INTRODUCTION

Top Abstract Introduction Materials and Methods Results Discussion References

Lipopolysaccharide (LPS) is a major nonprotein component of the outer membrane of enteric and nonenteric gram-negative bacteria (57). LPS is an amphipathic molecule consisting of lipid A and an oligosaccharide core domain. Some microorganisms also express a hydrophilic, surface-exposed O-specific polysaccharide that is found attached to the reducing end of the lipid A core (43, 57). In most cases, lipid A consists of five to seven saturated fatty acids attached to a glucosamine dimer and is responsible for the endotoxic activities of the LPS molecule (38). The core oligosaccharide can be further subdivided into inner and outer core domains. The outer core is generally made of hexoses, while the inner core oligosaccharide is composed of at least two residues of 3-deoxy-D-manno-octulosonic acid, and depending on the particular species of gram-negative bacteria, two or three residues of L-glycero-D-manno-heptose (LDHep) (19). The structure of the inner core has a high degree of conservation among enteric and nonenteric bacteria (19).

LPS plays an important role in maintaining the structural integrity of the bacterial outer membrane (35). Early studies by Tamaki et al. (47) have shown that Escherichia coli K-12 mutants lacking heptose in the LPS demonstrate hypersensitivity to hydrophobic antibiotics (such as novobiocin), detergents, and bile salts. Heptoseless E. coli K-12 mutants are also deficient in F plasmid conjugation (18) and transduction by the P1 bacteriophage (11). The collection of these phenotypes is referred to as the "deep rough" phenotype. The deep rough phenotype is related to a general defect in the assembly of outer membrane proteins, some of which are involved as receptors for conjugation and/or phage attachment (11, 18, 44), components of efflux systems (26), and F-pilus assembly (J. D. Klena, unpublished data). For other organisms, such as Haemophilus influenzae, a heptoseless mutant was found to be serum sensitive and displayed a reduced virulence in an animal model (20, 60).

Because of the structural conservation of the inner core in gram-negative bacteria, we have hypothesized that the biosynthesis pathway for LDHep is also conserved. From studies using Salmonella enterica, Eidels and Osborn (15, 16) proposed that LDHep is synthesized from sedoheptulose 7-phosphate via four steps: (i) conversion of sedoheptulose 7-phosphate into D-glycero-D-manno-heptose 7-phosphate by a phosphoheptose isomerase, (ii) formation of D-glycero-D-manno-heptose 1-phosphate by a mutase reaction, (iii) transfer of a nucleotide via a phosphodiester linkage, and (iv) epimerization of D-glycero-D-manno-heptose 1-phosphate residue of the sugar nucleotide to LDHep (Fig. 1). Further investigations resulted in the identification of ADP as the nucleotide sugar residue attached to glycero-manno-heptose in Shigella sonnei and S. enterica (24, 25). Verification of this pathway requires the identification of genes and gene products and the biochemical demonstration of their functions. The epimerase gene, gmhD (previously rfaD, see reference 40 for a discussion on the current gene nomenclature), of several microorganisms was isolated and characterized (10, 14, 34, 46, 55). Also, Brooke and Valvano (7, 8) recently identified gmhA (formerly lpcA) in E. coli and H. influenzae as the first gene of this pathway, confirming that it encodes a phosphoheptose isomerase activity. We have also shown that gmhA is highly conserved in Neisseria gonorrhoeae, Neisseria meningitidis, Helicobacter pylori, Vibrio cholerae, and Pseudomonas aeruginosa (49).

View larger version (30K): [in this window] [in a new window]   FIG. 1.   Pathway for the synthesis of ADP-L-glycero-D-manno-heptose as proposed by Eidels and Osborn (15, 16). gmhA and gmhD are the only genes whose functions have been established biochemically (7, 10).

In this study, we report the identification and molecular characterization of an E. coli gene encoding a bifunctional protein consisting of two distinct domains, both of which are required for the intermediate steps in the synthesis of ADP-LDHep. We also show that these domains function independently, and in some microorganisms they are encoded by separate genes. The predicted functions of these domains suggest a novel pathway for the synthesis of ADP-glycero-manno-heptose.

	MATERIALS AND METHODS

Top Abstract Introduction Materials and Methods Results Discussion References

Bacterial strains, plasmids, and media. Except when indicated, all chemicals used in this study were purchased from Sigma Chemical Co., St. Louis, Mo. Bacterial strains and plasmids used in this study are listed in Table 1. Bacteria were cultured in Luria broth (LB) (Sigma), supplemented when necessary with antibiotics at final concentrations of 50 µg/ml for novobiocin, 30 µg/ml for chloramphenicol, 100 µg/ml for ampicillin, and 20 µg/ml for tetracycline. S. enterica SL1102 was obtained by mutagenesis as described elsewhere (58). Gene libraries of E. coli VW187 (previously constructed in our laboratory by M. Handelsman) and VW194 were used to complement the rfaE mutants of E. coli K-12 KLC2926 and S. enterica SL1102. pMGC2 contained a 12.5-kb BamHI fragment from the E. coli VW194 chromosome cloned into the BamHI site of pACYC184. pTLS1 contained a 1.463-kb fragment from pMGC2, obtained by PCR with primers P54 and P56 (Table 2), cloned into the SmaI site of the expression vector pEX1 (37). pMAV47, containing Domain II from rfaE, was constructed by cloning a 0.5-kb fragment from pTLS1, obtained by PCR with primers PAB10542 and P56, into the SmaI site of pUC19. This strategy generates a protein fusion between Pro16 of -galactosidase with Pro305 of RfaE. pMAV51, containing only Domain I, was generated by deleting a 0.5-kb HindIII fragment of pTLS1. pMAV44 contains the chloramphenicol acetyltransferase (cat) gene from pBR325 inserted into the KpnI site of pTLS1. The cat gene was obtained by PCR amplification with primers P39 and P40. Before ligation with amplified DNA, the KpnI site in pTLS1 was treated with the Klenow fragment of DNA polymerase. The cat gene in pMAV44 is transcribed in the opposite direction from the rfaE gene. pMB10 is a 1.7-kb BamHI fragment containing the plac promoter and the rfaE gene from pTLS1 cloned into the BamHI site of pME6000. pMAV52 was obtained by subcloning a 1.2-kb BamHI/HindIII fragment containing the plac promoter and Domain I from rfaE into the BamHI/HindIII sites in pME6000. To generate pMAV53, a 0.5-kb amplification product obtained with primers P147 and P148 (both present in vector sequences flanking the DNA insert) and pMAV47 as a template was cloned into the SpeI site in pME6000. pCM200 and pCM202 were generated by PCR amplification of a 1.4-kb fragment containing the wild-type and mutated rfaE genes from S. enterica strains SL1027 and SL1102, respectively. The primers used in the reaction were P181 and P182, and the amplified fragments were cloned into the SmaI site of pEX1. Plasmid pCM204 was generated by digesting pCM200 with BamHI and inserting the linearized DNA into the BamHI site in pME6000. pCM203 was constructed in a similar manner, except that digestion was carried out with HindIII. Primers P91 (ptac promoter) and P182 were used to confirm that the inserts in these plasmids contained the S. enterica rfaE gene. pME6000 is a low-copy-number, broad-host-range cloning vector kindly provided by S. Heeb, Laboratoire de Biologie Microbienne, Université de Lausanne. pME6000 was constructed by replacing the cat gene of pBBR1MCS (27) with the tetRA genes of pVK100 (23).

Mating in R. eutrophus. Plasmids containing the different constructs in pME6000 were transferred into Ralstonia eutrophus HB3 by conjugation. The plasmids were first transformed by electroporation (13) into the E. coli K-12 strain BW19851. This strain contains an integrated RP4 plasmid carrying the necessary components to allow mobilization of recombinant plasmids containing the mob region present in vector pME6000. The donor strain BW19851, which carried the appropriate plasmids, and the recipient HB3 were mixed in equal proportions and washed once with prewarmed sterile LB. The bacterial mixture was spread on LB plates and incubated overnight at 30°C. The growth was resuspended and diluted in LB, and exconjugants were selected in LB plates containing kanamycin and tetracycline at final concentrations of 120 and 100 µg/ml, respectively.

Recombinant DNA methods. Small- and large-scale plasmid DNA extractions and DNA gel electrophoresis were conducted as described (31). Chromosomal DNA was prepared by a clear lysis method (51). Transformations were done by electroporation with a Gene Pulser apparatus (Bio-Rad Laboratories Ltd., Mississauga, Ontario, Canada) and 0.1-cm cuvettes, following the method described by Dower et al. (13). Restriction enzyme analysis and cloning were performed by following standard protocols (29). Restriction endonucleases were obtained from Roche Diagnostics, Dorval, Quebec, Canada, and Pharmacia Canada Inc., Baie d'Urfé, Quebec, Canada. Calf alkaline phosphatase, T4 DNA ligase, and polynucleotide kinase were from Roche Diagnostics. All enzymes were used as suggested by the manufacturers.

PCR. PCR were carried out with a Hybaid Omnigene temperature cycler (Interscience, Markham, Ontario, Canada) and PwoI DNA polymerase (Roche Diagnostics). The primers used in this study are listed in Table 2. Prior to cloning, amplicons were treated with polynucleotide kinase and purified by gel electrophoresis followed by extraction of the DNA band with the QIAquick gel extraction kit (Qiagen, Chatsworth, Calif.) according to the manufacturer's instructions.

LPS analysis. LPS was isolated as described by Marolda et al. (31) and analyzed by Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) as described by Schagger and von Jagow (42). Commercially cast 16% polyacrylamide gels were purchased from Novex, San Diego, Calif., and LPS bands were visualized by silver staining (31).

Bacteriophage U3. Aliquots of 400 µl of overnight E. coli cultures were added to 7 ml of warmed 0.8% LB agar containing the appropriate antibiotics. The mixture was applied as an overlay to LB media plates also containing 1.5% agar and antibiotics as necessary. Agar was allowed to harden, and a 20-µl drop of U3 bacteriophage lysate was carefully deposited on the center of the plate. When the drop was dry (after about 30 min at room temperature), plates were inverted and incubated at 37°C for 6 to 8 h. Lysis was evidenced by clear zones indicating growth inhibition.

Polypeptide analysis. In vitro transcription-translation was performed with the prokaryotic DNA-directed translation kit from Amersham with [³⁵S]methionine as recommended by the manufacturer. Polypeptides were separated by SDS-PAGE (31), followed by treatment with En³Hance (Dupont). Dried gels were exposed to Kodak X-Omat film at 80°C for 16 to 24 h.

DNA sequence analysis. DNA sequencing of plasmid constructs was carried out with an automated sequencer at MOBIX, McMaster's University, Hamilton, Ontario, Canada, and at the DNA Sequencing Facility of the Robarts Research Institute, London, Ontario, Canada. DNA and protein sequence analysis was carried out with the University of Wisconsin Genetics Computer Group package, version 9 (12). Nucleotide similarity searches were performed with the program BLAST2 (1) via the National Center for Biotechnology Information. Protein sequence alignments were produced with the program Clustal X (48) and edited with GeneDoc (K. B. Nicholas and H. B. Nicholas, Jr., 1997, GeneDoc: analysis and visualization of genetic variation [http://www.cris.com/~Ketchup/genedoc.shtml]).

Nucleotide sequence accession number. The DNA sequences of the S. enterica SL1027 rfaE and SL1102 rfae543 genes have been deposited in GenBank with the accession no. AF163661 and AF163662, respectively.

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

Isolation and characterization of the heptoseless mutant E. coli strain KLC2926. A transposon mutagenesis strategy using mini-Tn10 was utilized to isolate genes involved in the biosynthesis pathway of ADP-LDHep. Mutagenesis was conducted in the E. coli K-12 strain W1485. Since heptose-deficient mutants are usually nonviable in the presence of very low concentrations of novobiocin (49), 1,000 Tet^r colonies were screened for sensitivity to novobiocin at a concentration of 200 µg/ml. Seven Tet^r Nov^s colonies were identified. Only three of these colonies were also sensitive to novobiocin at a concentration of 50 µg/ml. One of them, KLC2926, had a mucoid appearance, a characteristic also found associated with the deep rough LPS phenotype (36). KLC2926 was also resistant to lysis by the core-specific bacteriophage U3, which recognizes a terminal galactose in the complete E. coli K-12 core (56). Therefore, mucoidity, phage U3 resistance, and high susceptibility to novobiocin suggested that KLC2926 makes a heptoseless LPS and consequently is unable to assemble the remaining sugar components of the outer core. This conclusion was supported by the electrophoretic analysis of KLC2926 LPS, which displayed a rapid migration in Tricine polyacrylamide gels comparable with that of the heptoseless LPS of strain 711 (Fig. 2A, lanes 2 and 3). None of the above-mentioned phenotypes of strain KLC2926 were complemented by transformation with plasmids pJB2, which contains the gmhA gene (7), or pJK2252, which contains the heptosyltransferases encoded by waaC and waaF (22) (data not shown). Furthermore, hybridization experiments with a core gene-specific probe indicated that mini-Tn10 was not located in the LPS core oligosaccharide gene cluster waa (data not shown), also ruling out that the mutation in KLC2926 was in gmhD or in any of the three heptosyltransferase genes mapping within this cluster (19, 43). Therefore, we concluded that the Tn10 insertion in KLC2926 defined a novel gene or genes for the synthesis of ADP-heptose.

View larger version (100K): [in this window] [in a new window]   FIG. 2.   Analysis of LPS in E. coli and R. eutrophus. (A) LPS profiles of E. coli strains. Lanes: 1, W1485 (wild type); 2, KLC2926 (rfaE::Tn10); 3, 711 (gmhA); 4, KLC2926(pTLS1); 5, KLC2926(pMAV51); 6, KLC2926(pMAV47); 7, KLC2926(pCM200); 8, KLC2926(pCM202). (B) LPS profiles of R. eutrophus strains. Lanes: 1, H16 (wild type); 2, HB3 (aut::Tn5); 3, E. coli 711 (gmhA); 4, HB3(pMB10); 5, HB3(pMAV52); 6, HB3(pMAV53); 7, HB3(pCM204); 8, HB3(pCM205). Arrows indicate the bands corresponding to polymeric O antigen. In both cases, cell lysates were separated by Tricine-SDS-PAGE and stained with silver.

The Tn10 insertion in KLC2926 is located within the rfaE gene. Preliminary attempts to clone the gene complementing the novobiocin-sensitive phenotype of KLC2926 from DNA fragments of strains W1485 and VW187 were unsuccessful. However, we were able to clone a 12.5-kb BamHI fragment from E. coli VW194, resulting in plasmid pMGC2 (Fig. 3A). KLC2926(pMGC2) was not only resistant to novobiocin but also became sensitive to bacteriophage U3 and displayed a complete lipid A core band in Tricine-SDS polyacrylamide gels, suggesting that the insert DNA carries gene or genes complementing the defect caused by the mini-Tn10 insertion in this strain (data not shown). pMGC2 in KLC2926 was unstable and yielded very little DNA upon plasmid purification. We speculated that the instability problems could be caused by recombination events involving sequences in the insert DNA near or within the locus defined by mini-Tn10. This appeared to be the case since pMGC2 was stable in the recA-defective strain DH5.

View larger version (15K): [in this window] [in a new window]   FIG. 3.   (A) Partial map of the chromosomal region cloned into pMGC2 that corresponds to the E. coli K-12 genetic map. Thick bars represent regions that were sequenced. The arrow indicates the direction of transcription of the putative ygiF-glnE-rfaE operon. (B) Map of the rfaE gene. Boxes correspond to the Domain I and Domain II regions. The positions of the various mutations are indicated: V70, the position of the Tn10 insertion in KLC2926 after the codon encoding valine 70 of the RfaE protein; TLAA, the location of the 12-bp deletion in SL1102 rfaE and the corresponding amino acids; GE, the glycine-to-glutamic acid substitution due to a base pair change (GGA to GAA). The partial maps show the various plasmids. The ColE1 vector was pEX1, and these plasmids were used in the experiments involving E. coli and S. enterica. The broad-host-range vector was pME6000, and these plasmids were used in the experiments involving R. eutrophus. The position and orientation (arrow) of the insertion of a cat cassette within the E. coli rfaE gene in pMAV44 is shown. pCM200 and pCM204 correspond to the cloned wild-type rfaE gene of S. enterica SL1027. pCM202 and pCM205 correspond to the cloned mutant rfaE543 gene of S. enterica SL1102. The box and triangle in pCM202/pCM205 denote the positions of the amino acid substitution and deletion, respectively.

The rfaE gene encodes a single protein with two distinct domains. The predicted RfaE polypeptide was compared with the BLAST program (1) to amino acid protein sequences deposited in GenBank. It was strikingly apparent that the predicted protein contained two distinct regions. The region from amino acid 1 to 318 (Fig. 3B Domain I) showed conservation with members of the ribokinase family (Table 3) (5). The amino acid conservation was especially strong in critical regions for enzymatic activity, such as an aspartic acid located in an anion "hole" in the E. coli ribokinase that is essential for catalysis (Fig. 4) (45). Other regions of the enzyme containing amino acids that make contact with the sugar substrate are also conserved in Domain I (Fig. 4). The C-terminal region of the RfaE protein (Fig. 3B) from amino acid 344 to 477, designated Domain II, had features in common with the cytidylyltransferase superfamily (Table 3) (4). This family is a member of a large group of diverse enzymes hydrolyzing the alpha-beta pyrophosphate bond in nucleoside triphosphates, which have been known to show the characteristic HXGH motif of the class I aminoacyl-tRNA synthetases (Fig. 5) (4, 54).

View larger version (88K): [in this window] [in a new window]   FIG. 4.   Amino acid sequencing alignment of Domain I of RfaE proteins from E. coli (RFAE_ECDI), S. enterica serovar Typhi (RFAE_TYPHI), S. enterica serovar Typhimurium strains SL11027 (RFAE_SL1027) and SL1102 (RFAE_SL1102), and E. coli ribokinase (RBSK_ECOLI). The alignment was produced with CLUSTAL W. Identical residues in all five proteins are boxed in black. *, residues that are also conserved in other members of the ribokinase family (45); r, a, and h, residues in the ribokinase that bind ribose (r) or ADP (a) via hydrogen bonds or that make van der Waals contacts with ribose and/or ADP (h) (for more details on the structure of ribokinase see reference 45). The arrow indicates the substitution of glycine (boxed in grey) by glutamic acid (bold) in the SL1102 RfaE mutant of SL1102. , the deletion of four amino acids in the SL1102 RfaE mutant.

View larger version (20K): [in this window] [in a new window]   FIG. 5.   Comparison of the known structure of the tyrosyl tRNA synthetase TyrTS (1tya_E) with Domain II. Secondary structure elements in TyrTS (6) are shown below the sequence (black bar, helix; gray bar, strand). The numbers indicate the helices and strands placed around the substrate binding site (according to reference 4).  and , conserved residues that line the ATP binding pocket. The flexible loop KFGKT, also involved in catalysis, and its corresponding segment in Domain II, STTNI (conserved in all members of cytidylyltransferases), are also boxed.

View larger version (73K): [in this window] [in a new window]   FIG. 6.   Autoradiograph showing rfaE expression by in vitro transcription-translation. Proteins were labeled with [14C]leucine as indicated by the supplier (Amersham) and separated by SDS-PAGE. M, [14C]methylated molecular weight markers (bovine serum albumin [69 kDa], ovalbumin [46 kDa], carbonic anhydrase [30 kDa], and lysozyme [14 kDa]). Lanes: 1, pMAV44; 2, pTLS1; 3, Kit's positive control. Arrowheads indicate unique polypeptides of ca. 55 and 38 kDa.

Expression of Domain I is sufficient to complement the Tn10 mutation in E. coli KLC2926. Since RfaE has two domains, it was important to investigate whether one or both of these domains are involved in the synthesis of ADP-heptose. For this purpose, plasmid pMAV51, containing a deletion eliminating Domain II, and pMAV47, expressing only Domain II, were constructed (Fig. 3B). In the case of pMAV47, a protein fusion of Domain II was generated by fusing this domain with Pro16 from the LacZ protein, thus adding 16 N-terminal amino acids of LacZ and driving the expression of Domain II by the lac promoter. Both plasmids were transformed into KLC2926 and the LPS phenotypes examined before. Only pMAV51 could complement the mutant phenotype as indicated by restoration of a wild-type lipid A-core band (Fig. 2A, lane 5) and the sensitivity to bacteriophage U3. In contrast, pMAV47 could not complement the heptoseless phenotype of KLC2926 (Fig. 2A, lane 6). These results indicate that Domain I is sufficient to complement the LPS-deficient phenotype. Therefore, Domain II is either not involved in lipid A-core synthesis or, alternatively, the Tn10 insertion is nonpolar on the expression of this domain. This may be due to the presence of at least three intragenic initiation codons preceded by potential ribosomal binding sites in the sequence downstream from the site of the Tn10 insertion.

The S. enterica rfaE mutation has an in-frame deletion and an amino acid substitution in Domain I. In a previous work, Lee et al. (28) reported that pHI3, a plasmid carrying the rfaE gene of H. influenzae, complements the rfaE543 mutation of S. enterica SL1102. The insert in pHI3 contains the entire Domain I homologue-encoding sequence but only a portion of that for Domain II. Following transformation of S. enterica SL1102 with pMAV51 and pMAV47, we determined that only pMAV51 could complement the deep rough LPS phenotype of this strain (data not shown). To establish the nature of the S. enterica rfaE mutation, we cloned and determined the DNA sequence of rfaE543. We took advantage of the recent genomic sequencing projects of S. enterica serovar Typhimurium and Serovar Typhi. A search of the available sequences released by the Sanger Centre resulted in the identification of the rfaE gene in S. enterica serovar Typhi, which we used to obtain primers P181 and P182 (Table 2). These primers amplified a 1,495-bp fragment from chromosomal DNA of S. enterica SL1102 and SL1027. Cloning of the SL1027 amplicon into pEX1 (pCM204) resulted in complementation of the heptoseless phenotype of E. coli KLC2926, while the corresponding plasmid containing the mutated rfaE543 allele from S. enterica SL1102 (pCM205) did not complement (Fig. 2A, lanes 7 and 8, respectively). Comparison of DNA sequences of both rfaE genes revealed an in-frame deletion of 12 bp within rfaE543, resulting in the loss of four amino acids, Thr-Leu-Ala-Ala, located after Ala272 and within Domain I of the mutated rfaE gene product. Additionally, there was one base pair change, which resulted in the replacement of Gly236 by glutamic acid. Leu274, absent in the mutated rfaE gene product, is a conserved residue present in -helix 8 of the ribokinase (Fig. 4), next to one of the two regions critical for substrate binding (45). Also, the replaced Gly236 is near another region for substrate binding. The fact that both mutations are localized within Domain I and the reading frame is unaltered strongly suggests that Domain II is expressed. At the same time, Domain I is important for the synthesis of ADP-heptose. This was consistent with the inability of pCM202, which carries the mutated rfaE gene of S. enterica, to complement the phenotype of E. coli KLC2926 (Fig. 2A, lane 8).

Domain II functions independently of Domain I in synthesis of LPS. Since the expression of Domain II was presumably not affected in the rfaE mutations studied, it was important to examine its role in LPS synthesis. For this purpose, we turned our attention to the Ralstonia (Alcaligenes) eutrophus strain HB3 containing a Tn5 insertion in the aut gene (17). This gene encodes a protein with strong amino acid conservation with Domain II. The aut::Tn5 mutation has been associated with pleiotropic effects, including autotrophic growth and changes in cell morphology and colony appearance (17). An examination of the available sequence flanking aut indicates no homologies with other known genes (data not shown). A comparison of the LPS electrophoretic profile of the aut::Tn5 mutant with that of the wild-type strain, H16 (Fig. 2B, lanes 1 and 2), revealed that the mutant has a rapid migrating lipid A-core band that comigrates with that of the E. coli K-12 heptoseless lipid A-core. H16 also exhibits slow migrating bands in the gel corresponding to polymeric O antigen which are not present in HB3. Furthermore, HB3 is novobiocin sensitive while H16 remains resistant to this antibiotic. Therefore, we concluded that the aut gene is involved in LPS synthesis. To confirm whether the E. coli rfaE gene can complement the function of aut in R. eutrophus, we subcloned the 1.7-kb BamHI fragment of pTLS1 into the vector pME6000 and mobilized this construct, pMB10, to strain HB3 by conjugation. The results indicate that this strain displayed a normal appearance, regained resistance to novobiocin, and expressed a complete LPS (Fig. 2B, lane 4). In contrast to the results with E. coli KLC2926, pMAV52 (encoding Domain I) failed to complement the aut mutation (Fig. 2B, lane 5), while pMAV53 (encoding Domain II) did complement, although a small proportion of LPS molecules remained heptoseless (Fig. 2B, lane 6). We concluded from these experiments that Domain II also functions in the synthesis of lipid A-core, and its function is independent from the activity mediated by Domain I.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

In this study, we have identified a mini-Tn10 mutation in E. coli K-12 associated with heptoseless LPS that corresponds to an insertion in the rfaE gene. The predicted polypeptide consisted of two distinct domains clearly localized to two different regions. The N-terminal portion of RfaE showed features described for the ribokinase family, and it was designated Domain I. The RfaE C-terminal region had features in common with the cytidylyltransferase superfamily, and it was designated Domain II.

The presence of two distinct domains in the RfaE protein prompted us to investigate their role in the synthesis of heptose. For this purpose, each domain was cloned and expressed separately. Complementation experiments were conducted with E. coli KLC2926 (rfaE::Tn10) and S. enterica SL1102 (rfaE543). Our data showed that expression of Domain I was sufficient to complement the heptoseless phenotype of both mutant strains and demonstrated that the mutations were located in the sequence encoding this domain. These experiments did not elucidate the possible involvement of Domain II in LPS synthesis. We approached this question by conducting another set of complementations with the aut::Tn5 mutation in R. eutrophus. Freter and Bowien (17) previously described mutations in a R. eutrophus locus, designated aut, which conferred a pleiotropic phenotype characterized by slow heterotrophic growth on substrates catabolized via the glycolytic pathway and by an altered colony morphology. We noticed that the predicted amino acid sequence of the Aut protein was highly conserved as compared to the sequence of Domain II of RfaE. However, the aut gene does not have an N-terminal portion encoding a protein similar to Domain I. Since, in contrast to the transparent and round appearance of the wild-type colonies, aut mutant colonies are whitish-opaque and have rough edges, we hypothesized that aut could be involved in LPS synthesis. A comparison of the LPS profile of the R. eutrophus strain HB3 (aut::Tn5) with that of the wild-type strain suggested that HB3 produced a heptoseless LPS core and also lacked O antigen. Furthermore, only plasmids expressing the E. coli rfaE Domain II were capable of complementing the LPS phenotype of the HB3 mutant. Taken together, our data demonstrate that the R. eutrophus aut gene encodes a protein involved in core LPS synthesis, which is functionally equivalent to the RfaE Domain II. More importantly, these experiments provided a biological demonstration that E. coli rfaE encodes a bifunctional protein. Unequivocal proof of the bifunctional nature of this protein will require in vitro biochemical experiments clearly demonstrating that RfaE can actually catalyze two separate reactions. The purification of the complete RfaE protein as well as each of its separate domains is underway in our laboratory.

The rfaE genes in E. coli and S. enterica were found downstream of glnE, and the same DNA strand contains both genes. Because of the sequence conservation and gene organization in both microorganisms, rfaE may have been acquired before E. coli and Salmonella diverged. This is in agreement with the G+C content of the rfaE DNA sequence, which is very similar to the average G+C content of E. coli and S. enterica. In contrast, the rfaE homologues in other nonenteric bacteria are not located in the vicinity of glnE. Also, in these cases the G+C content of rfaE is similar to that of the corresponding genus and species. Therefore, although it is difficult to trace the evolutionary path of rfaE, the available evidence suggests that this gene is evolutionarily older than the genes for the synthesis of outer core oligosaccharide components and O antigens (39, 43), in agreement with the conserved nature of the LPS inner core structure. A search for rfaE homologues in the available genomic databases of gram-negative bacteria showed that in Campylobacter jejuni, H. influenzae, H. pylori, P. aeruginosa, Streptomyces coelicolor, V. cholerae, and Yersinia pestis, the gene encodes Domain I and II. Separate genes encoding each domain may be found in N. gonorrhoeae, N. meningitidis, Bordetella pertussis, and Burkholderia pseudomallei, but the reasons for this separation are not obvious.

The intermediate steps of the biosynthesis pathway for ADP-LDHep are not completely elucidated. Eidels and Osborn (15, 16) postulated a pathway involving the isomerization of sedoheptulose 7-phosphate into glycero-manno-heptose-7 phosphate followed by the activity of a phosphomutase responsible for the transfer of the phosphate residue from carbon 7 to carbon 1, resulting in glycero-manno-heptose 1-phosphate (Fig. 1). Transfer of this sugar to ADP and a subsequent epimerization step complete this pathway. Since the genes involved in the isomerization and epimerization reactions have been previously identified and characterized (7, 8, 10) and the heptosyltransferase genes are also known (19), we conclude that rfaE must be involved in the intermediate steps of ADP-LDHep biosynthesis. However, the fact that Domain I is a putative kinase from the ribokinase family is difficult to reconcile with its role as a phosphomutase. The ribokinase family includes a large number of proteins whose function is the phosphorylation of sugars at positions 1 or 6 in the case of hexoses and at positions 1 or 5 in the case of riboses. None of the enzymes currently classified within the ribokinase family have phosphomutase activity. Recently, the crystal structure of the E. coli ribokinase in complex with ribose and dinucleotide has been determined (45), resulting in the identification of the critical amino acids for kinase activity. The majority of these residues are all conserved in Domain I, especially an aspartic acid located in an anion hole that is essential for catalysis (45). If Domain I is a sugar kinase, its role in the synthesis of ADP-LDHep as well as in the pathway of synthesis are not clear. One possibility is that the pathway lacks a phosphomutase reaction and a sugar kinase phosphorylates C1. This reaction, however, would result in glycero-manno-heptose-1,7-diphosphate, and there is no evidence of the existence of this sugar or its derivative ADP-glycero-manno-heptose 7-phosphate. Although some of the heptoses in the core are phosphorylated on carbon 7 (38), the phosphorylation appears to be due to the activity of enzyme(s) encoded by the waa cluster and probably occurs after the heptose residue has been incorporated onto the LPS core (19). Alternatively, Domain I may have phosphomutase activity, and the structural similarity to ribokinase may only be due to similar features involved in the recognition of the sugar phosphate. However, this would not explain the strong conservation of the kinase catalytic site in RfaE. If a sugar kinase step exists for the synthesis of glycero-manno heptose 1-phosphate, we have to postulate yet another step involving dephosphorylation of the 7-phosphate precursor. Thus, it is possible that instead of a phosphomutase, an unidentified sugar phosphate phosphatase acts to remove the phosphate in position 7 from either glycero-manno-heptose 7-phosphate or from a putative ADP-glycero-manno-heptose-1,7-diphosphate (49). We are currently exploring these possibilities by investigating a number of uncharacterized genes in E. coli K-12 with homologies to sugar phosphate phosphatases.

Domain II has strong homologies with the cytidylyltransferase superfamily. This family includes enzymes like glycerol-3-phosphate cytidylyltransferase from Bacillus subtilis and the eukaryotic choline phosphate cytidylyltransferase, which are involved in teichoic acid and phospholipid biosynthesis, respectively (4). Also, three members of this family, pantoate -alanine ligase, acetate:SH-citrate lyase, and phosphopantetheine adenylyltransferase, are ADP transferases (4, 21). Bork et al. (4) have shown that this family has structural conservation with the class I tRNA synthetases, all of which also use ATP. In addition, there are several bifunctional enzymes, such as FAD synthetase, PAPS synthase (forming 3'-phosphoadenosine 5'-phosphosulfate), and the transcriptional regulator NadR, that contain this domain (33). The crystal structure of one of the ADP transferases of this family, phosphopantetheine adenylyltransferase, has been recently established (21). Almost identical residues were found in the fold corresponding to the nucleotide-binding site (Fig. 5), which has the characteristics of a dinucleotide-binding fold (41). Therefore, it is possible that Domain II functions as a nucleotide sugar transferase. ADP-glycero-manno-heptose can be isolated from yeast and bacteria (24, 25), and in a previous study, Coleman has shown that the gmhD (formerly rfaD) gene codes for ADP-LDHep-6-epimerase (10), the last step prior to the transfer of LDHep to the core LPS. Therefore, it is likely that ADP is the nucleotide added in vivo to the phosphorylated D-glycero-D-manno-heptose. Since the sugar precursors for the biosynthesis of ADP-heptose are not readily available, it is difficult to determine the function of these two domains with certainty.

According to the current nomenclature for LPS biosynthesis genes (40), rfaE should be renamed with the prefix gmh (for glycero-manno-heptose synthesis). However, the lack of a clear functional assignment, as well as the presence of separate genes encoding each domain in some microorganisms, makes it difficult to reassign a meaningful new gene name to rfaE. To avoid more confusion, we prefer not to introduce a new gene name until further biochemical investigations clarify the intermediate steps of the biosynthesis pathway for ADP-heptose.