pathogenic Interactions
Long and Staskawicz 1993 proposed that in its broadest sense, the term plant parasite indicates any microorganism that lives together with plants and uses plant compounds as nutrients. A parasitic interaction can be symbiotic when the microorganism returns something to the plant or pathogenic when it harms the plant. Within the framework of this definition, it is easy to see that pathogenesis and symbiosis can have common aspects, especially those of plant control over invasion. Both pathogens and symbionts must have developed ways to avoid, suppress, or overcome defense responses of the host and then eventually elicit them. Furthermore, in both types of interactions, the host responds by deploying sets of similar tools, for instance, chitinases. Chitinases, first identified in defense responses, also play a role in autoregulation of nodulation (Vasse et al. 1993 ) and in normal nodule development; interestingly, specific chitinases recently have been shown to be transiently expressed during beneficial endomycorrhizal fungi interactions with plants (Slezack et al. 1996 ). These observations show that plants use a limited set of tools in versatile ways and under different controls.
There are many parallels between the endomycorrhizal and the rhizobial symbioses (Gianinazzi-Pearson 1996 , Gianinazzi-Pearson 1997 ), and the use of sophisticated chitin signaling and containment by chitinases in the Rhizobium–legume symbiosis may be another hint at a possible evolutionary relationship with the beneficial fungus–plant interactions. This interpretation does not exclude the fact that for some aspects of the interaction, such as nodule primordium initiation, Nod factors also mimic endogenous oligosaccharide plant signals.
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METHODS |
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Plant Material
Seeds (Sesbania rostrata) were sterilized as described by Goethals et al. 1989 and germinated on Petri dishes in the dark at 28°C for 2 days. The seedlings were then transferred to pots containing 50% potting soil and 50% sand. The plants were grown at 28°C with a 16-hr light period for 2 to 3 months. The root primordia on the stem were infected with different Azorhizobium caulinodans strains by painting the stems with the bacterial inoculum. The inoculum was prepared by growing bacteria overnight in YEB medium (0.5% beef extract, 0.5% peptone, 0.1% yeast extract, 0.5% sucrose, and 0.002 M MgSO4), centrifuging the cells, and resuspending them in half of the starting volume in water. The plant material was harvested by peeling off the root primordia from the stem and freezing the tissue in liquid nitrogen.
cDNA Library Screening
To isolate the full-length sequences, 105 plaques of a 
ZAP (Stratagene, La Jolla, CA) cDNA library, which was prepared from RNA of developing nodules (Goormachtig et al. 1997 ), were screened according to standard procedures (Sambrook et al. 1989 ). 32P-labeled probes of didi-13 differential display fragments were generated by the T7 Quick Prime kit (Pharmacia, Uppsala, Sweden). Three rounds of screening were performed to obtain single positive plaques. Phages from positive plaques were transferred to their corresponding plasmid forms, according to the manufacturer's protocol (Stratagene). Plasmid DNA was prepared according to standard procedures (Sambrook et al. 1989 ). A plasmid with the full-length Srchi13 sequence was named pdidi13fl25.
RNA Gel Blot Analysis
RNA from uninfected root primordia or developing stem nodules was prepared as described by Goormachtig et al. 1995 . Ten micrograms of RNA derived from the different tissue samples was separated on a 1% agarose gel containing 2% formaldehyde and transferred to Hybond-N filters (Amersham, Aylesbury, UK). Hybridization was performed as described by Goormachtig et al. 1995 . 32P-labeled probes from didi-13 differential display fragments were made by the T7 Quick Prime kit (Pharmacia). Hybridized filters were exposed to Fuji films overnight to several days. To control equal loading and transfer of RNA, we stained filters with methylene blue, according to Sambrook et al. 1989 .
DNA Gel Blot Analysis
S. rostrata genomic DNA was prepared from young leaves, as described by Dellaporta et al. 1983 . Ten micrograms of S. rostrata genomic DNA was digested with EcoRI, BamHI, XbaI, SphI, PstI, and HindIII, according to the protocols described by Sambrook et al. 1989 . The DNA fragments were separated on a 0.7% agarose gel. The DNA was transferred to Hybond-N filters (Amersham) and hybridized at 65°C, according to standard procedures (Sambrook et al. 1989 ; Goormachtig et al. 1995 ). 32P-labeled probes derived from the insert of pdidi13fl25 were generated as described above.
In Situ Hybridization
Fresh plant material was fixed overnight in 4% paraformaldehyde and 0.25% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.2, at 4°C, and it was embedded in paraffin according to Cox et al. 1984 . A microtome (Reichert-Jung, Nussloch, Germany) was used to obtain 10-µm sections that were mounted on Vectabond-coated slides (Sigma). The in situ hybridization procedure was performed as described by Angerer and Angerer 1991 . 35S-labeled sense and antisense probes were generated with T7 and T3 polymerase (Gibco BRL, Gaithersburg, MD). The plasmid pdidi13fl25 was digested with EcoRI or XhoI to produce antisense and sense probes, respectively. After hybridization (and RNAse treatment), slides were washed for 1 hr in 2 x SSC (1 x SSC is 150 mM NaCl, 15 mM sodium citrate, pH 7.0) at room temperature and 1 hr in 0.1 x SSC at 58°C. Photographs were taken with a Diaplan microscope (Leitz, Wetzlar, Germany) equipped with bright- and dark-field optics.
DNA Sequence Analysis
DNA sequence procedures were used as described by Sanger et al. 1977 . DNA sequence data were assembled and analyzed with the Genetics Computer Group package (version 7; Genetics Computer Group, Madison, WI). The percentage of similarity between sequences was determined with the GAP program (Genetics Computer Group).
Srchi13 Synthesis in Escherichia coli
Srchi13 was cloned in the expression vector pMal-c2 (New England Biolabs, Beverly, MA). First, the Srchi13-coding sequence was amplified by polymerase chain reaction (PCR) by using a sense primer (5'-AAAATATGAAT TCATGGCTCCC-3') containing an EcoRI restriction site at the correct position for in-frame cloning of Srchi13 with the sequence of the maltose binding protein (MBP) and an antisense primer (5'-GCATAT TCTAGAT T TCAATATG-3') containing an XbaI restriction site. The PCR consisted of 100 ng of template (pdidi13fl25), 100 ng of each primer, 2 µL of 10 mM deoxynucleotide triphosphates, 2 µL of Vent DNA polymerase (New England Biolabs), and 10 µL of 10 x Vent buffer in a total volume of 100 µL. The PCR conditions were as follows: denaturation for 2 min at 94°C, 30 cycles for 1 min at 94°C, 1 min at 50°C, and 2 min at 72°C, and an extension of 5 min at 72°C. The PCR product was blunt ended by using the Klenow fragment of DNA polymerase I (Boehringer Mannheim, Germany) and phosphorylated, according to the protocols described by Sambrook et al. 1989 . The fragment was then cloned into a pBluescript KS- vector (Stratagene). A few clones were controlled for eventual PCR mistakes by sequencing, and the insert of an intact clone was cloned directionally into the pMal-c2 by using the EcoRI and XbaI restriction sites and the protocols described by Sambrook et al. 1989 . The clone was called pMal-c2-Srchi13.
To express the MBP–Srchi13 fusion and as a control, MBP, E. coli (Mc1061) cells containing pMal-c2-Srchi13, or pMal-c2 were grown in Luria-Bertani medium (1% trypton, 1% NaCl, 0.5% yeast extract, and 0.1% glucose) at 37°C until cell density reached OD600 nm of 0.4 to 0.6. Next, isopropyl ß-D-thiogalactoside was added to a final concentration of 0.1 mM, and the cells were grown for another 2 hr at 30°C. The E. coli cells subsequently were fractionated by using ultrasonic waves, and the cell fraction and the supernatant were separated by centrifuging. The soluble protein fractions then were enriched for the MBP or MBP–Srchi13 by affinity chromatography according to the manufacturer's instructions (New England Biolabs). The proteins were eluted in PBS medium, pH 7.4.
Nod Factor–Degrading Activity in Developing Nodules
14C-labeled Nod factors of A. caulinodans ORS571 were prepared and purified as described by Mergaert et al. 1993 . Both Nod factors containing C18:1 fatty acid (peak I) and C18:0 fatty acid (peak II) were used for the experiment and gave similar results. Radioactive incorporation was measured by scintillation counting.
To obtain protein extracts of uninfected root primordia, the tissue was frozen in liquid nitrogen, ground, and resuspended in PBS buffer. For incubation, 20 µg of crude extract and 50 µg of enriched protein fractions were used.
Protein extracts and 14C-labeled Nod factors (3000 cpm) were mixed in 200 µL of 50 mM sodium acetate buffer, pH 5.2, and incubated at 37°C overnight. Nod factors and their degradation products (nonreducing end) then were extracted with butanol and analyzed on octadecyl silica thin-layer chromatography plates, as described by Mergaert et al. 1993 . The results were visualized and analyzed by using a PhosphorImager system (Molecular Dynamics, Sunnyvale, CA).
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ACKNOWLEDGMENTS |
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We thank Wim D'Haeze, Peter Mergaert, and Christa Verplancke for help with the Nod factors, Sylvia Herman for excellent technical assistance, Martine De Cock for help preparing the manuscript, and Karel Spruyt, Christiane Germonprez, and Rebecca Verbanck for art work. This work was supported by grants from the Belgian Programme on Interuniversity Poles of Attraction (Prime Minister's Office, Science Policy Programming, No. 38) and in part by the European Community's BIOTECH Programme, as part of the Project of Technological Priority 1993 to 1996. S.L. and M.H. are Research Fellow and Research Director of the Fund for Scientific Research (Flanders), respectively.
Received December 24, 1997; accepted March 25, 1998.
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REFERENCES |
|---|
Angerer, L.M., and Angerer, R.C. (1991) Localization of mRNAs by in situ hybridization. Methods Cell Biol. 35:37-71[Medline].
Baron, C., and Zambryski, P.C. (1995) The plant response in pathogenesis, symbiosis, and wounding: Variations on a common theme? Annu. Rev. Genet. 29:107-129[CrossRef][ISI][Medline].
Collinge, D.B., Kragh, K.M., Mikkelsen, J.D., Nielsen, K.K., Rasmussen, U., and Vad, K. (1993) Plant chitinases. Plant J. 3:31-40[CrossRef][ISI][Medline].
Cox, K., DeLeon, D.V., Angerer, L.M., and Angerer, R.C. (1984) Detection of mRNAs in sea urchin embryos by in situ hybridization using asymmetric RNA probes. Dev. Biol. 101:485-502[CrossRef][ISI][Medline].
De Jong, A.J., Cordewener, J., Lo Schiavo, F., Terzi, M., Vandekerckhove, J., Van Kammen, A., and De Vries, S.C. (1992) A carrot somatic embryo mutant is rescued by chitinase. Plant Cell 4:425-433[Abstract/Free Full Text].
Dellaporta, S.L., Wood, J., and Hicks, J.B. (1983) A plant DNA minipreparation: Version II. Plant Mol. Biol. Rep. 1:19-21.
Djordjevic, M.A., Gabriel, D.W., and Rolfe, B.G. (1987) Rhizobium—The refined parasite of legumes. Annu. Rev. Phytopathol. 25:145-168[CrossRef][ISI].
Duhoux, E. (1984) Ontogénèse des nodules caulinaires du Sesbania rostrata (légumineuses). Can. J. Bot. 62:982-994.
Franssen, H.J., Vijn, I., Yang, W.C., and Bisseling, T. (1992) Developmental aspects of the Rhizobium–legume symbiosis. Plant Mol. Biol. 19:89-107[CrossRef][ISI][Medline].
Gianinazzi-Pearson, V. (1996) Plant cell responses to arbuscular mycorrhizal fungi: Getting to the roots of the symbiosis. Plant Cell 8:1871-1883[CrossRef][ISI][Medline].
Gianinazzi-Pearson, V. (1997). Have common plant systems co-evolved in fungal and bacterial root symbioses? In Biological Fixation of Nitrogen for Ecology and Sustainable Agriculture, NATO ASI Series, Vol. G39, A. Legocki, H. Bothe, and A. Pühler, eds (Berlin: Springer-Verlag), pp. 321–324.
Goethals, K., Gao, M., Tomekpe, K., Van Montagu, M., and Holsters, M. (1989) Common nodABC genes in Nod locus 1 of Azorhizobium caulinodans: Nucleotide sequence and plant-inducible expression. Mol. Gen. Genet. 219:289-298[Medline].
Goethals, K., Leyman, B., Van den Eede, G., Van Montagu, M., and Holsters, M. (1994) An Azorhizobium caulinodans ORS571 locus involved in lipopolysaccharide production and nodule formation on Sesbania rostrata stems and roots. J. Bacteriol. 176:92-99[Abstract/Free Full Text].
Goormachtig, S., Valerio-Lepiniec, M., Szczyglowski, K., Van Montagu, M., Holsters, M., and de Bruijn, F.J. (1995) Use of differential display to identify novel Sesbania rostrata genes enhanced by Azorhizobium caulinodans infection. Mol. Plant-Microbe Interact. 8:816-824[ISI][Medline].
Goormachtig, S., Alves-Ferreira, M., Van Montagu, M., Engler, G., and Holsters, M. (1997) Expression of cell cycle genes during Sesbania rostrata stem nodule development. Mol. Plant-Microbe Interact. 10:316-325[ISI][Medline].
Hanfrey, C., Fife, M., and Buchanan-Wollaston, V. (1996) Leaf senescence in Brassica napus: Expression of genes encoding pathogenesis-related proteins. Plant Mol. Biol. 30:597-609[CrossRef][ISI][Medline].
Heidstra, R., and Bisseling, T. (1996) Nod factor–induced host responses and mechanisms of Nod factor perception. New Phytol. 133:25-43[CrossRef].
Heidstra, R., Geurts, R., Franssen, H., Spaink, H.P., Van Kammen, A., and Bisseling, T. (1994) Root hair deformation activity of nodulation factors and their fate on Vicia sativa.. Plant Physiol. 105:787-797[Abstract].
Kragh, K.M., Hendriks, T., de Jong, A.J., Lo Schiavo, F., Bucherna, N., Højrup, P., Mikkelsen, J.D., and de Vries, S.C. (1996) Characterization of chitinases able to rescue somatic embryos of the temperature-sensitive carrot variant ts11.. Plant Mol. Biol. 31:631-645[CrossRef][ISI][Medline].
Kuranda, M.J., and Robbins, P.W. (1991) Chitinase is required for cell separation during growth of Saccharomyces cerevisiae.. J. Biol. Chem. 266:19758-19767[Abstract/Free Full Text].
Lawton, K., Ward, E., Payne, G., Moyer, M., and Ryals, J. (1992) Acidic and basic class III chitinase mRNA accumulation in response to TMV infection of tobacco. Plant Mol. Biol. 19:735-743[CrossRef][ISI][Medline].
Levorson, J., and Chlan, C.A. (1997) Plant chitinase consensus sequences. Plant Mol. Biol. Rep. 15:122-133[ISI].
Long, S.R. (1996) Rhizobium symbiosis: Nod factors in perspective. Plant Cell 8:1885-1898[CrossRef][ISI][Medline].
Long, S.R., and Staskawicz, B.J. (1993) Prokaryotic plant parasites. Cell 73:921-935[CrossRef][ISI][Medline].
Lotan, T., Ori, N., and Fluhr, R. (1989) Pathogenesis-related proteins are developmentally regulated in tobacco flowers. Plant Cell 1:881-887[Abstract/Free Full Text].
Meins, F., Jr., Fritig, B., Linthorst, H.J.M., Mikkelsen, J.D., Neuhaus, J.-M., and Ryals, J. (1994) Plant chitinase genes. Plant Mol. Biol. Rep. 12(CPGN suppl.):22-28.
Mergaert, P., Van Montagu, M., Promé, J.-C., and Holsters, M. (1993) Three unusual modifications, a D-arabinosyl, an N-methyl, and a carbamoyl group, are present on the Nod factors of Azorhizobium caulinodans strain ORS571. Proc. Natl. Acad. Sci. USA 90:1551-1555[Abstract/Free Full Text].
Mergaert, P., Van Montagu, M., and Holsters, M. (1997) Molecular mechanisms of Nod factor diversity. Mol. Microbiol. 25:811-817[CrossRef][ISI][Medline].
Métraux, J.P., Streit, L., and Staub, T. (1988) A pathogenesis-related protein in cucumber is a chitinase. Physiol. Mol. Plant Pathol. 33:1-9.
Neale, A.D., Wahleithner, J.A., Lund, M., Bonnett, H.T., Kelly, A., Meeks-Wagner, D.R., Peacock, W.J., and Dennis, E.S. (1990) Chitinase, ß-1,3-glucanase, osmotin, and extensin are expressed in tobacco explants during flower formation. Plant Cell 2:673-684[Abstract/Free Full Text].
Nielsen, K.K., Mikkelsen, J.D., Kragh, K.M., and Bojsen, K. (1993) An acidic class III chitinase in sugar beet: Induction of Cercospora beticola, characterization, and expression in transgenic tobacco plants. Mol. Plant-Microbe Interact. 6:495-506[Medline].
Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual, 2nd ed. (Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press).
Sanger, F., Nicklen, S., and Coulson, A.R. (1977) DNA sequencing with chain-terminating inhibitors. Proc. Natl. Acad. Sci. USA 74:5463-5467[Abstract/Free Full Text].
Savouré, A., Sallaud, C., El-Turk, J., Zuanazzi, J., Ratet, P., Schultze, M., Kondorosi, A., Esnault, R., and Kondorosi, E. (1997) Distinct response of Medicago suspension cultures and roots to Nod factors and chitin oligomers in the elicitation of defense-related responses. Plant J. 11:277-287[CrossRef].
Schmidt, J., Röhrig, H., John, M., Wieneke, U., Stacey, G., Koncz, C., and Schell, J. (1993) Alteration of plant growth and development by Rhizobium nodA and nodB genes involved in the synthesis of oligosaccharide signal molecules. Plant J. 4:651-658.
Schultze, M., Kondorosi, E., Ratet, P., Buiré, M., and Kondorosi, A. (1994) Cell and molecular biology of Rhizobium–plant interactions. Int. Rev. Cytol. 156:1-75.
Slezack, S., Dassi, B., and Dumas-Gaudot, E. (1996). Arbuscular mycorrhiza-induced chitinase isoforms. In Chitin Enzymology, Vol. 2, R.A.A. Muzzarelli, ed (Grottammare, Italy: Atec Edizioni), pp. 339–347.
Staehelin, C., Müller, J., Mellor, R.B., Wiemken, A., and Boller, T. (1992) Chitinase and peroxidase in effective (fix+) and ineffective (fix-) soybean nodules. Planta 187:295-300.
Staehelin, C., Granado, J., Müller, J., Wiemken, A., Mellor, R.B., Felix, G., Regenass, M., Broughton, W.J., and Boller, T. (1994a) Perception of Rhizobium nodulation factors by tomato cells and inactivation by root chitinases. Proc. Natl. Acad. Sci. USA 91:2196-2200[Abstract/Free Full Text].
Staehelin, C., Schultze, M., Kondorosi, E., Mellor, R.B., Boller, T., and Kondorosi, A. (1994b) Structural modifications in Rhizobium meliloti Nod factors influence their stability against hydrolysis by root chitinases. Plant J. 5:319-330[CrossRef].
Staehelin, C., Schultze, M., Kondorosi, E., and Kondorosi, A. (1995) Lipo-chitooligosaccharide nodulation signals from Rhizobium meliloti induce their rapid degradation by the host plant alfalfa. Plant Physiol. 108:1607-1614[Abstract].
Tsien, H.C., Dreyfus, B.L., and Schmidt, E.L. (1983) Initial stages in the morphogenesis of nitrogen-fixing stem nodules of Sesbania rostrata.. J. Bacteriol. 156:888-897[Abstract/Free Full Text].
Tsujibo, H., Orikoshi, H., Imada, C., Okami, Y., Miyamoto, K., and Inamori, Y. (1993) Site-directed mutagenesis of chitinase from Alteromonas sp. strain O-7. Biosci. Biotechnol. Biochem. 57:1396-1397[Medline].
Vance, C.P. (1983) Rhizobium infection and nodulation: A beneficial plant disease? Annu. Rev. Microbiol. 37:399-424[CrossRef][ISI][Medline].
Van den Eede, G., Dreyfus, B., Goethals, K., Van Montagu, M., and Holsters, M. (1987) Identification and cloning of nodulation genes from the stem-nodulating bacterium ORS571. Mol. Gen. Genet. 206:291-299[CrossRef][ISI].
Vasse, J., de Billy, F., and Truchet, G. (1993) Abortion of infection during the Rhizobium meliloti–alfalfa symbiotic interaction is accompanied by a hypersensitive reaction. Plant J. 4:555-566[CrossRef][ISI].
Vijn, I., Yang, W.-C., Pallisgård, N., Jensen, E.Ø., Van Kammen, A., and Bisseling, T. (1995) VsENOD5, VsENOD12 and VsENOD40 expressing during Rhizobium-induced nodule formation on Vicia sativa roots. Plant Mol. Biol. 28:1111-1119[CrossRef][ISI][Medline].
Watanabe, T., Kobori, K., Miyashita, K., Fujii, T., Sakai, H., Uchida, M., and Tanaka, H. (1993) Identification of glutamic acid 204 and aspartic acid 200 in chitinase A1 of Bacillus circulans WL-12 as essential residues for chitinase activity. J. Biol. Chem. 268:18567-18572[Abstract/Free Full Text].
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ABSTRACT
INTRODUCTION
