The type and yield of lipopolysaccharide from symbiotically deficient Rhizobium lipopolysaccharide mutants vary depending on the extraction method

Brent L. Ridley, Benjamin S. Jeyaretnam and Russell W. Carlson1

Complex Carbohydrate Research Center, University of Georgia, 220 Riverbend Road, Athens, GA 3060 2, USA

Received on February 28, 2000; revised on April 26, 2000; accepted on April 27, 2000.


    Abstract
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
At least 18 lipopolysaccharide (LPS) extraction methods are available, and no single method is universally applicable. Here, the LPSs from four R.etli, one R.leguminosarum bv. trifolii mutant, 24AR, and the R.etli parent strain, CE3, were isolated by hot phenol/water ({phi}/W), and phenol/EDTA/triethylamine ({phi}/EDTA/TEA) extraction. The LPS in various preparations was quantified, analyzed by deoxycholate polyacrylamide gel electrophoresis (DOC-PAGE), and by immunoblotting. These rhizobia normally have two prominent LPS forms: LPS I, which has O-polysaccharide, and LPS II, which has none. The LPS forms obtained depend on the method of extraction and vary depending on the mutant that is extracted. Both methods extract LPS I and LPS II from CE3. The {phi}/EDTA/TEA, but not the {phi}/W, method extracts LPS I from mutants CE358 and CE359. Conversely, the {phi}/W but not the {phi}/EDTA/TEA method extracts CE359 LPS V, an LPS form with a truncated O-polysaccharide. {phi}/EDTA/TEA extraction of mutant CE406 gives good yields of LPS I and II, while {phi}/W extraction gives very small amounts of LPS I. The LPS yield from all the   strains using {phi}/EDTA/TEA extraction is fairly consistent (3-fold range), while the yields from {phi}/W extraction are highly variable (850-fold range). The {phi}/EDTA/TEA method extracts LPS I and LPS II from mutant 24AR, but the {phi}/W method partitions LPS II exclusively into the phenol phase, making its recovery difficult. Overall, {phi}/EDTA/TEA extraction yields more forms of LPS from the mutants and provides a simpler, faster, and less hazardous alternative to {phi}/W extraction. Nevertheless, it is concluded that careful analysis of any LPS mutant requires the use of more than one extraction method.

Key words: lipopolysaccharide/LPS/Rhizobium etli/Rhizobium leguminosarum bv. trifolii/mutants/extraction method/triethylamine/EDTA


    Introduction
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Endosymbiotic rhizobia participate in a complex developmental process with their host plants to produce nitrogen fixing root nodules (Sprent and Sprent, 1990Go; Hadri et al., 1998Go). Nodulating rhizobia enter the root in tube-shaped structures, called infection threads, and are conducted through the root tissue to the nodule meristem. At the nodule meristems the bacteria are enclosed in membrane from the host as they enter the plant cytoplasm in a process resembling endocytosis. Once in the host cytoplasm rhizobia develop into bacteroids which can fix nitrogen.

Nodulation is an intricately orchestrated process in which particular Rhizobium species can only nodulate particular host plants, suggesting that the symbiotic partners participate in many interorganismal recognition and signaling events (Spaink et al., 1998Go; Sprent and Sprent, 1990Go; Hirsch, 1992Go). Molecules located on the cell surface, such as the bacterial lipopolysaccharide (LPS), are particularly important in the nodulation process due to their proximity to the symbiotic partner’s cell surface and are likely to function in host–symbiont recognition or as signal molecules (Becker and Pühler, 1998Go; Kannenberg et al., 1998Go).

Two major lines of evidence support the idea that LPS has a role in the development of nitrogen fixing nodules. Studies using LPS recognizing monoclonal antibodies (mAbs) to study the process of nodule development provide the first line of evidence. These studies have shown that R.etli and R.leguminosarum bv. viciae LPS epitopes change during nodulation in a manner coordinated with nodule development (VandenBosch et al., 1989Go; Kannenberg et al., 1992Go, 1994; Tao et al., 1992Go). Furthermore, the nodulation and symbiotic nitrogen fixation processes are perturbed in another R.etli mutant which has a full length O-polysaccharide that does not undergo the normal nodulation induced O-polysaccharide epitope changes (Tao et al., 1992Go). This suggests that changes in the O-polysaccharide structure are needed for normal development of nitrogen-fixing nodules. Secondly, mutants of several Rhizobium, Bradyrhizobium, and Sinorhizobium species, which lack or have a modified O-polysaccharide, are symbiotically deficient (Kannenberg et al., 1998Go). In some cases, these mutant bacteria do not induce nodules on their host plants and in other cases they induce the formation of morphologically defective nodules which fix little or no nitrogen (VandenBosh et al., 1985Go; Noel et al., 1986Go; Perotto et al., 1994Go). For example, three R.etli strains with LPS mutations used in this study (CE166, CE358, and CE359) are able to induce nodule development in their host, black bean, but are almost entirely excluded from the nodule tissue (Noel et al., 1986Go). In the case of O-polysaccharide mutants of R.leguminosarum bv. viciae (a symbiont of pea), the normal nodulation process degenerates at both the infection thread and the bacteroid formation stages with what seems to be the induction of a host plant defense reaction (Perotto et al., 1994Go). This suggests that there may be a breakdown in the recognition process which causes the plant to recognize the mutant bacterium as a pathogen rather than as a symbiotic partner.

Efforts to characterize the LPS of symbiotically deficient R.etli mutants suggested that the results obtained could be affected by the extraction method used. To further investigate this phenomenon, LPS extracts obtained by two different methods from four R.etli LPS defective mutants (CE166, CE358, CE359, and CE406), the R.etli parent strain (CE3) and a R. leguminosarum bv. trifolii LPS mutant (AR24) were compared. The normal LPS structure from R.etli CE3 is one of the most extensively characterized of the Rhizobiacae (Nikaido and Vaara, 1987Go; Bhat and Carlson, 1992Go; Bhat et al., 1994Go; Forsberg and Carlson, 1998Go; Kannenberg et al., 1998Go; Forsberg et al., 2000Go, Que et al., 2000aGo, b). Like the LPS of enteric bacteria, CE3 LPS is composed of four regions; lipid A, the inner core oligosaccharide, the outer core oligosaccharide, and an O-polysaccharide with five trisaccharide repeating units (Figure 1). The wild type LPS from R.etli contains two major classes of molecules (Carlson et al., 1987Go; Kannenberg et al., 1998Go). One class, which is seen as a major fast migrating electrophoretic band (LPS II), has only lipid A and the inner core oligosaccharide; and the other class, which is seen as a major slow migrating electrophoretic band (LPS I), has all four regions.



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Fig. 1. The structure of R.etli LPS. Structural microheterogeneity occurs in this molecule due to alterations in the methylation of the O-polysaccharide fucosyl residues, and due to alterations in the lipid A fatty acylation profile.

 
The normal lipid A and core regions from R. leguminosarum bv. trifolii and R.etli have the same carbohydrate structure (Figure 1; Jeyaretnam, 1998Go; Kannenberg et al., 1998Go). However, the R.leguminosarum bv. trifolii mutant, 24AR produces a defective lipid A structure (called lipid-A24AR) in which the GalA residue is not added, and the reducing terminal GlcN is not reduced to form GlcN-onate (Figure 1; Russa et al., 1985Go; Jeyaretnam and Carlson, unpublished observations). In addition, lipid-A24AR retains the bisphosphate groups normally found only in lipid A precursor molecules since the mutant lacks the unique rhizobial phosphatase that removes these groups (Price et al., 1995Go). Carbohydrate composition and HPAEC analyses of 24AR LPS show that the core region of this mutant is also different from its parental strain (Jeyaretnam, 1998Go). R.leguminosarum bv. trifolii 24AR is able to add an O-polysaccharide to the defective lipid A- core structure LPS to produce ample LPS I (Jeyaretnam, 1998Go). However, the O-polysaccharide added differs from parental strain in its carbohydrate composition (Russa et al., 1985Go; Jeyaretnam, 1998Go).

The LPS of R.etli CE 3 consists of a heterogeneous pool of molecules which are variations of the basic structure (Bhat and Carlson, 1992Go; Forsberg and Carlson, 1998Go; Kannenberg et al., 1998Go). The bacteria can vary the composition of this pool in response to changes in environmental and physiological conditions. CE3 lipid A can vary due to changes in fatty acylation (Kannenberg and Carlson, unpublished observations), and the CE3 O-polysaccharide can vary due to changes in the number of repeating units, omission of glycosyl residues, and addition or loss of modifying groups including O-methyl and O-acetyl groups (Forsberg et al., 2000Go). The structural heterogeneity R.leguminosarum bv. trifolii LPS has not been investigated in detail.

Structural studies of the LPS produced by symbiotically deficient LPS mutants have contributed to our understanding of the features of LPS molecules which are important for successful symbiosis (Forsberg and Carlson, 1998Go; Carlson et al., 1995Go; Carlson et al., 1989Go). Nevertheless, the exact LPS structural features which are important in symbiosis have not been identified (Kannenberg et al., 1998Go). It is important that the LPS samples used for such studies contain a complete array of structures. This is a difficult task since different methods of extraction often extract different forms of LPS from the same bacterial strain (Galanos et al., 1969Go; Darveau and Hancock, 1983Go; Poxton and Brown, 1986Go; Bahrani and Oliver, 1991Go; Helander et al., 1992aGo,b; Kawahara et al., 1992Go; Guillorit and Samson, 1993Go; Chandan et al., 1994Go; DeLahooke et al., 1995Go; Russa et al., 1995Go; Alves and Lemos, 1996Go; Onoue et al., 1996Go; Valverde et al., 1997Go). In fact, all of the commonly used extraction procedures have, in some cases, selectively extracted particular forms of LPS (for examples, see Galanos et al., 1969Go; Darveau and Hancock, 1983Go; Helander et al., 1992aGo; Kannenberg et al., 1996Go; Onoue et al., 1996Go).

The structures, and, hence, the physical properties of LPS vary considerably between bacterial species, and are often heterogeneous within particular bacterial strains. Therefore, it should come as no surprise that at least eighteen different methods of LPS extraction have been reported to date, and that none of these methods has proven to be universally applicable (Ribi et al., 1959Go; Gray and Wilkinson, 1965Go; Morgan, 1965Go; Lieve, 1965Go; Staub, 1965Go; Westphal and Jann, 1965Go; Adams, 1967Go; Galanos et al., 1969Go; Johnson and Perry, 1976Go; Digat, 1978Go; Brade and Galanos, 1982Go; Darveau and Hancock, 1983Go; Hitchcock and Brown, 1983Go; Hoffmann and Houle, 1986Go; Uchida and Mizushima, 1987Go; Blais and Yamazaki, 1989Go; Domenico et al., 1989Go; Wood et al., 1989Go; Pettijean et al., 1990Go; Sugiyama et al., 1990Go; Eidhin and Mouton, 1993Go; Guillorit and Samson, 1993Go; Newton and Tiche, 1993Go; Apicella et al., 1994Go; Chandan et al., 1994Go; Roche et al., 1994Go; DeLahooke et al., 1995Go; Alves and Lemos, 1996Go; Nurminen and Vaara, 1996Go; Valverde et al., 1997Go). The wide variety of reagents that have been used successfully for LPS extraction is further evidence that the physical properties of the LPS from the different strains are quite variable. The reagents used include various solvents (e.g., light petroleum ether, methanol, chloroform, phenol, dimethylsulfoxide, diethylene glycol, triethylamine, and water), surfactants (e.g., sodium dodecylsulfate, Triton X-100, Zwittergent, and triethylamine), salts (e.g., NaCl, and sodium citrate), acids or bases (e.g., trichloroacetic acid, and NaOH), chelating agents (e.g., EDTA) and enzymes (e.g., DNase, RNase, proteinase, and lysozyme).

In this article a procedure is described which extracts the LPS from R.etli mutants more completely than the traditional hot phenol/water procedure ({phi}/W). This procedure (the {phi}/EDTA/TEA method) uses an extraction buffer with a single aqueous phase containing, phenol, EDTA, and triethylamine. The LPS in each R.etli extract was analyzed by DOC-PAGE with Alcian-blue silver staining and quantified by gas chromatography–mass spectrometric (GC–MS) determination of ß-hydoxymyristate, a fatty acid specific to LPS. As an alternative quantification method the LPS from R.leguminosarum bv. trifolii strain 24AR was labeled in vivo with 32P prior to extraction. The labeled LPS could be quantified by liquid scintillation counting and analyzed by DOC-PAGE with autoradiographic visualization. The LPS extracts from R.etli CE3 and three of the R.etli mutants were also characterized using mAbs that recognize epitopes which change during nodule development (Kannenberg et al., 1994Go). The {phi}/EDTA/TEA procedure is simpler, faster, and less hazardous than the traditional {phi}/W procedure, and with the R.etli LPS mutants studied, produces higher yields of LPS typically containing a more complete array of LPS structures.


    Results
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Electrophoretic analysis
The LPS from the wild type R.etli strain CE3 and strains with LPS defects were extracted using the {phi}/W and the {phi}/EDTA/TEA methods and analyzed by DOC-PAGE with Alcian-blue silver staining. In order to determine which DOC-PAGE bands in the crude extracts represent LPS, samples of the {phi}/EDTA/TEA extracts were affinity purified using polymyxin B-agarose and analyzed by DOC-PAGE (Figure 2). Most of the crude extracts contained a fairly prominent band migrating faster than LPS II. This band is not attributed to LPS since it does not bind to polymyxin B (for example compare Figure 2, CE3 lane 3 with 4). Previous work suggests that this band may contain the cyclic ß-linked glucan commonly found in rhizobial species (Breedveld and Miller, 1994Go; Kim et al., 1996Go). All the extracts contain diffusely staining material down the length of the gel which is not attributed to LPS, since it is also removed by polymyxin B affinity chromatography.



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Fig. 2. DOC-PAGE analysis of the parental R.etli strain (CE3) and the LPS mutants. Each panel depicts an Alcian-blue silver stained gel of extracts from the strain indicated. Lanes 1 contain the phenol layers form the {phi}/W extracts, lanes 2 contain the water layers from the {phi}/W extracts, lanes 3 contain the {phi}/EDTA/TEA extracts, and Lanes 4 contain polymyxin-affinity purified samples of the {phi}/EDTA/TEA extracts. I, Position of LPS I; II, position of LPS II. The amounts of each sample analyzed were adjusted to give equivalent staining whenever this was practical. *, Position of the prominent band that does not bind to polymyxin and is thought represent a rhizobial cyclic ß-linked glucan.

 
Both the phenol and water layers of the CE3 {phi}/W extracts have the two prominent LPS bands (LPS I and LPS II), as well as a number of minor LPS bands found in the high molecular weight region of the gel (Figure 2, CE3 lanes 1 and 2). Previous work has shown that the prominent LPS I band of CE3 is composed of molecules with a complete LPS structure that has an O-polysaccharide with five repeating units (Carlson et al., 1987Go; Forsberg et al., 2000Go). Monoclonal antibody binding studies have revealed immunological heterogeneity within CE3 LPS I indicating that there is subtle structural variation in the O-chain polysaccharide (Tao et al., 1992Go). The minor bands in the high molecular weight region are thought to consist of LPS molecules with O-polysaccharides having a number of repeating units other than five, or with additional or missing glycosyl residues, O-methyl, or O-acetyl groups (Forsberg et al., 2000Go). The {phi}/EDTA/TEA and the {phi}/W methods both extract the same forms of LPS from CE3, but the {phi}/EDTA/TEA extract has a greater proportion of LPS II than either layer of the {phi}/W extract (Figure 2, CE3). It is also interesting to note that after polymyxin B-affinity chromatography of the {phi}/EDTA/TEA extract there is a significant reduction in LPS II relative to LPS I. In fact, all the polymyxin B-affinity purified {phi}/EDTA/TEA R.etli extracts examined have a reduced proportion of LPS II compared to the starting material. This result suggests that polymyxin B-agarose binds R.etli LPS I in preference to LPS II.

The LPS mutant CE358 has been reported to produce a truncated LPS which is missing a particular galacturonosyl residue in the inner core, the entire outer core, and the entire O-polysaccharide (Forsberg and Carlson, 1998Go). Failure to add the outer core and O-polysaccharide is thought to be due to the inability of the truncated precursor to act as an acceptor (Forsberg and Carlson, 1998Go). As expected, LPS I was not detected in either layer of the {phi}/W extract, or in the {phi}/EDTA/TEA extract (Figure 2, CE358, lanes 1–3). Therefore, it was surprising to see a fairly intense band with the same electrophoretic mobility as CE3 LPS I in the {phi}/EDTA/TEA extract after purification by polymyxin B-agarose affinity chromatography (Figure 2, CE358, lane 4). It was subsequently discovered that a faint CE358 LPS I band can be detected by DOC-PAGE analysis of the crude {phi}/EDTA/TEA extract (i.e., prior to polymyxin B-agarose affinity purification) when an excess of sample is analyzed (data not shown). This result supports the previous conclusion that polymyxin B-affinity chromatography enriches for R.etli LPS I at the expense of LPS II. It is clear from these results that CE 358 does produce a small amount of LPS I despite the mutation which results in a defective LPS core structure.

The LPS mutant CE 359 has been reported to be defective in its ability to make an LPS I with a full-length O-polysaccharide (Forsberg and Carlson, 1998Go). In place of LPS I it has a significant amount of LPS V, which consists of lipid A, inner core, outer core, and a truncated O-polysaccharide containing only one oligosaccharide repeat unit (Tao et al., 1992Go; Forsberg and Carlson, 1998Go). A prominent band corresponding to LPS V is found in both layers of the CE359 {phi}/W extract, but not in the {phi}/EDTA/TEA extract (Figure 2 CE359, lanes 1, 2, and 3). Surprisingly, the CE359 {phi}/EDTA/TEA extract has an intense LPS I band with the same electrophoretic mobility as that of CE3 rather than the expected LPS V band (Figure 2 CE359, lanes 3 and 4). A faint LPS I band was observed in the water layer of CE359 {phi}/W extract, and a faint LPS V band was observed in the CE359 {phi}/EDTA/TEA extract when excess amounts of each were analyzed by DOC-PAGE (data not shown). It is clear from these data that, in the case of CE359, the {phi}/W method preferentially extracts LPS V whereas the {phi}/EDTA/TEA method preferentially extracts LPS I. This constitutes the only case in which the {phi}/W method efficiently extracted a form of LPS (LPS V) from the mutants that is not efficiently extracted by the {phi}/EDTA/TEA method.

Mutant CE166 was reported to make less LPS I, as well as a reduced total amount of LPS (LPS I plus LPS II) compared to the parent strain, CE3 (Cava et al., 1989Go). All of the CE166 extracts using either method have reduced amounts of LPS I relative to LPS II when compared to the equivalent CE3 extracts (Figure 2 compare CE166 with CE3). However, the {phi}/EDTA/TEA method extracted an abundance LPS II from CE166, suggesting that it produces about the same total amount of LPS as CE3 (Figure 2). This notion is supported by quantification of the LPS in the extracts since a nearly identical amount of total LPS is found in the {phi}/EDTA/TEA extracts from CE166 as from the parent strain, CE3 (Table I). The large LPS II to LPS I ratio in CE166 supports previous work by Cava et al. (1989)Go which shows that this mutant is partially defective in the addition of the outer core and O-polysaccharide.


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Table I. Yield of R.etli LPS
 
The mutant strain CE406 was isolated based on a change in the immunological properties of its LPS (the immunological features of this LPS mutant are discussed below). The {phi}/W extract of CE406 had no detectable LPS in the water layer, and a only trace of LPS I, but no detectable LPS II, in the phenol layer (Figure 2 CE406, lanes 1 and 2). In order to visualize any LPS, the phenol layer had to be analyzed using a concentration 100-fold higher than that of the {phi}/EDTA/TEA extract. In contrast, the {phi}/EDTA/TEA extract of CE406 contains an abundance of LPS consisting of both LPS I and II in a proportion similar to that of CE3 (Figure 2 CE406, lane 3). The {phi}/EDTA/TEA extract does have more non-LPS contaminating material (seen as a background smear) than the {phi}/EDTA/TEA extracts from the other strains (Figure 2 CE406, lane 3). However, it is clear that the {phi}/EDTA/TEA method is useful for extracting LPS from CE406 LPS, while the {phi}/W method is impractical due to low yields.

The results obtained when in vivo 32P-labeled extracts from R.leguminosarum bv. trifolii mutant 24AR were analyzed using DOC-PAGE and visualized by autoradiography were similar to those obtained with the R.etli extracts despite the use of a different rhizobial species and analysis method (Figure 3). The {phi}/W extract of 24AR had an abundance of LPS I but no detectable LPS II in the water layer, whereas both LPS I and LPS II were found in the phenol layer (Figure 3, lanes 1 and 2). The {phi}/EDTA/TEA extraction method recovered both LPS I and LPS II in a proportion similar to that of the phenol layer of the {phi}/W extract (Figure 3, lane 3). The {phi}/EDTA/TEA method provides an attractive alternative to {phi}/W extraction of LPS II from 24AR, since the recovery of LPS from phenol solutions is particularly laborious (see Materials and Methods).



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Fig. 3. Autoradiogram of 32P-labeled LPS from mutant 24AR subsequent to PAGE. Lane 1 contain the phenol layers form the {phi}W extracts, lane 2 contain the water layers from the {phi}/W extracts, lane 3 contain the {phi}/EDTA/TEA extracts, and lanes 4 contain polymyxin-affinity purified samples of the {phi}/EDTA/TEA extracts. The amounts of each sample analyzed were adjusted to give equivalent staining whenever this was practical.

 
Immunological characterization of LPS extracts
The immunological properties of the LPS in {phi}/W and {phi}/EDTA/TEA extracts from CE3, CE3 grown at pH 5, and three of the four LPS mutants discussed above (CE358, CE359, and CE406) were investigated. This investigation was initiated because DOC-PAGE analysis indicated that the {phi}/EDTA/TEA method extracted novel forms of LPS from the R.etli mutants. It was possible that these novel LPS forms might also have altered immunological properties. Three mAbs (JIM27, JIM28, and JIM29) which recognize the R.etli CE3 O-polysaccharide (Tao et al., 1992Go) were used for the DOC-PAGE immunoblot analysis. All of the immunological results were confirmed by the analysis of two independent extracts (one preparative scale and one analytical scale) from the same bacterial culture.

An earlier study using the SDS extraction procedure of Hitchcock and Brown (1983)Go showed that R.etli CE3 LPS I changes its expression of the JIM28 and JIM29 epitopes in response to changes in physiological conditions, while the expression of the JIM27 epitope remains unchanged (Tao et al., 1992Go). For example, CE3 expresses the JIM27, JIM28, and JIM29 epitopes after growth on rich media at the normal oxygen level, but expresses only the JIM27 and JIM29 epitopes after growth at a reduced oxygen concentration, and only the JIM27 epitope after growth at pH 5 and normal oxygen level. The CE3 LPS I from bacteroids has unaltered expression of the JIM27 epitope and diminished expression of the JIM28 and JIM29 epitopes (Tao et al., 1992Go). All three antibodies bind to either {phi}/W (data not shown) or {phi}/EDTA/TEA (Figure 4) extracted LPS I from R.etli CE3 grown on rich media at the normal oxygen level as expected. Furthermore, the LPS I in {phi}/EDTA/TEA extracts of CE3 grown at pH 5 binds to JIM27, but not to JIM28 or JIM29 as expected (Figure 4).



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Fig. 4. Immunoblot analysis of the {phi}/EDTA/TEA extracts from the parental R.etli strain (CE3) and the LPS mutants. Only the LPS I region is shown since that is the only region which binds the monoclonal antibodies used in this experiment. The panel labeled "stained gel" represents an Alcian-blue silver stained DOC-PAGE gel of the R.etli strains indicated. The remaining panels represent immunoblots of similar gels. The label above each of the panels denotes the mAbs each blot was probed with. The label above each lane indicates the sample analyzed. Each of the gels used for the immunoblots had the same amounts of the samples as the stained gel shown.

 
The forms of LPS I found in the polymyxin B-purified {phi}/EDTA/TEA extracts of CE358 and CE359 grown in rich media at the normal oxygen level are antigenically different from CE3 grown under the same conditions although they have the same electrophoretic mobility (Figure 4). The LPS I from CE358 binds JIM27 weakly, does not bind JIM28, and binds JIM29 normally (Figure 4). This LPS I constitutes the first instance where the binding of JIM27 to a form of R.etli LPS I is diminished but the binding of JIM29 is retained (Tao et al., 1992Go). The LPS I from CE359 binds JIM27 and JIM29, but does not bind JIM28. These results show that forms of LPS I being made by CE358 and CE359 are antigenically different from CE3 LPS I, and that the subtle structural modifications that lead to these antigenic differences are correlated with altered {phi}/W extraction properties (discussed in Electrophoretic analysis).

Mutant CE406 was isolated on the basis of its inability to bind JIM28 when grown in rich media under standard conditions. The LPS from CE406 cells grown under these conditions was isolated using the {phi}/EDTA/TEA method and analyzed by DOC-PAGE, and immunoblotting (Figure 4). Under these conditions CE406 produces LPS I with a normal sized O-polysaccharide which binds JIM27 and JIM29, but binds JIM28 very weakly. Therefore, the CE406 mutation has resulted in a relatively minor change in the structure of the O-polysaccharide affecting the binding of JIM28, but not affecting on the overall size or the binding of JIM27 and JIM29. This mutation also resulted in an LPS with dramatic changes in its extraction properties, i.e., it was not extracted by the {phi}/W method.

Yield and purity of LPS
The yield of LPS in all the extracts from CE3 and the four LPS mutants (Table I) was determined based on the ß-hydoxymyristate content (determined by GC-MS of its trimethysilyl (TMS) methylester derivative). Quantification of ß-hydoxymyristate provides a convenient means of estimating the amount of LPS in crude extracts since it is found constitutively in LPS and not in other known rhizobial glycolipids (Uchida and Mizushima, 1987Go; Bhat et al., 1994Go). Furthermore, quantitative determination of ß-hydroxymyristate is unaffected by the presence of the contaminating molecules found in these crude extracts. However, the results could be affected if there is variation in the number of ß-hydroxymyristate moieties per LPS molecule. There are typically three ß-hydroxymyristate moieties per R.etli LPS molecule, although some LPS molecules may have four, while others may have two (Forsberg et al., unpublished observations).

The total amount LPS extracted from CE3 by the conventional {phi}/W method was about two-fold more than that extracted by the {phi}/EDTA/TEA method. However, the {phi}/EDTA/TEA method was significantly more effective for extraction of the R.etli LPS mutants (Table I). Compared to the {phi}/W method, the {phi}/EDTA/TEA method extracted about 11-fold more LPS from CE358, 4-fold more LPS from CE359, 12-fold more LPS from CE166, and 270-fold more LPS from CE406. In addition, carbohydrate composition analysis of the crude R.etli {phi}/EDTA/TEA extracts did not detect ribose, while the {phi}/W extracts had a substantial amount of this residue (data not shown). This shows that the R.etli {phi}/EDTA/TEA LPS extracts are substantially free of RNA contamination.

It is also significant that the yield from all the R.etli strains was relatively constant using the {phi}/EDTA/TEA method (varying in a 3-fold range) compared to the {phi}/W method where the yield was highly variable (850-fold range). The CE406 mutant constitutes the most dramatic example of this variation. The LPS yield from this mutant was about 1.7-fold less than that for CE3 when the {phi}/EDTA/TEA method was used, but at least 680-fold less when the {phi}/W method was used (Table I). Mutant CE406 was also unusual in that no LPS was detected in the {phi}/W water layer, while the LPS from CE3, and the other mutants extracted mostly into the water layer. For example, 4-fold more LPS was found in the water than in the phenol layer of CE3 and about 8-fold more in the water layer of CE358 (Table I).

Liquid scintillation counting was used to determine the amount of LPS in the in vivo 32P-labeled extracts from R. leguminosarum bv. trifolii mutant AR24 (Table II). Less than 15% of radioactive material in the {phi}/EDTA/TEA extract was nuclease sensitive, whereas about 60% of the radioactive material in the water layer of the {phi}/W extract was nuclease sensitive (Table II). This demonstrates that the {phi}/W method extracts more contaminating nucleic acids than does the {phi}/EDTA/TEA method. Polymyxin-B affinity purification of the water layer of the {phi}/W extract or the {phi}/EDTA/TEA extract, in either case, removed about 20% of the radioactivity remaining after nuclease treatment (Table II). The amount of LPS in the phenol layer of the {phi}/W extract was not quantified by scintillation counting, but was estimated to be roughly one-fifth of the amount found in the water layer, based on the autoradiography of the DOC-PAGE gels. Therefore, the two methods extract similar amount of LPS form this strain.


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Table II. Yield of R.leguminosarum bv. trifolii 24AR LPS
 

    Discussion
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Extraction of the Rhizobium LPS mutants
It is clear from the electrophoretic, immunological, LPS quantification, and carbohydrate composition data that the {phi}/EDTA/TEA method yields a more complete array of reasonably pure LPS structures in greater abundance than the {phi}/W method. The yield of LPS obtained from each of the R.etli LPS mutants using the {phi}/EDTA/TEA method was similar to that obtained from the parental strain, whereas {phi}/W extraction of the mutants gave poor yields. On the other hand, the LPS yield from the R.leguminosarum bv. trifolii AR24 mutant was comparable using the two methods.

In the case of two of the mutants, CE359 and CE406, novel forms of LPS I were extracted using the {phi}/EDTA/TEA method but were poorly extracted by the {phi}/W method. Interestingly, these novel LPS I molecules had the same mAb binding pattern (both bind JIM27 and JIM29 but not JIM28), and this pattern differs from that of the parental strain, CE3 (LPS I binds all three mAbs). This result suggests that the O-polysaccharide structures of LPS I from CE359 and CE406 are altered in similar ways, and that the altered immunological and extraction properties are correlated. On the other hand, CE406 LPS II is poorly extracted using the {phi}/W method while CE359 LPS II is efficiently extracted. This difference suggests that the CE406 mutation causes a structural alteration in LPS II that is not found in CE359.

The LPSs from R.etli CE3 and R. leguminosarum bv. trifolii AR24 are found in both the water and phenol layer after {phi}/W extraction indicating that they are moderately hydrophobic. The hydrophobic character of R.etli CE3 LPS is probably due in part to the structure of its O-polysaccharide. The repeating unit of the CE3 O-polysaccharide contains the hydrophobic deoxyglycosyl residues, fucose and 3-methyl-6-deoxytalose (Forsberg et al., 2000Go; Bhat and Carlson, 1992Go). However, the CE3 O-polysaccharide repeating unit also contains the hydrophilic acidic glycosyl residue GlcA. Surprisingly, this O-polysaccharide, after being cleaved from the lipid A and core moieties, does not bind to anion exchange resin as expected for an acidic polysaccharide (Kannenberg et al., 1998Go). It is thought that the lack of binding may result from methyl esterification of the GlcA carboxyl groups which would increase the hydrophobicity of the O-polysaccharide (Forsberg et al., 2000Go). In any case, the hydrophobicity of CE3 LPS is at a threshold where it can partition into either the phenol or water layers during {phi}/W extractions. Therefore, it seems plausible that mutations causing relatively small changes in LPS structure could have dramatic effects on the outcome of {phi}/W extraction. This may explain why the {phi}/W extraction properties of the LPS I from the CE359 and CE406 mutants are so different from those of the parental strain. An alternative explanation is that a particular LPS mutant could be affected in other, non-LPS, bacterial surface components which alter the extraction properties of the LPS.

The phenol/triethylamine/EDTA ({phi}/EDTA/TEA) extraction method.
The {phi}/EDTA/TEA method employs an aqueous solution of 5% phenol, EDTA, and triethylamine to extract LPS on a preparative scale. Valverde et al. (1997)Go developed a similar method for analytical scale LPS extraction in which the bacterial cells are extracted with an aqueous solution EDTA/triethylamine followed by purification using polymyxin B-agarose affinity chromatography. The {phi}/EDTA/TEA method can also be used effectively on an analytical scale. The volume of {phi}/EDTA/TEA buffer added is simply reduced in proportion to the amount of bacterial cells to be extracted.

Like the {phi}/EDTA/TEA method, several previously used LPS extraction methods rely on reagents which disrupt the ionic, polar, and non-polar interactions that hold the LPS in the outer membrane, thereby releasing it in a form that is soluble in aqueous solution (Hitchcock and Brown, 1983Go; Uchida and Mizushima, 1987Go; Domenico et al., 1989Go; Wood et al., 1989Go; Pettijean et al., 1990Go; Apicella et al., 1994Go; DeLahooke et al., 1995Go; Valverde et al., 1997Go). These methods normally combine a chelating agent such as EDTA to disrupt ionic interactions involving divalent cations, and a surfactant such triethylamine to disrupt nonpolar interactions. The cationic surfactant used in the {phi}/EDTA/TEA method, triethylamine is particularly useful since it forms highly soluble salt complexes with LPS (Galanos and Lüderitz, 1975Go; Carlson et al., 1978Go). This surfactant has an added advantage in that it is easier to remove by dialysis than detergents such as sodium dodecylsulfate (SDS) or Triton-X-100. A small percentage of phenol (5%) is also used in the {phi}/EDTA/TEA extraction buffer since this reagent is known to disrupt LPS-protein interactions (Westphal and Jann, 1965Go; Alves and Lemos, 1996Go).

Aqueous solutions of EDTA can extract the LPS from the outer membranes of bacteria without cell rupture, avoiding the release of unwanted nucleic acids and proteins (Lieve, 1965Go). The {phi}/EDTA/TEA method uses a moderately hypertonic solution of EDTA and TEA in order to avoid osmotic rupture of the bacterial cells. The relatively low level of nucleic acid contamination in both the R.etli and R.leguminosarum bv. trifolii 24AR {phi}/EDTA/TEA extracts suggests that this method does not cause extensive cell rupture.

Unlike the {phi}/EDTA/TEA method, some established methods of LPS extraction rely on dissolution in organic solvents or on partition between water and organic solvent phases (Ribi et al., 1959Go; Morgan, 1965Go; Staub, 1965Go; Westphal and Jann, 1965Go; Galanos et al., 1969Go; Brade and Galanos, 1982Go; Hoffmann and Houle, 1986Go; Nurminen and Vaara, 1996Go). Organic solvent-based methods are inherently selective, since they separate the LPS from other macromolecules exclusively based on its solubility in particular organic solvent systems. The drawback of these methods is that different forms of LPS have different solubility properties. Of the preparative scale LPS extraction methods available, the hot phenol/water method ({phi}/W) of Westphal and Jann (1965)Go and the phenol/chloroform/petroleum ether method of Galanos et al. (1969)Go are the most commonly used. Both of these organic solvent based methods have, on a number of occasions, resulted in poor yields, excessive contamination by non-LPS molecules, or extracted some forms of LPS and excluded others (Galanos et al., 1969Go; Darveau and Hancock, 1983Go; Poxton and Brown, 1986Go; Uchida and Mizushima, 1987Go; Pettijean et al., 1990Go; Bahrani and Oliver, 1991Go; Helander et al., 1992aGo; Kawahara et al., 1992Go; Chandan et al., 1994Go; DeLahooke et al., 1995Go; Russa et al., 1995Go; Alves and Lemos, 1996Go; Kannenberg et al., 1996Go; Onoue et al., 1996Go; Valverde et al., 1997Go; this work). These methods have the additional disadvantage in that they involve exposure to concentrated phenol at elevated temperatures, which may facilitate the removal of fatty acids attached by liable ester linkages (Westphal and Jann, 1965Go; Tsang et al., 1974Go; Okuda et al., 1975Go; Brade and Galanos, 1982Go).

Initially, it was thought that the hydrophobic phenol/chloroform/petroleum ether method was selective for truncated forms of LPS (rough forms) without O-polysaccharide and the hydrophilic hot phenol/water method was more selective for forms of LPS (smooth forms) having the O-polysaccharide (Galanos et al., 1969Go). However, some O-polysaccharides, including that of R.etli, contain an abundance of hydrophobic deoxyglycosyl residues or other substituents which lead to increased hydrophobicity. Smooth forms of LPS with hydrophobic O-polysaccharides often extract into the phenol layer during {phi}/W extraction, and can be extracted with the phenol/chloroform/petroleum ether method (Galanos et al., 1969Go; Poxton and Brown, 1986Go; Uchida and Mizushima, 1987Go; Carrion et al., 1990Go; Pettijean et al., 1990Go; Kawahara et al., 1992Go; DeLahooke et al., 1995Go; Russa et al., 1995Go; Onoue et al., 1996Go).

Accurate elucidation of the immunological and structural effects of an LPS mutation depends on the ability to extract a complete array of LPS forms. In general the {phi}/EDTA/TEA method extracts a more complete array of LPS forms from the R.etli LPS mutants than does the {phi}/W method. However, the method extracts one form of LPS (CE 359, LPS V) much less efficiently than the {phi}/W method does. This result is in accord with the findings discussed above for the other LPS extraction methods. Any of the currently available LPS extraction procedures can miss particular forms of LPS. The results presented in this paper also demonstrate that it is very difficult to know when a complete array of LPS forms has been obtained. Therefore, more than one LPS extraction method should be employed when the characterization of a complete array of LPS forms is desired.


    Materials and methods
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Bacterial strains and culture
Rhizobium etli strains CE3, CE166, CE358, and CE359 were gifts from K. D. Noel and have been previously characterized (Noel et al., 1984Go; Cava et al., 1989Go, 1990; Tao et al., 1992Go). The R.etli strains were grown at normal or low pH, and at normal or low oxygen as described previously (Tao et al., 1992Go). Parental strain CE3 is a spontaneous streptomycin resistant isolate of the wild type R.etli strain, CFN42. All of the above mentioned R.etli LPS mutants were derived from CE3 by Tn5 mutagenesis (Simon et al., 1983Go; Cava et al., 1989Go, 1990; Carroll and Mathews, 1990Go; Tao et al., 1992Go). The LPS mutant CE406 was isolated by RLB in the laboratory of K.D.Noel at Marquette University, Milwaukee, WI, using Tn5 mutagenesis with the E.coli donor strain S17-1 carrying the broad host range transposon mutagenesis vector, pSUP2021 (Simon et al., 1983Go). The CE406 mutant was selected based on its lack of mAb JIM28 epitope expression, which is normally expressed on the O-polysaccharide during growth on TY media with normal oxygen concentration (Tao et al., 1992Go). Rhizobium leguminosarum bv. trifolii mutant, 24AR, is a spontaneous non-nodulating mutant lacking the symbiotic plasmid (Russa et al., 1985Go; Kowalczuk et al., 1981Go) and was obtained from R. Russa (Curie-Sklodoeska University, Lublin, Poland). This strain was grown at 29°C in liquid medium (79 CA) containing mannitol, yeast extract, casamino acids, and salts (Kowalczuk and Lorkienwicz, 1979Go).

Extraction and purification of R.etli LPS
R.etli cultures were grow at high oxygen levels in TY media (Tao et al., 1992Go) to late log phase in a 100 l fermenter at The University of Georgia fermentation plant. Typically 100 g wet weight of cells was used for preparative scale extraction and 3 g were used for analytical scale extraction.

When the {phi}/EDTA/TEA method was used the cell pellet was suspended in {phi}/EDTA/TEA extraction buffer (20% wet weight cells/buffer volume; 0.25 M EDTA, 5% phenol, pH adjusted to 6.9 with triethylamine), stirred at 37°C for 1 h, and then centrifuged at 10,000 x g for 1 h. The supernatant was dialyzed exhaustively (2,000 MWCO Spectra/Por 7 tubing, 3 days against running tap water, followed by 1 day against de-ionized water), concentrated by rotary evaporation, clarified by centrifugation (5000 x g, 20 min), and stored at –80°C.

For the {phi}/W method, the cell pellet was suspended in 45% aqueous phenol (20% wet weight cells/volume phenol solution) and, stirred at 65–70°C for 30 min. The water and phenol layers were separated by centrifugation (10,000 x g, 1 h at 10°C). The water phase was dialyzed, concentrated, clarified by centrifugation, and stored as described for the {phi}/EDTA/TEA method. The phenol phase was dialyzed in 2000 MWCO tubing against running tap water until all the liquid phenol was gone (about 5 days). The solid residue, which remained after dialysis, was removed by centrifugation (5000 x g, 20 min). The supernatant was dialyzed, concentrated, clarified, and stored as described for the {phi}/EDTA/TEA method. Sodium azide (0.02%) was added to all the extracts to inhibit microbial growth.

LPS extracts were affinity purified using polymyxin B-agarose as described by Forsberg and Carlson (1998)Go. Polymyxin B specifically binds to lipid A with a high affinity. The procedure was adapted for use with a 100 ml bed volume column by increasing the flow rate and elution volumes proportionally.

Electrophoresis and immunoblotting
Electrophoresis and Alcian-blue silver staining were performed according to the method of Reuhs et al. (1998)Go. Immunoblotting was preformed according to the method of Reuhs et al. (1998)Go with some modifications. Briefly, LPS containing gels were soaked (3 x 5 min.) in transfer buffer (48 mM Tris, 39 mM glycine, 20% methanol) and electrophoretically transferred to a Nytran Plus membrane (Schleicher & Schuell) using a Bio-Rad Transblot SD semi-dry transfer cell (10 V for 20 min.). The membranes were equilibrated for 15 min in TBS (0.2M NaCl, 20mM Tris pH 7.4), blocked (1 h in 5% nonfat dry milk added to the TBS), and then incubated with one of the primary mAbs (JIM27, JIM28 or JIM29 overnight in the blocking solution). Next, the membranes were washed (5 x 10 min in TBS), and then incubated (3 h in the blocking solution) with the appropriate alkaline phosphatase–conjugated secondary antibodies (Sigma). Finally, the membranes were washed (5 x 5 min in TBS), equilibrated in substrate buffer (5 min in 10 ml of 0.1 M Tris, 0.1 M NaCl, 5 mM MgCl2 pH 9.5), and developed by the addition of 33 µl of 2-amino-2-methyl-1,3-propanediol (50 mg/ml in dimethylformamide) and 66 µl of nitroblue tetrazolium (50 mg/ml in 70% dimethylformamide).

Quantification and carbohydrate analysis of LPS from R.etli
The relative amounts of LPS in each of the crude extracts were determined by quantification of its ß-hydoxymyristate content (Bhat et al., 1991Go) normalized to the wet weight of the bacterial pellet. Heptadecanoic acid was added to each LPS sample as an internal standard prior to ß-hydoxymyristate quantification. Methylesters of the fatty acids in the samples were prepared by transesterification in methanolic HCl. After the samples were dried under N2, the fatty acyl methylesters were extracted with hexane. Trimethylsilyl ether derivatives of hydroxy fatty acyl methylesters were prepared using Tri-Sil reagent (Pierce Chemical Co.). After evaporation of the solvent and addition of hexane, the TMS fatty acid methyl esters were analyzed by GC-MS using a 30 m DB-1 capillary column (J&W Scientific). The amount of ß-hydoxymyristate in each sample was calculated from the ratio of its peak area to that of the heptadecanoic acid standard peak. The GC-MS responses of the TMS fatty acid methyl esters and that of the internal standard are known to be equivalent on a mass basis (Bhat et al., 1991Go).

The carbohydrate composition of the LPS extracts was analyzed to determine if ribose or other contaminating non-LPS carbohydrates were present. The carbohydrate composition of the LPS extracts was determined by GC-MS analysis of TMS methylglycosides according the method of York et al. (1985)Go.

Labeling, extraction, and purification of R.leguminosarum bv. trifolii 24AR LPS
Four 2 ml cultures of 24AR were grown in sterile glass tubes with [32P]-phosphoric acid (250 µCi/tube). Cells were harvested by centrifugation and washed twice with water. The washed cell pellet was re-suspended in 6 ml of water and divided equally into two portions. One portion was used to extract LPS by the {phi}/W method using 3 ml of cell suspension and 3 ml of phenol (90%) that was preheated to 65°C. The mixture was maintained at 65°C for 20 min. During this time the {phi}/W solution was stirred continuously. After brief cooling, the phenol and water phases were separated by centrifugation. Two milliliters of the upper aqueous phase was withdrawn, and the phenol phase was extracted again using an additional 2 ml of water. The aqueous phases were pooled and dialyzed against water. The second portion of the cell pellet was extracted using the {phi}/EDTA/TEA method as described above.

The resulting crude LPS preparations were treated with ribonuclease (1 mg/ml, 100 Kunitz units) and deoxyribonuclease (1 mg/ml, 1000 Kunitz units) (both from Sigma, St. Louis, MO), and then dialyzed against water. The nuclease-treated LPS preparations were further purified by polymyxin-B affinity chromatography as described by Forsberg and Carlson, 1998Go. Each R.leguminosarum bv. trifolii 24AR LPS preparation was analyzed by DOC-PAGE and visualized using a Molecular Dynamics PhosphorImager equipped with IMAGEQUANT software. The amount of 32P label in the LPS extracts was quantified by liquid scintillation counting.


    Acknowledgments
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
We thank Michael Harvey for technical assistance. We are grateful to John Kim, Dr. L.Scott Forsberg and Dr. Bradley Reuhs for helpful discussions. We are also grateful to Dr. L.Scott Forsberg for sharing unpublished results. We thank Dr. Debra Mohnen for critical reading of the manuscript. This work was supported by NIH grant GM 39583 to R.W.C. and DOE grant DE-FG09-93ER20097 to the Complex Carbohydrate Research Center.


    Abbreviations
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
BCIP, 5-bromo-4-chloro-3-indolyl phosphate; DMF, dimethylformamide; DOC-PAGE, deoxycholate polyacrylamide gel electrophoresis; Fuc, fucose; GC–MS, gas chromatography– mass spectrometry; Gal, galactose; GalA, galacturonic acid; GlcA, glucuronic acid; GlcN, glucosamine; GlcN-onate, 2-amino-2-deoxygluconic acid; HPAEC, high performance anion-exchange chromatographic; Kdo, 3-deoxy-D-manno-2-octulosonic acid; LPS, lipopolysaccahride; Man, mannose; 3Me-dTal, 3-methyl-6-deoxytalose; NBT, nitroblue tetrazolium; QuiNAc, N-acetylquinovosamine (2-N-acetamido-2,6-dideoxyglucose); 2,3,4-tri-Me-Fuc, 2,3,4-tri-O-methyl fucose; TMS, trimethysilyl; mAb, monoclonal antibody; {phi}/EDTA/TEA; phenol EDTA triethylamine extraction method; {phi}/W; phenol water extraction method.


    Footnotes
 
1 To whom correspondence should be addressed Back


    References
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
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