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Department of Microbiology, Immunology, and Molecular Genetics, University of Kentucky, Lexington, Kentucky 40536-0084, USA
Correspondence
Robert D. Perry
rperry{at}pop.uky.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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-barrel structure with a large periplasmic domain while HmsF possesses polysaccharide deacetylase and COG1649 domains. HmsR is a putative glycosyltransferase while HmsS has no recognized domains. In this study, specific amino acids within conserved domains or within regions of high similarity in HmsH, HmsF, HmsR and HmsS proteins were selected for site-directed mutagenesis. Some but not all of the substitutions in HmsS and within the periplasmic domain of HmsH were critical for protein function. Substitutions within the glycosyltransferase domain of HmsR and the deacetylase domain of HmsF abolished biofilm formation in Y. pestis. Surprisingly, substitution of highly conserved residues within COG1649 did not affect HmsF function.
A table of primers is available with the online version of this paper.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Davey & O'Toole, 2000).
The Yersinia pestis Hms+ phenotype is a manifestation of biofilm formation causing massive adsorption of haemin or Congo red (CR) during growth at 26–34 °C but not at 37 °C. This is required for colonization and eventual blockage of the flea proventriculus – allowing efficient transmission of plague from some fleas, like the oriental rat flea, to mammals (Bacot & Martin, 1914; Bacot, 1915; Hinnebusch et al., 1996; Lillard et al., 1997, 1999; Pendrak & Perry, 1991, 1993; Perry et al., 1990, 1993, 2004; Perry & Fetherston, 1997; Pollitzer, 1954). Biofilm formation is not involved in mammalian virulence – an in-frame deletion in hmsR did not affect the LD50 in a bubonic plague model (Lillard et al., 1999). Recent studies have demonstrated Hms-dependent biofilm formation in vitro as well as in fleas and nematodes (Darby et al., 2002; Jarrett et al., 2004; Jones et al., 1999; Kirillina et al., 2004). CR binds basic and neutral polysaccharides while wheat germ agglutinin binds to N-acetylglucosamine, suggesting that the Hms-dependent biofilm contains an exopolysaccharide (EPS) component. In addition, dispersin, an enzyme from Actinobacillus actinomycetemcomitans which hydrolyses -1,6-N-acetyl-D-glucosamine, prevents plague biofilm formation but does not disrupt a preformed biofilm (Itoh et al., 2005; Spiers et al., 2002; Tan & Darby, 2004; Wood, 1980).
The 102 kb pgm locus in Y. pestis contains the high-pathogenicity island (Ybt) and the hmsHFRS operon encoding four of the six hms genes required for biofilm formation and its regulation. Escherichia coli K-12 pgaABCD (formerly ycdSRQP) encode proteins with similarities (ranging from 34 to 83 %) to Hms proteins and are required for the synthesis of an EPS containing poly--1,6-N-acetyl-D-glucosamine (Blattner et al., 1997; Deng et al., 2002; Fetherston et al., 1992; Jones et al., 1999; Lillard et al., 1997; Wang et al., 2004). HmsH and HmsF are outer-membrane proteins while HmsS, HmsR, and HmsT are located in the inner membrane. While HmsH is predicted to be a -barrel protein with a large periplasmic domain, its function and that of HmsS are unknown. HmsR, a putative glycosyltransferase, and HmsF, with a polysaccharide deacetylase domain, are likely involved in the synthesis and modification of the EPS component of the biofilm. HmsF also has a conserved but uncharacterized domain, COG1649, that is present in other HmsF-like proteins in Gram-negative bacteria (Kirillina et al., 2004; Pendrak & Perry, 1991; Perry et al., 2004). Proteins with this domain are assigned to the TIM barrel glycosyl hydrolase superfamily by Pfam (Bateman et al., 2004). Recently, gmhA, a gene encoding phosphoheptose isomerase, has been identified as important for biofilm formation (Darby et al., 2005).
Temperature regulation of plague biofilm formation does not involve transcriptional regulation or mRNA stability. Rather, this is achieved by proteolytic degradation of HmsR, HmsH and HmsT at 37 °C. HmsT, a putative diguanylate cyclase, may be the key to this temperature regulation. It is a positive regulator of biofilm formation and is involved in the synthesis of cyclic-di-GMP (c-di-GMP). HmsP, a negative regulator of biofilm formation, is an EAL phosphodiesterase and likely degrades c-di-GMP. Like other glycosyltransferases involved in EPS or cellulose productions, the enzymic activity of HmsR may depend upon c-di-GMP (Bobrov et al., 2005; Hare & McDonough, 1999; Jones et al., 1999; Kirillina et al., 2004; Perry et al., 2004; Simm et al., 2005).
Previously, we used site-directed amino acid substitutions to identify several critical residues of HmsT and HmsP (Kirillina et al., 2004). In this study, specific amino acids within conserved domains or within regions of high similarity in HmsH, HmsF, HmsR and HmsS were selected for site-directed mutagenesis. Substitutions within the glycosyltransferase domain of HmsR and the deacetylase domain of HmsF abolished biofilm formation in Y. pestis while substitutions in HmsS and HmsH had diverse effects. Surprisingly, substitution of highly conserved residues within COG1649 did not affect HmsF function.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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. E. coli cells were grown on Luria broth agar (LBA). Where appropriate, ampicillin (Ap; 100 µg ml–1), kanamycin (Km; 50 µg ml–1), streptomycin (Sm; 50 µg ml–1) or chloramphenicol (Cm; 20 µg ml–1) were added to cultures. Strains with arabinose-inducible expression were grown in the presence of 0.2 % (w/v) arabinose. Y. pestis cells were streaked onto Congo red (CR) plates from buffered glycerol stocks stored at –80 °C and incubated at 26 °C for 48 h. For most experiments, individual colonies were inoculated onto Tryptose Blood Agar Base (TBA; Difco) slants and incubated at 26 °C for 24–48 h. Cells were washed off TBA slants with Heart Infusion Broth (HIB; Difco) and grown overnight with aeration in HIB at the appropriate temperature. Cultures were transferred to fresh medium the next morning. For some experiments, Y. pestis cells were grown in TMH medium (Straley & Bowmer, 1986). Cell growth was monitored on a Spectronic Genesys5 spectrophotometer at 620 nm. For Western blot analyses, isolated colonies from TBA plates were spread onto CR plates and allowed to form a lawn (48 h). Cells were scraped off the plate and suspended in sample buffer.
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. Briefly, cells were grown overnight on TBA slants and used to inoculate TMH to an OD620 of 0.1. Cultures were grown for 16–18 h in a shaking water bath at 26 °C and then exposed to 0.01 % CV for 15–20 min. Cells attached to the culture tube were washed three times with distilled water and the CV staining the cells was solubilized with a mixture of 80 % ethanol/20 % acetone. The amount of dye solubilized, representing the mass of attached bacterial cells, was measured at 570 nm on a Spectronic Genesys5 spectrophotometer.
CR-binding assay.
The original CR-binding assay (Hartzell et al., 1999) was modified as previously described (Kirillina et al., 2004). Briefly, Y. pestis cells were grown overnight (
16 h) in HIB at 26 °C. Cells were pelleted and resuspended in HIB/CR medium [1 % (w/v) HIB containing 0.2 % galactose and 30 µg CR ml–1] such that all cultures had an equivalent wet weight of cells per µl of medium. An 20 mg wet cell weight aliquot of each culture was incubated for 3 h on a rocking platform at room temperature. The amount of CR bound by the cells was determined by measuring the absorbance of cell-free supernatants at 500 nm with a Spectronic Genesys5 spectrophotometer and subtracting this value from the reading obtained with uninoculated HIB/CR medium.
Plasmids and recombinant DNA techniques.
All the plasmids used in this study are listed in Table 1
. Plasmids were purified from overnight cultures by alkaline lysis (Birnboim & Doly, 1979) or with a Qiaprep spin Miniprep kit (Qiagen) and further purified when necessary by polyethylene glycol precipitation (Humphreys et al., 1975). A standard CaCl2 protocol was followed to introduce plasmids into E. coli (Sambrook & Russell, 2001). Y. pestis cells were transformed by electroporation as previously described (Fetherston et al., 1995). All plasmid DNA and PCR products used for construction of chromosomal deletion mutants or cloned amino acid substitution mutants were sequenced by Elim Biopharmaceuticals to ensure that the correct mutation had been introduced and no secondary mutations had occurred. In a few instances, aberrant amino acid substitution clones were identified and these were also characterized.
Construction of hmsH, hmsR, hmsF and hmsS mutants.
Suicide vector pKNG101 (Kaniga et al., 1991) was used to generate in-frame deletions in hmsH and hmsR. Fragments spanning the upstream and downstream regions of hmsH were obtained by NsiI/XmnI (1147 bp) and SwaI/BamHI (1149 bp) digestions of pNPM22 and cloned into the PstI–BamHI sites of pBluescript SK. The resulting construct, pBSSK
hmsH, and pKNG101 were both digested with SalI and XbaI followed by ligation of the resulting 2.4 kb fragment from pBSSKhmsH into pKNG101, yielding pKNGhmsH (Table 1). For the hmsR deletion, 1.1 kb SalI–NcoI and 1.1 kb BsrBI–BamHI fragments of the upstream and downstream region of hmsR in pHMS1.2 were cloned into the SalI–BamHI sites of pWSK29, yielding pWSK29hmsR. This plasmid was digested with SalI/XbaI and the resulting 2.2 kb fragment was cloned into the same sites in pKNG101, yielding pKNGhmsR (Table 1).
PCR and DNA sequencing (Elim Biopharmaceuticals) confirmed that the correct DNA sequences were present in pKNGhmsH and pKNGhmsR prior to electroporation into Y. pestis KIM6+. As previously described (Bearden & Perry, 1999), transformants were selected on TBA plates containing 50 µg Sm ml–1 and grown overnight in HIB without antibiotics to select for sucrose-resistant isolates that had completed the allelic exchange. Strains in which the wild-type gene was replaced by the deleted version were identified by PCR. One isolate of each mutation was selected and named KIM6-2115 (in-frame hmsH2115) and KIM6-2118 (non-polar hmsR2118). These deletion mutations remove 569 of 822 and 327 of 457 aa in HmsH and HmsR, respectively.
Red-mediated recombination (Datsenko & Wanner, 2000) was employed to construct non-polar deletions in hmsF and hmsS in Y. pestis KIM6(pKD46)+. Electrocompetent cells were made as previously described (Fetherston et al., 1995) from cultures grown at 26 °C in HIB to OD620 0.6 and then incubated for 1.5 h with 0.2 % arabinose to induce expression of the Red recombinase (Datsenko & Wanner, 2000). Gene-specific primers (see Table S1, available as supplementary material with the online version of this paper) were used to amplify the cam gene cassette from pKD3. Cells, transformed with 0.5–2 µg purified PCR product, were plated on TBA plates containing Cm to select for recombinant cells. To cure pKD46 from selected recombinants, colonies, grown at 37 °C on TBA without Ap, were isolated and screened for sensitivity to Ap. For selected mutants, the presence of the cam cassette was confirmed by PCR. Flippase-expressing pCP20 was electroporated into selected Cmr mutants and the transformation mixture was plated, after overnight incubation at room temperature, on TBA plates containing Ap. As described above, cells sensitive to Cm and Ap (excised Cm cassette and pCP20-cured) were isolated. Mutations were confirmed by PCR and the resulting strains designated KIM6-2116 (hmsF-2116) and KIM6-2119 (hmsS-2119). These non-polar mutations remove all of the coding region for the HmsF protein (673 aa) and 150 of 155 aa residues in the HmsS protein. The hmsF deletion in KIM6-2116 overlaps with hmsR and potentially removes four N-terminal amino acids of HmsR. Despite this elimination of the predicted start site (Lillard et al., 1997), expression of this version of HmsR is functional (see Results).
Construction of alanine substitutions in HmsH, HmsF, HmsR and HmsS.
Most of the amino acid substitutions were made by a two-step PCR cloning scheme (Fig. 1) using two outside primers (1, 2) and two overlapping interior primers (3, 4) carrying the desired mutation. This method has been used to avoid the occurrence of PCR mutations while amplifying a long template even with DNA polymerases with proof-reading activity. First, two smaller PCR products were generated (primers 1-3 and 4-2), overlapping at the region of interest by primers carrying the desired mutation (primers 3 and 4). PCR products were gel purified, with the exception of hmsR mutations, where a DpnI digestion was used to eliminate carryover template DNA. Second, the products of the first PCR (1-3, 4-2), hybridizing across the site of interest and with the introduced amino acid codon substitution (3-4), underwent a fill-in cycle to yield a full-length (1-2) double-stranded template. Third, the outside primers (1, 2) were added to the reaction to complete the second PCR and thus generate a gene fragment carrying the desired mutation. The fill-in PCR consisted of one 2 min cycle at 95 °C followed by two cycles at 95 °C for 30 s, 50 °C for 30 s and 68 °C for 4 min. After introducing the outside primers (1, 2), the PCR reaction continued with a 2 min cycle at 95 °C followed by 30 cycles at 95 °C for 30 s, 50 °C for 30 s and 68 °C for 4 min. A 4 min extension time was applied in the case of a 2 kb fragment (hmsF) and was modified accordingly for fragments of different sizes. The final PCR product was column purified and concentrated (Zymo Research), digested (site1, site2 in Fig. 1) and cloned into a target vector. ProofStart DNA polymerase (Qiagen) was used in most amplification reactions, unless stated otherwise. In some cases the QuickChange site-directed mutagenesis kit (Stratagene) was used, following the manufacturer's instructions. Sequences of primers specific for each amino acid substitution are given in Table S1.
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) express hmsH genes and a truncated hmsF from the native hmsHFRS promoter.
Plasmid pNPM11 was digested with SwaI/TatI to subclone hmsF into the KpnI–SmaI sites of pBAD30. An isolated EcoRI–XbaI fragment from this plasmid containing the hmsF gene was treated with DNase I (long Klenow fragment) and cloned into the SmaI site of pBAD30, yielding pBADHmsF. All HmsF substitutions were constructed in pBAD30 by insertion of the modified hmsF PCR fragments generated from pBADhmsF and digested with EcoRI and XbaI (Table 1).
Plasmid pHMS1.2 was used as a template to amplify wild-type hmsR and hmsS DNA. Both genes were cloned as EcoRI–XbaI fragments into the same sites in pBAD30, generating pBADHmsR and pBADHmsS (Table 1). HmsR D176A and D269A mutations were constructed with the QuickChange site-directed mutagenesis kit (Stratagene). Platinum Pfx DNA polymerase (Invitrogen) was used in the course of hmsR mutagenesis. Substitutions in hmsS were constructed in pBAD30 by insertion of the modified hmsS PCR fragments generated from pBADhmsS and digested with EcoRI and XbaI (Table 1).
Sequence analysis.
Amino acid sequences presented in protein alignments (Fig. 2) were obtained from the NCBI Entrez Genome Project website. All sequence alignments were generated in T-Coffee (version 1.6; http://www.ch.embnet.org/software/TCoffee.html) (Notredame et al., 2000).
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) was used for immunodetection. Briefly, PVDF membranes were blocked with 5 % non-fat dry milk or 1.5 % BSA in 10 mM Tris/HCl (pH 7.6), 137 mM NaCl (TBS) with 0.1 % Tween 20 (TBST) and then incubated with the appropriate antibody diluted in TBST. Antibodies, raised against portions of HmsH, HmsF and HmsS proteins, were generated in rabbits by Animal Pharm Services. Antiserum against a synthetic immunogenic peptide of HmsR was generated by Research Genetics. Following incubation with horseradish peroxidase-conjugated protein A (Amersham Pharmacia Biotech), the immunoreactive proteins were detected with the ECL enhanced chemiluminescence Western blotting detection reagent (Amersham Pharmacia Biotech). Alternatively, proteins interacting with alkaline phosphatase-labelled anti-rabbit IgG molecules were detected with the chemiluminescent substrate CSPD (Roche). Immunoblot results were visualized on Kodak Biomax light film. |
RESULTS |
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). To test the effect of these substitutions, in-frame deletions of individual hms genes in Y. pestis were constructed. These mutations yielded an Hms– phenotype at 26–30 °C: white colonies on CR plates and absence of aggregation in liquid broth cultures. The four cloned wild-type hms genes complemented their respective in-frame or non-polar chromosomal mutations. Variants of hms genes with single amino acid substitutions in pBAD30 with arabinose-inducible expression (hmsF, hmsR and hmsS) or in versions of pNPM22 (e.g. pHmsH-R134A) with expression of hmsH alleles from the native hmsH promoter were tested for the ability to bind CR or to attach to glass in CV-staining assays as parameters of biofilm formation. The cellular level of wild-type and variant Hms proteins was assessed by Western blot analysis (Table 2).
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, Pendrak & Perry, 1993; Perry et al., 2004) and have amino acid sequence identities/similarities of 41.1 %/58.2 % and 48.3 %/60.8 %, respectively, to E. coli proteins PgaA and PgaB (Fig. 2). HmsH is the largest Hms protein (822 aa) with a predicted -barrel structure typical of a transmembrane protein in the outer membrane. We have changed charged residues R134, E269, E281 and D289 to alanines in the N-terminal portion of HmsH. These residues are within a periplasmic domain predicted by the hidden Markov model (PRED_TMBB; http://bioinformatics.biol.uoa.gr/PRED-TMBB). Alanine substitutions E269A and D289A in HmsH had no significant effect on the Hms phenotype. The E281A change, where the stability of the resulting HmsH-E281A protein was affected (Table 2), apparently complemented the hmsH mutation on CR plates but gave significantly decreased values in our quantitative assays for CR adsorption and CV staining (Fig. 3A, B) compared to complementation with wild-type hmsH. The HmsH-E281A protein was defective in biofilm formation as assessed by CV staining (Fig. 3). The HmsH-R134A protein also appears defective in biofilm formation by the CV staining assay but failed to show a similar defect in quantitative CR binding. A neutral change, hmsH-A350V, fully complemented the hmsH mutation as expected (Fig. 3).
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). Several of the conserved residues in this group are not highly conserved in enzymes with proven deacetylase activity from a wide variety of organisms. Comparison of the deacetylase domains of these proteins, including Bacillus subtilis PdaA with known crystal structure (Blair & van Aalten, 2004), to Y. pestis HmsF (Fig. 2B2) indicated highly conserved residues related to enzymic activity. The highly conserved nature of all these residues (Fig. 2B1–3) suggests they might be essential.
The D114A alteration in HmsF, corresponding to the D73 residue of PdaA, abolished the HmsF protein function, confirming the importance of this residue in the deacetylase domain. Moreover, the D115A substitution in the ‘TFD(D)G’ conserved motif and alteration of the most conserved H184 residue in HmsF both caused biofilm-defective phenotypes (Fig. 3C, D). HmsF with a double substitution (D114A and D115A) retained the expected phenotypic loss as well. Another strongly conserved residue among proven deacetylases, K107, was not critical for HmsF function (Fig. 3C, D). However, the K107A change caused an 
50 % loss in CR binding. Among the residues conserved in HmsF-like proteins (R123, W143, R161, W167, Q169, E172 and E180), only the W143 substitution had an effect on HmsF function (Table 2). The W143A substitution (identical to W143 of E. coli PgaB but not present in other deacetylases) had an intermediate-pink phenotype on CR agar plates (Table 2) and an adverse effect in both quantitative assays we used to assess the Hms phenotype (Fig. 3C, D).
The C-terminal part of Y. pestis HmsF has a predicted domain COG1649 that is present in other HmsF-like proteins of Gram-negative bacteria (Fig. 2B2). None of the nine individual substitutions in conserved residues in the COG1649 region of HmsF caused a biofilm-defective phenotype (Table 2). An altered HmsF protein with a large deletion (aa 374–674) in COG1649 failed to complement our hmsF mutant. However, Western blots did not detect the truncated protein, indicating that either the stability of the protein was affected or our polyclonal antibody against HmsF does not detect the truncated protein (data not shown).
HmsR and HmsS amino acid substitutions
The similarities of the putative glycosyltransferase HmsR from Y. pestis to those of Staphylococcus epidermidis – IcaA and E. coli – PgaC (formerly YcdQ) and to the cellulose synthetase BscA from Salmonella typhimurium have been described previously (Jones et al., 1999; Kirillina et al., 2004; Lillard et al., 1997; Wang et al., 2004). The functionality of these proteins is critical for biofilm formation in all these bacteria (Heilmann et al., 1996; Lillard et al., 1997; Pendrak & Perry, 1993; Wang et al., 2004; Zogaj et al., 2001). Amino acid sequence comparison of the cellulose synthase BscA from Gluconobacter xylinus with many other putative glycosyltransferases, including HmsR, IcaA and PgaC, led to the identification of a conserved ‘D, D, D35QxxRW’ motif (Saxena & Brown, 1997) (Fig. 2C). We therefore tested alanine substitutions at positions D176, D269, Q305 and R308 as well as in non-conserved residues T114 and L324. Expression of HmsR-T114A and HmsR-L324A complemented the hmsR mutation as well as the wild-type hmsR gene in forming red colonies on CR agar and restored high levels of CR binding and CV staining (Fig. 4). However, substitutions D176A, D269A, Q305A or R308A in HmsR did not complement the hmsR mutant, giving rise to white colonies on CR plates and only low levels of CV staining. For all HmsR variants tested, the results of the quantitative CR-binding assays correlated with CV-staining assays (Fig. 4A, B). Hence, four conserved residues from the glycosyltransferase motif ‘D, D, D35QxxRW’ in HmsR, D176, D269, R305, R308, are critical for production of the biofilm extracellular matrix.
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) and was selected for mutagenesis. Expression of HmsS-N73A or HmsS-R87A in KIM6-2119 (hmsS) restored CR binding and biofilm formation. On the other hand, substitutions W80A and Y83A gave a strong biofilm-defective phenotype equal to that of KIM6 (hmsHFRS), with little CV staining or CR binding. Thus, the W80A and Y83A, but not the N73A and R87A, residues are critical to the function of HmsS. |
DISCUSSION |
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; Pendrak & Perry, 1993). In this study, we constructed non-polar deletion mutations in each gene of the operon and demonstrated loss of biofilm formation in each mutant via quantitative CR-binding and CV-staining assays. Moreover, biofilm formation was restored by complementation with the corresponding gene (Figs 3 and 4). This definitively confirms our previous conclusions that HmsH, HmsF, HmsR and HmsS are each essential for biofilm formation.
Genes encoding orthologues of each of these Y. pestis proteins have been identified in a number of diverse organisms (Bell et al., 2004; da Silva et al., 2002; Jones et al., 1999; Lee et al., 2005; Paulsen et al., 2005; Salanoubat et al., 2002; Wang et al., 2004). Of these, the E. coli PgaABCD system is involved in formation of an EPS containing a -1,6-N-acetyl-D-glucosamine polymer, and an EPS with a similar component is also part of the Y. pestis biofilm (Itoh et al., 2005; Wang et al., 2004). Several other proteins related to HmsH, HmsF, HmsR and HmsS are from genome sequence projects (Bao et al., 2002; Methé et al., 2003; Nascimento et al., 2004; Ren et al., 2003). We have used these in amino acid sequence alignments with the Hms proteins to identify highly conserved residues that might be critical to the function of these four Hms proteins. Alteration of selected conserved residues to alanine identified critical residues, residues with a moderate effect, and highly conserved residues that are not important for forming a functional Hms protein.
HmsH is predicted to form a -barrel protein with a large N-terminal periplasmic domain (Bagos et al., 2004; Kirillina et al., 2004). Our mutagenesis study concentrated on this periplasmic domain to avoid a loss of phenotype due to disruption of the -barrel structure. Two of four residues (E269 and D289) had no effect on biofilm formation while the other two (R134 and E281) had only a moderate effect on the composition of the usual Y. pestis KIM6+ biofilm. Complementation with both mutated HmsH proteins yielded CR+ colonies on CR plates, indicating no change in a long-term phenotype (CR binding over >48 h). However, both HmsH-E281A and HmsH-R134A had a significant reduction in CV staining compared to HmsH, indicating an alteration in biomass bound to glass over a period of 16 h. HmsH-E281A also caused a reduction in CR binding in liquid assay over a 3 h period. Thus, R134 or E281 of HmsH are not absolutely crucial to the function of HmsH but may moderately reduce its efficacy.
HmsF belongs to a large family of deacetylases that possess strong sequence similarities across a wide range of bacteria, with some variability due to the distinct carbohydrate substrates used. The role of B. subtilis PdaA in spore germination has been described in detail and in vitro enzymic studies have demonstrated PdaA deacetylase activity with an N-acetylmuramic acid substrate (Fukushima et al., 2002, 2005; Gilmore et al., 2004). PdaB deacetylase is involved in assembly of peptidoglycan during the course of B. subtilis sporulation (Fukushima et al., 2004). Within the HmsF deacetylase domain, we selected 11 highly conserved residues to mutate. Four residues are highly conserved in proven deacetylases (Fig. 2B2) while the other seven residues are highly conserved in HmsF-like proteins (Fig. 2B1) but not in proven deacetylases (Fig. 2B2). Alteration of the HmsF residues D114 and D115, corresponding to D73 and N74 of PdaA, completely prevented biofilm formation, possibly by affecting the putative deacetylase activity of HmsF. The H184A substitution (corresponding to H124 in PdaA) also abolished CR binding and CV staining. However, the HmsF containing the K107A alteration appears to produce another attenuated CR-binding mutant with reduced short-term binding at 3 h that does not affect the formation of CR colonies by 48 h of growth (Fig. 3, Table 2). The W143A substitution caused negligible CR binding and CV straining that led to an intermediate-pink colony formation on CR plates, suggesting this residue is not essential but does affect protein function – possibly enzymic activity. W143 was the only substitution in HmsF-like conserved residues that had any effect on the functionality of HmsF.
A second region of Y. pestis HmsF with high similarity to other proteins (Fig. 2B2) is COG1649; proteins with this domain of unknown function are assigned to the TIM barrel glycosyl hydrolase superfamily by Pfam (Bateman et al., 2004). Nine alanine substitutions of highly conserved residues within the COG1649 domain of HmsF did not affect HmsF – all altered proteins were able to fully restore an Hms+ phenotype (Table 2). However, expression of a truncated HmsF lacking the COG1649 domain failed to complement the hmsF mutant. Truncated HmsF protein was not detected by HmsF antiserum used in immunodetection of full-length HmsF proteins. The recombinant protein used to produce HmsF antiserum lacked the first 87 aa but contained both the deacetylase and COG1649 domains (Kirillina et al., 2004; Pendrak & Perry, 1991; Perry et al., 2004). This may indicate protein instability or lack of antigenic domains recognized by our HmsF antibody. Further studies will be required to determine whether the COG1649 domain is important to the function of HmsF.
There are several lines of evidence indicating that HmsR is a likely -1,6-N-acetyl-D-glucosamine transferase. First, HmsR has a high degree of similarity to IcaA and PgaC, which are both required for the synthesis -1,6-N-acetyl-D-glucosamine (Heilmann et al., 1996; Jones et al., 1999; Wang et al., 2004). Second, PgaC was able to restore the CR-binding phenotype of an hmsR mutant when expressed from a high-copy-number plasmid (Jones et al., 1999). Third, DspB, a -hexosaminidase and biofilm-dispersing enzyme of A. actinomycetemcomitans (Kaplan et al., 2003), shown to specifically hydrolyse the glycosidic linkages of poly--N-acetyl-D-glucosamine, caused complete inhibition of biofilm formation by Y. pestis KIM6+ (Itoh et al., 2005). Our results further support this conclusion. A non-polar hmsR deletion abrogated synthesis of an extracellular matrix which binds CR and expression of HmsR proteins with substitutions in conserved residues of the glycosyltransferase motif ‘D, D, D35QxxRW’, i.e. D176A, D269A, R305A and R308A, failed to restore CR binding and CV staining to this mutant (Fig. 2; Table 2). The substitutions D236Y, D333R, Q369M and R372A within this motif in the AcsAB cellulose synthase of G. xylinus impaired the enzymic activity in vitro (Saxena & Brown, 1997; Saxena et al., 2001). Consequently, we propose that the substitutions in HmsR at similar positions may abolish its enzymic activity in poly-N-acetylglucosamine production.
The essential HmsS protein, which is associated with the inner membrane like HmsR and HmsT (Lillard et al., 1997; Perry et al., 2004), has two predicted transmembrane regions but no other currently recognized domains. Alignment of HmsS-like proteins identified a conserved region [‘NA(V)xLIxW(A)x(Y)x(Q)x(R)’; Fig. 2D)]. N73A and R87A changes did not affect HmsS protein function. In contrast, a dramatic loss of HmsS function was related to W80A and Y83A alterations (Fig. 4A, B). Both W80 and Y83 are within a predicted cytoplasmic region of HmsS.
These site-directed mutagenesis studies have identified critical residues within predicted enzymic domains in HmsF and HmsR and in a conserved region of HmsS. Two residue changes in HmsH and two in HmsF (HmsH-R134A, HmsH-E281, HmsF-K107A, HmsF-W134A) had differential effects in our biofilm assays. The reason for this differential effect is obscure. The ability of cells expressing HmsF-K107A to achieve normal attachment levels while displaying an EPS-defective phenotype suggests that this mutant produces an EPS form that is not conducive to CR binding. In addition, these and some of the other amino acid residue substitution mutations may affect protein–protein interactions that may be important for biofilm formation.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 15 June 2006;
revised 28 July 2006;
accepted 9 August 2006.
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