Purification and Characterization of WaaP from Escherichia coli, a Lipopolysaccharide Kinase Essential for Outer Membrane Stability*

Jeremy A. YethonDagger and Chris Whitfield§

From the Department of Microbiology, University of Guelph, Guelph, Ontario N1G 2W1, Canada

Received for publication, September 8, 2000, and in revised form, October 25, 2000



    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

In Escherichia coli, Salmonella enterica, and Pseudomonas aeruginosa, the waaP (rfaP) gene product is required for the addition of phosphate to O-4 of the first heptose residue of the lipopolysaccharide (LPS) inner core region. This phosphate substitution is particularly important to the biology of these bacteria; it has previously been shown that WaaP is necessary for resistance to hydrophobic and polycationic antimicrobials in E. coli and that it is required for virulence in invasive strains of S. enterica. WaaP function is also known to be essential for the viability of P. aeruginosa. The predicted WaaP protein shows low levels of similarity (10-15% identity) to eukaryotic protein kinases, but its kinase activity has never been tested. Here we report the purification of WaaP and the reconstitution of its enzymatic activity in vitro. The purified enzyme catalyzes the incorporation of 33P from [gamma -33P]ATP into acceptor LPS purified from a defined E. coli waaP mutant. Enzymatic activity is dependent upon the presence of Mg2+ and is maximal from pH 8.0 to 9.0. The apparent Km (determined at saturating concentrations of the second substrate) is 0.13 mM for ATP and 76 µM for LPS. These data are the first proof that WaaP is indeed an LPS kinase. Further, site-directed mutagenesis of a predicted catalytic residue suggests that WaaP shares a common mechanism of action with eukaryotic protein kinases.



    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The outer membrane of a Gram-negative bacterium is a barrier to many antibiotics and host defense factors (1, 2). This barrier function is due in large part to structural features of the lipopolysaccharide (LPS)1 molecules that make up the outer leaflet of the outer membrane bilayer. In Escherichia coli, Salmonella enterica, and a variety of other Gram-negative pathogens including members of the families Pseudomonadaceae and Vibrionaceae, the LPS molecule is conceptually divided into three distinct regions: 1) a hydrophobic membrane anchor designated lipid A; 2) a short, branched chain of sugar residues with multiple phosphoryl substituents, referred to as the core oligosaccharide; and 3) a structurally diverse polysaccharide composed of repeating oligosaccharide units, termed the O antigen (3) (Fig. 1). The presence of phosphoryl substituents on the inner (lipid A proximal) region of the LPS core oligosaccharide is a key structural feature required for the formation of a stable outer membrane in these bacteria (4-6). These phosphoryl substituents are postulated to be critical to outer membrane integrity because their negative charge allows neighboring LPS molecules to be cross-linked by divalent cations (1, 2).

The genes involved in core phosphorylation have only recently been identified with certainty. In E. coli, waaP was shown to be required for phosphate addition to HepI (4), and this reaction was found to be a prerequisite for the addition of the HepIII residue by the waaQ gene product, which in turn was required for the waaY-mediated addition of a second phosphate at HepII (see Fig. 1). Core phosphorylation in S. enterica proceeds in similar fashion (5). Given the sequential action of the waaP, waaQ, and waaY gene products, mutation of waaP alone is enough to eliminate all phosphate from the heptose region of the LPS inner core, resulting in a strain that is hypersensitive to detergents and hydrophobic antibiotics. Although mutation of waaY does reduce the amount of core phosphate, this reduction is not sufficiently serious to cause hypersensitivity to such compounds (4).

Mutants of E. coli and S. enterica with highly truncated core oligosaccharides, such that they lack the inner core heptose residues that serve as the sites for phosphorylation (see Fig. 1), exhibit a pleiotropic phenotype called "deep-rough." Characteristics of the deep-rough phenotype include 1) hypersensitivity to detergents and hydrophobic antibiotics, 2) the appearance of phospholipid bilayer patches in the outer membrane, 3) leakage of periplasmic proteins into the culture medium, and 4) a marked decrease in the protein content of the outer membrane (reviewed in Refs. 7 and 8). It has also been shown that the LPS from deep-rough mutants cannot support the proper folding of some outer membrane proteins (9). It was thought originally that all these characteristics could be explained simply by loss of core phosphoryl substituents. However, this was recently shown to be somewhat of an oversimplification, at least in the case of the outer membrane protein defect; a defined waaP mutant was shown to have wild-type levels of outer membrane proteins despite a complete lack of core phosphate (10). Therefore, while the loss of core phosphate undoubtedly plays an important role in the manifestation of the deep-rough phenotype, clearly other factors must also be involved.

The importance of LPS core phosphorylation and of WaaP activity in particular extend beyond the obvious membrane defects. For example, mutation of waaP in S. enterica serovar Typhimurium has been shown to cause a complete loss of virulence in mouse infection models (5). In addition, WaaP activity is intuitively a prerequisite for the functioning of the currently unidentified enzyme responsible for 2-aminoethyl phosphate (PEtN) modification of the E. coli and S. enterica inner core heptose region (see Fig. 1). PEtN modification of the LPS inner core is correlated with resistance to polymyxin in these bacteria (11, 12) and may provide a means by which these pathogens can modulate their surface charge. Inhibition of WaaP would thus hinder the ability of these bacteria to adapt their surfaces to particular microenvironments during pathogenesis.



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Fig. 1.   Structure of the LPS from E. coli strains with an R1-type core. Core residues are designated by sugar abbreviation and number to facilitate identification. P, phosphate. All sugars are in the pyranose configuration, and the linkages are alpha  unless otherwise indicated. The assignment of function to genes encoding core glycosyltransferases and phosphotransferases has been reported previously (4, 32-34). The predicted activity of the waaP gene product is indicated by an asterisk.

The predicted WaaP protein shares limited similarity with eukaryotic protein kinases (4), but kinase activity has never been demonstrated. Here, we report the purification of a His6-tagged derivative of WaaP, and the development of an in vitro assay that demonstrates unequivocally that WaaP is an LPS core heptose kinase. We also give the first characterization of the catalytic properties of the WaaP enzyme and provide site-directed mutagenesis data that suggest that the enzyme's mechanism of action is similar to that of eukaryotic protein kinases.


    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Materials and Bacterial Strains-- Materials and kits were purchased from the following suppliers: PCR primers (Guelph Molecular Supercentre, University of Guelph); restriction enzymes (New England Biolabs and Roche Molecular Biochemicals); QIAprep Spin Miniprep Kit and QIAquick PCR Purification Kit (Qiagen); antibiotics (Sigma); nickel-nitrilotriacetic acid-agarose (Ni2+-NTA-agarose) (Qiagen); PD-10 desalting columns (Amersham Pharmacia Biotech); [gamma -33P]ATP (3000 Ci/mmol) (PerkinElmer Life Sciences); other assay reagents (Tris, MgCl2, Triton X-100, dithiothreitol) (Sigma); and EcoLite scintillation fluid (ICN). LPS acceptor for the WaaP assay was purified from defined waaP mutant strain CWG296 (4) by hot phenol/water extraction of cells (13). E. coli strain BL21(DE3) (F- ompT gal [dcm] [lon] hsdSB (rB-mB-) lambda DE3) is from Novagen, and E. coli strain DH5alpha (supE44 Delta lacU169 (phi 80lacZDelta M15) hsdR17 recA1 endA1 gyrA96 thi-1 relA1) is from Life Technologies, Inc.

Cloning and Expression of waaP-- Oligonucleotide primers for the amplification of waaP from the E. coli F470 chromosome (R1 core prototype strain; see Ref. 14) were designed to introduce appropriate restriction sites for cloning. The forward primer (5'-TGTGGATccAAATAGTGGGCACTCA-3') introduced a BamHI site (underlined) 32 base pairs downstream of the waaP stop codon, and the reverse primer (5'-GGGTGGTCcatATGGTTGAACTTAA-3') introduced an NdeI site (underlined) overlapping the waaP ATG start codon (bases in lowercase indicate mismatches between the primer and chromosomal sequences). PCR amplification was performed with PwoI DNA polymerase (Roche Molecular Biochemicals), used as recommended by the manufacturer. The coding region for waaP was subsequently isolated as a BamHI-NdeI fragment and then cloned between the BamHI and NdeI sites of pET-28a(+) (Novagen), generating plasmid pWQ910, and sequenced to ensure error-free amplification. Plasmid pWQ910 introduces an N-terminal His6 tag onto the WaaP protein and provides high-level, IPTG-inducible expression from the T7 promoter.

Cloning and Expression of the Chaperones Encoded by groES and groEL-- The GroES and GroEL chaperones of E. coli were overexpressed simultaneously with the recombinant His6-WaaP to increase the yield of soluble WaaP protein (based on the work of Ref. 15). The operon containing groES and groEL was PCR-amplified from the E. coli W3110 chromosome (16). Again, primers were designed to introduce appropriate restriction sites for cloning; the forward primer (5'-GGAGAGTTAcatATGAATATTCG-3') introduced an NdeI site (underlined) overlapping the groES start codon, and the complementary reverse primer (5'-AGGTGCAGGAAgcttACATCATG-3') introduced a HindIII site (underlined) overlapping the groEL stop codon (bases shown in lowercase indicate mismatches between the primer and chromosomal sequences). The coding region for groESL was subsequently isolated as an NdeI-HindIII fragment and cloned between the NdeI and HindIII sites of pET-30a(+) (Novagen). The groESL coding region was then excised from the pET-30a(+) derivative as an XbaI-HindIII fragment (along with the optimally positioned ribosome binding site) and cloned between the XbaI and HindIII sites of pBAD33 (17). The resulting plasmid, pWQ911, allows for high level chaperone expression from the arabinose-inducible PBAD promoter.

Growth Conditions and Preparation of Cell-free Extracts-- Luria-Bertani broth containing chloramphenicol (30 µg/ml) and kanamycin (50 µg/ml) was inoculated (1:20) from an overnight culture of BL21(DE3)pWQ910/pWQ911 and grown to an A600 of ~0.8 at 37 °C with shaking at 200 rpm. L-Arabinose (0.02%) was then added to induce GroES and GroEL expression, and cells were grown for an additional 45 min, at which point IPTG (0.5 mM) was added to induce expression of WaaP. After 4.5 h, cells were harvested by centrifugation (15 min, 5000 × g, 4 °C) and then frozen and stored overnight at -20 °C. The cell pellet was then thawed on ice and resuspended in a minimal volume (4 ml/g, wet weight) of lysis buffer (50 mM sodium phosphate, pH 8, 350 mM NaCl, 10 mM imidazole, 2% Triton X-100, 10 mM beta -mercaptoethanol). Lysozyme (1 mg/ml) was added to initiate lysis, and both DNase I (5 µg/ml) and RNase A (10 µg/ml) were added to reduce viscosity. After 45 min, the sample was sonicated on ice (six 10-s bursts, with 20-s cooling periods) to ensure complete lysis and then centrifuged (30 min, 25,000 × g, 4 °C) to remove cellular debris.

Immobilized Metal Affinity Chromatography-- One ml of Ni2+-NTA-agarose (50% slurry; Qiagen) was added to every 6 ml of cleared lysate, and batch binding was allowed to proceed for 1 h (4 °C, with shaking at 200 rpm). The lysate/Ni2+-NTA-agarose mixture was loaded into a disposable plastic column (5 ml), and, once settled, the cleared lysate was allowed to flow through. The column was washed with 3 × 5 ml of wash buffer (50 mM sodium phosphate, pH 8, 350 mM NaCl, 30 mM imidazole, 0.5% Triton X-100), and then the His6-tagged WaaP was eluted in 4 × 0.5 ml of elution buffer (50 mM sodium phosphate, pH 8, 250 mM NaCl, 200 mM imidazole, 0.5% Triton X-100). The second 0.5-ml elution typically contained the highest concentration of WaaP.

High Performance Anion Exchange Chromatography-- The enzyme eluted from the Ni2+-NTA-agarose column was further purified by using subtractive anion exchange chromatography to bind the remaining contaminating proteins. Protein eluted from the Ni2+-NTA-agarose column was first exchanged into a lower ionic strength loading buffer (25 mM sodium phosphate, pH 8, 150 mM NaCl, 0.5% Triton X-100) using PD-10 columns (containing Sephadex G-25) as recommended by the manufacturer (Amersham Pharmacia Biotech). The sample was then applied (0.8 ml/min) to a Beckman Q HyperD 20 column (equilibrated in 40 mM sodium phosphate, pH 8, 50 mM NaCl), and the WaaP-containing flow-through was collected in 0.5-ml fractions.

Assay Conditions and Analysis-- Unless otherwise indicated, reaction mixtures contained 50 mM Tris, pH 8.5, 17.5 mM MgCl2, 0.5% Triton, 0.5 mM dithiothreitol, 1 µCi of [gamma -33P]ATP, diluted to 1 µM with cold ATP, and 1 mM LPS. The enzyme source was added last to initiate the reaction, and comprised 10% of the reaction volume. Reactions were incubated at 35 °C for 30 min and then stopped by the addition of an equal volume of stop solution (1 M acetic acid, 0.1 M sodium phosphate monobasic, 25 mM MgCl2). LPS was collected in a pellet by centrifugation (15 min, 14,000 rpm) and washed three times in stop solution to remove unincorporated radiolabel. (For each wash, the LPS pellet was completely resuspended by vortexing for about 5 min.) After the third wash, the LPS pellet was resuspended in water, and the incorporated radiolabel was quantified in EcoLite scintillation fluid (ICN) using a Packard TRI-CARB Liquid Scintillation Analyzer (model 2000CA).

Analysis of Reaction Products by Gel Filtration Chromatography-- Radiolabeled LPS was pooled from 10 reaction mixtures and resuspended in 250 µl of 2% acetic acid. The LPS/acetic acid suspension was then incubated at 100 °C for 90 min to cleave the acid-labile ketosidic linkages involving the anomeric carbon of Kdo (i.e. between KdoI and the nonreducing GlcN of lipid A and between KdoII and KdoI). Insoluble lipid A was removed by centrifugation (15 min, 14,000 rpm). The water-soluble core oligosaccharides were then applied to a column of Sephadex G-25 (1 × 40 cm) with water as eluent. The column void volume and included volume were determined using blue dextran and KCl, respectively. Eluted fractions (0.85 ml) were collected and analyzed for radioactivity by liquid scintillation counting (described above).

Site-directed Mutagenesis-- The gene encoding the His6-tagged derivative of WaaP (together with its ribosome binding site) was excised from the WaaP expression plasmid pWQ910 (described above) as an XbaI-HindIII fragment and cloned between the XbaI and HindIII sites of pBAD18 (17). The resulting plasmid, pWQ912, was used as the template for PCR-based site-directed mutagenesis. It was necessary to move waaP out of plasmid pWQ910 for these experiments to allow for regulated expression of WaaP and its mutated versions in strain CWG296 (waaP) (4), which is not lysogenic for lambda DE3 expressing the T7 RNA polymerase. Complementary oligonucleotide primers were designed containing the desired mutation (resulting in the replacement of Asp162 with Ala in WaaP): 5'-CAGATATAACAGGCACGGTGGTTAATGCCC-3'; 5'-GGGCATTAACCACCGTGCCTGTTATATCTG-3' (the underlined bases indicate the mismatch between primer and template sequences). PCR amplification of the entire pWQ912 plasmid was performed with PwoI DNA polymerase (Roche Molecular Biochemicals) as follows: 1) one cycle at 95 °C for 2 min; 2) 16 cycles at 95 °C for 30 s, 55 °C for 1 min, and 68 °C for 11 min. The amplified plasmid was purified using the QIAquick PCR Purification Kit (Qiagen) and then digested with DpnI (specific for the methylated parental DNA template, thus selecting for the plasmid containing the mutation). The DpnI-digested sample was transformed into electrocompetent E. coli DH5alpha cells and plated on LB-agar plates containing ampicillin (100 µg/ml). Plasmid purified from the resulting colonies (pWQ913) was sequenced to ensure that only the desired mutation had been introduced into the waaP coding region.

In Vivo Complementation-- The plasmids encoding the parental and mutated (D162A) versions of WaaP (pWQ912 and pWQ913, respectively) were transformed into CWG296 (waaP) by electroporation, and WaaP expression was induced by the addition of 0.02% L-arabinose. To determine the ability of WaaP (D162A) to complement the antibiotic supersensitivity of the waaP mutation in CWG296, SDS and novobiocin sensitivity testing were performed as described previously (4).


    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Expression of WaaP-- The addition of an N-terminal His6 tag onto WaaP was shown to have no effect on its ability to complement the CWG296 waaP mutant phenotype (data not shown), thus validating the use of the His6 tag for purification. Expression of the His6-tagged WaaP derivative from plasmid pWQ910 resulted in a significant amount of protein, as seen by the appearance of an intense band located at ~33 kDa in Coomassie Blue-stained SDS-polyacrylamide gels (Fig. 2). The predicted molecular mass of WaaP is 31,049 Da, and the mass of His6-tagged WaaP is 33,212 Da, a difference that could be observed by SDS-PAGE (data not shown). The identity of the overexpressed protein was further confirmed by Western immunoblotting with monoclonal antibodies specific for the His6 tag (data not shown). Reactivity with the His6 tag-specific monoclonal antibody also demonstrated that the N terminus of the protein was not processed post-translation, consistent with the prediction from the waaP sequence. Unfortunately, when the WaaP-expressing cells were lysed and fractionated by differential centrifugation (into insoluble matter, membranes, and cytosol), the recombinant protein was found almost exclusively in insoluble inclusion bodies.



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Fig. 2.   Coomassie Blue-stained SDS-polyacrylamide gel of fractions generated during the purification of WaaP. Lane 1, molecular weight standards; lane 2, preinduced whole-cell lysate; lane 3, arabinose- and IPTG-induced whole-cell lysate; lane 4, insoluble material from cell lysate; lane 5, cleared lysate; lane 6, flow-through from Ni2+-NTA-agarose affinity chromatography column; lane 7, protein eluted from Ni2+-NTA-agarose affinity chromatography column; lane 8, flow-through from anion exchange chromatography column.

Many standard variables were tested to increase the yield of soluble WaaP (e.g. growth medium, growth temperature, time of induction, duration of induction, inducer concentration, removal of the His6 tag, expression from different promoters, and compatible solute accumulation); however, none of these treatments provided substantial improvement in the yield of soluble protein. We therefore created a plasmid that would allow for the coexpression of the E. coli GroES and GroEL chaperones (based on the work of Ref. 15), with the hope that this would increase the yield of soluble WaaP. Since the WaaP expression plasmid pWQ910 had a kanamycin resistance marker and a pBR origin of replication, we based the chaperone expression plasmid on pBAD33, which has a chloramphenicol resistance marker and a compatible origin of replication from pACYC (17). Not only can these two plasmids be maintained simultaneously in a bacterial cell, but expression from each is controlled independently (expression from pBAD plasmids requires L-arabinose induction, while expression from pET plasmids requires IPTG induction). The ability to express chaperones independently of WaaP allowed for accumulation of chaperones prior to the induction of WaaP. Using this coexpression system, it was possible to achieve sufficient soluble protein expression to allow for the purification of the protein under native conditions, although the majority of protein still remained insoluble (Fig. 2). Subcellular fractionation also indicated a tendency for the recombinant His6-tagged WaaP to associate with membranes, so Triton X-100 was added to the lysis buffer to help extract any membrane-associated WaaP and achieve the highest possible yield of soluble protein. The association of recombinant WaaP with membranes was not unexpected; indeed, the enzymes involved in LPS core assembly are predicted to function as peripheral membrane proteins at the cytoplasmic face of the inner membrane (8), where they have access to both their cytoplasmic substrates (sugar nucleotides or ATP) and their lipid acceptor molecules.

Purification of WaaP-- The incorporation of an N-terminal His6 tag onto the recombinant WaaP protein allowed for the use of immobilized metal affinity chromatography as an initial purification step (Fig. 2). Given that the predicted pI of the WaaP protein is very high (~9.8), the next logical step was to use cation exchange chromatography. However, the protein tended to precipitate rapidly during the buffer exchange required prior to loading on the cation exchange column. To circumvent this problem, the enzyme was further purified by using subtractive anion exchange chromatography to bind the remaining contaminating proteins, allowing WaaP to be collected in the flow-through (Fig. 2). In this way, the protein could be kept in a higher ionic strength buffer to minimize the precipitation problem. The purity of the protein collected in the flow-through from the anion-exchange column was over 95% as judged by SDS-PAGE (Fig. 2). After this final step, the specific activity of the His6 tagged WaaP was 25.8-fold higher than the crude extract, with a total activity yield of 0.53% (Table I).


                              
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Table I
Purification of His6-tagged WaaP from BL21(DE3)pWQ910/pWQ911

Of note, the presence of 10 mM beta -mercaptoethanol in the lysis buffer (see "Experimental Procedures") was found to greatly increase the activity of the purified protein (greater than 10-fold; data not shown). Further, the purified enzyme could be stored in 50% glycerol at -20 °C, without significant loss of activity for over 30 days.

Effect of Mg2+ Concentration and pH on Kinase Activity-- The kinase activity of purified WaaP was measured using a range of Mg2+ concentrations from 0 to 25 mM, as shown in Fig. 3A. The enzyme displays an absolute requirement for Mg2+ that cannot be substituted with Ca2+ (data not shown). Maximal kinase activity was observed at 17.5 mM MgCl2. The activity of WaaP was then tested over a pH range from 4.5 to 9.0, as shown in Fig. 3B. The kinase was shown to be inactive at pH 4.5, with maximal activity between pH 8.0 and 9.0. 



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Fig. 3.   Effect of Mg2+ concentration and pH on kinase activity. A, WaaP activity was measured under standard assay conditions over a range of MgCl2 concentrations, as indicated. Mg2+ was absolutely required for activity, and optimal activity was observed at 17.5 mM MgCl2. B, the pH dependence of the reaction was demonstrated under standard assay conditions using 75 mM sodium acetate (pH 4.5, 5.0, and 5.5), 75 mM sodium phosphate (pH 6.0, 6.5, 7.0, 7.5, and 8.0), or 75 mM diethanolamine-HCl (pH 8.0, 8.5, and 9.0). The enzyme was inactive at pH 4.5, and optimally active between pH 8.0 and 9.0.

Kinetic Properties of Purified WaaP-- The activity of WaaP was shown to be linearly dependent on both time (from 0 to 30 min) and protein concentration (from 0-200 µg/ml) (data not shown). When the concentration of ATP in the reactions was held constant at 2.5 mM and the concentration of LPS was varied (Fig. 4A), the Km for the LPS acceptor was calculated to be 76 µM. Likewise, when the concentration of LPS acceptor was held constant at 1 mM and the concentration of ATP was varied (Fig. 4B), the Km for ATP was determined to be 0.13 mM. In both cases, the apparent Vmax for the reaction was ~3.7 nmol/min/mg.



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Fig. 4.   Kinetic properties of purified WaaP. Standard assay conditions were used in these experiments, except that the substrate concentrations were varied as indicated. A, the concentration of ATP was held constant at 2.5 mM, and the concentration of LPS was varied. The apparent Km for the LPS acceptor was calculated to be 76 µM. B, the concentration of LPS acceptor was held constant at 1 mM, and the concentration of ATP was varied. The apparent Km for ATP was determined to be 0.13 mM. In both cases, the apparent Vmax for the reaction was ~3.7 nmol/min/mg. Values were determined using EnzFitter version 1.05 software (Biosoft), and lines were drawn using a nonlinear least squares fitting to the following equation: V = (Vmax × [S])/(Km + [S]).

Characterization of the Reaction Products by Gel Filtration Chromatography-- When the acceptor LPS purified from our defined waaP mutant strain, CWG296, is run on an SDS-polyacrylamide gel and stained with silver, two distinct molecular weight species are observed (see Fig. 5, inset). The structure of the larger of these two species is known to consist of complete lipid A-core lacking the usual modifications (phosphate, PEtN, and HepIII) on the heptose region of the core (see Fig. 1) (4). The smaller molecular weight species is a truncated form of the larger, terminated after GlcI (see Fig. 1) as determined by comigration with LPS from a defined waaO mutant on SDS-PAGE (data not shown). To determine whether WaaP was capable of phosphorylating both forms of LPS from the waaP mutant strain, the LPS reaction products were first hydrolyzed in mild acid to remove the insoluble lipid A portion of the molecule. (Treatment with 2% acetic acid at 100 °C for 2 h cleaves the acid-labile ketosidic linkages involving the anomeric carbon of Kdo: between KdoI and the nonreducing GlcN of lipid A and between KdoII and KdoI.) After hydrolysis and centrifugation, ~95% of the radioactivity was recovered in the supernatant as soluble core oligosaccharides (data not shown). The soluble core oligosaccharides were then separated on a column of Sephadex G-25 (Fig. 5). Two distinct peaks of radioactivity were observed within the fractionation range of the column (Mr of 100-5000 for dextrans), indicating that WaaP is indeed capable of phosphorylating both the larger and smaller molecular weight LPSs isolated from the waaP mutant strain (Fig. 5). The very small third peak that occurs at the column-included volume probably corresponds to a small amount of 33P being hydrolyzed from the reaction products during the acid hydrolysis.



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Fig. 5.   Separation of radiolabeled core oligosaccharides by chromatography on Sephadex G-25. Oligosaccharides were generated by mild acid hydrolysis of the LPS reaction products from assays performed under standard conditions. The two peaks of radioactivity correspond to the core oligosaccharides from the two major LPS species that are evident in SDS-PAGE profiles of waaP mutant LPS (inset).

Site-directed Mutagenesis-- Alignment of the predicted WaaP proteins from E. coli and S. enterica shows greater than 80% identity (90% similarity), while the Pseudomonas aeruginosa WaaP homolog is ~55% identical (70% similar) to those of E. coli and S. enterica (data not shown). Of particular note, residues 159-171 (NHRDCYICHFLLH) and 184-192 (SVIDLHRAQ) (E. coli residue numbers) are absolutely conserved among all of the WaaP homologs. On the other hand, alignment of the E. coli WaaP protein with eukaryotic protein kinases typically shows only about 10-15% identity (25-30% similarity) (Fig. 6). It is interesting, however, that the majority of conserved amino acids in Fig. 6 coincide with residues that are known in eukaryotic protein kinases to be important for catalysis, based on both sequence similarity (18) and crystal structure data (19, 20). To test the biological significance of these sequence similarities, we engineered a nucleic acid change into the waaP open reading frame in plasmid pWQ912 that resulted in the replacement of Asp162 with Ala (plasmid pWQ913). We chose Asp162 specifically because it is absolutely conserved among all of the known WaaP homologs (within the sequence NHRDCYICHFLLH; see above), and because sequence similarity predicts that it functions as the WaaP catalytic base. In this role, the WaaP Asp162 residue would accept a proton from the 4-OH of HepI, thus leaving an oxyanion capable of nucleophilic attack at the gamma -phosphate of ATP. Recent data have questioned the proposed general base role of this residue in eukaryotic protein kinases (21), but there is no doubt that it is essential for catalysis. The replacement of Asp162 with Ala would thus be expected to completely abrogate WaaP activity. When plasmid pWQ913 encoding WaaP (D162A) was introduced into our defined waaP mutant strain, CWG296, it was unable to complement the mutant's antibiotic and detergent supersensitivity (data not shown). Further, purified WaaP (D162A) showed negligible kinase activity in our in vitro assay system (data not shown), clearly indicating that Asp162 is an essential residue.



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Fig. 6.   Alignment of WaaP from E. coli to cAMP-dependent protein kinase (cAPK) from Mus musculus (mouse). Identical residues are indicated by single letter code, and similar residues are indicated by a plus sign. The WaaP and cAPK sequences show only 12% identity (23% similarity) over the length of the alignment; however, the following list denotes potentially important similarities. 1) cAPK residues 50-55 contain the nucleotide-binding motif GXGXXG, of which Gly 52 (Gly44 in WaaP) is an invariant residue among eukaryotic protein kinases. 2) cAPK residue Lys72 (Lys58 in WaaP) is an invariant residue that forms a salt bridge with the invariant residue Glu91 (Glu70 in WaaP). The cAPK Lys72 residue is also involved in hydrogen bonding to the alpha - and beta -phosphates of ATP. 3) cAPK residues Glu121 and Glu127 (Glu114 and Glu123 in WaaP) are involved in hydrogen bonding to adenosine. 4) cAPK residue Asp166 (Asp162 in WaaP) is an invariant residue that potentially functions as the enzyme's catalytic base and is shown in this study to be essential for WaaP activity. 5) cAPK residue Asp184 (Asp187 in WaaP) is an invariant residue required to orient the gamma -phosphate of ATP for efficient transfer to the substrate. The potential roles of conserved residues in cAPK were derived from crystal structure data (19, 20).



    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Phosphorylation of the LPS core region is essential for outer membrane stability in E. coli and S. enterica and has been studied over the course of many years (reviewed in Refs. 7, 8, and 22). Indeed, the activity of an LPS core phosphorylating enzyme was reported as early as 1969, using acceptor LPS from an undefined phosphate-deficient mutant of S. enterica and crude lysates from wild-type bacteria as the enzyme source (23). Unfortunately, the kinase implied by these early studies was never actually identified; nor was it confirmed that ATP was the direct donor of phosphate. More recently, a single gene called rfaP (now waaP) was implicated in all of the phosphoryl modifications of the E. coli LPS core (24). However, the conclusions from these studies were necessarily limited by their reliance on strains with polar mutations and also in hindsight by the strict sequential action of the two core kinases. The direct involvement of the waaP gene product in the phosphorylation of the E. coli LPS core at HepI was only recently established with confidence, based on the LPS core structures resulting from defined waaP, waaQ, and waaY mutations (4). Since then, mutation of waaP in S. enterica was shown to result in the same LPS defect (5), and the homolog of WaaP in P. aeruginosa was shown to complement a waaP defect in S. enterica (6).

The data presented here provide the first biochemical characterization of WaaP activity and prove unequivocally that WaaP is indeed an LPS core heptose kinase. Purified protein was used to determine the optimal assay conditions and catalytic properties of WaaP. Optimal activity was observed between pH 8.0 and 9.0, and like most kinases, the enzyme displayed an absolute requirement for Mg2+. The apparent Km values for the WaaP-catalyzed reaction were calculated to be 0.13 mM for ATP, and 76 µM for LPS (determined at saturating concentrations of the second substrate). In this study, the site of WaaP phosphorylation was only localized to the soluble core oligosaccharide portion of the LPS that was released by mild acid hydrolysis. However, given the known structure of the E. coli core (25, 26) and the core structures resulting from defined waaY and waaP mutations (loss of phosphate on HepII and loss of phosphate on both HepII and HepI, respectively) (4), it is possible to conclude that WaaP is in fact a heptose-specific LPS core kinase, and is responsible for phosphorylation at HepI. Further, the WaaP enzyme is known to catalyze the addition of phosphate to only HepI (and not HepII), since LPS purified from a defined waaY mutant strain could not serve as an acceptor for the WaaP enzyme (data not shown).

These data thus help to clarify the sequence of events in core biosynthesis that result in the observed heterogeneity of phosphoryl modifications on HepI (see Fig. 1). It was previously speculated that all LPS molecules might initially be modified by the addition of PPEtN to HepI, with a subset of these molecules subsequently undergoing cleavage to liberate PEtN and leave just phosphate (8). Now, however, with the assignment of WaaP as the HepI kinase, the more likely scenario is clearly that WaaP catalyzes the transfer of phosphate to HepI with a subset of molecules being further substituted at this phosphate residue with PEtN by a currently unidentified enzyme (see Fig. 1). The determination of the gene responsible for this PEtN modification is of particular interest, given the potential role of PEtN core modification in resistance to polymyxin and modulation of surface charge (11, 12).

Given the importance of WaaP to membrane stability (4) and its requirement for virulence in vivo (5), inhibitors to the WaaP-catalyzed reaction could potentially function as novel antimicrobial compounds. As a starting point for the design of inhibitors, it would be useful to know the minimal structure that is capable of being phosphorylated by WaaP. The finding that WaaP is capable of phosphorylating both the high and low molecular weight LPS species isolated from a waaP mutant strain (Fig. 6) helps to better define this minimal acceptor; apparently, the outer core glycoses distal to GlcI are not strictly required (see Fig. 1). This finding is supported by the recent report that LPS isolated from a defined waaG mutant strain (truncated after HepII; see Fig. 1) carries the phosphate modification on HepI, albeit at only 40% of wild-type levels (10). Interestingly, the HepII phosphate is completely absent in the waaG mutant LPS (see Fig. 1), indicating that the putative kinase encoded by waaY requires the presence of GlcI as part of its acceptor substrate (10).

Finally, it is noteworthy that WaaP was never assigned putative kinase function based on sequence homology alone, although the sequence of the E. coli waaP gene has been known for some time (27), and homologs have since been identified in S. enterica (28) and P. aeruginosa (6). The complicating factor has always been that the primary sequences of these predicted proteins bear such limited semblance to characterized kinases that similarities cannot be detected with typical BLAST searches (29). However, with the advent of the PSI-BLAST search tool (30) it was observed that WaaP did show homology to kinases when considering only key residues throughout the length of the entire protein (4). We tested the biological significance of one of these key similarities by specifically mutating the WaaP Asp162 residue, predicted by homology to function as the enzyme's catalytic base. As expected for mutation of a residue essential to catalysis, the WaaP (D162A) protein showed no activity in vivo or in vitro. It is suggested, therefore, given this conservation of catalytic residues, that WaaP may have a similar mechanism of action to other eukaryotic protein kinases. We are currently pursuing structural studies to garner further evidence in support of this hypothesis. Interestingly, the crystal structure of the APH(3')-IIIa aminoglycoside kinase from Enterococcus, which catalyzes the phosphorylation of a broad spectrum of aminoglycoside antibiotics, was recently solved (31). Like WaaP, the APH(3')-IIIa protein displays virtually no sequence similarity to eukaryotic kinases except at key residues, such as the conserved Asp mutated in this study. Yet despite this lack of sequence homology, the APH(3')-IIIa three-dimensional structure is strikingly similar to that of eukaryotic protein kinases (31).


    FOOTNOTES

* This work was supported in part through funding by the Canadian Bacterial Diseases Network (Network of Centers of Excellence) (to C. W.).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 Recipient of a Doctoral Research Award from the Canadian Institutes of Health Research.

§ A Canadian Institutes of Health Research Senior Scientist. To whom correspondence should be addressed. Tel.: 519-824-4120 (ext. 3478); Fax: 519-837-1802; E-mail: cwhitfie@uoguelph.ca.

Published, JBC Papers in Press, November 7, 2000, DOI 10.1074/jbc.M008255200


    ABBREVIATIONS

The abbreviations used are: LPS, lipopolysaccharide; PEtN, 2-aminoethyl phosphate; PPEtN, 2-aminoethyl disphosphate; Kdo, 3-deoxy-D-manno-oct-2-ulosonic acid; Hep, L-glycero-D-manno-heptose; IPTG, isopropyl-1-thio-beta -D-galactopyranoside; NTA, nitrilotriacetic acid; PAGE, polyacrylamide gel electro- phoresis; PCR, polymerase chain reaction.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES


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