A Deacylase in Rhizobium leguminosarum Membranes That Cleaves the 3-O-Linked beta -Hydroxymyristoyl Moiety of Lipid A Precursors*

Shib Sankar Basu, Kimberly A. White, Nanette L. S. Que, and Christian R. H. RaetzDagger

From the Department of Biochemistry, Duke University Medical Center, Durham, North Carolina 27710

    ABSTRACT
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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Lipid A from the nitrogen-fixing bacterium Rhizobium leguminosarum displays many structural differences compared with lipid A of Escherichia coli. R. leguminosarum lipid A lacks the usual 1- and 4'-phosphate groups but is derivatized with a galacturonic acid substituent at position 4'. R. leguminosarum lipid A often contains an aminogluconic acid moiety in place of the proximal glucosamine 1-phosphate unit. Striking differences also exist in the secondary acyl chains attached to E. coli versus R. leguminosarum lipid A, specifically the presence of 27-hydroxyoctacosanoate and the absence of laurate and myristate in R. leguminosarum. Recently, we have found that lipid A isolated by pH 4.5 hydrolysis of R. leguminosarum cells is more heterogeneous than previously reported (Que, N. L. S., Basu, S. S., White, K. A., and Raetz, C. R. H. (1998) FASEB J. 12, A1284 (abstr.)). Lipid A species lacking the 3-O-linked beta -hydroxymyristoyl residue on the proximal unit contribute to this heterogeneity. We now describe a membrane-bound deacylase from R. leguminosarum that removes a single ester-linked beta -hydroxymyristoyl moiety from some lipid A precursors, including lipid X, lipid IVA, and (3-deoxy-D-manno-octulosonic acid)2-lipid IVA. The enzyme does not cleave E. coli lipid A or lipid A precursors containing an acyloxyacyl moiety on the distal glucosamine unit. The enzyme is not present in extracts of E. coli or Rhizobium meliloti, but it is readily demonstrable in membranes of Pseudomonas aeruginosa, which also contains a significant proportion of 3-O-deacylated lipid A species. Optimal reaction rates are seen between pH 5.5 and 6.5. The enzyme requires a nonionic detergent and divalent metal ions for activity. It cleaves the monosaccharide lipid X at about 5% the rate of lipid IVA and (3-deoxy-D-manno-octulosonic acid)2-lipid IVA. 1H NMR spectroscopy of the deacylase reaction product, generated with lipid IVA as the substrate, confirms unequivocally that the enzyme cleaves only the ester-linked beta -hydroxymyristoyl residue at the 3-position of the glucosamine disaccharide.

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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
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DISCUSSION
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Lipopolysaccharide, a macromolecular glycolipid found in the outer membranes of Gram-negative bacteria (1-6), is anchored to the outer leaflet of the outer membrane by its lipid A moiety (Fig. 1). The biosynthesis of the lipid A portion of Escherichia coli lipopolysaccharide is required for cell viability (3, 7, 8). Additionally, lipid A (endotoxin) causes extreme stimulation of the innate immune system of animals, resulting in the overproduction of diverse cytokines, which can cause the syndrome of Gram-negative sepsis (3, 4, 9, 10). Pharmacological studies have shown that both phosphate groups, the glucosamine disaccharide, and the correct number of fatty acyl chains (Fig. 1) are crucial for the cytokine-inducing activities of lipid A (3, 4, 9).

The structure of lipid A varies slightly among different Gram-negative bacterial pathogens (1, 11), such as E. coli versus Pseudomonas aeruginosa (Fig. 1), but most of the distinguishing structural features are conserved. However, the lipid A from the nitrogen-fixing bacterium Rhizobium leguminosarum differs strikingly from that of E. coli (Fig. 1) (12-15). Both phosphate groups are missing, a galacturonic acid residue is attached at the 4'-position, and the glucosamine 1-phosphate unit of E. coli lipid A is largely replaced with an aminogluconate moiety (Fig. 1) (12, 13). In the initial structural studies by Carlson and co-workers (12, 13), it was further suggested that R. leguminosarum lipid A does not possess any acyloxyacyl residues and that it contains a peculiar long fatty acid, 27-hydroxyoctacosanoic acid (Fig. 1) (16). R. leguminosarum lipid A therefore lacks many of the features thought to be necessary for stimulation of innate immunity in animals (1, 3, 4, 9). Conceivably, the unique structure of R. leguminosarum lipid A might be important for the establishment of successful symbiosis in plants (17, 18).

Despite the structural diversity of their lipid A moieties, both E. coli and R. leguminosarum employ the same seven enzymes to synthesize the key, phosphate-containing lipid A precursor, Kdo2-lipid IVA1 (19). A number of distinct R. leguminosarum enzymes are then required for the alternative processing of Kdo2-lipid IVA to generate R. leguminosarum lipid A. We have previously identified a 4'-phosphatase (20), a 1-phosphatase (21), a long chain acyl transferase (22), a mannosyl transferase (23, 24), a galactosyl transferase (21, 24), and a special Kdo transferase (24) that are involved in the unique metabolism of Kdo2-lipid IVA in extracts of R. leguminosarum. The biosynthetic origins of the galacturonic acid and the aminogluconate moieties are unknown.

We have recently discovered that lipid A of R. leguminosarum can be separated into five related molecular species (14, 15),2 two of which are shown in Fig. 1A (dashed bond at position 3). Structural studies have revealed that some of this heterogeneity can be attributed to lipid A variants lacking the equivalent of the ester-linked beta -hydroxymyristoyl moiety that is usually attached to the 3-position of lipid A disaccharides (Fig. 1) (14, 15). Unexpectedly, our reevaluation of the structure of R. leguminosarum lipid A also indicates the presence of a single acyloxyacyl moiety in all five molecular species (14, 15),2 as illustrated in Fig. 1A for two of the subtypes.

We now describe a divalent cation-dependent deacylase from R. leguminosarum membranes that selectively removes a single 3-O-linked beta -hydroxyacyl chain from certain precursors of lipid A that are common to both E. coli and R. leguminosarum. The enzyme removes only the ester-linked beta -hydroxymyristoyl residue that is attached to the 3-position of the proximal glucosamine unit (3) of precursors like lipid IVA (Fig. 1B) and Kdo2-lipid IVA. It is also capable of cleaving the monosaccharide precursor lipid X at a slow rate. A similar deacylase is found in membranes of P. aeruginosa, in which the presence of 3-O-deacylated lipid A species is well established (Fig. 1A) (25, 26). E. coli K-12 and Rhizobium meliloti do not contain the deacylase. The enzyme may therefore account for the presence of the 3-O-deacylated lipid A subtypes found in R. leguminosarum (Fig. 1A), and it may be a useful reagent for the preparation of novel endotoxin analogs with which to study innate immunity.

    EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
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REFERENCES

Chemicals and Materials-- [gamma -32P]ATP and 32Pi were obtained from NEN Life Science Products. Silica gel 60 (0.25-mm) thin layer plates were purchased from EM Separation Technologies. DEAE-cellulose (DE52) was obtained from Whatman. BAKERBOND octadecyl (C18) reverse phase resin was from J. T. Baker, and Silica Gel DavisilTM (grade 638, 100-200 mesh, 60 Å) was from Aldrich. Triton X-100 and bicinchoninic acid were from Pierce. Yeast extract and tryptone were purchased from Difco. All other chemicals were of reagent grade, and were obtained from Sigma or Mallinckrodt. Deuterated solvents were purchased from Aldrich.

Bacterial Strains and Growth Conditions-- R. leguminosarum biovar phaseoli CE3 (recently reclassified as Rhizobium etli) was a gift of K. D. Noel (Marquette University, Milwaukee, WI) (27). R. leguminosarum biovar viciae 8401 (20, 21) was obtained from J. A. Downie (John Innes Institute, Norwich, United Kingdom), and mutant 24AR of R. leguminosarum biovar trifolii was obtained from R. Russa via R. Carlson (Marie Curie Sklodowska University, Lubin, Poland) (28). R. meliloti 1021 was from S. Long (Stanford University). All other strains of Rhizobium were purchased from the American Type Culture collection (ATCC). E. coli strain W3110 was obtained from the E. coli Genetic Stock Center of Yale University. P. aeruginosa strain PAO1 was a gift of G. Pier (Harvard), and P. aeruginosa strain 27853 was obtained from the American Type Culture Collection.

All Rhizobium strains were grown at 30 °C on TY medium, which contains 5 g/liter tryptone, 3 g/liter yeast extract, 10 mM CaCl2, and 20 µg/ml nalidixic acid (19). In addition, 200 µg/ml streptomycin was also added to the medium for the growth of CE3. E. coli W3110 and P. aeruginosa (PAO1 and 27853) were grown at 30 °C in LB broth, consisting of 10 g of NaCl, 10 g of tryptone, and 5 g of yeast extract per liter (29).

Preparation of Radiolabeled Substrates-- [4'-32P]Lipid IVA was generated from [gamma -32P]ATP and the appropriate tetraacyldisaccharide 1-phosphate acceptor by using the overexpressed 4'-kinase present in membranes of E. coli BLR(DE3)pLysS/pJK2 (30). Kdo2-4'-32P-lipid IVA was then prepared from [4'-32P]lipid IVA by the action of the purified E. coli Kdo transferase (31, 32). The [4'-32P]lipid IVA and the Kdo2-[4'-32P]lipid IVA were purified by thin layer chromatography (31, 32) and were stored as aqueous dispersions at -20 °C in 10 mM Tris chloride, pH 7.8, containing 1 mM EDTA and 1 mM EGTA. Prior to use, all lipid substrates were dispersed by sonic irradiation for 1 min in a bath sonicator.

The substrate 32P-lipid X was prepared from 32Pi labeled cells of E. coli strain MN7, as described previously (33, 34). Tetraacyldisaccharide-1-32P was made from 32P-lipid X and UDP-2,3-diacylglucosamine using a highly purified preparation of E. coli disaccharide synthase (34). Nonradioactive tetraacyldisaccharide 1-phosphate carrier was prepared in the same way (34). The [4'-32P]lipid IVA (100 µM, 20,000 cpm/nmol) was hydrolyzed for 90 min at 100 °C in 0.2 M HCl to make tetraacyldisaccharide-4-32P.

Gal-Man-Kdo2-[4'-32P]lipid IVA was prepared from Kdo2-[4'-32P]lipid IVA (20,000 cpm/nmol, 20 µM) with membranes of R. meliloti 1021/pIJ 1848, which overexpresses both the mannosyl transferase and the galactosyl transferase (24). The reaction mixture was incubated with 0.75 mg/ml membrane protein at 30 °C for 90 min. The membranes were then inactivated by heating at 65 °C for 10 min. The deacylase reaction (see below) was subsequently carried out in the same reaction tube without further purification of the substrate by adding the additional necessary components.

Lauroyl-Kdo2-[4'-32P]lipid IVA was synthesized from Kdo2-[4'-32P]lipid IVA with Kdo-dependent lauroyl transferase (HtrB), partially purified from MLK1067/pKS12 (35, 36). The reaction conditions for lauroyl transferase were as follows: 0.35 µg/ml lauroyl transferase, Kdo2-[4'-32P]lipid IVA (20,000 cpm/nmol, 20 µM), lauroyl-ACP (30 µM), Triton X-100 (0.2%), MgCl2 (5 mM), and NaCl (50 mM) at pH 7.5 in 50 mM Hepes. The reaction mixture was incubated at 30 °C for 60 min. The lauroyl transferase was inactivated by heating the reaction mixture at 65 °C for 10 min. The heat-inactivated reaction mixture was then used as the substrate for the deacylase reaction without further purification.

Preparation of Cell-free Extracts and Membranes-- Two liters of mid-logarithmic phase cells (A550 = 0.6-0.8) were harvested by centrifugation (7,000 × g for 15 min at 4 °C) and were resuspended in 50 mM Hepes, pH 7.5, to give a final protein concentration of 5-10 mg/ml. Cells were broken by passage through a French pressure cell at 18,000 p.s.i. Remaining intact cells and large debris were removed by centrifugation at 7000 × g for 15 min. Membranes were prepared by ultracentrifugation at 149,000 × g for 60 min. Membrane pellets were resuspended in 50 mM Hepes, pH 7.5, at a protein concentration of ~10 mg/ml. All preparations were carried out at 4 °C, and samples were stored frozen in aliquots at -80 °C. Protein concentrations were determined with bicinchoninic acid (37), using bovine serum albumin as the standard.

Deacylase Assay Conditions and Thin Layer Chromatography-- Optimized standard assay conditions for the deacylase were as follows. The reaction mixture (10-20 µl) contained 50 mM MES, pH 6.25, 1.0% Triton X-100, 2 mM dithiothreitol, 2 mM EDTA, 20 mM calcium chloride, and 10 µM [4'-32P]lipid IVA (20,000 cpm/nmol). The reactions were incubated at 30 °C for 30 min or as indicated. The reactions were terminated by spotting 2-5-µl samples onto silica gel 60 thin layer chromatography plates. The spots were allowed to dry, and the plates were developed in the solvent chloroform/pyridine/88% formic acid/water (50:50:16:5, v/v/v/v). When substrates other than lipid IVA or tetraacyldisaccharide 4'-phosphate were used, a different solvent system was employed, consisting of chloroform/pyridine/88% formic acid/water (30:70:16:10, v/v/v/v). These alternative substrates were Kdo2-[4'-32P]lipid IVA, Gal-Man-Kdo2-[4'-32P]lipid IVA or lauroyl-Kdo2-[4'-32P]lipid IVA. The solvent chloroform/methanol/water/acetic acid (25:15:4:2, v/v/v/v) was used for deacylase reactions in which 32P-lipid X or tetraacyldisaccharide-1-32P was the substrate.

Radiochemical analyses of thin layer chromatography plates were carried out with a Molecular Dynamics PhosphorImager 425S, equipped with ImageQuant software. The percentage of conversion of unreacted 32P-labeled substrates to enzymatic products was calculated for each reaction tube and could be converted to specific activity (nmol/min/mg) based on the chemical concentration of the substrate in the assay.

Base Hydrolysis of the Deacylase Product Generated from [4'-32P]lipid IVA-- Two incubations (designated 1 and 2) were set up using the standard optimized assay conditions for the deacylase with 10 µM [4'-32P]lipid IVA as the substrate. Incubation 1 contained no enzyme, while incubation 2 contained 1 mg/ml R. leguminosarum (8401) membranes. Both incubations 1 and 2 were incubated for 16 h at 30 °C. Under these conditions, the conversion of [4'-32P]lipid IVA to the slower migrating deacylation product was almost complete in incubation 2, whereas no change was seen in incubation 1. For mild base hydrolysis of the products, a 2-µl portion of each incubation was mixed with 3 µl of triethylamine (TEA) and 5 µl of H2O, and the material was then incubated for various times at 37 °C. After 0, 2, 5, and 120 min, 2-µl samples of the TEA hydrolysis mixture were added to 2 µl of H2O and spotted onto a silica gel 60 thin layer plate. For strong base hydrolysis of the products, 2-µl portions of 1 or 2 were mixed with 18 µl of chloroform/methanol (2:1, v/v) and 0.4 µl of 10 M NaOH. This NaOH hydrolysis was carried out at room temperature for 30 min with occasional mixing of the phases. Next, each mixture was further diluted 3-fold with water, and 4-µl portions were spotted onto a silica gel 60 thin layer plate. The plates were developed with choloroform/pyridine/88% formic acid/water (50:50:16:5, v/v/v/v), dried, and exposed overnight to a PhosphorImager screen.

Large Scale Isolation of the Deacylase Product-- R. leguminosarum 8401 membranes, prepared by ultracentrifugation as described above, were enriched for the deacylase activity before being used for the large scale preparation of the reaction product. Membrane preparations (25 ml, 10 mg/ml protein) were mixed at 4 °C with 2.5% Triton X-100 for 90 min, followed by ultracentrifugation at 149,000 × g for 60 min. The pellet, containing the deacylase activity, was resuspended in 10 ml of 50 mM Hepes buffer (pH 7.5) to a protein concentration of 7.0 mg/ml. The detergent extraction process was repeated another time with 1% Triton X-100. In the pellet (resuspended in 10 ml of 50 mM Hepes buffer, pH 7.5; 5.5 mg/ml protein), more than 90% of the deacylase activity was recovered. The activity was enriched 2-fold (final specific activity of 0.1 nmol/mg/min), while most of the interfering 4'-phosphatase activity (>80%) was solubilized by the detergent. The procedure was also effective in removing about half of the membrane lipids.

Three 10-ml deacylase reaction mixtures were prepared using 50 µM lipid IVA substrate and 2 mg/ml of Triton X-100-extracted membrane protein under conditions otherwise similar to the standard assay. The reaction mixtures were initially incubated at 30 °C for 24 h. Then additional membranes were added to yield a final protein concentration of 3 mg/ml, and the reactions were continued for another 24 h. The progress of the reaction was monitored by thin layer chromatography, as described above for the assays, but the deacylase product was detected by charring the plates after spraying with 10% sulfuric acid in ethanol. More than 90% of the substrate was deacylated under these conditions. Prior to product isolation, the reaction mixtures were stored at -20 °C. After thawing, the reactions were distributed equally into two 150-ml Corex bottles. The reactions were diluted with water to yield a final volume of 20 ml/bottle. The proteins were precipitated by adding 1.25 ml of CHCl3, 2.5 ml of methanol, and 0.04 ml of concentrated HCl per ml of the diluted reaction mixtures. The samples were thoroughly mixed, and then centrifuged at 3,000 × g for 20 min at room temperature. The supernatant was decanted, and it was converted to a two-phase system by adding 0.263 ml of CHCl3 and 0.263 ml of water per ml of supernatant. After mixing, the phases were separated by centrifugation, as above. The CHCl3-rich lower phase was removed, and the upper phase was washed twice with fresh, preequilibrated lower phase (i.e. a lower phase generated by mixing chloroform, methanol, and 0.1 M HCl in a ratio of 2:2:1.8, v/v/v). The lower phases were pooled, 0.5 ml of pyridine was added to neutralize residual HCl, and the solvent was removed by rotary evaporation.

The residue was redissolved in ~15 ml of chloroform/pyridine/88% formic acid/water (70:60:16:3, v/v/v/v) and was loaded onto a 9.5-ml silicic acid column, equilibrated in the same solvent. The column was washed with another 20 ml of the same solvent, followed by 60 ml of chloroform/methanol (95:5, v/v). The lipid IVA-derived deacylase product was then eluted with ~20 ml of an acidic single phase Bligh and Dyer mixture, consisting of chloroform, methanol, and 0.1 M HCl (1:2:0.8, v/v/v) (38). The fractions containing the desired product were identified by thin layer chromatography, followed by charring as described above. The pertinent fractions were pooled and converted to a two-phase system by the addition of 0.263 ml of CHCl3 and 0.263 ml of water per ml. The solution was mixed, and the phases were separated by centrifugation. The lower phase was collected, and the upper phase was washed twice with preequilibrated acidic lower phase (see above). The lower phases were pooled, 5-10 µl of high pressure liquid chromatography grade pyridine was added, and the solvents were removed by rotary evaporation.

A small amount of contaminating lipid IVA was removed from the deacylase product by the reverse phase chromatography procedure described by Hampton et al. with minor modifications (39). The method utilizes a two-solvent system for the resolution of lipid IVA derivatives with octadecylsilane silica gel (C18 silica). Solvent A was 50% (v/v) acetonitrile in water, and solvent B was 85% isopropyl alcohol in water. Both solvents contained 5 mM tetrabutylammonium phosphate. The dried compound was redissolved in 3 ml of a 1:1 (v/v) solvent mixture of A and B. The compound was loaded onto a 0.5-ml C18 silica column equilibrated in the same solvent ratio. The column was washed with 3 ml of the 1:1 (v/v) solvent ratio. The column was then washed with 1.5 ml of solvent consisting of a 1:2 (v/v) ratio of A to B. Finally, the column was washed with 1.5 ml of solvent consisting of 1:4 (v/v) ratio of A to B. During the chromatography, 0.5-ml fractions were collected. Fractions containing the deacylase product were identified by charring. The relevant fractions were pooled and diluted 1:1 with chloroform/methanol/water (2:3:1, v/v/v). The diluted pool was loaded directly onto a 1-ml DE52 column equilibrated in chloroform/methanol/water (2:3:1, v/v) to remove the tetrabutyl-ammonium phosphate. The column was washed with 6 ml of chloroform/methanol/water (2:3:1, v/v/v), and the compound was then eluted with ~10 ml of chloroform, methanol, and 480 mM ammonium acetate (2:3:1, v/v/v). The compound-containing fractions were again identified by charring and were pooled. The pooled fractions were converted to a two-phase Bligh and Dyer system by the addition of 0.167 ml of CHCl3 and 0.283 ml of water per ml of pool. The solution was mixed, and the phases were separated by centrifugation. The lower phase was collected, and the upper phase was washed twice with preequilibrated acidic lower phase (see above). The lower phases were pooled, and the solvents were removed by rotary evaporation. The isolated deacylase product was stored at -20 °C prior to further analysis.

1H NMR Analysis of the Deacylase Reaction Product-- The purified enzymatic reaction product (2 mg) was dissolved in 0.6 ml of CDCl3/CD3OD/D2O (2:3:1, v/v/v). Its 1H NMR spectrum was recorded on a Varian 600 Unity spectrometer using a 3779.29-Hz spectral window with the 5-mm probe at 25 °C. Chemical shifts were referenced to the methyl protons of internal tetramethylsilane (0.00 ppm). A line broadening of 0.2 Hz before Fourier transformation was used to process the data. The water signal at 4.6 ppm was suppressed by presaturation (satpower = 0).

Two-dimensional 1H correlation (COSY) spectra were recorded in the absolute value mode over the same spectral region used in the one-dimensional 1H NMR spectrum. Four hundred time increments were collected and zero-filled to 2048 points with sine-bell weighting along both f1 and f2 dimensions. Three hundred twenty scans were collected per increment, and the relaxation delay was 1 s. Presaturation of the water line was also included in the pulse sequence.

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EXPERIMENTAL PROCEDURES
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A Novel Deacylase in Membranes of R. leguminosarum and P. aeruginosa-- As shown in Fig. 2, membranes of R. leguminosarum 8401 and R. leguminosarum/etli CE3 can convert [4'-32P]lipid IVA to a more hydrophilic metabolite in the presence of 1% Triton X-100 and 20 mM CaCl2. As will be demonstrated below, this product, designated metabolite A in Fig. 2, corresponds to a deacylated derivative of lipid IVA specifically lacking the 3-O-linked beta -hydroxymyristoyl residue (Fig. 1B). Membranes of the nodulation-deficient mutant 24AR of R. leguminosarum biovar trifolii, which were previously shown to lack the 4'- and 1-phosphatases of the R. leguminosarum lipid A pathway (20), do contain this deacylase. A similar deacylase is also detected in cell extracts and membranes (Fig. 2) of two wild type strains of P. aeruginosa (PA1O and 27853). However, deacylation of [4'-32P]lipid IVA is not observed in extracts or membranes (Fig. 2) of either E. coli or R. meliloti, as judged by comparison with the no enzyme control.


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Fig. 1.   Structures of lipid A from three diverse Gram-negative bacteria and their relationship to the conserved precursor lipid IVA. A, predominant species of lipid A found in E. coli K-12 (3), R. leguminosarum (12), and P. aeruginosa (25, 26). The presence of an acyloxyacyl moiety involving the C28 acyl chain and the 3-O-deacylated forms of R. leguminosarum lipid A was discovered recently in our laboratory based on new isolation techniques (14, 15). Molecular species of R. leguminosarum and P. aeruginosa lipid A may differ by the presence or absence of a hydroxyacyl chain at position 3, as indicated by the dashed bond. B, proposed reaction catalyzed by the 3-O-deacylase of R. leguminosarum with lipid IVA as the substrate. Key hydrogen atoms used to assign the structure of the product by 1H NMR spectroscopy are labeled in this representation.


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Fig. 2.   Deacylation of [4'-32P]lipid IVA in membranes of R. leguminosarum and P. aeruginosa but not of E. coli or R. meliloti. Membranes of the indicated strains were assayed for deacylase activity using the standard conditions. The protein concentration was 1.0 mg/ml, and the incubations were carried out for 60 min at 30 °C. The products generated from [4'-32P]lipid IVA were separated by thin layer chromatography and detected with a PhosphorImager. Lanes 1, no membrane control; lane 2, E. coli W3110; lane 3, R. leguminosarum biovar phaseoli CE3; lane 4, R. leguminosarum biovar viciae 8401; lane 5, R. leguminosarum biovar trifolii 24AR; lane 6, R. meliloti 1021; lane 7, P. aeruginosa PAO1; lane 8, P. aeruginosa 27853.

R. leguminosarum biovar viciae 8401 was used as the source of the enzyme in all subsequent experiments. Strain 8401 (20) lacks the pSym plasmid that carries many of the nodulation genes. All of the deacylase activity in R. leguminosarum 8401 extracts is membrane-associated (Fig. 3). Similarly, in P. aeruginosa PAO1 and 27853, all of the deacylase also is membrane-bound (not shown).


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Fig. 3.   Deacylase activity is associated with the membrane fraction of R. leguminosarum. Deacylase was assayed under standard conditions with 10 µM [4'-32P]lipid IVA and crude extract, cytosol, or membranes as the enzyme source. Reactions were analyzed after the indicated times by thin layer chromatography and PhosphorImager analysis, as described under "Experimental Procedures".

Assay and Catalytic Properties of the R. leguminosarum Deacylase-- Deacylation of [4'-32P]lipid IVA by R. leguminosarum membranes proceeds in a linear fashion for up to 4 h at 30 °C with 1.0 mg/ml protein (Fig. 4A). After prolonged incubation, deacylation is nearly complete. Deacylation activity also increases with increasing membrane protein concentrations, but the effect is not linear above 0.5 mg/ml (Fig. 4B), perhaps reflecting the presence of inhibitors in R. leguminosarum membranes. Optimal deacylase activity is observed between pH 5.5 and 6.5 (not shown).


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Fig. 4.   Time course and protein concentration dependence of the R. leguminosarum deacylase. A, membranes of R. leguminosarum 8401 were used at 1 mg/ml. The deacylase reaction was performed under standard conditions in 30 µl. At each time point, a 2-µl portion was withdrawn and analyzed by thin layer chromatography and PhosphorImager analysis. B, the deacylase reaction was performed under standard conditions in 10 µl. Membranes of R. leguminosarum 8401 were used as the enzyme source at the indicated protein concentrations. Reaction mixtures were incubated for 20 or 40 min at 30 °C.

The deacylation reaction is absolutely dependent upon the presence of a nonionic detergent. Maximal activity is seen in the presence of 1% Triton X-100 at membrane protein concentrations below 2 mg/ml. Nonidet P-40 (1% also) supports deacylase activity, but Tween 20, deoxycholate, CHAPS, and dodecylmannoside are inhibitory (not shown).

The presence of EDTA or EGTA at 2 mM completely inhibits the deacylase activity, suggesting a divalent metal requirement. Accordingly, the deacylase was assayed in the presence of varying concentrations of calcium, magnesium, or manganese ions (Fig. 5). In this experiment, the concentration of EDTA was held constant at 2 mM. Among these three divalent metal ions, calcium is the most effective. Enzymatic activity increases with increasing concentrations of calcium ions up to 20 mM. Partial stimulation of the activity is also seen with magnesium or manganese chloride (Fig. 5). However, 5 mM ferrous and zinc ions completely inhibit the activity (not shown), and monovalent cations (like sodium and potassium) have no effect.


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Fig. 5.   Dependence of the deacylase reaction on divalent metal ions. The deacylase was assayed under standard conditions, except that the indicated concentration of CaCl2, MgCl2, or MnCl2 was added together with 2 mM EDTA to the reaction mixtures. R. leguminosarum 8401 membranes were used at 1.0 mg/ml. Reaction mixtures were incubated for 40 min at 30 °C, after which product formation was analyzed by thin layer chromatography and PhosphorImager analysis.

Under standardized assay conditions with R. leguminosarum 8401 membranes, partial dephosphorylation of the 4'-phosphate of lipid IVA by the 4'-phosphatase is observed in parallel with deacylation as judged by the appearance of 32Pi (Fig. 2, lane 4), but the 4'-phosphatase is partially inhibited by the presence of 20 mM CaCl2 and 1% Triton X-100. Since Kdo2-lipid IVA is strongly preferred over lipid IVA by the 4'-phosphatase (20), lipid IVA was generally employed for the deacylase assay. The 1-phosphatase of the R. leguminosarum lipid A pathway (21) was inhibited completely by 20 mM CaCl2; therefore, its activity was not apparent under the assay conditions used for the deacylase. Furthermore, the lipid product that is generated by the 4'-phosphatase (20) is not seen under these assay conditions, since it is not radioactive after the removal of the 4'-phosphate residue.

With 25 µM [4'-32P]lipid IVA under optimized conditions, the specific activity of the deacylase in crude extracts and membrane preparations of R. leguminosarum 8401 is 0.019 and 0.045 nmol/min/mg protein, respectively. The apparent Km for lipid IVA in this mixed micelle system (40) is estimated as 17.9 µM (Fig. 6).


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Fig. 6.   Effect of lipid IVA concentration on deacylase specific activity. The deacylase was assayed under standard conditions with membranes of R. leguminosarum 8401 at 1.0 mg/ml. The lipid IVA concentration was varied, as indicated. Reaction mixtures were incubated for 40 min, after which product formation was analyzed by thin layer chromatography and PhosphorImager analysis. The inset shows a double reciprocal plot of the same data.

Substrate Specificity of the Deacylase-- As shown in Fig. 7, the deacylase utilizes lipid IVA and Kdo2-lipid IVA at about the same rate. Unlike the 4'-phosphatase, the deacylase is not dependent upon the Kdo domain. However, the rate of deacylation of the monosaccharide precursor lipid X is about 20 times slower than that of lipid IVA (Fig. 7), indicating that the distal diacylglucosamine unit of lipid IVA somehow enhances catalytic efficiency. The biosynthetic precursor tetraacyldisaccharide-1-32P (3) and the analog tetraacyldisaccharide-[4'-32P] (prepared by acid hydrolysis of [4'-32P]lipid IVA) are both deacylated efficiently (data not shown), indicating that both phosphates are not required for efficient turnover. Extra core sugars (mannose and galactose) (21, 24) attached to Kdo2-lipid IVA do not interfere with deacylation. However, the presence of an acyloxyacyl group on the distal glucosamine residue of Kdo2-lipid IVA, as in lauroyl-Kdo2-lipid IVA generated by HtrB (35) or in lipid A generated from compound 505 by the 4'-kinase (30), prevents deacylation (data not shown).


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Fig. 7.   Substrate specificity of the deacylase. The deacylase was assayed under standard conditions with 10 µM [4'-32P]lipid IVA, Kdo2-[4'-32P]lipid IVA, or 32P-lipid X (each at about 20,000 cpm/nmol). Membranes (1.0 mg/ml) of R. leguminosarum 8401 were used as the enzyme source. Product formation was analyzed by thin layer chromatography and PhosphorImager analysis.

Chromatographic Characterization of the Deacylase Product Generated from Lipid IVA-- To characterize the structure of the material generated by the enzymatic deacylation of lipid IVA, the reaction product was subjected to mild alkaline hydrolysis. The substrate [4'-32P]lipid IVA (not treated with enzyme) (Fig. 8) was processed in parallel (see "Experimental Procedures"). Treatment with TEA removes both ester-linked beta -hydroxyacyl chains from lipid IVA. When carried out at 30 °C, TEA hydrolysis of [4'-32P]lipid IVA proceeds via two distinct and separable intermediates (designated intermediates 1 and 2 in lanes 3 and 5 of Fig. 8). These eventually collapse to form the same limiting alkaline hydrolysis product, which lacks both ester-linked beta -hydroxyacyl chains (Fig. 8, lane 7). Intermediates 1 and 2 arise by the loss of either the 3- or the 3'-beta -hydroxyacyl chains of lipid IVA. Stronger base (NaOH) hydrolysis of lipid IVA (Fig. 8, lane 9) rapidly removes both O-linked beta -hydroxyacyl chains without accumulation of intermediates 1 and 2 under the conditions employed.


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Fig. 8.   Mild alkaline hydrolysis and thin layer analysis of [4'-32P]lipid IVA and of the deacylase reaction product. As described under "Experimental Procedures," two incubations (designated 1 and 2) were set up using the standard optimized assay conditions with 10 µM [4'-32P]lipid IVA as the substrate. Incubation 1 contained no enzyme, while incubation 2 contained 1 mg/ml R. leguminosarum 8401 membranes. After 16 h at 30 °C, portions of these incubations were treated with TEA or 0.1 M NaOH for the times indicated in the figure, after which a portion of each treated sample was analyzed by thin layer chromatography and PhosphorImager analysis.

The more rapidly migrating intermediate 1 derived by TEA treatment of [4'-32P]lipid IVA (Fig. 8, lanes 3 and 5) is the same as the product made by the R. leguminosarum deacylase (Fig. 8, lane 2, metabolite A). Consequently, treatment of the deacylase reaction product with TEA (Fig. 8, lanes 4, 6, and 8) or with NaOH (Fig. 8, lane 10) results in the direct conversion of the deacylase reaction product to a compound that migrates with the limiting NaOH hydrolysis product of [4'-32P]lipid IVA. This finding demonstrates that the deacylase does not remove an N-linked beta -hydroxyacyl chain from lipid IVA, and it also shows that the deacylase removes only one specific O-linked beta -hydroxyacyl chain from [4'-32P]lipid IVA. Last, it can be inferred that the glucosamine disaccharide backbone structure of the deacylase reaction product is likely to be the same as that of lipid IVA.

1H NMR Spectroscopy of the Deacylase Reaction Product Generated from Lipid IVA-- A definitive assignment of the position (3 or 3') that is attacked by the deacylase in lipid IVA cannot be made by thin layer chromatography analysis alone. Accordingly, a 2-mg sample of the deacylase reaction product was isolated and subjected to 1H NMR spectroscopy under conditions reported previously (41).

Characterization of the purified reaction product by COSY spectroscopy established unequivocally that lipid IVA is deacylated by the enzyme exclusively at the 3-position. While the 1H NMR spectrum of the substrate, lipid IVA (not shown), displays two downfield overlapping triplets at 5.18 ppm, which are attributed to H-3 and H-3' of the acylated glucosamine disaccharide (42, 43), the deacylase product retains only one of these two triplets at 5.17 ppm (Fig. 9). Based on the COSY spectrum of the deacylase product, this remaining downfield triplet is assigned to an axial H-3' (Fig. 9), indicating that the 3'-position is still acylated in the product (33, 45). However, a new triplet at 3.69 ppm (Fig. 9), which is not detected in the substrate lipid IVA (42, 43), was observed in the product (Fig. 9). By tracing the cross-peaks of the COSY spectrum (Fig. 9), the new resonance at 3.69 ppm could be assigned to the glucosamine H-3 of the product, which is shifted upfield by about 1.5 ppm compared with lipid IVA, because of the loss of the 3-O-acyl moiety. Deacylation at the 3-position was also indicated by the slight (~0.2 ppm) upfield shifts of the H-2 and H-4 signals of the product in comparison with the H-2 and H-4 resonances of lipid IVA (42, 43). Thus, the deacylase selectively removes the ester-linked beta -hydroxymyristate chain at the 3-position of the proximal glucosamine unit of lipid IVA.


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Fig. 9.   Partial COSY spectrum of the deacylase reaction product generated from lipid IVA. The one-dimensional spectrum in the region of the relevant carbohydrate proton resonances is shown along the edges. Proton cross-peak assignments are indicated, and the atom labeling of the glucosamine disaccharide protons follows the numbering scheme shown in Fig. 1B. The distal glucosamine 1H chemical shifts of the deacylase product are nearly the same as those in the lipid IVA substrate (43). The most pronounced difference between lipid IVA and the deacylase reaction product is seen with H-3 of the proximal glucosamine residue, which is shifted upfield by 1.5 ppm relative to H-3 of lipid IVA (43) because of the loss of the acyl chain.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

In previous studies, we described six enzymes unique to R. leguminosarum extracts that convert Kdo2-lipid IVA (20-24), an intermediate made both by E. coli and R. leguminosarum, to novel compounds that are precursors of the unusual lipid A of R. leguminosarum (Fig. 1). We have now identified a novel deacylase specific to R. leguminosarum membranes that selectively removes the O-linked beta -hydroxyacyl chain from the 3-position of Kdo2-lipid IVA as well as from the precursors lipid IVA and lipid X (Figs. 1, 7, and 9). This membrane-associated deacylase is present in extracts of all R. leguminosarum strains tested so far, but not of E. coli or R. meliloti (Fig. 2).

Although the deacylase is not dependent upon the Kdo domain of Kdo2-lipid IVA for activity, it displays a strong kinetic preference for substrates containing a distal diacylglucosamine unit (Fig. 7). However, the enzyme selectively cleaves only the beta -hydroxymyristoyl moiety attached to position 3 of the proximal glucosamine moiety (Figs. 1 and 9). 1H NMR spectroscopy (Fig. 9) provided unequivocal evidence for the specificity of the enzyme, as demonstrated by the 1.5-ppm upfield shift of the H-3 of the proximal glucosamine residue in the product compared with the substrate. The NMR spectrum also revealed that the proton chemical shifts of the distal glucosamine unit of the product were virtually unchanged relative to those of lipid IVA (43).

The deacylase does not require the presence of a phosphate group at either the 1- or the 4'-position for activity, and the attachment of the inner core sugars (mannose and galactose) to Kdo2-lipid IVA does not interfere with the deacylation reaction (data not shown). Interestingly, the enzyme does not deacylate lauroyl-Kdo2-lipid IVA (in which an acyloxyacyl group is present on the N-linked hydroxymyristate group of the distal glucosamine unit) (35). These findings suggest that the deacylase functions after lipid A disaccharide formation but before the Kdo-dependent acylation of the distal unit to generate an acyloxyacyl residue (3). The observation that the deacylase strongly prefers glucosamine disaccharides as substrates (Fig. 7) is reasonable given that the disaccharide synthase of E. coli (34) is highly specific for diacylated monosaccharide precursors, like lipid X. If the deacylase were to catalyze rapid cleavage of these monosaccharide precursors, it would interfere with the functioning of the disaccharide synthase.

Like many of the lipases that deacylate glycerophospholipids (46), the R. leguminosarum deacylase requires Triton X-100 and divalent metal ions for activity (Fig. 5). Whether or not the R. leguminosarum deacylase also utilizes glycerophospholipids as substrates can only be assessed once the enzyme is purified to homogeneity. However, the remarkable specificity of the deacylase for the 3-position of lipid IVA argues against a more general role for this enzyme as a phospholipase. Furthermore, when commercially available lipases from Rhizopus and Pseudomonas or bovine pancreatic phospholipase A2 was tested under the conditions optimized for the R. leguminosarum deacylase (or under their own optimal conditions for the hydrolysis of glycerophospholipids) (46), no deacylation of [4'-32P]lipid IVA was observed (data not shown).

The present work therefore represents the first description of a lipase from a procaryotic organism that deacylates lipid A precursors. While several deacylases that attack certain lipid A-like molecules have been reported in eucaryotic systems, these appear to be distinct from the R. leguminosarum deacylase. For instance, the acyloxyacyl hydrolase of human leukocytes (47) removes the secondary acyl chains from the lipid A residues of intact lipopolysaccharide, thereby reducing the immunostimulatory activity and the toxicity of the lipid A moiety. Since the R. leguminosarum deacylase does not attack Kdo2-(lauroyl)-[4'-32P]lipid IVA, it is obviously not an acyloxyacyl hydrolase (data not shown). Rosner et al. (48) and Verret et al. (49, 50) identified two distinct amidases in extracts of the slime mold, Dictyostelium discoideum, an organism that is likely to scavenge E. coli in nature. These enzymes remove the two amide-linked beta -hydroxymyristoyl residues of lipid A, but only after complete O-deacylation by prior base treatment. Amidase I of D. discoideum is inhibited by chitobiose and N-acetyl-alpha -glucosamine 1-phosphate (49, 50). The R. leguminosarum deacylase does not cleave either of the amide-linked acyl chains (Fig. 8), and it is not affected by the above compounds up to 1.0 mM (data not shown). The specificity of our deacylase is thus completely different from those of the previously reported lipid A hydrolases (49, 50) of D. discoideum. Finally, Drozanski et al. (51) reported deacylation of lipopolysaccharide in Acanthamoeba castellanii crude extracts with release of both O-linked and N-linked acyl chains. These early studies were carried out without the benefit of structurally defined substrates. The putative enzymes described by Drozanski et al. (51) were not further characterized. Their specificity and function remain unclear.

The physiological function of the R. leguminosarum deacylase is not yet known. Elucidation of its biological roles will require genetic studies. It is very likely that the R. leguminosarum deacylase generates the lipid A species lacking the 3-O-linked beta -hydroxyacyl chain that we have recently identified in R. leguminosarum (14, 15) (Fig. 1A). The related deacylase found in P. aeruginosa membranes (Fig. 2) may likewise explain the structure of the predominant pentaacylated form of lipid A present in P. aeruginosa lipopolysaccharide (25, 26), in which the 3-O-linked beta -hydroxydecanoate group is missing in the proximal glucosamine unit. Mass spectrometry of lipid A variants isolated from diverse Gram-negative bacteria suggests that enzymatic cleavage of 3-O-linked beta -hydroxyacyl chains may actually be a widespread phenomenon (52, 53).

Many studies have confirmed the importance of the structure and composition of the acyl chains attached to lipid A for biological activity in the stimulation of mammalian immune cells (1, 3, 4, 9). The deacylase of R. leguminosarum may provide a new tool for the selective modification and preparation of interesting lipid A analogs. Given the structure-activity relationships of known lipid A derivatives, one would expect that many of the unusual chemical features of R. leguminosarum lipid A (Fig. 1A), including the partial 3-O-deacylation, would reduce immune stimulation in animal systems (4, 9).

The distinctive structure of R. leguminosarum lipid A and its possible lack of immunostimulatory activity might also play a role the establishment of symbiosis in plants (17, 18). Although not yet characterized in terms of their ability to respond to lipid A, some plants have recently been shown to possess systems of innate immunity (54-56) and to synthesize antibacterial peptides in a manner that is reminiscent of insects infected with bacteria or fungi (44). The unusual lipid A of R. leguminosarum might therefore help bacteroids evade the innate immune response of plants during symbiosis in root cells, while still allowing the plant to defend itself against Gram-negative pathogens containing the more typical, phosphorylated lipid A disaccharide. Isolation of R. leguminosarum mutants that specifically lack the 3-O-deacylase and the other unique enzymes recently identified in our laboratory (20, 21, 24) will be required to validate this hypothesis.

    FOOTNOTES

* This work was supported by National Institutes of Health Grant GM-51796 (to C. R. H. R.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed. Tel.: 919-684-5326; Fax: 919-684-8885; E-mail: raetz{at}biochem.duke.edu.

2 N. L. S. Que and C. R. H. Raetz, manuscript in preparation.

    ABBREVIATIONS

The abbreviations used are: Kdo, 3-deoxy-D-manno-octulosonic acid; TEA, triethylamine; MES, 4-morpholineethanesulfonic acid; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid; COSY, two-dimensional 1H correlation spectroscopy.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
  1. Raetz, C. R. H. (1990) Annu. Rev. Biochem. 59, 129-170[CrossRef][Medline] [Order article via Infotrieve]
  2. Raetz, C. R. H. (1993) J. Bacteriol. 175, 5745-5753[Medline] [Order article via Infotrieve]
  3. Raetz, C. R. H. (1996) in Escherichia coli and Salmonella: Cellular and Molecular Biology (Neidhardt, F. C., ed), 2nd Ed., Vol. 1, pp. 1035-1063, American Society for Microbiology, Washington, D. C.
  4. Rietschel, E. T., Kirikae, T., Schade, F. U., Mamat, U., Schmidt, G., Loppnow, H., Ulmer, A. J., Zähringer, U., Seydel, U., Di Padova, F., Schreier, M., and Brade, H. (1994) FASEB J. 8, 217-225[Abstract/Free Full Text]
  5. Schnaitman, C. A., and Klena, J. D. (1993) Microbiol. Rev. 57, 655-682[Abstract]
  6. Reeves, P. (1994) in Bacterial Cell Wall: New Comprehensive Biochemistry (Neuberger, A., and van Deenen, L. L. M., eds), Vol. 27, pp. 281-314, Elsevier Science Publishers, New York
  7. Galloway, S. M., and Raetz, C. R. H. (1990) J. Biol. Chem. 265, 6394-6402[Abstract/Free Full Text]
  8. Onishi, H. R., Pelak, B. A., Gerckens, L. S., Silver, L. L., Kahan, F. M., Chen, M. H., Patchett, A. A., Galloway, S. M., Hyland, S. A., Anderson, M. S., and Raetz, C. R. H. (1996) Science 274, 980-982[Abstract/Free Full Text]
  9. Levin, J., Alving, C. R., Munford, R. S., and Stütz, P. L. (eds) (1993) Bacterial Endotoxin: Recognition and Effector Mechanisms, Excerpta Medica, Amsterdam
  10. Ulevitch, R. J., and Tobias, P. S. (1995) Annu. Rev. Immunol. 13, 437-457[CrossRef][Medline] [Order article via Infotrieve]
  11. Morrison, D. C., and Ryan, J. L. (eds) (1992) Bacterial Endotoxic Lipopolysaccharides, Vol. I, CRC Press, Inc., Boca Raton, FL
  12. Bhat, U. R., Forsberg, L. S., and Carlson, R. W. (1994) J. Biol. Chem. 269, 14402-14410[Abstract/Free Full Text]
  13. Forsberg, L. S., and Carlson, R. W. (1998) J. Biol. Chem. 273, 2747-2757[Abstract/Free Full Text]
  14. Que, N. L. S., Basu, S. S., White, K. A., and Raetz, C. R. H. (1998) FASEB J. 12, 1284 (abstr.)
  15. Lin, S., Woods, A. S., Cotter, R. J., Raetz, C. R. H., and Que, N. L. S. (1998) FASEB J. 12, A14LB56 (abstr.)
  16. Bhat, U. R., Mayer, H., Yokota, A., Hollingsworth, R. I., and Carlson, R. (1991) J. Bacteriol. 173, 2155-2159[Medline] [Order article via Infotrieve]
  17. Long, S. R., and Staskawicz, B. J. (1993) Cell 73, 921-935[Medline] [Order article via Infotrieve]
  18. Whitehead, L. F., and Day, D. A. (1997) Physiol. Plant. 100, 30-44[CrossRef]
  19. Price, N. J. P., Kelly, T. M., Raetz, C. R. H., and Carlson, R. W. (1994) J. Bacteriol. 176, 4646-4655[Abstract]
  20. Price, N. J. P., Jeyaretnam, B., Carlson, R. W., Kadrmas, J. L., Raetz, C. R. H., and Brozek, K. A. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 7352-7356[Abstract]
  21. Brozek, K. A., Kadrmas, J. L., and Raetz, C. R. H. (1996) J. Biol. Chem. 271, 32112-32118[Abstract/Free Full Text]
  22. Brozek, K. A., Carlson, R. W., and Raetz, C. R. H. (1996) J. Biol. Chem. 271, 32126-32136[Abstract/Free Full Text]
  23. Kadrmas, J. L., Brozek, K. A., and Raetz, C. R. H. (1996) J. Biol. Chem. 271, 32119-32125[Abstract/Free Full Text]
  24. Kadrmas, J. L., Allaway, D., Studholme, R. E., Sullivan, J. T., Ronson, C. W., Poole, P. S., and Raetz, C. R. H. (1998) J. Biol. Chem. 273, 26432-26440[Abstract/Free Full Text]
  25. Kulshin, V. A., Zahringer, U., Lindner, B., Jager, K. E., Dmitriev, B. A., and Rietschel, E. T. (1991) Eur. J. Biochem. 198, 697-704[Abstract]
  26. Karunaratne, D. N., Richards, J. C., and Hancock, R. E. (1992) Arch. Biochem. Biophys. 299, 368-376[Medline] [Order article via Infotrieve]
  27. Cava, J. R., Elias, P. M., Turowski, D. A., and Noel, K. D. (1989) J. Bacteriol. 171, 8-15[Medline] [Order article via Infotrieve]
  28. Russa, R., Lüderitz, O., and Rietschel, E. T. (1985) Arch. Microbiol. 141, 284-289
  29. Miller, J. R. (1972) Experiments in Molecular Genetics, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  30. Garrett, T. A., Kadrmas, J. L., and Raetz, C. R. H. (1997) J. Biol. Chem. 272, 21855-21864[Abstract/Free Full Text]
  31. Belunis, C. J., and Raetz, C. R. H. (1992) J. Biol. Chem. 267, 9988-9997[Abstract/Free Full Text]
  32. Brozek, K. A., Hosaka, K., Robertson, A. D., and Raetz, C. R. H. (1989) J. Biol. Chem. 264, 6956-6966[Abstract/Free Full Text]
  33. Takayama, K., Qureshi, N., Mascagni, P., Nashed, M. A., Anderson, L., and Raetz, C. R. H. (1983) J. Biol. Chem. 258, 7379-7385[Abstract/Free Full Text]
  34. Radika, K., and Raetz, C. R. H. (1988) J. Biol. Chem. 263, 14859-14867[Abstract/Free Full Text]
  35. Clementz, T., Bednarski, J. J., and Raetz, C. R. H. (1996) J. Biol. Chem. 271, 12095-12102[Abstract/Free Full Text]
  36. Clementz, T., Zhou, Z., and Raetz, C. R. H. (1997) J. Biol. Chem. 272, 10353-10360[Abstract/Free Full Text]
  37. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985) Anal. Biochem. 150, 76-85[Medline] [Order article via Infotrieve]
  38. Bligh, E. G., and Dyer, J. J. (1959) Can. J. Biochem. Physiol. 37, 911-918
  39. Hampton, R. Y., Golenbock, D. T., and Raetz, C. R. H. (1988) J. Biol. Chem. 263, 14802-14807[Abstract/Free Full Text]
  40. Carman, G. M., Deems, R. A., and Dennis, E. A. (1995) J. Biol. Chem. 270, 18711-18714[Free Full Text]
  41. Garrett, T. A., Que, N. L., and Raetz, C. R. H. (1998) J. Biol. Chem. 273, 12457-12465[Abstract/Free Full Text]
  42. Strain, S. M., Armitage, I. M., Anderson, L., Takayama, K., Qureshi, N., and Raetz, C. R. H. (1985) J. Biol. Chem. 260, 16089-16098[Abstract/Free Full Text]
  43. Ray, B. L., and Raetz, C. R. H. (1987) J. Biol. Chem. 262, 1122-1128[Abstract/Free Full Text]
  44. Lemaitre, B., Reichhart, J. M., and Hoffmann, J. A. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 14614-14619[Abstract/Free Full Text]
  45. Ray, B. L., Painter, G., and Raetz, C. R. H. (1984) J. Biol. Chem. 259, 4852-4859[Abstract/Free Full Text]
  46. Reynolds, L. J., Washburn, W. N., Deems, R. A., and Dennis, E. A. (1991) Methods Enzymol. 197, 3-23[Medline] [Order article via Infotrieve]
  47. Hagen, F. S., Grant, F. J., Kuijper, J. L., Slaughter, C. A., Moomaw, C. R., Orth, K., O'Hara, P. J., and Munford, R. S. (1991) Biochemistry 30, 8415-8423[Medline] [Order article via Infotrieve]
  48. Rosner, M. R., Verret, R. C., and Khorana, H. G. (1979) J. Biol. Chem. 254, 5926-5933[Abstract]
  49. Verret, C. R., Rosner, M. R., and Khorana, H. G. (1982) J. Biol. Chem. 257, 10228-10234[Abstract/Free Full Text]
  50. Verret, C. R., Rosner, M. R., and Khorana, H. G. (1982) J. Biol. Chem. 257, 10222-10227[Abstract/Free Full Text]
  51. Drozanski, W., Galanos, C., Schlecht, S., and Lüderitz, O. (1986) Eur. J. Biochem. 155, 433-437[Abstract]
  52. Karibian, D., Deprun, C., Szabó, L., Le Beyee, Y., and Caroff, M. (1991) Int. J. Mass Spectrom. Ion Processes 111, 273-286[CrossRef]
  53. Zarrouk, H., Karibian, D., Bodie, S., Perry, M. B., Richards, J. C., and Caroff, M. (1997) J. Bacteriol. 179, 3756-3760[Abstract]
  54. Medzhitov, R., and Janeway, C. A., Jr. (1998) Curr. Opin. Immunol. 10, 12-15[CrossRef][Medline] [Order article via Infotrieve]
  55. Boman, H. G. (1998) Scand. J. Immunol. 48, 15-25[CrossRef][Medline] [Order article via Infotrieve]
  56. O'Neill, L. A., and Greene, C. (1998) J. Leukocyte Biol. 63, 650-657[Abstract]


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