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Journal of Bacteriology, July 2007, p. 4603-4613, Vol. 189, No. 13
0021-9193/07/$08.00+0     doi:10.1128/JB.00236-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Genetic Dissection of Specificity Determinants in the Interaction of HPr with Enzymes II of the Bacterial Phosphoenolpyruvate:Sugar Phosphotransferase System in Escherichia coli{triangledown}

Birte Reichenbach,1 Daniel A. Breustedt,2,{dagger} Jörg Stülke,1 Bodo Rak,2 and Boris Görke1*

Department of General Microbiology, Institute of Microbiology and Genetics, Georg-August University, Grisebachstrasse 8, D-37077 Göttingen, Germany,1 Faculty of Biology, Biology III, University of Freiburg, Schänzlestrasse 1, D-79104 Freiburg, Germany2

Received 12 February 2007/ Accepted 12 April 2007


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ABSTRACT
 
The histidine protein (HPr) is the energy-coupling protein of the phosphoenolpyruvate (PEP)-dependent carbohydrate:phosphotransferase system (PTS), which catalyzes sugar transport in many bacteria. In its functions, HPr interacts with a number of evolutionarily unrelated proteins. Mainly, it delivers phosphoryl groups from enzyme I (EI) to the sugar-specific transporters (EIIs). HPr proteins of different bacteria exhibit almost identical structures, and, where known, they use similar surfaces to interact with their target proteins. Here we studied the in vivo effects of the replacement of HPr and EI of Escherichia coli with the homologous proteins from Bacillus subtilis, a gram-positive bacterium. This replacement resulted in severe growth defects on PTS sugars, suggesting that HPr of B. subtilis cannot efficiently phosphorylate the EIIs of E. coli. In contrast, activation of the E. coli BglG regulatory protein by HPr-catalyzed phosphorylation works well with the B. subtilis HPr protein. Random mutations were introduced into B. subtilis HPr, and a screen for improved growth on PTS sugars yielded amino acid changes in positions 12, 16, 17, 20, 24, 27, 47, and 51, located in the interaction surface of HPr. Most of the changes restore intermolecular hydrophobic interactions and salt bridges normally formed by the corresponding residues in E. coli HPr. The residues present at the targeted positions differ between HPrs of gram-positive and -negative bacteria, but within each group they are highly conserved. Therefore, they may constitute a signature motif that determines the specificity of HPr for either gram-negative or -positive EIIs.


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INTRODUCTION
 
The carbohydrate:phosphotransferase system (PTS) is the predominant carbohydrate uptake system in many bacteria (7, 20). It comprises a chain of phosphoryl transfer reactions, starting with the phosphoenolpyruvate (PEP)-dependent autophosphorylation of enzyme I (EI), which subsequently phosphorylates histidine protein (HPr) at its His15 residue (HPr~P). HPr serves as the central phosphocarrier protein and delivers the phosphoryl groups to the IIA domains of the sugar-specific enzymes II (EIIs). Subsequently, the phosphoryl group is transferred to a residue in the IIB domain of the EIIs and from there to the substrate during transport through the membrane-bound domain(s) IIC and sometimes IID. Based on their phylogeny, EIIs are grouped into seven families (26). Members of one family share more than 25% sequence identity over the entire molecule, and functional complementation between equivalent domains is often possible within a family (17). The A, B, and C domains of EIIs of different families usually do not share structural similarity with one another, supporting the idea that they are unrelated (25).

In addition, in many bacteria HPr regulates the activities and the expression of enzymes involved in the utilization of carbon sources (7). This is achieved by protein-protein interaction, e.g., the activation of glycogen phosphorylase by binding to HPr in Escherichia coli (31) or by HPr-dependent phosphorylation of the target protein, as shown for the glycerol kinase GlpK in gram-positive bacteria (34). Many bacteria possess antiterminator proteins of the BglG/SacY family and other transcriptional regulators containing PTS regulatory domains (PRDs), which require phosphorylation by HPr to be active (7). In low-GC gram-positive bacteria, HPr can be phosphorylated by the HPr kinase/phosphorylase (HPrK/P) at a second site, Ser46. HPr(Ser)-P subsequently forms a complex with catabolite control protein A (CcpA), and binding of this complex to operator sites on the DNA triggers the main mechanism of carbon catabolite control in these bacteria (34, 40). Altogether, in its respective host, HPr must be able to interact specifically with a large number of structurally and evolutionarily unrelated proteins.

HPrs of different organisms are at least 35% identical, being most conserved around the active-site His15 (14). The three-dimensional structures of the HPrs of E. coli, Bacillus subtilis, and of a few other species have been determined, and they all exhibit the same overall fold, that of an open-faced ß-sandwich consisting of three {alpha}-helices on top of a four-stranded ß-sheet (for reviews, see references 2 and 43). The solution structures of E. coli HPr in complex with several of its different partner proteins have been resolved. These are EI (9), IIAGlc (41), IIAMtl (6), IIAMan (44), and glycogen phosphorylase (42). In all these structures HPr uses essentially the same narrow convex surface for interaction. No large conformational changes in HPr or its partners occur upon complex formation. The key interacting residues of HPr are located in {alpha}-helices 1 and 2 and in the loops preceding {alpha}1 and following {alpha}2. The central portion of the interacting protein surface in HPr is predominantly hydrophobic and surrounded by polar and positively charged residues which are involved in electrostatic interactions. Several salt bridges are formed involving the side chains of Arg and Lys residues at positions 17, 24, 27, and 49 in E. coli HPr. The structures of the complexes of HPr of B. subtilis with its partner proteins IIAGlc, HPrK/P, and CcpA were also solved and revealed an interaction surface in HPr very similar to that of its E. coli homologue (5, 8, 13, 30). Residues within the {alpha}-helices {alpha}1 and {alpha}2 in HPr participate in hydrophobic and/or electrostatic interactions comparable to the roles of the corresponding residues in HPr of E. coli.

In view of the very similar structures of the HPrs of B. subtilis and E. coli, their conserved interaction surfaces (6), and the many unrelated proteins with which they interact, one could imagine that HPr is flexible enough to interact with the homologous partner proteins of the respective other species. However, interaction studies showed that, in vitro, E. coli EI has a 29-fold-lower affinity for B. subtilis HPr than E. coli HPr (24). Moreover, in vitro, the affinity of E. coli IIAGlc for B. subtilis HPr is 300-fold lower than for E. coli HPr (24), a result that was confirmed by nuclear magnetic resonance chemical shift mapping experiments (22).

In the present work, we studied the in vivo interaction of B. subtilis HPr with its heterologous partner proteins in E. coli. We show that there is no or just weak functional interaction with various E. coli EIIs of different families, suggesting that impaired interaction with EIIs of E. coli is a general phenomenon. In contrast, activation of the E. coli BglG regulatory protein by B. subtilis HPr-catalyzed phosphorylation was more efficient. To understand the reasons for the impaired interactions with the E. coli EIIs, we identified mutants of B. subtilis HPr with improved in vivo interaction properties. These mutants carried amino acid changes located almost exclusively in the interaction surface of HPr, and the changes have the potential to restore the intermolecular interactions carried out by the corresponding residues in E. coli HPr. Our results yield insight into the reasons for the species specificity of HPr.


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MATERIALS AND METHODS
 
Plasmids, strains, and growth conditions. The strains and plasmids used and their relevant characteristics are listed in Table 1. T4GT7 transduction was used to move the {Delta}[ptsH ptsI crr]::neo allele of E. coli strain TP2811 into strain R1967 (12), resulting in strain R1969. T4GT7 (T4 generalized transducer no. 7) is a mutant of E. coli bacteriophage T4 that allows high-frequency generalized transduction of selectable alleles present in the E. coli genome (45). For DNA cloning, strain DH5{alpha} was used by following standard procedures (27). Bacteria were routinely grown in Luria-Bertani broth supplemented with the appropriate antibiotics (kanamycin at 30 µg/ml, tetracycline at 12.5 µg/ml, and ampicillin at 100 µg/ml). For in vivo phosphorylation assays and growth rate studies in M9 minimal medium, the concentrations of tetracycline and ampicillin were reduced to 10 µg/ml and 50 µg/ml, respectively.


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  		TABLE 1. Strains and plasmids used in this study

Construction of isopropyl-ß-D-thiogalactopyranoside (IPTG)-inducible ptsHI operons. Plasmid pFDX3851 contains the p15A origin of replication and a tetracycline resistance gene and carries genes ptsH and ptsI, both from E. coli, in tandem under tacOP control. For its construction, E. coli-derived ptsH (ptsHEc) was amplified by using primers 532 and 533 and plasmid pFDX3153 (12) as the template (see Table 2 for oligonucleotides used in this study). The PCR fragment was digested at the SspI sites within the primers and inserted into the DraI site located between tacOP and ptsIEc of plasmid pFDX3161 (12). Similarly, for construction of an artificial B. subtilis-derived ptsH (ptsHBs), the ptsIEc operon under tacOP control, ptsHBs was amplified with primers 534 and 535 and plasmid pTS22 (10) as the template. The PCR fragment was digested with SspI and inserted into the DraI site of plasmid pFDX3161, resulting in plasmid pFDX3853. Plasmid pFDX3877 carrying ptsH and ptsI, both from B. subtilis, in tandem under tacOP control was constructed by a three-fragment ligation combining the ClaI-PstI and the PstI-SmaI fragments of plasmid pFDX3853 with a PCR fragment encompassing ptsIBs, which was amplified with primers 588 and 589 from plasmid pTS20 (10) as the template and digested with SspI and ClaI. Note that the above constructions provided genes ptsH and ptsI irrespective of their species origin with the untranslated leader sequences naturally found in front of the E. coli ptsH and ptsI genes, respectively. In addition, an NdeI site overlapping the ptsH ATG start codon was introduced in these constructs by primers 532 and 534. To construct plasmid pFDX3852, which is isogenic with plasmid pFDX3851 but carries an H15A mutation in ptsHEc, ptsHEc-H15A was amplified by using primers 532 and 533 and plasmid pFDX3223 (12) as the template. The PCR fragment was digested at the SspI sites within the primers and inserted into the DraI site of plasmid pFDX3161. For construction of plasmid pFDX3854, which is isogenic with plasmid pFDX3853 but encodes an H15A change in the ptsHBs gene, ptsHBs-H15A was amplified with primers 534 and 535 and plasmid pLUM1104 (35) as the template. After digestion with SspI, the PCR fragment was inserted into the DraI site of plasmid pFDX3161.


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  		TABLE 2. Oligonucleotides used in this study

Growth monitoring and calculation of generation times. Precultures were grown overnight in Luria broth supplemented with the appropriate antibiotics. The cells were washed and inoculated to an optical density at 600 nm (OD600) of 0.1 in M9 minimal medium supplemented with proline (40 µg/ml), thiamine (1 µg/ml), the appropriate antibiotics, and a PTS carbohydrate (1% [wt/vol]) as the single carbon source as indicated. IPTG was added as indicated. The bacteria were incubated at 37°C at 200 rpm, and the turbidity at 600 nm was recorded periodically. The doubling times (G) were determined from the exponential phases of the growth curves using the formula G = log2 x [({Delta}t)/(logb – logB)], where {Delta}t is the time interval in minutes, b is the OD600 of the culture at the end, and B is the OD600 at the beginning of this time interval.

ß-Galactosidase assays. Determination of ß-galactosidase activities was carried out as described previously (12). Enzyme activities are expressed in Miller units. Standard deviations were below 15%.

Detection of B. subtilis HPr by Western blotting. E. coli cells were grown in M9 minimal medium containing 1 mM IPTG and either 1% glycerol or 1% N-acetyl-D-glucosamine (GlcNAc) as a single carbon source to an OD600 of ~0.5. Of each culture, one OD600 was harvested and resuspended in sodium dodecyl sulfate (SDS) sample buffer. Of these lysates, 1 µg total protein of each was loaded on SDS-15% polyacrylamide gels. After separation, the proteins were blotted to a polyvinyl difluoride (PVDF) membrane and HPr was detected using polyclonal rabbit antibodies directed against B. subtilis HPr as described previously (33). These experiments were carried out twice.

Analysis of the phosphorylation state of HPr in vivo. The phosphorylation state of B. subtilis HPr in vivo was assayed by Western blot analyses. E. coli cells were grown in M9 minimal medium supplemented with IPTG as indicated in Fig. 3. The cells were harvested in the exponential growth phase, washed, and resuspended in 50 mM Tris-HCl, pH 7.5, 200 mM NaCl. Crude cell extracts were prepared by sonication, and after removal of cell debris, 1 µg total protein was loaded on 10% native polyacrylamide gels, allowing the separation of phosphorylated and nonphosphorylated HPr. To demonstrate the (heat-labile) phosphorylation of His15 of HPr, a second aliquot of each crude extract was incubated at 70°C for 10 min before gel electrophoresis. Proteins were blotted, and HPr species were detected as described above.


Figure 3
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  		FIG. 3. Determination of the phosphorylation state of B. subtilis HPr in E. coli. Strains R1279 (pts+ fru+; lane 1), R1969 ({Delta}pts {Delta}fru; lane 2), and the transformant R1969/pFDX3877 carrying ptsI and ptsH of B. subtilis on a plasmid (lanes 3 to 18) were grown in M9 minimal medium supplemented with IPTG and the carbon source as indicated. Protein extracts (1 µg each) were separated on a 10% native polyacrylamide gel, and B. subtilis HPr was detected by Western blotting. Prior to being loaded, an aliquot of each cell extract was heated (70°C, 10 min) to cause the conversion of the phosphorylated into the nonphosphorylated form of HPr.

Random mutagenesis of the B. subtilis ptsH gene and isolation of ptsH mutants. For the introduction of random mutations in ptsHBs, several independent error-prone PCRs were performed as described previously (3) using primers BG151 and BG166 and plasmid pFDX3877 as the template. The PCR products were digested with NdeI (overlapping the ptsH start codon) and SacI (located in the 5' end of ptsI) and subsequently ligated to the NdeI-SacI vector moiety of plasmid pFDX3877. Strain R1969 was transformed with the ligation mixtures, and recombinants were selected either on LB-tetracycline or directly on MacConkey-tetracycline plates supplemented with 1% PTS sugar (D-mannose, D-mannitol, glucitol, or N-acetyl-D-glucosamine). Colonies that exhibited a more intense red coloration were isolated and restreaked on the same type of plates to verify the altered phenotype. The colonies obtained on the LB plates were subsequently replica plated on M9 minimal medium plates supplemented with proline (40 µg/ml), thiamine (1 µg/ml), tetracycline, and either 1% D-mannose, D-mannitol, D-glucitol, or N-acetyl-D-glucosamine as a single carbon source. The plates were incubated at 37°C, and colonies that resumed growth were restreaked on the same type of minimal plate alongside strain R1969 transformed with unmutated plasmid pFDX3877 as a control. Plasmid DNAs from mutants that passed this test and showed improved growth on a PTS substrate were isolated and subjected to HincII restriction enzyme analysis. Plasmids with restriction patterns indistinguishable from that of the original plasmid pFDX3877 were sequenced using primer BG151 (annealing in the tacOP sequence). Thereafter, the plasmids were reintroduced into strain R1969 for subsequent growth monitoring in liquid M9 minimal medium. Due to the cloning strategy, some of these plasmids also carried mutations in the 5' end of the ptsI gene. Those mutants were excluded from further analyses. Sequencing of the error-prone PCR-derived ptsHBs alleles was performed by the Göttingen Genomics Laboratory (G2L) at the Institute of Microbiology and Genetics of the University of Göttingen.


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RESULTS AND DISCUSSION
 
A system for HPr complementation studies in E. coli. In order to study the functional interaction of B. subtilis HPr with proteins of the E. coli PTS in vivo, E. coli strain R1969 was constructed. This strain lacks the ptsH-ptsI-crr operon, coding for HPr, EI, and IIAGlc, as well as the fruB-fruK-fruA operon, coding for the diphosphoryltransfer protein (DTP), fructose-1-P kinase, and EIIFru. DTP is a chimeric protein containing a subdomain homologous to HPr and was shown to substitute for a defect in HPr in PTS sugar utilization and other HPr-controlled processes (12, 32, 36). As expected, the strain was unable to grow on any substrate of the PTS (Table 3). However, on mannose, residual growth with a doubling time of >1,500 min still occurred, and therefore transport of D-mannose at a very low rate might still be possible in this mutant. The component responsible for this phenomenon remains to be determined. Transformation of the {Delta}pts {Delta}fru mutant with the low-copy plasmid pFDX3851, encoding HPr and EI from E. coli (Fig. 1, top), restored growth on N-acetyl-D-glucosamine (GlcNAc), D-mannose, D-mannitol, and D-glucitol. These substrates were chosen because the corresponding PTS transporters EIINag, EIIMan, EIIMtl, and EIIGut possess their own IIA domains for the interaction with HPr rather than relying on IIAGlc (20), which is absent in this strain. On plasmid pFDX3851, ptsH and ptsI are transcribed as an operon from the artificial tac promoter, which is repressed by the LacI repressor protein. The additional presence of the compatible plasmid pFDY226, carrying the lacI gene, rendered growth of the transformant IPTG inducible on the various PTS substrates. The addition of increasing IPTG concentrations led to increasingly faster growth and thus to decreasing generation times (Table 3 and Fig. 2A). The highest growth rates were obtained in the presence of 0.1 mM IPTG, with generation times of ~70 min on GlcNAc, D-mannitol, and D-mannose and of 119 min on D-glucitol. These growth rates were comparable to the rates obtained with the parent wild-type strain (pts+ fru+) of strain R1969. The doubling times of the parent strain (R1279) were 75 min on GlcNAc, 91 min on D-mannitol, and 76 min on D-mannose. Full induction of expression by the addition of 1 mM IPTG slightly diminished the growth rates, suggesting that overexpression of HPr and/or EI slightly impairs growth. The data suggest that the cellular amounts of the HPr and EI phosphotransferases are growth limiting when the cultures are induced with IPTG concentrations below 0.1 mM. Substitution of the active-site His15 in HPr by Ala abolished growth on the PTS substrates (Table 3), verifying that none of the four remaining HPr paralogues (37) is able to substitute for HPr in PTS sugar utilization under the conditions used. Assuming that the differences in generation times determined in this complementation system reflect different phosphoryl transfer rates towards the tested substrates, the system appeared to be suited for studying the in vivo interspecies cross-phosphotransfer between PTS components of E. coli and HPr of B. subtilis.


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  		TABLE 3. HPr from B. subtilis cannot replace its homologue in E. colia


Figure 1
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  		FIG. 1. Relevant structures of the basic plasmid constructs used for the in vivo complementation studies. Genes ptsH and ptsI, either from E. coli or from B. subtilis, were placed on low-copy plasmids under the control of the IPTG-inducible Ptac promoter. All plasmids carried the translation initiation sequences of the respective E. coli genes in front of ptsH and ptsI. The sequence between ptsH and ptsI corresponds to the intergenic region naturally found between these genes in E. coli.


Figure 2
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  		FIG. 2. HPr of B. subtilis is unable to substitute for the function of its homologue in E. coli in the utilization of PTS sugars. Strain R1969 ({Delta}[ptsH-ptsI-crr] {Delta}[fruB-fruK-fruA]) was transformed with (A) plasmid pFDX3851 carrying E. coli ptsH and ptsI, (B) plasmid pFDX3853 carrying B. subtilis ptsH and E. coli ptsI, and (C) plasmid pFDX3877 carrying B. subtilis ptsH and ptsI. In addition, plasmid pFDY226 delivering the LacI repressor for the IPTG-inducible expression of HPr and EI was present. Growth tests were performed with these transformants in minimal medium containing the indicated PTS sugar as the sole carbon source. Expression of ptsH and ptsI was induced with different concentrations of IPTG as indicated. The empty strain R1969 is represented in panel A by filled squares.

Replacement of E. coli HPr by its counterpart from B. subtilis results in severe growth defects on PTS-sugars. To study the functional interaction of B. subtilis HPr with the various E. coli PTS partner proteins in vivo, two additional plasmid constructs were tested. Both are isogenic with plasmid pFDX3851 carrying the E. coli ptsH and ptsI genes, but in plasmid pFDX3853 ptsH was replaced by its homologue from B. subtilis, whereas in plasmid pFDX3877 both genes were from B. subtilis (Fig. 1). All plasmids carried the translation initiation sequences of the respective E. coli genes to keep the expression levels comparable. Strain R1969 ({Delta}pts {Delta}fru) was cotransformed with the different ptsH-ptsI expression constructs and the Lac repressor delivery plasmid pFDY226, and growth of the transformants was monitored as before. The transformant which expressed B. subtilis HPr and E. coli EI was unable to grow on D-mannitol and D-glucitol regardless of the presence or absence of IPTG, and it exhibited only slow growth on GlcNAc and D-mannose (Table 3; Fig. 2B). These growth rates were strictly IPTG dependent, with the shortest generation times of ~200 min in the presence of 0.1 and 1 mM IPTG. In comparison to the wild-type E. coli strain R1279 and the transformant carrying both genes from E. coli, the generation times were ~3-fold higher on these substrates (Table 3). Taking these data together, B. subtilis HPr cannot substitute for or only very inefficiently substitutes for its homologue in E. coli in the uptake of the various PTS substrates. In principle, this result could be attributed to a less efficient phosphorylation of B. subtilis HPr by the E. coli EI, as suggested by previous in vitro experiments (24), and/or to a weaker interaction of B. subtilis HPr with the various E. coli EIIs compared to the homologous interactions. In an attempt to distinguish between these two possibilities, growth tests were performed using the transformant carrying ptsH and ptsI from B. subtilis. This arrangement should allow for unrestricted phosphoryl transfer between HPr and EI, leaving the heterologous phosphoryl transfer interactions of HPr with the various E. coli EIIs as a bottleneck. As can be seen (Table 3; Fig. 2C), the growth properties of this transformant were very similar to those of the transformant carrying ptsH from B. subtilis and ptsI from E. coli. Again, no growth on D-glucitol and D-mannitol could be observed, and growth was slow on GlcNAc and D-mannose, exhibiting the shortest generation times of 202 and 217 min, respectively, in the presence of 1 mM IPTG. These data suggest that B. subtilis HPr cannot efficiently phosphorylate the IIA domains of EIINag, EIIMan, EIIMtl, and EIIGut and that in the last two cases, phosphorylation by HPr is too weak to promote growth.

Enzyme I of B. subtilis autophosphorylates in E. coli and efficiently transfers phosphoryl groups to B. subtilis HPr. In principle, it appeared possible that the growth defects resulting from the substitution of EI and HPr of E. coli by the homologous proteins of B. subtilis are due to a weak supply of B. subtilis HPr with phosphoryl groups. This could be caused either by a less efficient autophosphorylation of B. subtilis EI or by an impaired subsequent transfer of the phospho groups to B. subtilis HPr.

In vivo protein phosphorylation experiments revealed that EI of E. coli and EI of B. subtilis autophosphorylate with comparable activities in E. coli. (These experiments and their descriptions are available at http://wwwuser.gwdg.de/~genmibio/goerke/supplemental/reichenbach_2007.pdf). To test whether B. subtilis HPr is readily phosphorylated by its cognate EI in E. coli in vivo, we performed Western blotting experiments. The transformant of strain R1969 expressing EI and HPr of B. subtilis was grown in minimal medium containing either glycerol (a non-PTS carbohydrate) or GlcNAc as a carbon source in the presence of various IPTG concentrations. Subsequently, proteins were extracted and separated on native polyacrylamide gels, and B. subtilis HPr was detected by immunoblotting (Fig. 3). In native gels HPr~P migrates faster than nonphosphorylated HPr. The phosphorylation of HPr at its His15 residue can be easily demonstrated by heating an aliquot of the cell extracts prior to loading on the gel, which causes loss of the phosphoryl group. No signal was detectable in the untransformed mutant strain R1969 ({Delta}pts {Delta}fru) or its parent, wild-type R1279 (pts+ fru+), demonstrating that the antibody used does not cross-react with E. coli HPr or other E. coli proteins (Fig. 3, lanes 1 and 2). In contrast, in the transformant of strain R1969 which expressed EI and HPr of B. subtilis, both phosphorylated and nonphosphorylated HPr were detected in the unheated extracts and signal intensities increased with increasing concentrations of IPTG, as expected (Fig. 3, odd-numbered lanes 3 to 17). Heating caused the conversion of the faster- into the slower-migrating form of HPr, verifying that this fraction of HPr was phosphorylated at His15 (Fig. 3, even numbers of lanes 4 to 18). Quantification of the data revealed that under the various conditions about 70% of the total HPr amount was phosphorylated. No large differences in the HPr phosphorylation patterns were detectable between cells grown on either glycerol or GlcNAc (Fig. 3, compare lanes 7 to 12 with lanes 13 to 18), indicating that the slow PTS-dependent growth on GlcNAc consumes only a small amount of the phosphoryl groups provided by HPr (note that due to the growth impairments in GlcNAc, IPTG concentrations lower than 0.05 mM could not be tested). Taking these findings together, we conclude that EI and HPr of B. subtilis are efficiently phosphorylated in E. coli. Therefore, the growth defects seen in the presence of the B. subtilis PTS enzymes (Table 3) are attributable to the inefficient interaction of B. subtilis HPr with the various E. coli EIIs.

HPr of B. subtilis can functionally replace its homologue in the regulation of antiterminator protein BglG activity in E. coli. Next, we wanted to determine whether the impaired interaction of B. subtilis HPr with the E. coli EIIs also extends to interaction partners other than the EIIs. HPr of E. coli phosphorylates the transcriptional antiterminator protein BglG, leading to its activation (12). BglG positively controls expression of the bgl operon which codes for BglG itself, the ß-glucoside-specific EIIBgl, and other functions required for the utilization of aryl-ß-glucosides (29). The activity of BglG is in turn antagonistically controlled by dual EIIBgl- and HPr-catalyzed phosphorylations at its two PTS regulatory domains (12). In the absence of a substrate, EIIBgl phosphorylates BglG at PRD1, leading to its inactivation (4). The presence of a substrate results in the reversal of the process. In addition to the EIIBgl-catalyzed dephosphorylation, BglG requires phosphorylation by HPr at PRD2 for activation (11).

To test whether HPr of B. subtilis is able to regulate activity of BglG, we made use of a reporter plasmid that carries the bgl-t2 terminator downstream of a constitutive promoter and upstream of the lacZ reporter gene (Fig. 4) (12). The ß-galactosidase activity produced by this plasmid reflects activity of BglG which is required for inactivation of the bglt2 terminator and thus for expression of lacZ. BglG and its negative regulator EIIBgl are encoded in trans on the chromosome, and their expression is inducible by IPTG. Therefore, only low ß-galactosidase activities are detectable in the absence of IPTG (Fig. 4, white bars). Addition of IPTG did not result in higher activities unless salicin, a substrate of EIIBgl, was also added, reflecting the known negative regulation of BglG by EIIBgl (Fig. 4, compare black and gray bars in line 1). Deletion of the ptsHI-crr and fruBKA operons led to low ß-galactosidase activities due to lack of activation of BglG by HPr or its paralogue DTP (Fig. 4, bars 2) (12). Transformation of this strain with plasmid pFDX3851 carrying the genes coding for E. coli HPr and EI (Fig. 1) completely restored the regulation of BglG activity; i.e., BglG was kept repressed by EIIBgl in the absence of salicin, and it became fully active in its presence (Fig. 4, bars 3). Substitution of the active-site His15 in HPr completely abolished BglG activity, since it could no longer be phosphorylated by HPr (Fig. 4, black bar, line 4). Transformation of the {Delta}pts {Delta}fru mutant with plasmid pFDX3853, encoding HPr of B. subtilis and EI of E. coli, resulted in 40% of the activity obtained with the transformant expressing the cognate E. coli HPr protein (Fig. 4, compare gray bars in lines 5 and 3). In the absence of salicin (but the presence of IPTG), low BglG activity was measured (Fig. 4, black bar in line 5), suggesting that EIIBgl is sufficiently provided with phosphoryl groups by B. subtilis HPr to keep BglG fully inactive. In the transformant expressing HPr and EI of B. subtilis, activation of BglG was considerably improved and reached 64% of the activity level measured in the transformant expressing the cognate E. coli proteins (Fig. 4, compare the gray bar in line 7 with that of lines 5 and 3).


Figure 4
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  		FIG. 4. HPr of B. subtilis is able to regulate antiterminator protein BglG activity in E. coli. Genes bglG and bglF were cloned under tacOP control and subsequently integrated into the chromosomes of the {Delta}[ptsH-ptsI-crr] {Delta}[fruB-fruK-fruA] mutant strain (strain R1977; bars 2 to 7) and of the corresponding wild-type (wt) strain (strain R1752; bar 1). These strains were transformed with the different ptsH and ptsI expression plasmids schematically shown in Fig. 1 and their derivatives carrying H15A changes in ptsH. In addition, antitermination reporter plasmid pFDX3158 was present. Inducers were added as indicated, and the ß-galactosidase activities were determined.

These results suggested that B. subtilis HPr is able to substitute for its E. coli counterpart in the regulation of BglG activity, and hence its interaction with BglG appears to be more effective in comparison to the interactions with the EIIs. B. subtilis possesses a system for aryl-ß-glucoside utilization that is very similar to the E. coli bgl operon. In this system the antiterminator protein LicT of the BglG/SacY family positively controls expression of the bglPH operon via a transcriptional antitermination mechanism reminiscent of the control of the E. coli bgl operon (28). LicT is the closest relative of BglG in gram-positive bacteria (42% amino acid sequence identity) and, similarly to BglG, its activity is antagonistically controlled by EIIBgl and HPr-catalyzed phosphorylations at its PRDs (19, 38). Interestingly, the E. coli bgl operon is located on a genomic island and is not present in all E. coli strains (K. Schnetz, personal communication). The codon usage of the E. coli bgl genes is similar to that of a low-GC gram-positive bacterium (29) (http://www.kazusa.or.jp/codon/), and the GC content of the bglG gene is almost identical to that of licT (42.9% and 42.5%, respectively) and is typical for genes of low-GC gram-positive bacteria. This raises the possibility that the bgl operon was acquired late in proteobacterial evolution by a horizontal gene transfer event from a low-GC gram-positive bacterium like B. subtilis. Therefore, BglG might have kept more of the characteristics required for an effective interaction with HPr of a gram-positive bacterium.

Identification of amino acid changes in HPr of B. subtilis which improve the phosphoryl transfer towards the E. coli enzymes II. Our data suggest that HPr of B. subtilis is unable to efficiently phosphorylate EIIs of E. coli in vivo (Table 3). The EIIs tested here belong to different families, i.e., the glucose/glucoside family (EIINag), the mannose family (EIIMan), the fructose/mannitol family (EIIMtl), and the glucitol family (EIIGut). They are evolutionary unrelated, and their IIA domains share only weak homology. Homologues of these EIIs with identical substrate specificities also do exist in B. subtilis, except for EIIGut (23). The IIA domains of the mannitol-specific EIIMtl and of the mannose-specific EIILev of B. subtilis share 42% and 37% homology with the corresponding IIA domains of the E. coli EIIMtl and EIIMan proteins, respectively. Therefore, it is a paradox that in its natural host, B. subtilis HPr is flexible enough to interact with a number of proteins sharing no or only limited homology and that at the same time it is unable to phosphorylate EIIs of E. coli for which highly homologous counterparts exist in B. subtilis. Hence, species-specific determinants must exist in HPr of E. coli that are required for its interactions with the cognate EIIs, and HPr of B. subtilis lacks these features.

In order to elucidate these specificity determinants, we introduced random mutations into ptsH of B. subtilis and screened for variants of HPr that improved growth of the E. coli {Delta}pts {Delta}fru mutant on the different PTS substrates. The strategy for mutagenesis was to amplify B. subtilis ptsH by error-prone PCR and to subsequently replace the wild-type gene in plasmid pFDX3877 (carrying ptsH and ptsI from B. subtilis). Several thousand recombinants were screened, and candidates were isolated which exhibited higher growth rates or better fermentation responses on the different PTS substrates than the majority of the colonies. The plasmids of clones that reproducibly showed improved growth properties were isolated and sequenced. Altogether, 26 plasmids were isolated, of which 19 carried a single mutation in ptsH, while 7 plasmids had two or more (Table 4). All mutations resulted in amino acid changes. The single mutations resulted in 10 different amino acid changes in altogether nine positions (Fig. 5A). These were residues Ala16, Arg17, Thr20, Gln24, Ser27, Ile47, Met51, Asp72, and Asn75. For Met51, two different replacements were found. Of the 10 different amino acid changes, 3 were found at least twice, supporting their importance for the interaction with the EIIs of E. coli. Of the seven ptsH alleles carrying more than one change, five had a mutation that was also detected as a single mutation, i.e., the changes Q24R, S27R, and M51L. The Q24R change was present in three of the multiple mutants: twice in combination with a G87S change and once in combination with an S12Y mutation. The latter change was additionally found in combination with a K28N change, indicating that it contributes to the improved interaction with the EIIs. Except for the D72G and N75D changes, all other changes identified in the single mutants clustered in helix {alpha}1 or its vicinity (S12Y) and in helix {alpha}2 (Fig. 5A and 6). These are the regions in HPr known to interact with the various PTS partner proteins. In E. coli HPr the conserved Arg17 residue and the lysines present at positions 24 and 27 form salt bridges to negatively charged Asp or Glu residues present in the various IIA domains. The side chains of residues Asn12, Thr16, Ala20, Leu47, and Gln51 all contribute to the hydrophobic core of the interaction surface, and the methyl group of Thr16 was identified as the central component of this core. In addition, Asn12 and Gln51 form H bonds with residues within the IIA domains (6, 41, 44). Obviously, B. subtilis HPr is incapable of forming these interactions, and the changes identified here may restore them (see below). However, in principle it is also possible that the identified mutations (Table 4) increased the stability or expression of B. subtilis HPr, which could result in a better supply of the various EIIs with phosphoryl groups and hence in better growth on PTS substrates. To investigate this possibility, we grew the various transformants expressing B. subtilis HPr or its mutants in minimal medium in the presence of 1 mM IPTG and performed Western blotting experiments using antiserum directed against B. subtilis HPr. As revealed by two independent experiments, the mutant HPr proteins were not produced at higher amounts than the wild type, irrespective of whether glycerol or GlcNAc was present as a single carbon source (Fig. 5B and data not shown).


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  		TABLE 4. Mutations identified in the screen for B. subtilis HPr variants with an improved interaction with the E. coli enzymes II and their effects on growth on PTS sugarsa


Figure 5
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  		FIG. 5. Amino acid substitutions identified in the screen for B. subtilis HPr variants with an improved interaction with E. coli EIIs. (A) Amino acid sequence alignments of HPr proteins of gram-negative and -positive bacteria. The positions of the {alpha}-helices and ß-sheets in HPr are indicated at the top. The mutations in B. subtilis HPr resulting in a better interaction with the E. coli EIIs are indicated by arrows. Residues conserved in all HPr proteins are in boldface. Residues which differ between gram-negative and -positive bacteria but are conserved within each group are highlighted in light and dark gray, respectively. (B) Comparison of the cellular amounts of wild-type B. subtilis HPr (lane 2 and 11) and its various mutants by Western blotting. The transformants from Table 4 were grown in M9-glycerol supplemented with 1 mM IPTG, and total cell protein was separated by SDS-PAGE. The proteins were blotted on PVDF membranes and probed with antiserum directed against B. subtilis HPr. As a control, the empty strain R1279 was employed in lanes 1 and 10 ("none").


Figure 6
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  		FIG. 6. Amino acid substitutions enhancing the interaction of B. subtilis HPr with the E. coli EIIs and their location within the structure of HPr. A ribbon representation of the structure of HPr of B. subtilis (according to reference 15) is shown. Except for changes D72G and N75D (not shown), all other mutations that improve interaction with the E. coli EIIs are located in the {alpha}-helices 1 and 2 and in the loop preceding {alpha}1. These regions form the interaction surface of HPr. The active-site H15 is shown in boldface. The figure was generated using Pymol software.

Impact of the individual amino acid changes in helices {alpha}1 and {alpha}2 of B. subtilis HPr on the interaction with the various E. coli EIIs. Next, we wanted to determine quantitatively the impact of the amino acid changes in B. subtilis HPr on its functional interaction with the different E. coli EIIs. To address these questions, strain R1969 was transformed with the various mutant plasmids and the generation times were determined (Table 4). The data show that each of the individual mutations in HPr stimulated growth on at least one of the four different PTS substrates. However, not all mutations improved growth on all substrates simultaneously to similar degrees. The individual mutations had comparable effects on the growth on GlcNAc and D-mannose. Globally, the changes S12Y, A16V, T20A, Q24R, and S27R, all located in helix {alpha}1 or in its vicinity (Fig. 5A and 6), had the most pronounced stimulatory effect on the growth velocity on these substrates, with the T20A change yielding the strongest positive effect. The Q24R and S27R changes introduce positively charged Arg residues able to substitute in salt bridge formation for the lysines present in E. coli HPr. Similarly, the A16V change introduces a methyl group in the correct position and at a distance capable of forming the hydrophobic interactions normally carried out by a threonine in E. coli HPr. The positive effect of the T20A change is obvious, since it introduces the residue naturally found at this position in E. coli HPr. The side chain of the Ser12 residue in B. subtilis HPr is presumably too short, and the S12Y change might introduce a residue with the features required to form the interactions carried out by Asn12 in E. coli HPr. Of the changes identified in helix {alpha}2, only the M51L substitution accelerated growth on GlcNAc and D-mannose. A leucine should be able to perform the hydrophobic interactions carried out by Gln51 in E. coli HPr and for which a Met residue is unfavorable. The changes D72G and N75D are located outside of the interaction surface of HPr (Fig. 5A) and caused less pronounced growth improvements than other mutations (Table 4). The N75D mutation introduces a negative charge where in E. coli HPr a negatively charged glutamate is present, and the D72G change removes a negative charge where E. coli HPr contains a positively charged lysine. Hence, these mutations bring the charges in line with the charges present at the corresponding positions in E. coli HPr. A previous work showed that mutation of residue Asp69 or Glu70 located in the vicinity impairs the phosphorylation of EIIs by E. coli HPr (16). Presumably, mutations in this region affect the binding interface of HPr, perhaps by distortion of its overall structure.

Regarding the utilization of mannitol, a different picture was obtained. On this substrate the changes A16V and R17H had the most pronounced positive effect on growth (Table 4). Interestingly, the R17H change completely abolished growth of the transformant on GlcNAc and mannose and had no growth-enhancing effect on glucitol, suggesting that it exclusively improves the phosphoryl group transfer towards EIIMtl and at the same time impairs interaction with the other EIIs (Table 4). This finding is in agreement with studies demonstrating that an R17H mutation in E. coli HPr reduced the phosphorylation of EIINag and EIIMan more than 100-fold in vitro, whereas phosphorylation of EIIMtl was unimpaired (1). As an exception, the Arg17 residue is not involved in salt bridge formation in the HPr/EIIMtl complex (6), which explains why the R17H mutation inhibits interaction of B. subtilis HPr with all E. coli EIIs except EIIMtl. In addition, H bonds formed by the backbone amide protons of Arg17 have a role in orienting the phosphoryl group at His15 to allow for favorable electrostatic interaction with the target proteins (15, 21, 39). The R17H change in B. subtilis HPr might alter this orientation in a way that favors the transfer of the phospho group to the E. coli IIAMtl domain. On glucitol the mutations T20A, Q24R/G87S, I47V, and M51I yielded the strongest growth improvements. Analogously with the effects of the R17H change (see above), the changes I47V and M51I improved growth on glucitol but simultaneously delayed growth on GlcNAc and mannose.

We conclude from these experiments that (i) subtle differences in the composition of the interaction surfaces of the HPr proteins of E. coli and B. subtilis are primarily responsible for the inefficient phosphoryl transfer from B. subtilis HPr to the E. coli EIIs and (ii) EIIMan and EIINag have similar requirements for an effective interaction with HPr, whereas (iii) these requirements should be somewhat different for EIIMtl and EIIGut. With the exception of the salt bridges formed by Lys24 and Lys27 of E. coli HPr, many features of the interaction surface of HPr seem to be preserved between gram-negative and -positive bacteria (6). Examples are the mainly hydrophobic character of the side chains of the residues 16, 20, 47, and 51 or the hydrogen bonding carried out by residue 12 (Fig. 5A). However, our results suggest that the residues present at these positions in B. subtilis HPr cannot form the required interactions. The residues at these positions are highly conserved in gram-negative and in gram-positive bacteria, but they differ between these two groups of bacteria (Fig. 5A). Therefore, they may constitute a signature motif that determines the specificity of HPr for either gram-negative or gram-positive EIIs. It is evident that the species specificity of HPr provides a barrier for the horizontal transfer of genes coding for EIIs. An EII acquired by a gram-positive host from a gram-negative donor would initially be inactive or only weakly active and require mutations in the interaction interface to become functional. However, phylogenetic analyses suggest that the multiple EIIs present in one organism were obtained by extensive lateral gene transfer rather than by gene duplications (46). It is tempting to speculate that some of the EIIs of unknown functions encoded in the genome of, e.g., E. coli (37) are derived from recent horizontal gene transfer events and are still in transition to functional transporters.


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ACKNOWLEDGMENTS
 
This work was supported by a grant of the Deutsche Forschungsgemeinschaft (GO 1355/2-1) to Boris Görke and by the Freiburg Graduiertenkolleg Biochemie der Enzyme.

We thank José Luis Neira for helpful comments on the manuscript and Kalpana D. Singh for advice on the detection of P~HPr.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of General Microbiology, Institute of Microbiology and Genetics, Georg-August University, Grisebachstrasse 8, D-37077 Göttingen, Germany. Phone: (49) 551 393796. Fax: (49) 551 393808. E-mail: bgoerke{at}gwdg.de Back

{triangledown} Published ahead of print on 20 April 2007. Back

{dagger} Present address: F. Hoffmann-La Roche Ltd., CH-4070 Basel, Switzerland. Back


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Journal of Bacteriology, July 2007, p. 4603-4613, Vol. 189, No. 13
0021-9193/07/$08.00+0     doi:10.1128/JB.00236-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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