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. 2002 Jul;184(14):3992–4002. doi: 10.1128/JB.184.14.3992-4002.2002

A Repressor Protein, PhaR, Regulates Polyhydroxyalkanoate (PHA) Synthesis via Its Direct Interaction with PHA

Akira Maehara 1,†, Seiichi Taguchi 1,*, Tatsuaki Nishiyama 2, Tsuneo Yamane 2, Yoshiharu Doi 1,3
PMCID: PMC135160  PMID: 12081972

Abstract

Phasins (PhaP) are predominantly polyhydroxyalkanoate (PHA) granule-associated proteins that positively affect PHA synthesis. Recently, we reported that the phaR gene, which is located downstream of phaP in Paracoccus denitrificans, codes for a negative regulator involved in PhaP expression. In this study, DNase I footprinting revealed that PhaR specifically binds to two regions located upstream of phaP and phaR, suggesting that PhaR plays a role in the regulation of phaP expression as well as autoregulation. Many TGC-rich sequences were found in upstream elements recognized by PhaR. PhaR in the crude lysate of recombinant Escherichia coli was able to rebind specifically to poly[(R)-3-hydroxybutyrate] [P(3HB)] granules. Furthermore, artificial P(3HB) granules and 3HB oligomers caused the dissociation of PhaR from PhaR-DNA complexes, but native PHA granules, which were covered with PhaP or other nonspecific proteins, did not cause the dissociation. These results suggest that PhaR is able to sense both the onset of PHA synthesis and the enlargement of the granules through direct binding to PHA. However, free PhaR is probably unable to sense the mature PHA granules which are already covered sufficiently with PhaP and/or other proteins. An in vitro expression experiment revealed that phaP expression was repressed by the addition of PhaR and was derepressed by the addition of P(3HB). Based on these findings, we present here a possible model accounting for the PhaR-mediated mechanism of PHA synthesis. Widespread distribution of PhaR homologs in short-chain-length PHA-producing bacteria suggests a common and important role of PhaR-mediated regulation of PHA synthesis.

Polyhydroxyalkanoates (PHAs) are intracellular carbon and energy storage materials produced in various microorganisms under nutrient-limited conditions (2, 7, 20, 23). PHAs can be classified into two groups, short chain length (SCL) (3 to 5 carbons per monomer) and medium chain length (MCL) (6 to 14 carbons per monomer) (43). Paracoccus denitrificans, a facultative methylotrophic bacterium, accumulates SCL-PHAs from several alcohols (54). Two gene clusters involved in PHA synthesis and degradation in P. denitrificans have been cloned and characterized. The first is the phaA-phaB gene cluster, which encodes β-ketothiolase and acetoacetyl-coenzyme A (CoA) reductase (50), respectively, and the second is the gene cluster phaZ-phaC-phaP-phaR, which encodes PHA depolymerase (9), PHA synthase (47), PHA granule-associated phasin (26), and a DNA-binding regulatory protein (24), respectively.

Phasins are the most dominant proteins of relatively small molecular size that are associated with PHA granules. It has been proposed that phasins form an amphiphilic layer between the PHA granule and cytoplasm in a manner similar to that of oleosins, which form a layer at the surface of triacylglycerol inclusion in oilseed plants (32, 33, 42, 49). These lipid-body-associated proteins exist in several organisms (for a recent review, see reference 30). Phasins also affect the size and the number of PHA granules (15, 16, 49) and positively affect PHA synthesis (8, 15, 26, 32, 33, 36, 40, 42, 49, 53). The production of phasin in Ralstonia eutropha (formerly designated Alcaligenes eutrophus) is suggested to be dependent on the presence of an intact PHA synthesis apparatus, although the mechanism of their regulation is unknown (49). This phenomenon was confirmed rigorously by using several phaC and phaP deletion-replacement mutant strains of R. eutropha (52). The amount of phasin in the cells was also found to be proportional to the content of PHA in P. denitrificans (26). In a heterologous expression experiment in Escherichia coli, overexpression of PhaP occurred in the absence of phaR, while PhaP expression was strongly repressed in the presence of phaR without poly[(R)-3-hydroxybutyrate] [P(3HB)] production. However, expression of PhaP was observed in E. coli even in the presence of phaR when P(3HB) was produced under carbon-rich conditions (26).

Genes that encode proteins homologous to PhaR have been found in pha loci from many SCL PHA-producing bacteria (24, 26). In our previous studies, we found that PhaR is a 22-kDa protein which is able to bind to the upstream regions of both phaP and phaR of P. denitrificans (24). The ability of PhaR to bind to DNA and its regulatory function for PhaP expression in vitro were analyzed by using purified recombinant PhaR produced by E. coli (24).

However, little is known about the detailed regulatory mechanism of PhaR in PHA metabolism (17, 24, 26). Most recently, it was shown that PhaR was involved in P(3HB) synthesis via PhaP expression in vivo, using a phaR knockout mutant of R. eutropha (54). To gain a new insight into the role of PhaR in PHA synthesis, we have analyzed the function of the PhaR protein in more detail. In this paper, we determined the distinct target DNA sequences for PhaR binding and further demonstrated that PhaR binds to not only DNA but also PHA. We found evidence that P(3HB) acts as an inducer for phaP expression in a PhaR-mediated regulatory system. This report also presents direct evidence that PhaR interacts with PHA via direct binding. Furthermore, our observations may raise the possibility that the PhaR-mediated regulatory mechanism in response to PHA accumulation in cells is common in SCL PHA-producing bacteria.

MATERIALS AND METHODS

Bacterial strains, plasmids, and growth conditions.

The bacterial strains used in this study were P. denitrificans (ATCC 17741), E. coli XL1-Blue, E. coli BL21(DE3), and R. eutropha H16 (DSM428). The recombinant plasmid pTV119N::phaR was used for P. denitrificans PhaR production as previously described (24). In addition, we used plasmids pPDPK1.7 (26), pTVC (26), pTVCP4 (26), and pBBRKmAB (26). P. denitrificans was grown aerobically at 30°C in inorganic salt medium (25) containing 1% (vol/vol) methanol or 0.1% (vol/vol) n-pentanol. E. coli was grown at 37°C in Luria-Bertani (LB) medium (47). R. eutropha was cultivated at 30°C in a nutrient-rich medium (28). When needed, ampicillin (100 μg/ml) and kanamycin (50 μg/ml) were added to the media.

Production and purification of PhaR.

PhaR was purified as previously described (24), with some modifications. E. coli BL21(DE3) cells harboring recombinant plasmid pTV119N::phaR were grown at 37°C for 13 h in 500 ml of LB medium. Cells were harvested by centrifugation (8,000 × g; 5 min; 4°C), washed, suspended in T buffer (20 mM Tris-HCl [pH 7.5]), recentrifuged, resuspended in 10 volumes of TEP buffer (20 mM Tris-HCl [pH 7.5]), 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride), and then disrupted by two passages through a French press (138 MPa). Unbroken cells and cellular debris were removed by centrifugation (10,000 × g; 60 min; 4°C), and solid ammonium sulfate was added to the supernatant to 10% saturation. After removing the precipitate, ammonium sulfate was added again to the supernatant to 30% saturation. The precipitates were collected by centrifugation (10,000 × g; 20 min; 4°C), dissolved in 6 ml of P buffer (20 mM potassium phosphate [pH 7.5]), and dialyzed against the buffer. The sample was overloaded onto a column (26 by 100 mm) of phosphocellulose (Whatman P11) previously equilibrated with P buffer. The tailing fractions followed by a compact peak containing most proteins were collected, added to ammonium sulfate (20% saturation), and loaded onto a butyl-Sepharose column (25 ml; Amersham Pharmacia Biotech, Piscataway, N.J.) previously equilibrated with PA buffer (20 mM potassium phosphate [pH 7.5], 1 M ammonium sulfate). After washing the column with PA buffer, PhaR was eluted within a stepwise gradient of 1 to 0 M ammonium sulfate. Fractions containing PhaR were collected, diluted 10-fold with 10 mM Tris-HCl (pH 7.5), and loaded onto a Resource Q column (6 ml; Amersham Pharmacia Biotech) previously equilibrated with T buffer. After washing the column with T buffer, PhaR was eluted within a linear gradient of 0 to 0.5 M NaCl. The purified fractions were combined, dialyzed against T buffer, and stored. Protein concentrations were determined from the absorbance at 280 nm. The molar extinction coefficient at 280 nm (ɛ280; 12,090 M−1 cm−1 for PhaR) was determined by the method of Gill and von Hippel (10), with the amino acid composition derived from the nucleotide sequence. Proteins were separated by sodium dodecyl sulfate (SDS)-12.5% or 15% polyacrylamide gel electrophoresis (PAGE) and were stained with Coomassie brilliant blue (CBB) R-250 as described by Laemmli (19).

Analytical procedures for PhaR.

Matrix-assisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF MS) analysis of PhaR was performed on a REFLEX apparatus (Bruker-Franzen Analytik, Bremen, Germany) with sinapinic acid as a matrix. Dynamic light scattering (DLS) was performed in a DynaPro-801 DLS instrument (Protein Solutions Inc., Charlottesville, Va.) at 25°C using purified PhaR (1.12 mg/ml in 20 mM Tris-HCl [pH 7.5]).

DNase I footprinting experiment.

DNase I footprinting using infrared dye was performed by the method described by Machida et al. (22). DNA fragments containing PhaR-binding sites were prepared by PCR using 50 ng of plasmid pTVCP4, containing the phaP and phaR promoter regions, as template. IRD800 dye-labeled custom primers were made by Aloka (Tokyo, Japan). A combination of 5′-biotinylated and 5′-IRD800-labeled primers were used to introduce biotin and IRD800 fluorescent dye at different ends of the DNA fragments. The fragments were analyzed by electrophoresis on an agarose gel before use and were densitometrically quantitated after ethidium bromide staining with an ATTO Lane & Spot Analyzer equipped with a UV transilluminator (Atto). After purification by gel filtration (MicroSpin S-400 HR; Amersham Pharmacia Biotech), the biotinylated IRD800-labeled DNA fragments were immobilized on streptavidin-coated paramagnetic beads (Dynabeads M-280-Streptavidin; Dynal, Oslo, Norway) according to the manufacturer's specifications. The immobilized fragment (ca. 25 to 200 fmol) was mixed with PhaR (ca. 1 to 25 μg) in 100 μl of buffer (25 mM HEPES-NaOH [pH 7.8], 50 mM KCl, 0.05 mM EDTA, 0.5 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride, and 5% glycerol) at 25°C for 30 min. After addition of MgCl2 (final concentration, 5 mM) and DNase I (7.5 × 10−3 U), the reaction mixture was incubated for 1 min at 25°C and then quenched by addition of 1 volume of ice-cold stop buffer (4 M NaCl, 0.1 M EDTA [pH 8.0]). After washing the beads with BW buffer (10 mM Tris-HCl [pH 7.5], 1 mM EDTA, 2 M NaCl), the beads were suspended in 1.5 μl of loading buffer (95% formamide, 10 mM EDTA [pH 7.6], 0.1% bromophenol blue). Samples for a DNA ladder were prepared by the dideoxy chain termination method using 5′-IRD800-labeled primers with a Thermo Sequenase fluorescent-labeled primer cycle sequencing kit with 7-deaza-dGTP (Amersham Pharmacia Biotech). The samples were denatured at 95°C for 2 min, loaded on a sequencing gel (25-cm length, containing 5.5% Long Ranger [FMC BioProducts], 7 M urea, 0.6× Tris-borate-EDTA [TBE] buffer, 0.5× TBE running buffer), and electrophoresed by using a LI-COR 4000L sequencer with BaseImagIR version 4.0 (LI-COR, Lincoln, Nebr.).

Preparation of native PHA granules.

Poly[(R)-3-hydroxyvalerate] [P(3HV)] granules from P. denitrificans were obtained from cells that were cultivated for 24 h at 30°C in a 300-ml flask in inorganic salt medium containing 0.1% (vol/vol) n-pentanol. P(3HB) granules from E. coli XL1-Blue(pBBRKmAB pTVC) or XL1-Blue(pBBRKmAB pTVCP4) were obtained from cells that were cultivated for 30 h at 37°C in a 100-ml flask culture in LB medium supplemented with 2% sodium lactate. PHA [P(3HB-co-3HV)] granules from R. eutropha were prepared from cells that were cultivated for 55 h at 30°C in a 100-ml flask in a nitrogen-limited mineral salt medium (28) supplemented with 1% fructose and 0.5% sodium pentanoate (0.1% sodium pentanoate was added to the medium five times). The cells were harvested by centrifugation (8,000 × g; 10 min; 4°C), washed, resuspended in 3 volumes of 50 mM Tris-HCl (pH 7.5), and then disrupted by two passages through a French press (138 MPa). After centrifugation (10,000 × g; 30 min; 4°C), the precipitate was suspended in 50 mM Tris buffer. Approximately 200 to 500 μl of the suspension was layered on a discontinuous sucrose gradient consisting of 1 ml each of 2.00, 1.67, 1.33, and 1.00 M sucrose in 50 mM Tris buffer. After centrifugation (210,000 × g; 2.5 h; 4°C), the PHA granule condensed layer was isolated. The native granules were then washed twice with 50 mM Tris buffer, resuspended in the same buffer, and stored at −20°C.

Preparation of artificial PHA granules.

Crystalline P(3HB) granules prepared by the hypochlorite detergent method were purchased from ICI. Artificial amorphous P(3HB) granules were prepared from crystalline P(3HB) by the method described by Horowitz and Sanders (12), using sodium oleate as a surfactant (37).

Preparation of 3HB oligomers by alkaline hydrolysis.

The single crystals of P(3HB) were grown from dilute solution according to a method derived from that of Marchessault et al. (27). For 3HB oligomer preparation, the single crystals of P(3HB) were hydrolyzed by sodium hydroxide solution according to a method previously described (14). For alkaline hydrolysis, P(3HB) single crystals were collected by centrifugation, washed once with distilled water, and resuspended in 0.1 N sodium hydroxide solution. The crystals were hydrolyzed at 37°C for about 16 to 24 h. This solution was not shaken, in order to prevent the single crystals from breaking. The degraded single crystals consisting of 3HB oligomers were washed three times with distilled water to remove the sodium hydroxide solution. The 3HB oligomers were then redispersed in methanol, washed twice by centrifugation, and resuspended in dimethyl sulfoxide. The number-average molecular weight of the prepared 3HB oligomers was about 2 × 103 to 3 × 103, indicating that the 3HB oligomers were approximately 30-mers.

Preparation of antibody.

The purified PhaR protein (about 1 mg) was mixed with Freund's complete adjuvant and injected into a rabbit. Before sensitization to PhaR, a small quantity of serum was prepared from the rabbit. Two weeks after the first sensitization to PhaR, the antigen was again injected into the rabbit, with Freund's incomplete adjuvant. After 3 weeks, immune serum with high immunity toward PhaR antigen was prepared from the rabbit and used for several immunological assays.

Western blot analysis.

Western blotting was done as described by Burnette (5) with cellulose nitrate membrane or polyvinylidene difluoride membrane. In the immunoblot analysis, peroxidase-conjugated anti-rabbit immunogloblin G (Bio-Rad Laboratories) was used as the secondary antibody. The blot was developed with 4-chloro-1-naphthol (39).

Estimation of the ratios of PhaP and PhaR to total PHA granule-associated proteins from P. denitrificans.

The ratios of PhaR and PhaP to total PHA granule-associated proteins from P. denitrificans were determined by densitometric scanning of SDS-PAGE gels stained with CBB. When large amounts of granule-associated proteins were applied to the gel for staining PhaR with CBB, the intensity of the band corresponding to PhaP was already saturated. Because of this saturation, other SDS-PAGE gels handled with exponentially increasing amounts of granule-associated proteins were compared to one another. The contents of PhaR and PhaP were estimated by extrapolating to the obtained intensities.

Rebinding of PhaR to two different P(3HB) granules.

The two kinds of P(3HB) granules (5 mg), crystalline and artificial amorphous, were independently incubated for 30 min at 4°C with supernatant of crude lysate [in 1 ml of TE buffer (10 mM Tris-HCl, pH 8.0; 1 mM EDTA)], corresponding to 10 ml of cells of an E. coli BL21(DE3) strain culture expressing phaR. A control experiment was carried out in parallel with crude lysate prepared from the strain bearing the plasmid vector only (pTV119N). After the incubation, granules were collected by centrifugation, washed twice with 200 μl of TE buffer, and resuspended in denaturing buffer. P(3HB) granule-associated proteins and the crude lysates were analyzed by SDS-15% PAGE.

Gel mobility shift assay using candidate effector molecules.

To determine the effector molecule(s) for PhaR, a gel mobility shift assay in the presence of substances related to P(3HB) metabolism was carried out by the method described previously (24) with some modifications. The DNA fragments used for the gel mobility shift assay were prepared by digesting plasmid pTVCP4 with SalI and SmaI to give 225-, 276-, 407-, 639-, and 4,873-bp fragments (see Fig. 4, lane 1). The resulting DNA fragments (2 μg) were mixed with PhaR (158 ng) in binding buffer (10 mM Tris-HCl [pH 7.5], 1 mM EDTA, 80 mM NaCl, 4% glycerol) in a total volume of 20 μl. The water-soluble metabolites free CoA, acetyl-CoA, acetoacetyl-CoA, (R)-3-hydroxybutyryl-CoA, (S)-3-hydroxybutyryl-CoA, (rac)-3-hydroxybutyric acid (3HB), (R)-3HB-dimer, NAD+, NADH, NADP+, NADPH, acetyl phosphate, citric acid, and phosphoenolpyruvate were each used at a 1 mM concentration. Polyphosphate (final concentration, 0.2 μg/μl) was also tested. Insoluble P(3HB) derivatives, namely, crystalline P(3HB) granules, artificial amorphous P(3HB) granules, and 3HB oligomers (approximately 30-mer), were also tested in this assay. Native PHA granules and partially purified PhaP were also used.

FIG. 4.

FIG. 4.

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Gel mobility shift assay with potential effector molecules for PhaR. The DNA fragments used were generated by SalI/SmaI digestion of plasmid pTVCP4 (24), and the numbers on the left side of the gel indicate the length of the DNA fragments before formation of PhaR-DNA complexes. The 225-bp DNA fragment (DNA225) includes PhaR-binding sites in IRCP (Fig. 1). The 639-bp fragment includes the PhaR-binding site in IRPR (Fig. 1). PhaR (0.158 μg) was added to the mixtures (lanes 2 to 12). An aliquot (5, 10, 20, and 100 μg/lane) of crystalline P(3HB) granules was added to the mixtures (final volume, 20 μl) before incubation for 30 min (lanes 3 to 6). Parentheses indicate that the crystalline P(3HB) granules (100 μg/lane) were added after formation of PhaR-DNA complexes for 20 min (lane 7). Other additives before incubation were as follows: lane 8, 3HB oligomers (20 μg/lane); lane 9, artificial amorphous P(3HB) granules (100 μg/lane); lane 10, native P(3HB) granules from E. coli XL1-Blue(pBBRKmAB pTVCP4) (91 μg/lane); lane 11, native P(3HV) granules from P. denitrificans (79 μg/lane).

After incubation for 30 min at 25°C, the DNA-protein complexes were separated from the unbound DNA fragments on a 5% native polyacrylamide gel, using 0.5 × TBE buffer (39) as the electrophoresis buffer. After electrophoresis, DNA fragments were stained with ethidium bromide and subjected to UV monitoring.

Cell-free protein synthesis.

Cell-free protein synthesis using the E. coli S30 extract system for circular DNA (Promega, Madison, Wis.) was carried out as recommended by the manufacturer, with some modifications. The template DNA vectors used were pPDPK1.7 carrying the phaP gene and pTV119N (as a negative control) (24). PhaR was added to the reaction mixtures before addition of template DNA. A 1-μl suspension of crystalline P(3HB) granules (0.1 mg/μl) was added to the reaction mixture in the presence of P(3HB). Determination of cell-free protein synthesis was performed by adding 4 μg of plasmid DNA to the total volume of 50 μl at 37°C for 60 min and stopping synthesis by adding 200 μl of cold acetone. The precipitates were collected by centrifugation (10,000 × g; 10 min; 4°C) and dissolved in 200 μl of 1× SDS-PAGE sample buffer. After denaturation of proteins (100°C, 5 min), the translation products were separated by an SDS-12.5% PAGE and analyzed by Western blotting.

Database searches and sequence alignments.

Nucleotide sequences potentially coding for proteins similar to PhaR were searched among both GenBank and the unfinished microbial genomes database available on the National Center for Biotechnology Information web page (http://www.ncbi.nlm.nih.gov/Microb_blast/unfinishedgenome.html) using the BLAST alignment program (1). Preliminary sequence data for Burkholderia fungorum, Chloroflexus aurantiacus, Desulfitobacterium hafniense, Magnetococcus sp. MC-1, Magnetospirillum magnetotacticum, Ralstonia metallidurans, Rhodobacter sphaeroides 2.4.1, Rhodopseudomonas palustris, and Sphingomonas aromaticivorans were obtained from the DOE Joint Genome Institute at http://www.jgi.doe.gov/JGI_microbial/html/index.html. Bordetella bronchiseptica, Bordetella parapertussis, Bordetella pertussis, and Burkholderia pseudomallei sequences were produced by the respective sequencing groups at The Sanger Institute (http://www.sanger.ac.uk/). Preliminary sequence data for Burkholderia mallei were obtained from The Institute for Genomic Research website at http://www.tigr.org. The Legionella pneumophila sequence was obtained from the Legionella Genome Project (http://genome3.cpmc.columbia.edu/∼legion/index.html). Protein sequences were aligned by using CLUSTAL W, version 1.8 (45). Based on the alignment of PhaR homologs, a phylogenetic tree was made using TreeView (31).

RESULTS

Purification and structural properties of recombinant PhaR protein.

To facilitate studies on the interaction of PhaR with the phaP and phaR promoter regions, recombinant PhaR was expressed and purified from E. coli BL21(DE3)(pTV119N::phaR) (24) as described in Materials and Methods. The molecular mass of PhaR was determined to be 21,796 Da by MALDI-TOF MS analysis, which is consistent with the deduced amino acid sequence minus the N-terminal methionine. The PhaR native molecular mass was estimated as 96,000 Da by DLS (data not shown), consistent with previous gel filtration chromatography analyses results (93,000 Da) (24), indicating that PhaR forms a homotetramer in the native state.

Identification of PhaR-binding regions of DNA.

Previously, it was demonstrated that PhaR is a DNA-binding protein which blocks phaP expression in vitro (24). The binding sites of PhaR were suggested to be both the phaC-phaP intergenic region (designated IRCP) and the phaP-phaR intergenic region (designated IRPR) (24). To more precisely determine PhaR-binding sites, DNase I footprinting experiments were performed on both IRCP and IRPR, as shown in Fig. 1. The PhaR-binding site in IRCP was shown to have a 56-bp core sequence which overlaps the putative phaP promoter region (Fig. 1A, B, and E). PhaR appeared to protect a 23-bp sequence which partially overlaps the putative phaR promoter region in IRPR (Fig. 1C, D, and F). Several repeated TGC sequences were shared by both intergenic regions and protected against DNase I digestion. These results suggested the presence of a common motif which includes the repeated TGC sequences (designated the TGC-rich region). In IRCP, two TGC-rich regions (TGC I and TGC II) were found (Fig. 1E), whereas one TGC-rich region (TGC III) was found in IRPR (Fig. 1F). A gel mobility shift assay revealed that PhaR binding affinity toward IRCP was higher than that towards IRPR by about 1 order of magnitude (data not shown). This difference in the binding affinities may be due to the difference in the number of TGC-rich regions within each motif.

FIG. 1.

FIG. 1.

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DNase I footprinting analysis of the phaP and phaR promoter regions with PhaR. The fluorescent (IRD800)-labeled biotinylated fragments containing IRCP or IRPR were generated by PCR using IRD800-labeled primers and biotinylated primers. DNase I digestion was performed in the presence (+) or absence (−) of PhaR. Numbers at the left side of the footprints indicate nucleotide positions from the first letter A of the translational start codon (ATG). DNA ladders for sequence analysis were prepared with the same fluorescence-labeled primers used for footprinting analysis. The fluorescence-labeled fragments were incubated in separate reaction mixtures with 0 or 1.18 μg of PhaR for IRCP and with 0 or 23 μg of PhaR for IRPR and then subjected to DNase I digestion. (A) Footprinting of the coding (sense) strand on IRCP. (B) Footprinting of the noncoding (antisense) strand on IRCP. (C) Footprinting of the coding (sense) strand on IRPR. (D) Footprinting of the noncoding (antisense) strand on IRPR. (E and F) Positions of the PhaR-binding sites for IRCP and IRPR, respectively, as deduced from DNase I footprints. The boxed sequences indicate the DNase I-protected regions. −35 and −10 indicate putative σ70 recognizable promoter regions. Thick lines indicate TGC repeat sequences. Arrows denote an inverted repeat sequence capable of forming a stem-loop structure (47).

To examine whether these regions are common in PHA-producing bacteria, DNA sequences of the upstream region of phaP genes from other bacteria were compared. A TGC-rich region was found approximately 90 bp upstream of the phaP-encoding region of R. sphaeroides 2.4.1 (http://www.jgi.doe.gov/tempweb/JGI_microbial/html/index.html) (GCAACGCAAGGCGTTTCGGGCCAGGTGCGAAAAAATGCTGCGGTGC), but no apparent TGC-rich region was found upstream of phaP of R. eutropha (AF314206). If regulatory sequences for other PhaR homologs are determined, then it will be demonstrated that common sequences recognized by PhaR homologs may be found. Another possibility is that each PhaR homolog may recognize a specific, unique target sequence.

Existence of PhaR on native PHA granules in vivo.

We assumed that PhaR is a PHA-responsive repressor, based on the following findings: (i) PhaR homologs have been found in many SCL PHA-producing bacteria (24); (ii) PhaP production seems to be regulated by the presence of phaR in response to P(3HB) production in vivo (26); (iii) PhaR can bind the upstream regulatory sequences for phaP and phaR (Fig. 1; see also reference 24). To examine the localization of PhaR in vivo, SDS-PAGE and Western blotting were carried out for soluble proteins and PHA granule-associated proteins from P. denitrificans, R. eutropha, and two PHA-producing recombinant E. coli XL1-Blue strains carrying phaAB-phaC or phaAB-phaCPR. To further address the function of PhaR, two strains of recombinant E. coli were prepared. One carries pBBRKmAB and pTVC [only for P(3HB) generation], and the other carries pBBRKmAB and pTVCP4 (in coexistence with the regulated PhaP production system) (26). These two E. coli strains were individually cultivated in LB medium (with ampicillin and kanamycin) supplemented with sodium lactate as an excess carbon source to produce P(3HB). P. denitrificans was cultivated in inorganic medium supplemented with n-pentanol to accumulate P(3HV). R. eutropha was grown in mineral salts medium supplemented with both fructose and sodium pentanoate to produce P(3HB-co-3HV). The results are shown in Fig. 2.

FIG. 2.

FIG. 2.

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Localization of PhaR. Proteins were separated in an SDS-15% polyacrylamide gel, stained with CBB R-250 (A, C, and E), and subjected to Western blot analysis employing antibody raised against the recombinant P. denitrificans PhaR (PhaRPd) (B and D). Lanes: 1, molecular mass standard proteins (in kilodaltons); 2 to 4, proteins from P. denitrificans; 5 to 7, proteins from E. coli XL1-Blue(pBBRKmAB pTVCP4) (phaAB and phaCPR are introduced); 8 to 10, proteins from E. coli XL1-Blue(pBBRKmAB pTVC) (phaAB and phaC are introduced); 11 to 13, proteins from R. eutropha; 14, proteins from E. coli XL1-Blue that overproduces PhaR of R. eutropha (PhaRRe). (The detailed construction of expression vector for PhaRRe in E. coli would be published elsewhere.) T, total cellular proteins; S, soluble proteins fraction; G, PHA granule-associated proteins. (E) PHA granule-associated proteins from P. denitrificans. The sample is the same as that in lane 4 of panel A, but a large amount of proteins was applied to the gel and stained with CBB R-250. The positions of PhaR (PhaRPd and PhaRRe) are indicated by black arrowheads. White arrowheads indicate PhaP, PhaC, or PhaZ. A proteolytic degradation product of PhaP [designated PhaP(deg)] was detected on the gel.

In P. denitrificans, PhaR was detected in all fractions of proteins (Fig. 2, lanes 2 to 4). PhaR was detected in the soluble fraction (lane 3) at a much lower level than in the P(3HV) granule fraction (lane 4). The XL1-Blue(pBBRKmAB pTVCP4) strain also produced PhaR (lanes 5 to 7). However, in E. coli organisms without phaPR [XL1-Blue(pBBRKmAB pTVC)], no signal for PhaR was detected in any fractions (lanes 8 to 10), confirming that the 22-kDa signal corresponds to PhaR (lanes 2 to 7). PhaR bound to both P(3HV) granules (lane 4) and P(3HB) granules (lane 7) in vivo, suggesting that the recognition of PhaR for PHA polymer is independent of the structural differences between P(3HB) and P(3HV).

Interestingly, it was demonstrated that a protein (≈19 kDa) of R. eutropha could be detected in total cellular proteins (lane 11) and PHA granule-associated proteins (lane 13) by using polyclonal antibody raised against PhaR from P. denitrificans. This indicates that R. eutropha possesses a ≈19-kDa protein which is immunologically and functionally similar to PhaR of P. denitrificans. A plausible candidate for the ≈19-kDa protein is a PhaR homolog of R. eutropha. This homolog is encoded by a gene (temporarily designated as ORF1) which is located downstream of the phbCAB genes encoding PHA synthesis enzymes (41). The deduced amino acid sequence of the PhaR homolog of R. eutropha (PhaRRe) consists of 183 amino acid residues with a molecular mass of about 21 kDa, and it exhibits high homology to that of P. denitrificans (36.2% identity). The immunological cross-reaction observed with a recombinant protein of PhaRRe and anti-PhaRPd antibody (Fig. 2C and D, lanes 14) demonstrates that the PhaRPd and PhaRRe proteins are closely related to each other in function (binding to PHA granules), localization, and structure.

PHA granule-associated proteins from P. denitrificans.

The ratios of PhaR and PhaP to total PHA granule-associated protein from P. denitrificans were estimated to be 0.3% ± 0.1% and 96%, respectively, by densitometric scanning of SDS-PAGE gels stained with CBB (Fig. 2E). The presence of PHA synthase (PhaC) and PHA depolymerase (PhaZ) in the PHA granule-associated proteins was also revealed by the immunological analysis for PhaC and N-terminal amino acid sequence analysis for PhaZ (data not shown). Figure 2E shows that PhaR exists on native PHA granules from P. denitrificans, although PhaR is not a predominant PHA granule-associated protein.

Rebinding of PhaR to P(3HB) granules.

It was of interest to investigate how PhaR recognizes and tightly binds to PHA granule. A PhaR binding experiment was performed in vitro using two different P(3HB) granules, crystalline and artificial amorphous P(3HB)s. It is known that P(3HB) exists in an amorphous state within the cell but that the extracted P(3HB) proceeds to crystallize (3, 4, 16). Therefore, artificial amorphous P(3HB) granules were prepared and tested for the rebinding experiments in addition to crystalline P(3HB) granules.

Figure 3 shows that PhaR has a high binding affinity toward P(3HB), because PhaR was enriched in the P(3HB) granule-associated fractions (lanes 3 and 4) from the lysate containing PhaR (lane 2). Although increased concentrations of other proteins were detected for amorphous granules compared with the crystalline granules, the overall concentration of these other proteins was relatively low compared with that of PhaR. Even though the lysate without PhaR was used (lane 5), some proteins bound weakly to only the crystalline granules (lane 6). Cytoplasmic proteins seemed to bind more tightly to amorphous granules than to crystalline granules (lane 6 and 7). These observations suggest that PhaR would bind to P(3HB) granules without discriminating between the two different biophysical states of P(3HB) (crystalline or amorphous). The ability of PhaR to bind P(3HB) was further confirmed by using the purified PhaR (data not shown). Results from Fig. 1 to 3 clearly demonstrate that PhaR binds not only to DNA but also to P(3HB).

FIG. 3.

FIG. 3.

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Binding of PhaR and cytoplasmic proteins of E. coli to crystalline or amorphous P(3HB) granules. Proteins were separated in an SDS-15% polyacrylamide gel and stained with CBB R-250. The supernatants of lysates used were prepared from E. coli XL1-Blue(pTV119N::phaR) isolates that expressed phaR (lane 2) or XL1-Blue(pTV119N), or that did not have phaR (lane 5). Crystalline or amorphous P(3HB) granules were incubated for 30 min with the supernatants of the crude lysates. After incubation, the P(3HB) granules were collected and washed twice with TE buffer and resuspended with denaturing buffer. Results are for supernatant (S), crystalline P(3HB) granule-associated proteins (C) from the lysate of XL1-Blue(pTV119N::phaR) (lane 3) or XL1-Blue(pTV119N) (lane 6), and amorphous P(3HB) granule-associated proteins (A) from the lysate of XL1-Blue(pTV119N::phaR) (lane 4) or from that of XL1-Blue(pTV119N) (lane 7).

P(3HB) causes the dissociation of PhaR from PhaR-DNA complexes.

To determine whether P(3HB) affects the DNA binding ability of PhaR, gel mobility shift assays were performed in the absence and presence of P(3HB) or other potential effector molecules (Fig. 4). DNA fragments were prepared by digestion of plasmid pTVCP4 as previously described (24). The 225-bp and 639-bp fragments (DNA225 and DNA639) contain the target sequences for PhaR binding (Fig. 1). The 3HB oligomers were also used as a reference for the high-molecular-mass P(3HB) to examine whether PhaR has chain-size dependency in recognizing P(3HB).

Addition of PhaR to the assay mixture changed the electrophoretic mobility of PhaR-DNA225 and PhaR-DNA639 templates. The PhaR-DNA template complexes were seen as shifted bands in the upper region of the gel (Fig. 4, lane 2). An increase in the amount of crystalline P(3HB) granules in the mixture disrupted the complexes, resulting in free DNA (lanes 3 to 6). After PhaR-DNA complexes were formed, subsequent addition of crystalline P(3HB) granules to the mixture also caused dissociation of PhaR from the PhaR-DNA complexes (lane 7). The 3HB oligomers (approximately 30-mer) and artificial amorphous P(3HB) granules also inhibited the formation of PhaR-DNA complexes (lanes 8 to 9). Interestingly, neither native P(3HB) granules purified from E. coli XL1-Blue(pBBRKmAB pTVCP4) (harboring phaAB and phaCPR) nor native P(3HV) granules from P. denitrificans, which are predominantly covered with PhaP (Fig. 2A, lanes 7 and 10), inhibited the formation of PhaR-DNA complexes (lanes 10 to 11).

These results suggest that PhaR is able to sense both the onset of PHA synthesis and the growing size of the granules through the direct binding of PhaR to PHA, and that free PhaR does not sufficiently sense the mature PHA granules which are already covered with PhaP. P(3HB) granules purified from E. coli XL1-Blue(pBBRKmAB pTVC) (harboring phaAB and phaC) also caused no effect on the formation of PhaR-DNA complexes (data not shown). The granules were covered nonspecifically with numerous endogenous cellular proteins (Fig. 2A, lane 4). These results indicate that a naked PHA polymer chain and/or surface is needed to dissociate the PhaR-DNA complexes.

To test whether other metabolites could be effector molecules for PhaR, the following substances possibly related to PHA synthesis were applied to the gel mobility shift assay: CoA-SH, acetyl-CoA, acetoacetyl-CoA, (R)-3-hydroxybutyryl-CoA, (S)-3-hydroxybutyryl-CoA, (rac)-3HB, (R)-3HB dimer, NAD+, NADH, NADP+, NADPH, acetyl phosphate (29), polyphosphate (6, 13, 37), citric acid, and phosphoenolpyruvate. Partially purified PhaP was also used in this assay. However, none of these substances had an effect on the binding of PhaR to DNA (results not shown), indicating that these metabolites and PhaP do not participate in the PhaR-mediated regulation as effector molecule for PhaR. Therefore, it was concluded that the effector of PhaR is P(3HB).

PhaR is a repressor protein that derepresses phaP expression in response to P(3HB).

The effect of P(3HB) on phaP expression in the presence of PhaR was investigated by cell-free protein synthesis using E. coli S30 extract. As shown in Fig. 5, PhaP production (≈18 kDa) was immunologically observed on the blotting membrane when pPDPK1.7, which includes IRCP and phaP, was used as the template DNA (lane 3). However, no PhaP production was detected when pTV119N lacking phaP was used (lane 2). Addition of PhaR (0.69 μg) to the reaction system caused the repression of phaP expression (lane 5); however, addition of P(3HB) (100 μg) caused the derepression of phaP expression in the presence of PhaR (lane 6). These results suggest that PhaR-mediated regulation of phaP expression is associated with the presence of P(3HB). Considering the fact that PhaR binds both P(3HB) and P(3HV) granules in vivo, it was concluded that SCL PHA is an effector molecule that triggers derepression of phaP expression repressed by PhaR.

FIG. 5.

FIG. 5.

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Effect of crystalline P(3HB) granules on expression of the phaP gene in the presence of PhaR, determined using in vitro cell-free protein synthesis. Plasmid pTV119N (not carrying phaP as a control, lane 2) and pPDPK1.7 (carrying phaP, lanes 3 to 8) were used as DNA templates. Proteins were separated in an SDS-12.5% polyacrylamide gel and subjected to Western blot analysis employing antibody raised against PhaP. Lane 1 shows molecular mass standards (in kilodaltons). Additives were as follows: lane 4, PhaR (0.25 μg); lane 5, PhaR (0.70 μg); lane 6, PhaR (0.70 μg) and crystalline P(3HB) granules (100 μg); lane 7, PhaR (0.70 μg) and crystalline P(3HB) granules (300 μg); lane 8, PhaR (0.70 μg) and crystalline P(3HB) granules (600 μg). The arrow indicates PhaP.

Furthermore, inhibition of transcription-translation reactions was caused by increasing the amount of P(3HB) (Fig. 5, lanes 7 and 8). This suggests that the surface of P(3HB) without PhaP would serve as a place for nonspecific interaction with proteins indispensable for the protein synthesis system. If so, binding of PhaP onto the P(3HB) granule surface would be physiologically important for maintenance of P(3HB) within a bacterial cell. Previously, it was demonstrated that an increased amount of P(3HB) was observed in the presence of PhaP alone or PhaP/R when PhaA, PhaB, and PhaC were present (26). Additionally, in E. coli that has been engineered to produce P(3HB) but lacks a phaP gene, production of P(3HB) triggers expression of heat shock proteins (11).

DISCUSSION

According to the conventional classification of PHA granule-associated proteins proposed by Steinbüchel et al. (42), the following three distinct proteins can be defined functionally: (i) class I comprises the PHA synthases, which catalyze the polymerization of the monomers of hydroxyacyl-CoA; (ii) class II comprises the PHA depolymerases, which are responsible for the intracellular degradation and mobilization of PHA; (iii) class III comprises the phasins (designated as PhaP), which probably form a protein layer at the surface of the PHA granule with phospholipids, lipids, and other proteins; and (iv) class IV comprises all other proteins.

PhaR, which we have characterized here, is a new type of protein belonging to class IV. In addition to our previous studies (24, 26), we provide here insights into important aspects of the regulatory mechanism for PHA synthesis through PhaR-mediated PhaP expression. First, PhaP expression in vivo is repressed in the presence of phaR without P(3HB) production (26). Second, PhaR is a DNA-binding protein that represses PhaP expression in vitro (24). Third, we demonstrated in this study that PhaR is a PHA-responsive protein that can bind PHA both in vivo and in vitro. These data indicate that PhaR has bifunctional characteristics, namely, binding abilities toward both PHA and DNA. Furthermore, we could argue that P(3HB) itself is an effector molecule for PhaR-mediated PhaP expression in a cell-free protein synthesis system (Fig. 5). PhaR, to our knowledge, is the first regulator protein that interacts directly with the PHA polymer. The recognition requirement for this interaction was relatively nonspecific, because PhaR bound to all forms of P(3HB) used—crystalline, amorphous, and 3HB oligomers.

Based on these findings, a plausible model of the involvement of PhaR in PHA biosynthesis can be drawn (Fig. 6). The phaC locus of P. denitrificans consists of four genes, phaZ, phaC, phaP, and phaR, that encode an intracellular PHA depolymerase, a PHA synthase, a phasin, and a PHA-responsive repressor PhaR, respectively (Fig. 6A). Although the regulatory mechanisms for phaC and phaZ expression are not clear, phaC seems to be expressed constitutively (data not shown). When an insufficient amount of substrates are provided for PHA biosynthesis (Fig. 6B), the cells are not able to accumulate PHA even if PhaC is produced sufficiently. At the same time, the expression of phaP and phaR remains at basal level, and PhaR binds to the upstream elements in IRCP and IRPR. Since PhaP is a predominant PHA granule-associated protein, PhaP production is not needed for the cells without PHA accumulation. Under this condition, PhaP production is repressed by PhaR through the direct binding of PhaR to the upstream element for phaP. Excessive PhaR production is also repressed by PhaR autoregulation. If sufficient substrates for PHA synthase are supplied (Fig. 6C), the cells begin to accumulate PHA. PhaR recognizes and binds directly to the PHA polymer chains being synthesized, and then the expression of PhaP is initiated at the onset of dissociation of PhaR from the upstream element for phaP. During the elongation of PHA polymer chains, the PHA granules enlarge in size, and then the surfaces of PHA granules become covered with PhaP and other specific proteins before the other nonspecific proteins bind to the PHA granules. Based on the fact that the phaP promoter is very strong (24), a number of PhaP molecules are expected to be produced under these conditions. When PHA synthesis is stopped or when PHA degradation occurs predominantly (Fig. 6D), free PhaR molecules bind to the upstream elements and repress both phaP expression and phaR expression. Accordingly, we assume that PhaR functions as a sensor for PHA synthesis in the cells. In the PhaR-mediated regulatory mechanism, it remains unclear whether there is any other factor(s) that forces PhaR to dissociate from the PhaR-PHA complex.

FIG. 6.

FIG. 6.

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Hypothetical model of PhaR-mediated phaP expression in P. denitrificans. Z, C, P, and R indicate the names of pha genes in P. denitrificans. (A) Gene organization of a pha locus and produced proteins. (B) Repression of the expression of phaP and phaR under non-PHA accumulation conditions. PhaR that is produced at a basal level binds to both IRCP and IRPR, and therefore phaP expression is repressed. (C) Derepression of expression of phaP and phaR under PHA accumulation conditions. Once PHA accumulation is initiated, PhaR is released from DNA by the binding of PhaR to both oligomeric and polymeric forms of 3HB, and then the phaP expression is initiated at the onset of dissociation of PhaR from the upstream element for phaP. (D) Repression of the expression of phaP and phaR under offset-PHA accumulation conditions or PHA degradation conditions. Since PHA granules accumulated are already covered with proteins (the predominant protein is PhaP), PhaR that is newly produced at a basal level or that is released from PHA granules by degradation of PHA binds to both IRCP and IRPR, and therefore phaP expression is repressed by the binding of PhaR to the upstream element for phaP.

Our current model of PHA production regulation has been established through in vitro experiments (this study and reference 24) and in vivo experiments using an E. coli heterologous expression system (26). York et al. (54) recently reported that PhaP expression of R. eutropha is negatively regulated by PhaR, based on complementation experiments using both a phaR deletion mutant of R. eutropha and a heterologous E. coli system introducing phaP, phaR, and phaCAB of R. eutropha. They demonstrated that PhaR directly links PhaP accumulation to the presence of P(3HB) in vivo, consistent with our model for a PhaR-mediated regulatory system.

In order to examine whether PhaR homologs are widely distributed among SCL PHA-producing bacteria, we searched for the nucleotide sequences potentially coding for PhaR homologs among GenBank, EMBL, and DDBJ and several unfinished microbial genome databases available on the internet (see Material and Methods) by using the BLAST program (1). To estimate whether a bacterium potentially carrying a PhaR homolog is a PHA producer, the genes coding for proteins similar to PHA synthases were searched simultaneously. At present, 31 structural genes for PhaR homologs have been found. Molecular phylogenetic analysis of PhaR homologs has revealed that the PhaR family has two main clusters, the first consisting of bacteria belonging to the α-subdivision of Proteobacteria, including P. denitrificans, and another cluster consisting of bacteria belonging to the β- and γ-subdivisions of Proteobacteria, including R. eutropha (41) and Allochromatium vinosum (21), respectively (data not shown). Most of the bacteria that possess a PhaR homolog also have genes for PHA biosynthesis, and genes for PhaR homologs are often in close proximity to phaC, phaA, and/or phaB. The PhaCs of these bacteria belong to the type I or type III PHA synthase families. These two types of PHA synthases can produce SCL PHA (43), which suggests that the PhaR-mediated regulatory mechanism recognizing PHA accumulation may be widespread in SCL PHA-producing bacteria.

Prieto et al. (35) reported that two major MCL PHA granule-associated proteins, PhaF and PhaI, were found in Pseudomonas oleovorans, which possesses type II PHA synthases (43). PhaF (35 kDa) is a nonenzymatic protein considered to have bifunctional characteristics. PhaF, like PhaR, is able to bind to both PHA granules and DNA. PhaF has been shown to be involved in transcriptional regulation of phaC genes and phaI (35), although direct evidence for binding of PhaF with DNA in vitro has not been obtained. PhaI (18 kDa) has also been confirmed to have PHA granule-binding ability in vivo. However, the deduced amino acid sequence of PhaR showed no similarity to that of either PhaF or PhaI. In addition, PhaD, which is not associated with PHA granules, was identified as a regulator protein positively affecting MCL PHA synthesis and PhaI production (18). Identification of a characteristic gene cluster, phaC1ZC2DFI, in two other strains of Pseudomonas (P. putida [48] and P. aeruginosa [44]) suggests that the PHA synthesis regulation governed by these genes is commonly distributed in MCL PHA-producing pseudomonads. The regulatory system of MCL PHA synthesis thus seems to be more complicated than that of SCL-PHA synthesis.

Povolo and Casella (34) showed that a mutant of Sinorhizobium meliloti that was defective in a phaR homolog aniA produced a larger amount of extracellular polymers, such as polysaccharides, than the wild-type strain under anoxic condition. The aniA gene is located upstream of phaAB in the opposite orientation (34, 46). They demonstrated that aniA is inducible under anaerobic conditions but that the defect of aniA causes little effect on P(3HB) synthesis. Therefore, a direct role of aniA in the P(3HB) biosynthetic pathway is not clear (34). York et al. (54) also recently reported the possibility that the PhaR from R. eutropha is involved in the regulation of PHA synthesis through PhaP-independent pathways. The findings obtained by us, Povolo et al., and York et al. raise the possibility that the PhaR-mediated regulatory system recognizing PHA accumulated in cells is involved not only in the expression of phaP but also in the expression of genes related to other metabolic pathways, namely, that PhaR may act as a PHA-responsive global repressor.

Acknowledgments

Preliminary sequence data were obtained from the DOE Joint Genome Institute at http://www.jgi.doe.gov/JGI_microbial/html/index.html, from the respective sequencing groups at The Sanger Institute (http://www.sanger.ac.uk/), or from The Institute for Genomic Research website at http://www.tigr.org. We thank T. Iwata for the preparation of 3HB oligomers, S. Ueda for the preparation of 3HB dimer, Y. Dohmae for MALDI-TOF MS analysis, M. Chijimatsu for the amino acid sequence analysis, and H. Nakano for useful discussions.

This work was supported by a grant for Ecomolecular Science Research, by a Grant-in-aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan, and by the Special Postdoctoral Researchers Program of the RIKEN Institute.

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