INTRODUCTION

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Osmoregulated periplasmic glucans (OPGs) are general constituents of the envelopes of gram-negative bacteria (4). Glucose is the sole constitutive sugar, and their abundance in the periplasmic compartment is osmoregulated, with the highest levels synthesized during growth at very low osmolarity. Four families of OPGs have been described on the basis of structural features of the polyglucose backbone. In family I, OPGs appear to range from 5 to 12 glucose residues, with the principal species containing 8 or 9 glucose residues. The structure is highly branched, the backbone consisting of -1,2-linked glucose units to which the branches are attached by -1,6 linkages. In family II, OPGs are composed of a cyclic -1,2-glucan backbone containing 17 to 25 glucose residues. In family III, OPGs are -1,6 and -1,3 cyclic glucans containing 10 to 13 glucose units per ring. In family IV, OPGs are cyclic and have a unique degree of polymerization (DP, = 13, 16, or 18). One linkage is -1,6 whereas all the other glucose residues are linked by -1,2 linkages. Depending on the species considered, OPGs can be modified to various extents by a variety of substituents. Mutations at the loci ndvA and ndvB in Sinorhizobium meliloti (8), ndvB and ndvC in Bradyrhizobium japonicum (3), chvA and chvB in Agrobacterium tumefaciens (17), and hrpM in Pseudomonas syringae (11, 13) impair OPG biosynthesis, and these mutants fail to interact properly with a host plant as a symbiont or a pathogen. However, beyond this functional homology, the OPGs synthesized by these different bacteria are very different in structure. The OPGs of S. meliloti and A. tumefaciens are cyclic structures of family II that may be modified with anionic substituents such as phosphoglycerol and/or succinyl moieties (5), the OPGs of B. japonicum are cyclic structures of family III that may be modified by substitution with phosphocholine (18), while the OPGs of P. syringae are linear and highly branched and devoid of any substituents (24).

Except for substitution, the OPGs of P. syringae are very similar to the OPGs synthesized by the enterobacterium Escherichia coli, which colonizes animals. In this bacterium, OPGs are a heterogeneous family of oligosaccharides ranging from 5 to 12 glucose residues (9). The structure is highly branched and consists of a backbone of -1,2-linked glucose units to which the branches are attached by -1,6 linkages. They may be substituted to various degrees with sn-1-phosphoglycerol, phosphoethanolamine, and O-succinyl ester residues.

Erwinia chrysanthemi is a pathogenic enterobacterium responsible for the soft rot disease of a wide range of plants. The pathogenicity of the bacterium is due in part to its ability to produce extracellular enzymes, such as pectinases, cellulases, and proteases, which are able to degrade constituents of the plant cell wall (1, 19).

Recently, the opgGH operon of E. chrysanthemi was cloned and inactivated on the chromosome (15). The mutants, defective in OPG synthesis, exhibit a pleiotropic phenotype and a complete loss of virulence. In an attempt to understand the function of OPGs at the molecular level in the plant-bacterium interaction, we examined the structures of the OPGs synthesized by E. chrysanthemi. OPGs produced by E. chrysanthemi are heterogeneous in size and possess structures similar to those of E. coli and P. syringae. In addition, they may be substituted by O-acetyl and O-succinyl ester residues.

	MATERIALS AND METHODS

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Bacterial strains and growth. E. chrysanthemi 3937 was grown on a rotary shaker at 30°C in LOS (low osmolarity) medium [4 g of casein hydrolysate, 0.5 mg of FeSO₄, 18 mg of MgCl₂, 200 mg of (NH₄)₂SO₄, and 175 mg of K₂HPO₄ per liter (pH 7.2)] supplemented with 2 mg of thiamine and 2 g of glycerol. The LOS medium osmolarity is 85 mosM. To obtain high-osmolarity medium, NaCl was added to LOS medium up to 0.3 M.

Cellular compartmentalization of OPGs. One hundred-milliliter cultures were harvested during the stationary phase of growth. After centrifugation at 4°C for 20 min at 4,000 × g, the supernatant was extracted with 50% ethanol, and cells from the pellet were gently resuspended at 4°C in 3 ml of 200 mM Tris-HCl (pH 8)-8.5% sucrose-10 µg of lysozyme per ml-20 mM EDTA and centrifuged at 4°C. Both the supernatant, containing the EDTA-released material, and the cell pellet, containing the retained fraction, were extracted with 50% ethanol. After concentration in a rotary evaporator, the extracts were fractionated by gel filtration on a Bio-Gel P-4 (Bio-Rad). The column (1.6 cm in cross section, 55 cm in height) was eluted at room temperature with 0.5% acetic acid at a flow rate of 15 ml/h, and fractions of 2.5 ml were collected. The oligosaccharides emerged in a peak of intermediate weight detected by the phenol-sulfuric acid procedure (6).

Large-scale isolation and purification of OPGs. Bacteria were collected during the exponential phase of growth by centrifugation at 4°C for 15 min at 8,000 × g. Cell pellets were extracted with 5% trichloroacetic acid (TCA), and the TCA extracts were neutralized with ammonium hydroxide and desalted on a Sephadex G-15 column. The desalted material was then fractionated by gel filtration on a Bio-Gel P-4 as described above. Fractions containing oligosaccharides were pooled and lyophilized.

Determination of neutral and anionic characteristics of OPGs. Glucans extracted as described above were desalted on a Bio-Gel P-2 column (Bio-Rad), lyophilized, and resuspended in 2 ml of 10 mM Tris-HCl (pH 7.4) buffer. OPG-containing fractions were pooled and chromatographed on a DEAE-Sephacel column (1.5 cm in cross section, 38 cm in height; Pharmacia) equilibrated with 10 mM Tris-HCl (pH 7.4) and eluted with the same buffer containing increasing concentrations of NaCl ranging from 0 to 0.2 M in steps of 0.05 M. A volume of 60 ml was used for each NaCl concentration, and the volume of each collected fraction was 4 ml.

Determination of succinate and acetate content from OPGs. OPGs were prepared as described above except that formic acid (0.5%) was used in place of acetic acid for Bio-Gel P-4 elution. OPGs were then desalted on Bio-Gel P-2 and separated by DEAE-Sephacel chromatography as described above. Fractions containing oligosaccharides were pooled, desalted, and lyophilized. One milligram of OPGs was dissolved in 0.2 ml of 0.5 M NaOH and incubated at 100°C for 30 min to liberate the succinyl and acetyl residues from OPGs. Glucosidic backbones were removed by absorption on 50 mg of charcoal suspended in 0.3 ml of water, and the charcoal was then washed three times with 0.5 ml of water. The four supernatants were pooled (2 ml) and neutralized with Dowex AG 50W-X8 (Bio-Rad) on H⁺ form. Succinic acid and acetic acid contents were determined with a succinic acid kit and an acetic acid kit, respectively (Boehringer Mannheim).

Deesterification of OPGs. OPGs were treated with 0.1 M KOH at 37°C for 1 h. After neutralization with AG 50W-X8 (Bio-Rad) on H⁺ form, the samples were desalted on a Bio-Gel P-2 column.

HPAEC-PAD. For high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD), analysis and preparation of oligosaccharides were performed on a CarboPac PA-100 anion-exchange column (4 by 250 mm; Dionex, Sunnyvale, Calif.) equipped with a CarboPac PA guard column (3 by 25 mm; Dionex). Oligosaccharides were detected with a PAD II pulsed amperometric detector with a gold electrode (Dionex). The chromatographic data were integrated and plotted with an SP 4270 integrator (Spectra-Physic, San Jose, Calif.). Oligosaccharides were eluted at a flow rate of 1 ml/min by a two-step procedure consisting of (i) 0.05 M sodium acetate in 0.15 M NaOH for 5 min and (ii) a linear gradient of 0.05 to 0.2 M sodium acetate in 0.15 M NaOH for 35 min. After every run, the column was reequilibrated in 0.15 M NaOH for 15 min.

MALDI-MS. For matrix-assisted laser desorption-ionization (MALDI)-mass spectrometry (MS), experiments were carried out on a Vision 2000 (Finnigan MAT, Bremen, Germany) time-of-flight mass spectrometer equipped with a nitrogen laser (337-nm wavelength and 3-ns pulse width). After selection of the appropriate site on the target by a microscope, the laser light was focused onto the sample-matrix mixture at an angle of 15° and a power level of 10⁶ to 10⁷ W/cm². Positive ions were extracted by a 5- to 10-keV acceleration potential, focused by a lens, and the masses were separated by a Reflectron time-of-flight instrument. At the detector, ions were postaccelerated to 20 keV for maximum detection efficiency. The resulting signals were recorded with a fast transient digitizer with a maximum of 2.5 ns channel resolution and transferred to a personal computer for accumulation, calibration, and storage. All MALDI mass spectra are the result of 20 single-shot accumulations.

Methylation analysis. The oligosaccharides were methylated according to Paz-Parente et al. (16). The methyl ethers were obtained after methanolysis (0.5 M HCl in methanol at 80°C for 24 h) and analyzed as partially methylated methyl glycosides by gas-liquid chromatography (GLC)-MS (7). GLC was performed using a Delsi apparatus with a capillary column (25 m by 0.2 mm) coated with DB-1 (0.5-µm film thickness), applying a temperature gradient of 110 to 240°C at 3°C/min and a helium pressure of 40 kPa. The mass spectra were recorded on a Nermag 10-10B mass spectrometer (Rueil-Malmaison, France) using an electron energy of 70 eV and an ionizing current of 0.2 mA.

NMR spectroscopy. Prior to nuclear magnetic resonance (NMR) spectroscopic analysis, the oligosaccharides were twice treated with ²H₂O at room temperature. After each exchange treatment, the materials were lyophilized. Finally, each sample was redissolved in 0.5 ml of ²H₂O (99.96 atom% ²H; Aldrich). The NMR experiments were performed on a Bruker AM-400 spectrometer controlled by an Aspect 3000 computer with an array processor and equipped with a 5-mm mixed ¹H-¹³C probe head at a temperature of 25°C. Chemical shifts () were referenced to acetone, the internal standard. Two-dimensional homonuclear COSY (correlation spectroscopy) 90, relayed and double-relayed COSY experiments, and two-dimensional heteronuclear multiple quantum coherence (HMQC) were performed by using standard Bruker pulse programs.

Other methods. Protein concentrations were determined by the method of Lowry et al. (12) with bovine serum albumin as the reference protein. Total carbohydrate concentrations were determined by the anthrone method of Spiro (22) with D-glucose as the standard. Sugar analysis was carried out by GLC of trimethylsilyl derivatives of methyl glycosides formed by methanolysis in 0.5 M HCl in methanol at 80°C for 24 h (14). Reducing sugars were measured by the same method after reduction of the oligosaccharides with NaBH₄.

	RESULTS

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Isolation and characterization of OPGs. OPGs were extracted from cells of E. chrysanthemi by previously described procedures which involved TCA extraction. Fractionation on a Bio-Gel P-4 column allowed the separation of two main sugar-containing compounds (Fig. 1). The compounds eluting in the void volume of the column were probably high-molecular-weight lipo- or exopolysaccharides, as previously observed (24). The second peak represented the OPGs. When cells of E. chrysanthemi were grown in LOS medium supplemented with 0.3 M NaCl, the amount of cell-associated oligosaccharides was reduced (Fig. 1). With no addition of NaCl, the OPG content (micrograms of glucose per milligram of cell protein) was 76 ± 5. With the addition of 0.3 M NaCl, the OPG content was 7 ± 3. Thus, cells grown in a medium of low osmolarity synthesized approximately 10 times more OPGs than cells grown in the same medium with 0.3 M NaCl.

View larger version (23K): [in this window] [in a new window]   FIG. 1.   Bio-Gel P-4 elution profile of OPGs of E. chrysanthemi grown in LOS medium () or in LOS medium supplemented with 0.3 M NaCl (). The column (1.6 by 55 cm) was eluted with 0.5% acetic acid, and aliquots were analyzed for total carbohydrates (see Materials and Methods). Results are expressed as glucose equivalents per milliliter of eluant. In both cases, fractions indicated by the horizontal bar were lyophilized.

Cellular compartmentalization of OPGs. Export of OPGs to the extracellular medium has been observed for members of the family Rhizobiaceae, and extracellular glucans may play a fundamental role in plant-bacterium interactions (3, 5). However, this export varies greatly among different species and strains and is dependent on growth stage and culture conditions. Thus, it was important to determine whether the OPGs produced by E. chrysanthemi can be recovered in the external medium, at least under certain circumstances. Actually, 75% of the OPGs were found in the growth medium from stationary-phase cultures at low osmolarity (see Materials and Methods). Of the remaining 25%, 80% were liberated by EDTA treatment, which is known to release periplasmic content. Similar results were obtained when cells were separated by filtration.

OPG substitution. Thin-layer chromatography analysis showed different patterns for native and KOH-treated OPGs (see Materials and Methods; data not shown). This was indicative of a substitution of OPGs by ester-linked residues (alkaline-sensitive substitution). Therefore, OPGs were analyzed by anion-exchange chromatography (DEAE-Sephacel). When the OPGs were extracted from cells grown at low osmolarity, practically no material was retained by the column (Fig. 2), but when the OPGs were extracted from cells grown at high osmolarity, 20% of the total amount was retained and subsequently eluted with low-ionic-strength buffer (10 mM Tris-HCl [pH 7.4], 50 mM NaCl).

View larger version (17K): [in this window] [in a new window]   FIG. 2.   DEAE-Sephacel anion-exchange column chromatography profiles of OPGs from E. chrysanthemi grown in LOS medium (A) or LOS medium with 0.3 M NaCl (B). Ionic strength was increased by steps of 0.05 M NaCl at the fractions indicated by the arrows. Fractions (4 ml) were collected, and total carbohydrate concentrations were determined by the anthrone method (see Materials and Methods). The amount of glucans in each fraction is expressed as a percentage of the total amount loaded on the column.

¹H-NMR analysis. The OPGs of E. chrysanthemi grown in the high-osmolarity medium (Fig. 3) were analyzed by ¹H-NMR analysis. Peaks at 5.4 ppm are indicative of the H-1 of reducing glucose residues with the -anomeric configuration. Peaks near 4.9 and 4.55 ppm are indicative of the H-1 of glucose residues engaged in -1,2 and -1,6 linkages, respectively. Those at 4.2 ppm were assigned to the H-6 of glucose residues linked by a -1,6 linkage. The two triplets present between 2.4 and 2.8 ppm confirmed the presence of succinyl substituents.

View larger version (17K): [in this window] [in a new window]   FIG. 3.   1H-NMR analysis of OPGs of E. chrysanthemi grown in LOS with 0.3 M NaCl. NMR spectra were recorded as described in Materials and Methods. HOD, partially deuterated water.

Methylation analysis. OPGs extracted from cells grown at low or high osmolarity were first subjected to alkaline treatment to remove all succinic and acetic substituents. Then, the OPGs were methanolized and, after acetylation, subjected to GLC-MS analysis. In both cases, the results of the methylation analysis revealed the presence of 3,4-di-, 3,4,6-tri-, and 2,3,4,6-tetra-O-methylglucose (Table 1). This indicated that the glucans were branched structures, with branch points doubly substituted in positions 2 and 6. The ratio of 2-linked Glc to 2,6-linked Glc was approximately 2. This suggested that about two glucose residues were linked through positions 1 and 2 for every internal glucose residue linked through positions 1, 2, and 6. This indicated that the glucans were highly branched structures.

¹³C-NMR analysis. KOH-treated OPGs of E. chrysanthemi grown in low-osmolarity medium were analyzed by ¹³C-NMR analysis (Fig. 4A). This analysis revealed that the C-1 resonance peaks all clustered near 104 ppm. This shift is indicative of the -glycosidic linkage. The peaks near 70 ppm are indicative of -1,6 linkages and represent the resonances for both C-4 and C-6 (-1,6) carbons. The C-6 assignment was confirmed by a nonselective polarization transfer technique, showing the characteristic CH₂ negative signal. The resonances near 83 ppm were assigned to C-2 in -1,2 glycosidic linkages. Peaks near 62 ppm are indicative of C-6 carbons not involved in glycosidic linkages, while those at 74.5 ppm are indicative of C-2 carbons not involved in glycosidic linkages. The resonances near 77.5 ppm may be assigned to both C-5 and C-3 carbons not involved in glycosidic linkages. The acyclic nature of the glucans was confirmed by the presence of resonances at 96 and 92.5 ppm, corresponding to the  and  anomeric carbons, respectively, of the reducing-glucose residues. Furthermore, the resonances at 72.7 ppm could be assigned to C-3 and C-5 of the  anomer of the reducing glucose, while the corresponding signals of the  anomer were part of the complex of signals at 75 ppm. All these assignments were confirmed by a ¹³C-decoupled, ¹H-detected HMQC analysis.

View larger version (23K): [in this window] [in a new window]   FIG. 4.   13C-NMR analysis of KOH-treated OPGs of E. chrysanthemi (A) and P. syringae (B) grown in LOS medium. NMR spectra were recorded as described in Materials and Methods.

MALDI-MS. Quasimolecular ions were obtained by the MALDI-MS method for the OPGs of E. chrysanthemi grown in LOS medium without NaCl. This analysis revealed the presence of six sodium-cationized molecular ions, [M + Na]⁺, at approximately m/z 1,013, 1,175, 1,337, 1,499, 1,661, and 1,823 (Fig. 5). These molecular ion species had masses identical to those expected for linear glucans composed of 6 to 11 glucose residues, with the principal species containing 8 glucose residues. Four sodium-cationized molecular ions, [M + Na]⁺, at approximately m/z 1,055, 1,217, 1,379, and 1,541, were detected and correspond to glucans of 6, 7, 8, and 9 degrees of polymerization, respectively, with a mass increment of 42, which is expected for an O-ester-linked acetyl residue (Fig. 5). Peaks corresponding to minor species with m/z decreased by 18 below the values of the main molecular ions were due to in-source loss of water occurring during ionization, a phenomenon frequently observed in MS of carbohydrates. Peaks corresponding to the [M + K]⁺ ions were also present, with m/z increased by 16 above the masses of the corresponding sodiated ions. This analysis confirmed that each species of OPGs can be substituted by one acetyl residue.

View larger version (23K): [in this window] [in a new window]   FIG. 5.   Positive-ion MALDI mass spectra of OPGs of E. chrysanthemi grown in LOS medium without addition. Mass assignments are based on an external calibration.

HPAEC-PAD analysis. In HPAEC-PAD analysis of KOH-treated OPGs (Fig. 6), the number of peaks was greater than the number of carbohydrate-based signals determined by MALDI-MS. In order to understand this phenomenon, we performed HPAEC separation and subsequent MALDI-MS and methylation analyses of the predominant fractions. The MALDI-MS analysis (Table 2) revealed a molecular weight distribution slightly larger (5 to 12 glucose residues) than observed before separation (Fig. 5) and indicated that the retention volume increased as the degree of polymerization increased. Moreover, the presence of several isomeric oligomers was also revealed, as the component of fractions 3 and 4 contained 7 glucose units (Fig. 7A and B) and components of fractions 5 and 6 contained 8 glucose units (Fig. 7C and D). The presence of minor signals in the spectra (Fig. 7) could result from some contamination of the neighboring peaks or from coelution with other compounds. Based on the MALDI-MS data, methylation analyses were performed on the components of fractions 3, 4, 5, and 6 (Table 3). These analyses clearly indicate that the OPGs of fraction 3 possess a higher number of -1,6 branches than the OPGs of fraction 4 (Table 3). The same result was obtained for the OPGs of fractions 5 and 6 (Table 3). This clearly indicates that for a given isomeric OPG, the retention volume decreased when the number of -1,6 branches increased.

View larger version (19K): [in this window] [in a new window]   FIG. 6.   HPAEC elution profile of KOH-treated OPGs of E. chrysanthemi. Oligosaccharide peaks were detected by PAD. The number above each peak indicates the fraction number. NaOAc, sodium acetate.

View larger version (14K): [in this window] [in a new window]   FIG. 7.   Positive-ion MALDI mass spectra of OPGs of E. chrysanthemi HPAEC-PAD fractions 3 (A), 4 (B), 5 (C), and 6 (D).

	DISCUSSION

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OPGs of E. chrysanthemi are found in larger amounts when the osmolarity of the growth medium decreases. The ratio observed (10-fold more in low osmolerity) is similar to those already reported for E. coli and P. syringae (12, 14, 24). The structure determination of E. chrysanthemi OPGs showed that they are small glucans ranging in size from 5 to 12 glucose residues, made of a -1,2-linked glucose backbone to which branches are attached by -1,6 linkages. Thus, the OPG backbone structures synthesized by the two Enterobacteriaceae and the pseudomonad can be considered identical. One noticeable difference is at the level of cell concentration (about 30, 50, and 75 µg of glucose per mg of cell protein for E. coli, P. syringae, and E. chrysanthemi, respectively). But the main differences reside in the number and variety of substituents, from the no-substituted OPGs of P. syringae to the highly substituted anionic OPGs of E. coli.

Further analyses by HPAEC-PAD revealed the high level of heterogeneity of these OPGs: a variable number of glucose residues are arranged with a variable number of branches whose position on the backbone is random. In no way can the OPGs of this family be described as repetitions of a conserved unit. This characteristic marks the demarcation between OPGs and other polysaccharidic structures found in bacteria.

Recently, the E. chrysanthemi genes involved in OPG biosynthesis (opgGH, accession number AJ294718) have been cloned. They exhibit a very high degree of similarity (86%) to their homologues mdoGH and hrpM, respectively, in E. coli and P. syringae (15). The fact that highly similar genes control the biosynthesis of OPGs belonging to the same structural family may seem obvious. However, the Rhodobacter sphaeroides opgGH genes (accession number AF016298) were recently isolated and characterized (A. Puskas, V. Cogez, E. Gak, J.-P. Bohin, and S. Kaplan, unpublished data). They show a significant, although lower (55%), level of similarity to mdoGH, but they control the biosynthesis of OPGs belonging to family IV (Talaga et al., unpublished). Thus, structural analysis of OPGs will remain necessary even when gene sequences are known.

Moreover, this structural analysis revealed differences at the level of backbone substitution. E. chrysanthemi OPGs can be substituted by acetyl and succinyl residues, and the presence of succinyl depends on the growth conditions. Thus, while P. syringae OPGs are neutral and E. coli OPGs are highly anionic (10), E. chrysanthemi OPGs could have a mild anionic character when growth occurs at high osmolarity. The function of glucan substitution, which varies to a large extent from species to species, remains obscure. A highly attenuated Salmonella enterica mutant with a high immunogenic and protective potential in mice was found to have a transposon insertion in a gene similar to mdoB (25). In E. coli, the product of this gene is a membrane-bound P-glycerol transferase, which transfers residues from phospholipids to OPGs. Thus, this observation supports the conclusion that OPG substitution could play a role in bacterial virulence. E. chrysanthemi opgG and opgH mutants were recently obtained by cassette insertion into the chromosome (15). These mutants, which are defective in OPG synthesis, exhibit a pleiotropic phenotype and a complete loss of virulence (15). In an attempt to isolate mutants defective in OPG substitution, we screened several collections of Tn5 insertions in the strain 3937, as previously described for E. coli (10). However, despite several attempts, we were unable to obtain such mutants.

A large fraction of the OPGs synthesized by E. chrysanthemi were recovered in the supernatant of centrifuged cultures when the cultures were collected in the stationary phase after growth at low osmolarity. However, it was not possible to determine the fraction of OPGs released (or secreted) during growth and the fraction released during the separation procedure. OPG detection in the medium is not possible without cell separation, and whatever the treatment, if one considers that outer membrane stability may be weakened under certain circumstances, shearing forces could release OPGs from the periplasmic space. No similar phenomenon was observed with E. coli cultures, but E. chrysanthemi envelope properties are clearly different, since, for example, OPG-defective mutants exhibit bile salt hypersensitivity, while similar mutants of E. coli have normal bile salt resistance (15). Similarly, strain T83 of Erwinia amylovora released a tightly complexed mixture of oligosaccharides and lipopolysaccharide in place of extracellular polysaccharide (20). The oligosaccharides, being branched structures with an average of 11 -1,2- and -1,6-linked glucose units, are most probably the OPGs produced by this particular strain. These observations open the questions of the actual organization of extracellular OPGs and of their role in pathogenesis. Membrane vesicles containing outer membrane protein, lipopolysaccharide, phospholipids, and periplasmic constituents are constantly extruded from the surface of a number of gram-negative bacteria (for a recent review, see Beveridge [2]). It was possible that OPGs released in the medium by E. chrysanthemi were included in such membrane vesicles. Nevertheless, we were unable to separate any fraction of the released OPGs when culture supernatants were further ultracentrifuged (100,000 × g, 3 h; data not shown). Thus, released OPGs are probably in a soluble form.

Growth conditions in plant tissues are certainly very different from those in a liquid low-osmolarity medium on a rotary shaker. Thus, any extrapolation should be taken with caution. However, if OPGs are released during the infection of plants by E. chrysanthemi, they could play a fundamental role in pathogenesis. To test this possibility, potato tubers were inoculated with a mixture of Opg⁺ and Opg bacteria (15), and growth of the different bacteria was compared. E. chrysanthemi strains defective in OPG synthesis were unable to grow in planta, and growth could not be restored by the OPGs eventually provided by the neighboring wild-type bacteria. OPGs must be present in the periplasmic space of E. chrysanthemi to allow its growth in the plant host.