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Journal of Bacteriology, February 2003, p. 929-937, Vol. 185, No. 3
0021-9193/03/$08.00+0     DOI: 10.1128/JB.185.3.929-937.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

In Vivo Analysis of HPr Reveals a Fructose-Specific Phosphotransferase System That Confers High-Affinity Uptake in Streptomyces coelicolor

Harald Nothaft, Stephan Parche,{dagger} Annette Kamionka, and Fritz Titgemeyer*

Lehrstuhl für Mikrobiologie, Friedrich-Alexander-Universität Erlangen-Nürnberg, D-91058 Erlangen, Germany

Received 26 August 2002/ Accepted 30 October 2002


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ABSTRACT
 
HPr, the histidine-containing phosphocarrier protein of the bacterial phosphotransferase system (PTS), serves multiple functions in carbohydrate uptake and carbon source regulation in low-G+C-content gram-positive bacteria and in gram-negative bacteria. To assess the role of HPr in the high-G+C-content gram-positive organism Streptomyces coelicolor, the encoding gene, ptsH, was deleted. The ptsH mutant BAP1 was impaired in fructose utilization, while growth on other carbon sources was not affected. Uptake assays revealed that BAP1 could not transport appreciable amounts of fructose, while the wild type showed inducible high-affinity fructose transport with an apparent Km of 2 µM. Complementation and reconstitution experiments demonstrated that HPr is indispensable for a fructose-specific PTS activity. Investigation of the putative fruKA gene locus led to identification of the fructose-specific enzyme II permease encoded by the fruA gene. Synthesis of HPr was not specifically enhanced in fructose-grown cells and occurred also in the presence of non-PTS carbon sources. Transcriptional analysis of ptsH revealed two promoters that are carbon source regulated. In contrast to what happens in other bacteria, glucose repression of glycerol kinase was still operative in a ptsH background, which suggests that HPr is not involved in general carbon regulation. However, fructose repression of glycerol kinase was lost in BAP1, indicating that the fructose-PTS is required for transduction of the signal. This study provides the first molecular genetic evidence of a physiological role of the PTS in S. coelicolor.


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INTRODUCTION
 
Bacteria have developed mechanisms to regulate the utilization of carbon sources in a coordinated manner so that preferred carbon sources are utilized first. This has been well documented for Escherichia coli and other gram-negative bacteria, as well as for low-G+C-content gram-positive bacteria, including Bacillus subtilis (33, 41). In both bacterial groups, proteins of the phosphotransferase system (PTS) catalyze the uptake of more than 20 different carbohydrates by sequential phosphoryl group transfer from phosphoenolpyruvate (PEP) to histidyl residues of the general phosphotransferases enzyme I (EI) and from HPr to the IIA domain of a carbohydrate-specific enzyme IIABC(D) complex. The IIA domain in turn phosphorylates the IIB domain, which phosphorylates the substrates that enter via the membrane-embedded IIC(D) transport channel. The phosphorylation state of PTS proteins further reflects the nutritional state of the cell, which leads to global carbon regulation by mechanisms known as carbon catabolite repression (CCR), inducer exclusion, and chemotaxis. In low-G+C-content gram-positive bacteria the HPr protein plays a major role. Under certain conditions HPr becomes phosphorylated by ATP at a regulatory serine site, a reaction that is catalyzed by the metabolite-activated HPr kinase/phosphatase. Seryl-phosphorylated HPr operates in CCR as a corepressor of catabolite control protein CcpA and in inducer exclusion by inhibition of sugar permeases (4). The histidyl-phosphorylated form triggers sugar-specific regulators and catabolic enzymes, such as glycerol kinase.

Carbon metabolism and regulation are much less well understood in high-G+C-content gram-positive bacteria, including Streptomyces coelicolor (8). As soil bacteria, streptomycetes are able to degrade many abundant biopolymers, like cellulose, chitin, and xylan. When S. coelicolor grows on glucose or other preferred low-molecular-weight carbohydrates, it down-regulates degradation of these biopolymers or generally less favored carbon sources by CCR to control catabolic gene activity (8). Mutations in several bld genes, in ccrA, and in glkA, encoding glucose kinase, cause a loss or alteration of CCR (6, 13, 17). bld mutants are also blocked in early stages of differentiation, antibiotic production, and cell-cell signaling, indicating that there is a relationship between carbon metabolism and fundamental cellular processes (31). How signals of carbon source regulation are transmitted is not known, and furthermore, not a single uptake system that causes transport of a carbohydrate that exerts CCR has been characterized.

Using a biochemical approach, we previously found that Streptomyces species possess a fructose 1-phosphate-forming PTS, while they do not possess a glucose-specific PTS (43, 44). Genome analysis of the S. coelicolor PTS complement revealed the presence of four potential enzyme II permeases and the general phosphotransferases EI and HPr (www.sanger.ac.uk/Projects/S_coelicolor/) (2, 28). The results of in vitro experiments showed that HPr encoded by the ptsH gene has the capacity of a typical general HPr phosphotransferase, which was demonstrated by EI-dependent HPr phosphorylation at a histidine residue and subsequent phosphotransfer to various enzyme II proteins (19, 29). HPr of S. coelicolor was also phosphorylated by ATP by using B. subtilis HPr kinase/phosphatase, suggesting that it plays a potential regulatory role. Butler et al. reported construction of a ptsH deletion mutant of S. coelicolor (5). However, no in vivo function of HPr was inferred with respect to carbon utilization or CCR. In this paper, we describe in vivo analyses of a similarly constructed ptsH mutant that has a fructose-negative phenotype. We provide evidence that HPr is essential for high-affinity fructose uptake via enzyme IIFru encoded by the fruA gene. We show that the fructose-PTS is the major uptake system for this sugar and that inactivation of this PTS eliminates fructose repression. We also explain why this HPr function was not detected in the previously constructed ptsH mutant strain.


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MATERIALS AND METHODS
 
Bacterial strains and growth conditions. S. coelicolor A3(2) M145 (SCP1- SCP2- prototroph) was used as the wild-type strain (14). S. coelicolor M701 ({Delta}ptsH::hyg) was kindly provided by Mike Butler and Merv Bibb (5). Escherichia coli DH5{alpha} was the host strain used for subcloning experiments (1). E. coli FT1 {Delta}ptsHIcrr Kanr(pLysS Cmr) harboring pFT3 (ptsH+) and E. coli M15(pREP4) harboring pAG3 (Bacillus subtilis ptsI+) or pFT35 (S. coelicolor ptsI+) were used to produce histidine-tagged S. coelicolor HPr and histidine-tagged EI, respectively (11, 29). E. coli LR2-175 fruA was used for heterologous expression of S. coelicolor fruA (21). B. subtilis QB5350 ptsH-H15A was the source of the complementing cell extract required for the S. coelicolor HPr activity assay (42). Cultures of S. coelicolor were grown for 30 to 72 h with vigorous shaking at 28°C by using tryptic soy broth without dextrose (TSB) (Difco) as a complex medium or mineral medium (MM) (28). Carbohydrates and Casamino Acids (CAA) were added to final concentrations of 50 to 100 mM and 0.1%, respectively. MM agar plates were prepared with only 0.6% Noble agar (Difco) to reduce growth on agar as a carbon source. S. coelicolor strains were grown in 500 ml-baffled flasks containing 200 ml of MM to obtain growth curves. The medium was inoculated by using 2 x 105 spores ml-1. To determine the dry weight, 10 ml of a mycelium-containing culture was filtered through an NC45 filter (Schleicher & Schuell) and dried in a microwave oven for 3 min at 800 W. B. subtilis and E. coli cultures were grown in Luria-Bertani broth at 37°C.

Construction of plasmids and isolation of a ptsH mutant and a fruA mutant. A ptsH gene disruption mutant was derived as follows. A 1,557-bp fragment containing 56 bp of the 3' end of ptsH was subcloned into the SmaI site of the low-copy vector pSU2718, yielding plasmid pFT21 (26). The fragment was generated by PCR with Taq DNA polymerase by using cosmid SC9B10 DNA as the template and oligonucleotides HN1 (AGCATATGGCGAAGCTGGTCGCCG) and HN2 (CCCATATGCAGCTGGCCGCGGACG). The region upstream of ptsH was cloned downstream of the apramycin resistance gene (aacC4) of plasmid pHLW1 as a 3.3-kb XbaI-KpnI fragment from cosmid SC9B10, resulting in plasmid pFT22 (3). The XbaI site was located 58 bp upstream of ptsH. The 4.7-kb KpnI-NdeI fragment of pFT22 comprising a 3.3-kb upstream region of ptsH and the apramycin resistance gene was filled in by using T4 DNA polymerase and was transferred into the PstI site of pFT21 filled in with T4 DNA polymerase, resulting in pFT23. E. coli ET12567 was used to generate unmethylated DNA (25). Plasmid pFT23 was linearized with NcoI, denatured by alkaline treatment, and transformed into strain M145 (14, 27). Protoplasts were plated onto R2YE plates, which were overlaid with soft agar containing apramycin (25 mg/liter) (14). After 20 h of incubation at 28°C, transformant colonies were identified and clonally isolated. One isolate was designated BAP1 and was used for further analyses. The ptsH mutation of BAP1 was verified by Southern blotting. Chromosomal DNA was isolated from S. coelicolor strains by the method of Pospiech and Neumann (32). Twenty micrograms of restricted DNA was separated on a 1% agarose gel and blotted onto a nylon membrane (Qiabrane; Qiagen) (1). Plasmid pFT23 served as the probe and was biotinylated with a translation kit for biotin-7-dATP (BRL). A Photo Gene kit (BRL) was used for signal detection. Only one fragment exhibited an increase in size of about 1.4 kb when wild-type and BAP1 DNA fragments were digested with KpnI, FspI, and PvuII. This size matched perfectly the size of an alteration introduced at the ptsH locus by insertion of the apramycin resistance gene cassette.

A fruA mutant was derived as follows. Cosmid SCE22 DNA was digested with MscI, and a 2,110-bp fragment containing the 3' end of fruK and the first 1,687 bp of fruA was cloned into the EcoRV site of pBluescript SK(+) (pFT131). The apramycin resistance gene (accC4) was prepared from pHLW1 cut with BamHI and treated with T4 polymerase to obtain blunt ends. The fragment was inserted into plasmid pFT131 linearized with SrfI, which yielded plasmid pFT132 ('fruA::aacC4). pFT132 was digested with EcoRI and HindIII, and the fruA::accC4 fragment was cloned into plasmid pWHM3 (10). The resulting plasmid, pFT133, was transformed into M145. fruA mutants were isolated by the procedure described by Fink and coworkers, except that the initial transformant colonies were selected in the presence of both thiostrepton and apramycin (10). After three rounds of sporulation on soy mannitol plates, fruA mutants were identified by replica plating as Aprr Tsrs colonies. The insertion of accC4 into the fruA locus was confirmed by several PCRs. While no pWHM3-specific product was detectable, 2.3- and 1.8-kb amplification products were obtained when an accC4-specific primer (positions 267 and 620 in accC4) was used together with a primer located on the chromosome 13 bp upstream and 17 bp downstream of the 'fruK-fruA' MscI fragment. This demonstrated that the accC4 gene was present at the desired chromosomal position. One mutant, in which the accC4 gene was inserted at bp 585 in the orientation opposite that of the 2,100-bp fruA gene, was designated BAP7.

A ptsH expression plasmid for S. coelicolor, pFT24, was constructed by insertion of a 1,124-bp EcoRI/HindIII fragment from cosmid SC9B10 into plasmid pUWL-KS (45). For constitutive expression of the fruA gene in E. coli, a 2,143-bp fragment was generated by PCR that introduced an optimized E. coli ribosome binding site and stop codons in all reading frames. Cosmid SCE22 DNA served as the template, and oligonucleotides fruA1 (ACTCTAAGTGAGGGGCGATGAGCGAGAGGAGGAGCCCGCGATGAGCG; the start codon is underlined) and fruA2 (GAGTCGCGCTTTACGCGGCCACCGCCTGC; the stop codon is underlined) served as primers for the PCR. The amplification product was cloned into the EcoRV site of pBluescript SK(+), which yielded pFT49, in which the fruA gene was transcribed from the lacZ promoter. All PCR-based constructs were confirmed by DNA sequencing.

Protein purification and membrane preparation. Recombinant His-tagged HPr of S. coelicolor A3(2) M145, His-tagged EI of S. coelicolor and B. subtilis, and enzyme IIFru-containing membranes were purified as described previously (29). Plasmid pFT35 for overexpression of S. coelicolor His-tagged EI was derived by insertion of a 1,692-bp ptsI SacI/HindIII fragment into plasmid pQE30 (Qiagen). The ptsI fragment was amplified from chromosomal S. coelicolor M145 DNA by using primers ptsI1 (CGGGTGAGCTCGAGACAACGCTGCG) and ptsI2 (ACCAAAGCTTGAGCAGCAGGCCGCACCCGGCCCTACTCGC; restriction sites are underlined). The ptsI DNA sequence was identical to the ptsI sequence determined by the genome sequencing project. Protein concentrations were determined by the Bio-Rad protein assay.

Enzyme assays and transport assays. HPr activity was assayed by examining complementation of the glucose-PTS of B. subtilis by measuring the PEP-dependent phosphorylation of methyl [{alpha}-14C]glucoside in the presence of mutant extract of B. subtilis QB5350 (ptsH-H15A) as described previously (29). PEP-dependent phosphorylation of [14C]fructose was assayed with dialyzed cell extracts. Enzyme IIFru specific activity was determined by combining rate-limiting amounts of enzyme IIFru-containing membrane vesicles (10 µg of protein) with an excess of B. subtilis His-tagged EI (1 µg) plus S. coelicolor His-tagged HPr (5 µg). The assay was carried out at 30°C by using a 0.1-ml mixture containing 12 µM (final concentration) [14C]fructose (3 mCi mmol-1). The reaction was linear within the first 45 min. Data were routinely collected after 30 min. Enzyme IIFru was also assayed by examining complementation of the fructose-PTS of E. coli LR2-175 fruA transformed with pFT49 (fruA+) or plasmid pBluescript SK(+). Assays were carried out with dialyzed crude cell extracts in the presence and absence of purified His-tagged HPr and His-tagged EI of S. coelicolor as described above. No activity (controls) was detected in the absence of PEP and/or membrane fractions, indicating the specificity of the PTS assay.

Transport assays were performed by using the method described by Schlösser and Schrempf (39). Strains were grown in 2-liter Erlenmeyer flasks under static growth conditions in 100 ml of MM. Mycelia were harvested during the exponential growth phase (40 h) by centrifugation, washed three times in transport buffer (50 mM Tris-HCl [pH 7.5], 50 mM NaCl, 10 mM KCl), and adjusted to a concentration of 0.4 to 1 mg (dry weight) ml-1. Mycelia prepared for transport assays could be stored at room temperature for up to 4 h. Uptake was initiated by addition of [14C]fructose at a final concentration of 20 µM (5 mCi mmol-1). Samples (1 ml) were taken at different times (10 s to 10 min), rapidly filtered through nitrocellulose filters (NC45), and washed with 0.1 M LiCl. Radioactivity was determined by scintillation oscillography. The initial velocity of [14C]fructose incorporation, which was linear within the first 60 s, was determined by using fructose at different concentrations (0.5 to 200 µM; 20 to 2 mCi mmol-1) by withdrawing 1-ml samples after 1 min of incubation.

Growth and preparation of cell extracts to determine glycerol kinase activity were performed as described previously, with the following modifications (20). Cell debris and membranes were removed by centrifugation for 1 h at 110,000 x g at 4°C. Glycerol kinase activity was assayed by the pyruvate kinase/lactate dehydrogenase assay, except that 50 mM NADH and 10 mM glycerol were used to start the reaction (40). Glucose repression and fructose repression were monitored after growth on 100 mM glycerol for 3 h in the presence or absence of 50 mM fructose or for 20 h in the presence or absence of 50 mM glucose. Data were collected from three independent experiments. The glycerol kinase levels of the wild type and BAP1 were comparable in each experiment, and the activities were between 0.11 and 0.16 U/mg of protein after 3 h of growth on glycerol and between 0.66 and 0.74 U/mg of protein after 20 h of growth on glycerol. Repression factors were within an error range of less than 10%.

Western blotting. Western blot analyses were carried out as described by Parche et al. (29), except that HPr was detected with rabbit polyclonal antibodies raised against His-tagged HPr of S. coelicolor (Eurogentec) (29). Western blot signals were quantified by performing a densitometric gray scale analysis with TINA software (version 2.08; Raytest).

Primer extension. Mycelia of S. coelicolor were pregrown on TSB for 12 h, washed in MM, and then grown for 48 h in MM supplemented with either 50 mM glucose or 50 mM fructose. Mycelia were harvested and resuspended in 100 µl of Tris-EDTA buffer (pH 8.0). Lysozyme was added to a final concentration of 10 mg ml-1, and the suspension was incubated for 1 h at 37°C. Lysis of cells and preparation of total RNA were performed with an RNeasy minikit from Qiagen. Primer extension experiments were carried out with avian myeloblastosis virus reverse transcriptase (Stratagene) and oligonucleotide ptsH-PE (GGCCTTGGCGATCGTCACCG). The oligonucleotide was 5' labeled with T4 polynucleotide kinase. In primer extension reactions 500 fmol of labeled primer was used with 10 µg of total cellular RNA. Reverse transcripts were resolved on 6% polyacrylamide-urea gels. Standard DNA sequencing reactions with Sequenase (U.S. Biochemical Corp.) in which the same oligonucleotide and plasmid pFT5 [pBluescript SK(+) containing 714 bp upstream of ptsH plus the first 114 bp of ptsH] were used were performed to determine the sizes of the primer extension products.

RNA dot blotting. Total RNA of S. coelicolor A3(2) M145 was prepared by using an RNeasy minikit (Qiagen), followed by DNase I (Roche Diagnostics) digestion for 1 h at 37°C. Total RNA was precipitated by addition of 0.1 volume of 3 M sodium acetate and 1 volume of ice-cold isopropanol. RNA concentrations were determined spectrophotometrically and adjusted to 1 µg/µl. The quality of RNA samples was further checked by reverse transcription-PCR to demonstrate that the amounts of 16S rRNA were constant. For detection of fruA mRNA, an antisense RNA (Riboprobe) was generated; an internal fruA fragment (bp 936 to 1352) that was 416 bp long was prepared by PCR and cloned into pBluescript SK(+) digested with KpnI and XbaI by using oligonucleotides fruAint1 (CCCGTCTAGAGCGGCGGCGAGTCCGGCGAGG) and fruAint2 (TATCGGTACCCGCTGGATCGCCAGCACCACC) (restriction sites are underlined), resulting in plasmid pFT108. A T7 RNA polymerase reaction was performed by using a digoxigenin-UTP in vitro transcription kit (Boeringer Mannheim) with an 18-µl (total volume) reaction mixture for 4 h at 37°C; 1 µg of plasmid pFT108 was used as the template. The reaction was stopped by adding 2 µl of EDTA (0.2 M). The riboprobe was precipitated with 2.5 µl of LiCl (4 M) and 75 µl of ice-cold ethanol (96%) overnight at -20°C, followed by centrifugation for 15 min at 4°C. The pellet was washed twice with ice-cold ethanol, dried under a vacuum, and resuspended in 100 µl of diethyl pyrocarbonate-treated Millipore H2O for 30 min at 37°C. The concentration of the probe was 100 ng/µl. To eliminate the possibility that there was contamination with chromosomal DNA, PCR experiments performed with oligonucleotides fruAint1 and fruAint2 resulted in no signal. RNA dot blot experiments were performed by spotting and cross-linking (twice, at 120 kJ) several concentrations of RNA onto a positively charged nylon membrane (Qiabrane; Qiagen) by using an SRC96D dot blot apparatus (Schleicher & Schuell). Hybridization of digoxigenin-labeled RNA probes was detected with Kodak X-OMAT X-ray film by using alkaline phosphatase-conjugated anti-digoxigenin Fab fragments and CSPD* (disodium 3-(4-methoxyspiro{1,2-dioxetane-3,2-(5-chloro)tricyclo[3.3.1.13,7]decan}-4-yl)phenylphosphate; Roche Diagnostics) as a light-emitting substrate. RNA dot blot signals were quantified by performing a densitometric gray scale analysis with TINA software (version 2.08; Raytest).


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RESULTS
 
To investigate the role of HPr in vivo, the ptsH gene was inactivated by gene replacement with the apramycin resistance gene aacC4. The derived mutant was verified by Southern blotting and was designated BAP1 ({Delta}ptsH::aacC4) (see Materials and Methods). Western blotting was performed to determine whether the ptsH gene product was absent. An HPr-specific immunosignal corresponding to a 14-kDa protein was observed in protein extracts of the wild-type strain (Fig. 1a). This signal was not present in extracts of BAP1, showing that no HPr was detectable in the ptsH deletion strain. Loss of HPr activity was monitored by performing a complementation assay of the B. subtilis glucose-PTS. Cell extracts from S. coelicolor wild-type and BAP1 cultures grown on TSB supplemented with fructose were added to B. subtilis QB5350 (ptsH-H15A) cell extract, which lacked a functional HPr. Addition of S. coelicolor wild-type cell extract resulted in the formation of methyl {alpha}-glucoside phosphate (8 nmol min-1 mg of protein-1), demonstrating that there was HPr activity. No HPr activity was detectable when BAP1 cell extract was used instead. Hence, the ptsH gene was inactivated in strain BAP1, resulting in a complete loss of HPr activity.



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  		FIG. 1. Verification, complementation, and growth of the ptsH mutant. (a) Western blot of a sodium dodecyl sulfate-15% polyacrylamide gel to detect HPr protein. Twenty micrograms of protein from a cell extract of M145 (lane 1) or BAP1 (lane 2) grown in TSB in the presence of 50 mM fructose was applied to the gel. Lane 3 contained 10 ng of purified His-tagged HPr. (b) Growth on different carbon sources. Spores of M145, BAP1, and BAP1(pFT24 ptsH+) were streaked on MM plates supplemented with 50 mM fructose (Fru), 50 mM glucose (Glc), or 50 mM glycerol (Gly). Photographs were taken after 72 h of incubation at 28°C. No obvious growth differences weredetected when galactose, glucose, glutamate, lactose, maltose, mannitol, mannose, sucrose, trehalose, or xylose served as the sole carbon source. (c) Growth curves for M145 (•) and BAP1 ({circ}) on MM supplemented with 0.1% CAA and 50 mM fructose.

Growth, pigment production, and sporulation of BAP1 were indistinguishable from growth, pigment production, and sporulation of the congenic wild-type strain M145 when the strains were grown on TSB and on soy mannitol agar plates. Because inactivation of HPr causes a pleiotropic negative phenotype for utilization of many carbon sources in other bacteria, we compared the growth behavior of BAP1 and the growth behavior of M145 on MM supplemented with typical PTS substrates and a number of non-PTS carbohydrates. BAP1 did not grow on fructose, while growth on glucose, mannitol, glycerol, glutamate, and other carbon sources tested was unchanged (Fig. 1b). The fructose-negative phenotype was restored when BAP1 was transformed with plasmid pFT24, which carried an intact ptsH gene. Growth on fructose was monitored over time in MM supplemented with 50 mM fructose and limited amounts of CAA, which were included to support initiation of growth (Fig. 1c). Within the first 35 h, both strains gained similar amounts of biomass. BAP1 then entered the stationary phase, while M145 continued growth for another 20 h. No differences in the growth curves were observed when glucose or glycerol served as the sole carbon source (data not shown).

Sugar uptake experiments were conducted to investigate whether loss of HPr affected transport of fructose. Figure 2 shows the time-dependent fructose uptake of M145 and BAP1 mycelia grown on glycerol and on glycerol plus fructose. BAP1 showed impaired uptake of fructose, whereas M145 incorporated fructose at significant rates. The initial uptake rates were 240 pmol of fructose/mg (dry weight) of mycelia/min when cells were grown on glycerol and about fourfold higher (880 pmol of fructose/mg [dry weight] of mycelia/min) when cells were grown on glycerol plus fructose. It is noteworthy that BAP1 mycelia could incorporate some residual fructose (<40 pmol of fructose/mg [dry weight] of mycelia/min), which was obviously not sufficient to support growth (Fig. 2b) (see below). Figure 2c shows the apparent Km (2 µM) and the Vmax (0.6 nmol/mg [dry weight]/min), which demonstrated that fructose is transported by a high-affinity uptake system. To rule out the possibility that BAP1 is not generally impaired in sugar transport, we performed glucose uptake experiments. The wild type and the ptsH mutant incorporated glucose at similar rates (data not shown).



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  		FIG. 2. Fructose transport. (a) Time course of fructose uptake. Mycelia of M145 and BAP1 were grown in MM supplemented with 50 mM fructose and 100 mM glycerol ({blacksquare} and •) or with 50 mM glycerol ({diamondsuit} and {blacktriangledown}). The uptake experiment was performed with 20 µM [14C]fructose. (b) Magnified version of the lower part of panel a, providing better visualization of the low level of residual fructose transport in BAP1. Values were determined in triplicate. Standard deviations are indicated by error bars. (c) Lineweaver-Burk plot for determination of the Km value of the fructose transport system of M145. Values ({blacksquare}) were determined in triplicate at fructose concentrations ranging from 0.5 to 200 µM. Data were derived from three independent experiments.

The fructose-negative phenotype of BAP1 contrasted with the results of a phenotype analysis of the previously constructed S. coelicolor ptsH mutant M701 (5). Comparative growth analysis revealed that the two strains, although designated S. coelicolor A3(2) M145, exhibited different growth rates on fructose. The wild type used by Butler and coworkers grew markedly slower on fructose, which made it hardly possible to detect a difference from the isogenic ptsH mutant. Transport assays, however, gave clear and consistent results. The two wild-type strains showed identical kinetics for fructose uptake, and both ptsH mutants had lost this ability.

To determine whether HPr is directly required for fructose transport via a fructose-specific PTS, PEP-dependent phosphorylation assays were conducted (Table 1). Phosphorylation of fructose was not detectable in extracts of BAP1 grown in the presence or absence of fructose. When purified HPr was added to cell extracts of BAP1 grown in the presence of fructose, the activity was restored to about wild-type levels (91 nmol of fructose-P/mg of protein/min). This result indicated that residual fructose uptake in BAP1 was sufficient to induce the components of the fructose-PTS but was not sufficient for growth.


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  		TABLE 1. PEP-dependent fructose phosphorylation of ptsH mutant BAP1a

We monitored cellular HPr levels to examine whether synthesis of HPr is stimulated by fructose. HPr-specific immunosignals were readily detectable in cell extracts grown in the presence of fructose or glycerol (Fig. 3). Interestingly, growth on glucose, which is not a PTS substrate (43), yielded a signal that was threefold stronger than the signal present in fructose-grown cells. Transcriptional analysis of the ptsH gene by primer extension experiments revealed two promoter sites (Fig. 4). A leaderless mRNA was detected in glucose-grown cell extracts with a transcription start at a T located four nucleotides upstream of the start codon (18). Growth on fructose led to a transcript that started at a G which was situated 125 nucleotides in front of the ptsH start codon or at a T which was situated 126 nucleotides in front of the ptsH start codon. Thus, transcription of ptsH occurs from two promoter sites in a carbon source-dependent manner.



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  		FIG. 3. Determination of HPr protein levels. The Western blot of a Tricine-15% polyacrylamide gel (38) shows the immunoreactive signal of HPr. Twenty micrograms of cellular protein was subjected to gel electrophoresis. Extracts were prepared from cells grown on MM containing 50 mM glycerol (Gly), 50 mM glucose (Glc), or 50 mM fructose (Fru). Similar results were obtained in three independent experiments. The bar graph shows the relative signal intensities; the amount of HPr found in glucose-grown mycelia was defined as 100%.



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  		FIG. 4. Primer extension and genetic organization of ptsH. (a) Transcriptional start sites of ptsH mRNA determined with mycelia grown on glucose (Glc) and fructose (Fru). Sequence interpretations of mRNA start sites of promoters P1 and P2 are indicated by asterisks. (b) Genetic organization of the ptsH promoter region. The sizes and orientations of open reading frames, promoters P1 and P2, and a 14-bp tandem repeat are indicated by arrows. The ptsH mRNA start sites (asterisks), the ribosome binding site (RBS), and the N-terminal protein sequences of ptsH and orf1039 are indicated. A possible terminator of transcription is indicated by a stem-loop symbol.

The observation that the level of HPr protein was highest in glucose-grown cells, together with the fact that HPr is a global factor in CCR in other bacteria, motivated us to examine whether HPr is involved in glucose repression of glycerol kinase. This enzyme is subject to CCR in diverse bacteria, including S. coelicolor, E. coli, and B. subtilis (33, 36). Mycelia were grown in glycerol in the presence and absence of glucose and fructose. Glycerol kinase activity was reduced in the wild type to 16% of the level observed in the presence of glycerol alone by glucose and to 62% of the level observed in the presence of glycerol alone by fructose (Table 2). In the ptsH mutant there was a comparable glucose effect, while fructose did not reduce glycerol kinase activity. This indicated (i) that HPr is not involved in general CCR and (ii) that the fructose-PTS is required for fructose repression.


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  		TABLE 2. Glycerol kinase activities in the wild type and a ptsH mutant

To further characterize the HPr-dependent fructose-PTS, we determined whether enzyme IIFru activity was stimulated by fructose. Purified EI of B. subtilis and HPr of S. coelicolor were combined with rate-limiting amounts of enzyme IIFru. An enzyme IIFru activity of 9.8 nmol of fructose phosphate min-1 was detected in membrane vesicles of glucose-grown mycelia, compared to the increased activity (28.1 nmol of fructose phosphate min-1) when membrane vesicles from fructose-grown cells were used. Addition of an equimolar amount of glucose to the growth medium did not result in decreased enzyme IIFru activity (27.8 nmol of fructose phosphate min-1). Thus, synthesis of enzyme IIFru was enhanced in the presence of fructose and was not subject to glucose repression.

We previously described an in silico analysis of a putative fruKA operon that encodes homologues of fructose 1-phosphate kinase and enzyme IIFru (28). To demonstrate that fruA is the structural gene for enzyme IIFru, we cloned the corresponding open reading frame in E. coli. Plasmid pFT49 (fruA+) was transformed into the fruA mutant LR2-175, which, however, did not lead to complementation of the fructose-negative phenotype when tests were performed on MacConkey agar plates supplemented with fructose. This could be explained by inefficient expression of FruA or by inefficient interaction of FruA with E. coli HPr. We then prepared cell extracts from LR2-175(pFT49) and measured PEP-dependent fructose phosphorylation (Table 3). While extracts of control strain LR2-175[pBluescript SK(+)] showed no activity, fructose-PTS activity was detected in LR2-175(pFT49) extracts by the formation of 10.1 nmol of fructose phosphate min-1 mg of protein-1. The activity increased to 82.5 nmol of fructose phosphate min-1 mg of protein-1 when purified HPr and EI of S. coelicolor were added.


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  		TABLE 3. PTS-dependent fructose phosphorylation of S. coelicolor enzyme IIFru expressed in E. colia

Transcription of fruA in S. coelicolor was monitored by performing RNA dot blot experiments. When fruA mRNA levels for mycelia grown on glycerol, fructose, or glucose were compared, the level of the fruA transcript was three- to fourfold higher in fructose-grown mycelia (Fig. 5). These data are in good agreement with the results obtained in the transport assay and in the enzyme IIFru activity assay.



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  		FIG. 5. RNA dot blot of fruA mRNA. The hybridization signals of fruA mRNA from mycelia grown on glycerol (Gly), fructose (Fru), and glucose (Glc) are shown. The amounts of total RNA applied are indicated on the left. The bar graph shows a densitometric quantification of the relative signal intensities derived from the 5-µg RNA dot blot signals. The data are the means of three independent experiments. Standard deviations are indicated by error bars.

The fruA gene was disrupted to obtain strain BAP7 fruA::aacC4 (see Materials and Methods). BAP7 exhibited a fructose-negative growth phenotype. An examination of fructose uptake by mycelia grown in mineral medium supplemented with 0.1% CAA and 50 mM fructose gave an uptake rate of 812 ± 28 pmol of fructose mg (dry weight)-1 min-1 for wild-type strain M145 and no detectable uptake for BAP7 (<10 pmol of fructose mg [dry weight]-1 min-1). Fructose-PTS assays were conducted with extracts of mycelia grown on CAA plus fructose. While wild-type extracts formed fructose phosphate (116 ± 9 nmol min-1 mg of protein-1), no activity was detectable in extracts of the fruA mutant. We concluded from the data presented above that enzyme IIFru is encoded by the fructose-inducible fruA gene.


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DISCUSSION
 
In this study we aimed to assess the function of the S. coelicolor HPr protein, a phosphotransferase that plays a central role in carbon metabolism and regulation in many bacteria. Deletion of the ptsH gene revealed a fructose-negative phenotype and showed that HPr is essential for phosphorylation of fructose by the fructose-specific enzyme II permease encoded by the fruA gene. The fructose-PTS was characterized as the major fructose uptake system, which is also required for fructose repression. The ptsH gene is transcribed from two promoter sites in a carbon-source-dependent manner. In contrast to the enzymes in gram-negative bacteria and low-G+C-content gram-positive bacteria (7, 33), HPr of S. coelicolor seems not to be involved in general carbon regulation (5).

Butler et al. have reported the construction of ptsH mutants of S. coelicolor and Streptomyces lividans. They did not detect a fructose-negative phenotype and showed that glucose repression of galactokinase and agarase was not triggered by HPr (5). We confirmed that it was impossible to detect a fructose-negative phenotype in their strain background, because the corresponding wild-type strain grew poorly on fructose. Fructose uptake assays, however, revealed an identical HPr-dependent phenotype in both pairs of strains. The reason for the differences in fructose fermentation is, therefore, not related to differences in the uptake systems and remains to be elucidated. A similar variation in fructose utilization among S. coelicolor strains was also reported in previous publications in which fructose was classified either as a good carbon source or as a poor carbon source (13, 20, 40, 44).

Since E. coli possesses several HPr paralogues, including a fructose-specific HPr, it was speculated that S. coelicolor might have more than one ptsH-like gene (5, 29). Our data, together with the genome analysis data, indicate that S. coelicolor has only one HPr-encoding gene (28).

The analyses of fructose transport indicate that the fructose-PTS is the essential uptake system for fructose. We found that incorporation of some residual fructose was still detectable in the ptsH mutant, while mutation of fruA completely eliminated transport. This suggests that fructose may enter the cell at a low rate via enzyme IIFru in the absence of HPr, a feature that has been reported for fructose- and mannitol-specific enzyme II permeases (22, 23). It should be mentioned that constitutive low-affinity fructose transport systems have been described for S. coelicolor and the twin species Streptomyces violaceoruber, while the inducible high-affinity PTS remained undetected (13, 37). This was probably due to the fact that millimolar sugar concentrations were used in the transport assays, which did not allow detection of high-affinity permeases with Km values in the low micromolar range (15, 33). The uptake data, however, were collected at 10-min intervals, making it impossible to monitor true initial transport rates that occur in the range of seconds. When we performed fructose uptake assays with concentrations in the millimolar range, we could not detect a low-affinity fructose permease (unpublished results).

The fructose-PTS requires EI, HPr, and enzyme IIFru. The corresponding genes are located at three distinct chromosomal loci (28, 29). While ptsH forms a monocistronic operon, ptsI, encoding EI, is linked to the crr gene, which encodes an enzyme IIAGlc-like protein (19, 28). The fruA gene is found downstream of fruK. It encodes an enzyme II belonging to the fructose-mannitol subfamily of PTS permeases with the domain order IIABC (28). Similar fructose-specific enzymes II have been postulated based on in silico analyses of other high- and low-G+C-content gram-positive bacteria and of cell wall-less mycoplasmas (30, 34, 35). Transposon insertions in the putative fruA genes of Mycoplasma genitalium and Mycoplasma pneumoniae resulted in a fructose-negative phenotype, showing that these bacteria also possess just one fructose uptake system (16). A different type of fructose-PTS is present in gram-negative bacteria. These organisms do not use the general HPr but use a second fructose-specific HPr domain that is fused to a IIA domain and in some cases also to an EI domain. The corresponding enzyme II permease has a duplicated IIB domain in the order IIB'BC (9, 12, 24, 46). In all cases the fruA genes are linked to a fruK gene. fruK of S. coelicolor belongs to a family that consists only of fructose 1-phosphate kinases, which are not related to any other type of fructose-converting enzymes. This supports our previous finding that Streptomyces species internalize fructose by an enzyme IIFru that forms fructose 1-phosphate, which subsequently is converted to fructose 1,6-bisphosphate by FruK (44).

Does HPr have additional functions? Our data from the transcriptional analysis and the measurement of HPr protein levels support this possibility. We recently demonstrated that HPr phosphorylates enzyme IIACrr (19), which could be the corresponding IIA protein for three putative enzyme II permeases of the glucose-sucrose subfamily present in S. coelicolor (28). It was suggested that two of these permeases, NagE1 and NagE2, transport N-acetylglucosamine, while the third permease, MalX, could transport a glucose-containing saccharide (28). Hence, it is likely that HPr is required for the functioning of these systems. A general role for HPr in carbon regulation seems unlikely, because no HPr effect was detected in glucose repression of glycerol kinase, agarase, and galactokinase (5). However, the observation that transcription of ptsH is carbon source regulated and involves two promoter sites and the observation that HPr levels are higher in mycelia grown on the non-PTS substrate glucose than in mycelia grown on the PTS substrate fructose may be good reasons to explore another, as-yet-unknown function of HPr in S. coelicolor.


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ACKNOWLEDGMENTS
 
We thank Merv Bibb, Mike Butler, Anke Engels, Knut Jahreis, Matthias Redenbach, and Andreas Schlösser for many discussions, for helpful suggestions, and for gifts of strains and cosmids. This study was carried out in the laboratory of Wolfgang Hillen, whose support is greatly appreciated.

This work was supported by grants from the Graduiertenkolleg Kontrolle der RNA Synthese and by grant SFB473 from the Deutsche Forschungsgemeinschaft.

H.N. and S.P. contributed equally to this work.


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FOOTNOTES
 
* Corresponding author. Mailing address: Lehrstuhl für Mikrobiologie, Friedrich-Alexander-Universität Erlangen-Nürnberg, Staudtstr. 5, D-91058 Erlangen, Germany. Phone: 49 (9131) 8528084. Fax: 49 (9131) 8528082. E-mail: ftitgem{at}biologie.uni-erlangen.de. Back

{dagger} Present address: Nestlé Research Centre, Vers-chez-les-Blanc, CH-1000 Lausanne 26, Switzerland. Back


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Journal of Bacteriology, February 2003, p. 929-937, Vol. 185, No. 3
0021-9193/03/$08.00+0     DOI: 10.1128/JB.185.3.929-937.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.



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