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Journal of Bacteriology, February 2007, p. 1118-1127, Vol. 189, No. 3
0021-9193/07/$08.00+0     doi:10.1128/JB.01550-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Autoregulator Protein PhaR for Biosynthesis of Polyhydroxybutyrate [P(3HB)] Possibly Has Two Separate Domains That Bind to the Target DNA and P(3HB): Functional Mapping of Amino Acid Residues Responsible for DNA Binding{triangledown}

Miwa Yamada,1 Koichi Yamashita,2 Akiko Wakuda,1 Kazuyoshi Ichimura,3 Akira Maehara,4 Michihisa Maeda,3 and Seiichi Taguchi1*

Division of Biotechnology and Macromolecular Chemistry, Graduate School of Engineering, Hokkaido University, N13W8, Kita-ku, Sapporo, Hokkaido 060-8628,1 Chemical Analysis Team and Polymer Chemistry Laboratory, RIKEN Institute, 2-1 Hirosawa, Wako-shi, Saitama 351-0198,2 School of Agriculture, Meiji University, Kawasaki, Kanagawa 214-8571,3 Niigata Research Laboratory, Mitsubishi Gas Chemical Company, Inc., Niigata 950-3112, Japan4

Received 5 October 2006/ Accepted 14 November 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
PhaR from Paracoccus denitrificans functions as a repressor or autoregulator of the expression of genes encoding phasin protein (PhaP) and PhaR itself, both of which are components of polyhydroxyalkanoate (PHA) granules (A. Maehara, S. Taguchi, T. Nishiyama, T. Yamane, and Y. Doi, J. Bacteriol. 184:3992-4002, 2002). PhaR is a unique regulatory protein in that it also has the ability to bind tightly to an effector molecule, PHA polyester. In this study, by using a quartz crystal microbalance, we obtained direct evidence that PhaR binds to the target DNA and poly[(R)-3-hydroxybutyrate] [P(3HB)], one of the PHAs, at the same time. To identify the PhaR amino acid residues responsible for DNA binding, deletion and PCR-mediated random point mutation experiments were carried out with the gene encoding the PhaR protein. PhaR point mutants with decreased DNA-binding abilities were efficiently screened by an in vivo monitoring assay system coupled with gene expression of green fluorescent protein in Escherichia coli. DNA-binding abilities of the wild-type and mutants of recombinant PhaR expressed in E. coli were evaluated using a gel shift assay and a surface plasmon resonance analysis. These experiments revealed that basic amino acids and a tyrosine in the N-terminal region, which is highly conserved among PhaR homologs, are responsible for DNA binding. However, most of the mutants with decreased DNA-binding abilities were unaffected in their ability to bind P(3HB), strongly suggesting that PhaR has two separate domains capable of binding to the target DNA and P(3HB).


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Polyhydroxyalkanoates (PHAs) constitute a family of natural polyesters that various microorganisms synthesize as intracellular granules to store carbon and reducing equivalents (2, 4, 20, 22, 39). When nutrient supplies are starved or imbalanced, it is advantageous for microorganisms to store excess nutrients in the form of PHAs for survival at a relatively low maintenance cost. Many microorganisms have evolved the ability to degrade and utilize these storage materials within their cells. PHAs extracted from microbial cells have attractive practical applications, since their properties can be similar to those of polypropylene. Because of their thermoprocessibility, biodegradability, and biocompatibility, they have received much attention as prime candidates for commodity and specialty production of commercial bioplastics (19).

PHAs can be classified into those with short-chain-length (SCL) repeating units (3 to 5 carbons per monomer) and those with medium-chain-length repeating units (6 to 14 carbons per monomer) (37). Paracoccus denitrificans, a facultative methylotrophic bacterium, is able to synthesize SCL PHAs from several alcohols (45, 46). A typical SCL PHA is a poly[(R)-3-hydroxybutyrate] [P(3HB)]. Two gene clusters associated with the synthesis, regulation, and degradation of P(3HB) in P. denitrificans have been cloned and characterized (8, 24, 25, 43, 44). One cluster is phaA-phaB, which encodes ß-ketothiolase and NADPH-dependent acetoacetyl coenzyme A reductase (44), respectively, and another is phaZ-phaC-phaP-phaR, which encodes P(3HB) depolymerase (8), P(3HB) synthase (43), P(3HB) granule-associated phasin (25), and P(3HB) synthesis regulatory protein (24), respectively. P(3HB) synthase (PhaC) and phasin (PhaP) are proteins that play important roles in the production and granule formation of P(3HB) (2, 4, 7, 9, 14, 25, 26, 32). Specifically, PhaP forms a boundary layer on the P(3HB) surface to sequester hydrophobic P(3HB) from the cytoplasmic substances (28, 29, 38). Thus, PhaP is able to prevent individual granules from coalescing and promotes P(3HB) synthesis by regulating the ratio of surface area to volume of P(3HB) granules. PhaP may also have a protective function in reducing the passive attachment of cytoplasmic proteins to the P(3HB) surface. In this sense, it is generally thought that the synthesis of PhaP protein is highly regulated (30-32, 48, 49). Our previous studies have shown that expression of phaP is negatively controlled by the autoregulated repressor PhaR in P. denitrificans (23-25). phaR, which is located immediately downstream of phaP, encodes a repressor that regulates the gene expression of phaP and phaR itself. PhaR can also sense the presence of P(3HB) and interact with nascent P(3HB) granules, resulting in the derepression of phaP expression.

Previously, the DNA-binding ability of PhaR and its regulatory function in phaP expression were analyzed in vitro using purified recombinant PhaR (24, 25). PhaR was also able to bind to P(3HB) granules and 3HB oligomers, accompanied by dissociation of the PhaR-DNA complex. In other words, P(3HB) acts as an inducer or effector of phaP expression in a PhaR-mediated regulatory system: once P(3HB) accumulation begins, PhaR dissociates from the target DNA by binding to P(3HB), allowing the production of PhaP and PhaR. Under P(3HB) accumulation conditions or P(3HB) degradation conditions, phaP and phaR expressions are again repressed by binding of PhaR to the upstream elements for phaP and phaR. This prominent PhaR-mediated regulatory mechanism in response to P(3HB) accumulation is consequently attributed to the unique property of PhaR that enables it to bind to both the target DNA and P(3HB).

We recently developed a quartz crystal microbalance (QCM) technique to study the binding of PhaR to P(3HB). The QCM measurement revealed that binding of PhaR onto the bare P(3HB) surfaces proceeds in a diffusion-controlled manner, mainly by nonspecific hydrophobic interactions (47). Because of this finding, we designed an experiment in which the PhaR-recognizable target DNA was added to the PhaR-P(3HB) complex in the QCM system. It is of interest to know if these three molecules are capable of forming a complex in the QCM simple system without PhaP. This study presents the first direct evidence that PhaR is capable of binding to both the target DNA and P(3HB) at the same time. In addition, functional mapping of amino acid residues responsible for the DNA-binding ability of PhaR was carried out using a combination of random point mutagenesis and a green fluorescent protein (GFP) reporter-based in vivo monitoring system developed here.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Bacterial strains, plasmids, and growth conditions. The bacterial strains and plasmids used in this study are listed in Table 1. Escherichia coli strains DH5{alpha} and BL21(DE3) were grown in Luria-Bertani (LB) medium (34) at 37°C. When needed, ampicillin (100 µg/ml) and/or kanamycin (50 µg/ml) was added to the medium.


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  		TABLE 1. Bacterial strains and plasmids used in this study

 
QCM analyses. A commercially available QCM setup of QCA-922 (SEIKO EG&G) was used by equipping with a homemade cell with an inner volume of ca. 0.2 ml. The Au-deposited oscillator of a polished 9-MHz AT-cut quartz crystal was supplied by SEIKO EG&G as a commercial product. The area size of the electrodes was 0.196 cm2 x 2. The mean roughness of the electrode surface was estimated to be 1.2 nm (30 x 30 µm) by atomic force microscopy (AFM). A frequency change of 1.0 Hz corresponds to a mass change of 1.0 ng on the electrode (0.196 cm2) according to the Sauerbrey equation (12, 35). A thin film of P(3HB) was prepared on the QCM oscillator by casting chloroform solution of the polymer on a spin coater (ME-300; MOC Co. Ltd.) as described previously (47). The mean surface roughness and thickness of the P(3HB) film were estimated by AFM to be 2.0 nm and ca. 100 nm in a 1- by 1-µm area, respectively. All the QCM data were obtained in batch reactions by stopping introduction of the sample solution into the cell just after replacement. The recombinant PhaR was purified as previously described (47). The 56-bp DNA fragment containing the promoter region of phaP was prepared by following PCR. As described in our previous paper, the target DNA used here was specifically recognized and bound by PhaR (23). The molecular weight of the 56-bp DNA is 34,396. The forward primer 5'-AGCGGGCCTAAAATTTTTCTG-3' and the reverse primer 5'-TGTCCAACATATACTTTCTGCA-3' were used. pTVCP (25) was used as a template for PCR. PCR was carried out using a program of 30 cycles of 94°C for 2 min, 56°C for 30 s, and 72°C for 30 s with an iCycler thermal cycler (Bio-Rad Laboratories, Hercules, CA). The PCR product was purified by use of a PCR clean-up system kit (Promega, WI). Calf thymus DNAs with similar sizes (70 to 100 bp) were prepared by sonication and supplied to QCM analyses as a negative control.

During analysis, temperature was maintained at 25.0 ± 0.1°C by circulating water. Sample solutions (containing PhaR and the target DNA or control DNA) and buffer substitution solution were preincubated for 10 min in the sample loop to equilibrate at the measuring temperature before injection. After stabilization of the QCM in 10 mM HEPES buffer solution (pH 7.4) containing 150 mM NaCl, the reaction was initiated by replacing the buffer solution with the PhaR solution. The total volume used for replacement was 1.25 ml. Resonant frequency and resonant resistance were monitored simultaneously to evaluate the influence of frictional changes. Since the resonant resistance did not change during the measurements (data not shown), the effects of the frictional changes of the QCM on addition of PhaR were negligibly small.

Construction of plasmid vectors. Standard recombinant DNA manipulation was used for isolation of plasmid DNA (34). All of the restriction endonucleases and modification enzymes for genetic engineering were purchased from TaKaRa Shuzo Co., Ltd. (Kyoto, Japan) and used under conditions recommended by the supplier. All other chemicals were of analytical grade for biochemical use and were used without further purification. The plasmid vectors constructed are listed in Table 1. To analyze the deletion effects of the N-terminal region of PhaR, the three plasmid vectors containing deleted phaR (pTV119N::N{Delta}2-5, pTV119N::N{Delta}2-16, and pTV119N::N{Delta}2-30) were prepared by following PCR. The forward primers of 5'-AACACGACCCCGCTTC-3' for N{Delta}2-5, 5'-AGCCGCCGCCTCTACAAT-3' for N{Delta}2-16, and 5'-CTTGAGGATATCGCGGGCTT-3' for N{Delta}2-30 and the common reverse primer of 5'-CATGGTCTGTTTCCTGTGTG-3' for N{Delta}2-5, N{Delta}2-16, and N{Delta}2-30 were used. pTV119N::phaR containing wild-type phaR was used as a template for PCR. PCR was carried out using a program of 25 cycles of 94°C for 2 min, 55°C for 30 s, and 72°C for 3 min. The PCR products were purified, phosphorylated by polynucleotide kinase, and self-ligated (34).

On the other hand, the expression plasmids for the deleted derivatives of phaR, C{Delta}73-195, C{Delta}121-195, C{Delta}164-195, and C{Delta}173-195, were constructed by inserting a stop codon into the corresponding deletion terminal site for each deletion mutant. For this purpose, PCR was conducted using the common forward primer and the reverse primers containing XbaI sites and stop codons. The forward primer was 5'-CCAGGCTTTACACTTTATGC-3', and the reverse primers were 5'-AATCTAGAttaGTTCTCGCCCCGGCCCTCATGCT-3' for C{Delta}73-195, 5'-AATCTAGAttaCTCGTCCGTCTCGGTCTCCT-3' for C{Delta}121-195, 5'-CCTCTAGAttaGTTGGGAAAGGTCGACAGATTC-3' for C{Delta}164-195, and 5'-GATCTAGAttaAGGAGCCCTTCTGGCCGCCACGGC-3' for C{Delta}173-195 (the underlined sequences and the lowercase sequences [tta] indicate XbaI sites and stop codons, respectively). PCR was carried out using pTV119N::phaR as the template with a program of 25 cycles of 95°C for 1 min, 58°C for 1 min, and 72°C for 2 min. The PCR products were purified, digested with NcoI and XbaI, and subcloned into the same reaction sites of pTV119N. The resulting plasmids were termed pTV119N::C{Delta}73-195, pTV119N::C{Delta}121-195, pTV119N::C{Delta}164-195, and pTV119N::C{Delta}173-195, respectively.

A plasmid vector termed pTV-R, which allows us to estimate the mutational effects of the PhaR on DNA-binding ability based on the intensity of the specific cell luminescence caused by gfp expression, is shown below (see Fig. 3). Basically, this vector was made by locating the expression elements in the following order from upstream: phaR, the T7 RNA polymerase promoter, the PhaR-binding DNA region (containing the phaP promoter), and gfp. First, the plasmid vector pGEFP (Promega, WI), containing the T7 RNA polymerase promoter and gfp, was digested with SphI and NcoI, blunted at both sites with T4 DNA polymerase, and ligated with the 0.2 kb of the SmaI-SalI (blunted) fragment of pPDPK1.7 (25) containing the phaP promoter. Next, the resultant vector was digested with PvuII and SphI and ligated with the 4.1-kb HindIII (blunted)-SphI fragment containing the rrnB terminator derived from E. coli. Finally, the resulting vector was digested with AseI and BamHI and blunted at both sites. Then, the 1.8 kb of the fragment containing the rrnB terminator, the phaP promoter, the T7 RNA polymerase promoter, and gfp was purified and ligated into pTV119N::phaR at the blunted EcoRI site and EcoT22I site to make pTV-R.


Figure 3
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  		FIG. 3. Schematic flow diagram of the in vivo assay system for analysis of mutational effects of PhaR on the ability to bind to target DNA. +, fluorescent exposure of GFP by recombinant E. coli.

 
Random mutagenesis by error-prone PCR. To introduce random mutations into the entire region of the phaR by error-prone PCR, forward and reverse primers were designed based on the nucleotide sequence outside of the structural gene (approximately 0.6 kb). The forward primer was 5'-CACACAGGAAACAGACCATGGCT-3' (the underlined sequence shows an NcoI site) and the reverse primer was 5'-GTCGACGGACCGCGGGGC-3' (the underlined sequence shows a SacII site). pTV119N::phaR was used as a template for PCR mutagenesis with the two primers under error-prone conditions (40): the addition of an excess 5 mM MgCl2 and 10% dimethyl sulfoxide to the 50 µl of reaction solution containing 2.5 nM pTV119N::phaR, 0.025 U of rTaq polymerase, 0.1 µM each of the two primers, 0.2 mM each deoxynucleotide triphosphate, 10 mM Tris-HCl (rather high pH value of 8.8 at 25°C), 50 mM KCl, and 0.1 mg/ml bovine serum albumin. PCR was carried out using a program of 25 cycles of 94°C for 1 min, 50°C for 1 min, and 72°C for 2 min.

Screened mutants leading to reduced DNA-binding ability. PCR mutagenesis of the entire region in the phaR was carried out by error-prone PCR. After amplification, a mixture of the NcoI-SacII fragments including the randomly mutagenized phaR religated into the same restriction sites of pTV-R to generate a mutant library. The resultant plasmids were introduced into E. coli DH5{alpha} cells, and transformants were then spread on the LB plates containing ampicillin (100 µg/ml). The change in DNA-binding ability resulting from the introduction of mutations into the phaR was judged on the basis of the intensity of specific luminescence of the cells caused by gfp expression with the FluorImager 595 (Amersham Biosciences, NJ). The emission wavelength of GFP was 510 nm.

Gel shift assay. To detect decreases in DNA-binding ability for PhaR, a gel shift assay was carried out by the method described previously (23) with some modifications. The target DNA fragments used for the gel shift assay were prepared to amplify the promoter region of phaP by PCR using plasmid pTVCP as a template and the following primers: PDPPR1 (5'-GGTGTTGGCCATTTGCTCTC-3') and PDPPF1 (5'-TTGCATGCCCGCGACCC-3'). PCR was carried out using a program of 30 cycles of 94°C for 2 min, 55°C for 30 s, and 72°C for 2 min. The PCR product was purified by use of a PCR clean-up system kit (Promega, WI). The DNA fragments (400 ng) were mixed with the wild type and each deletion or single-point mutant of PhaR in binding buffer (10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 80 mM NaCl, 10 mM ß-mercaptoethanol, and 4% glycerol) in a total volume of 20 µl and incubated for 30 min at 25°C. The DNA-PhaR complex was separated from the unbound DNA fragment on a 5% native polyacrylamide gel, using 0.5x Tris-borate-EDTA (34) as the electrophoresis buffer. DNA fragments were stained with ethidium bromide and detected by UV exposure.

DNA sequencing. DNA sequencing to confirm the new plasmid constructs and to analyze mutation points was carried out by the dideoxynucleotide chain-termination method with a Prism 310 genetic analyzer (Applied Biosystems, CA) and a BigDye Terminator cycle sequencing ready reaction kit (Applied Biosystems, CA). The nucleotide sequence was analyzed with GENETYX genetic information-processing software (Software Development Co., Tokyo, Japan).

Electrophoresis of proteins. Sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) was preformed as described by Laemmli (18). Proteins were separated by SDS-PAGE and stained with Coomassie brilliant blue (CBB).

Binding of PhaR to P(3HB) granules. In order to detect the P(3HB)-binding ability of PhaR, the artificial P(3HB) granules (Biopol) and water (1.5% [wt/vol]) were incubated with each of the crude lysates of E. coli that produce wild-type and each mutant of PhaR for 90 min on ice. After this incubation, the granules were collected by centrifugation, washed twice with 200 µl of 50 mM Tris-HCl (pH 7.0), and resuspended. All fractions throughout the steps consisting of mixing of PhaR with P(3HB) and washing were subjected to SDS-PAGE.

SPR detection of interaction between PhaR and its target DNA. The surface plasmon resonance (SPR) experiments were performed with a Biacore J biosensor system (Biacore AB, Uppsala, Sweden) at a sensor temperature of 25°C. The 5'-biotinylated DNA fragment of 56 bp containing the promoter region of phaP was prepared by PCR from the pTVCP (25) by use of the following primers: PproF (5'Bio-AGCGGGCCTAAAATTTTTCTG-3') and PDPPF1 (5'-TGTCCAACATATACTTTCTGCA-3'). PCR was carried out using a program of 30 cycles of 94°C for 3 min, 56°C for 30 s, and 72°C for 30 s. The PCR product was purified and coupled to the sensor chip (SA sensor chip) under conditions recommended by the supplier. HBS-EP buffer (10 mM HEPES, pH 7.4, 150 mM NaCl, 3 mM EDTA, and 0.005% surfactant P-20) was used for the mobile-phase buffer. Recombinant PhaR was purified as previously described (47). The purified wild-type PhaR and mutants of PhaR (S17R, R18C, and R18H) were injected into the flow cells at a flow rate of 35 µl/min. For observing the association of PhaR with the target DNA, the time of sample injection was set to 2 min. The dissociation of PhaR from the target DNA was monitored after sample injection under the general flow buffer conditions. Dissociation measurement was done for 5 to 10 min before injection and at 1 min after the addition of 0.5 M NaCl and 2 M MgCl2. The subsequent removal of MgCl2 from the system was ensured by washing for more than 1 min with 200 mM NaCl and 5 mM EDTA followed by washing with flow buffer prior to running a new cycle. The samples were injected at various concentrations, i.e., 0.45 µM, 0.67 µM, 0.9 µM, and 1.12 µM. The dissociation constants for wild-type PhaR and mutants were calculated from the sensorgrams with BIAevaluation software, version 3.2 (Biacore AB, Uppsala, Sweden).

CD spectroscopy. Circular dichroism (CD) spectra were measured in a 1-mm path length with a J-720 spectropolarimeter (Jasco; MD). Data points were collected every 0.5 nm with a 3-second averaging time. Far-UV CD spectra were recorded from 200 to 250 nm. Protein concentration was determined using a Bio-Rad protein assay kit with bovine serum albumin as a standard. The measurements were performed in 600 µl of 50 mM Tris-HCl (pH 7.5).


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
PhaR possibly has two separate domains that bind to the target DNA and P(3HB). PhaR is the autoregulator of its gene, phaR, itself as well as the repressor for the expression of phaP. After the surface of the P(3HB) granule is occupied at the saturated state with large amounts of PhaP molecules and a small amount of PhaR molecules, unbound free PhaR molecules rebind to the target DNA for repression of the gene expression. To evaluate the binding mode of PhaR to the target DNA and P(3HB), we set up a QCM-based analytical solution system consisting of these three molecules in the absence of PhaP. The binding and desorption interactions among these three molecules were monitored as the changes in frequency following stepwise operation. Figure 1 shows the profiles of frequency changes for all the following reaction steps: introduction to the P(3HB)-mounted QCM system of buffer containing PhaR (first step), substitution with buffer alone (second step), substitution with buffer containing DNA (third step), and substitution with buffer alone (fourth step), all in a batch mode. Clear and reproducible binding of PhaR onto P(3HB) was observed, as demonstrated in a previous study (47). No desorption of PhaR was detected upon introduction of the substitution buffer when the binding of PhaR to P(3HB) reached an equilibrium state, which took about 6 h. This result strongly suggested that homogeneous formation of a single layer of PhaR protein occurred on the surface of P(3HB), as was previously revealed using AFM (47). We observed the binding of the target DNA to PhaR adsorbed on the P(3HB) surface as an additional negative frequency shift of about –15 Hz when the solution system was substituted with buffer solution containing 10 µg/ml of a PCR-amplified 50 bp-target DNA fragment. Binding specificity between PhaR and the target DNA was confirmed by the absence of a frequency change for a control DNA fragment containing a sequence that was not recognizable by PhaR (Fig. 1). Also, we confirmed that no change in frequency was observed upon addition of the target DNA alone into the QCM system, in which the surface was not treated with PhaR (data not shown). Furthermore, when the QCM solution system was resubstituted with a buffer solution containing no PhaR, the frequency returned to a level (–26 Hz) almost the same as that observed before addition of the target DNA to the solution. This clearly implies a reversible binding reaction between PhaR and the target DNA, in contrast with what was observed for the irreversible interaction of PhaR with P(3HB) (47).


Figure 1
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  		FIG. 1. Frequency changes observed for PhaR binding to P(3HB) film followed by binding of the target DNA and control DNA to the PhaR layers in 10 mM HEPES buffer (pH 7.4) containing 150 mM NaCl. Arrows 1 to 4 indicate injections of solution containing PhaR (4 µg/ml), substitution buffer, solution containing DNA (10 µg/ml), and substitution buffer, respectively, and correspond to the lower illustrations explaining the behaviors of PhaR and the target DNA. Complex formation of three molecules, target DNA, PhaR, and P(3HB), takes place at the stage between 3 and 4. The target DNA fragment of 56 bp, including the specific PhaR-binding site in the phaP promoter (23), was amplified by PCR, and its molecular weight was 34,396. As a negative control, calf thymus DNA of about the same size as the target DNA was prepared by sonication. DF, frequency shift from the frequency before reactions.

 
For examination of the DNA-binding ability of PhaR, the following refined stoichiometry was obtained using QCM analysis. The amount of PhaR molecules bound onto the P(3HB) was estimated to be 30 ng/0.196 cm2 of electrode, which corresponds to 3.2 x 10–13 moles per 0.196 cm2 of electrode. This was calculated by assuming that PhaR protein molecules existed as homotetramers and by estimating the molecular mass of PhaR to be 93 kDa (24). The binding amount (15 ng) of the target DNA (molecular weight of 34,396) was calculated from the frequency change (15 Hz) in a similar manner and was estimated to be 4.3 x 10–13 moles per 0.196 cm2 of electrode. Assuming that PhaR binding onto the P(3HB) would take place by formation of a monolayer, as demonstrated in a similar experiment (47), the molar binding ratio of PhaR to the target DNA was estimated to be approximately 1 to 1. This stoichiometric binding data suggested that the PhaR protein, noncovalently immobilized on P(3HB), binds to the target DNA in the QCM system. It is also likely that the PhaR molecule possibly has two separate domains that bind to the target DNA and P(3HB) (Fig. 1).

Deletion effects of PhaR on the binding abilities to the target DNA and P(3HB). To identify the region of PhaR responsible for binding to the target DNA and P(3HB), a deletion experiment was carried out. Four mutants with deletions in the C-terminal region of PhaR were made based on secondary structure units predicted by DISOPRED software (Fig. 2A). When expressing the four deletion mutant proteins, three types of recombinant cell (those expressing C{Delta}73-195, C{Delta}164-195, and C{Delta}173-195) expressed PhaR as soluble forms, as revealed by SDS-PAGE (Fig. 2B). In the gel shift assay (Fig. 2C), two mutant proteins (C{Delta}164-195 and C{Delta}173-195) retained DNA-binding abilities, and one mutant (C{Delta}73-195) lost it. This suggested that the 32 C-terminal amino acid residues are dispensable for full DNA-binding ability. The same result was obtained for the P(3HB)-binding assay, as shown in Fig. 2D. The C terminally 32-residue-truncated form of PhaR exhibited binding ability for both the target DNA and P(3HB). On the other hand, three N-terminal deletion constructs, N{Delta}2-5, N{Delta}2-16, and N{Delta}2-30, resulted in the expression of insoluble PhaR protein (data not shown), suggesting that the N-terminal region is crucial for correct protein folding.


Figure 2
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  		FIG. 2. Deletion effects of PhaR on the ability to bind to target DNA and P(3HB). (A) The secondary structure units of PhaR predicted by DISOPRED (http://bioinf.cs.ucl.ac.uk/disopred) and schematic illustration of various C-terminal deletion mutants. (B) Expression of the wild type and of deletion mutants of phaR in recombinant E. coli BL21(DE3). Soluble proteins were subjected to electrophoresis in an SDS-18% polyacrylamide gel and stained with CBB. Lanes: 1, molecular mass standard proteins; 2, wild-type PhaR; 3, C{Delta}173-195; 4, C{Delta}164-195; 5, C{Delta}73-195. (C) Gel shift assay with PhaR deletion mutants. The target DNA fragment of about 200 bp includes the PhaR-binding site in the phaP promoter. The DNA fragments (2 µg) were mixed with crude lysates from recombinant E. coli that produces the wild-type PhaR or the deleted mutants in binding buffer (10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 80 mM NaCl, 10 mM ß-mercaptoethanol, 4% glycerol) in a total volume of 20 µl (lanes 2 to 5). Additives were as follows: for lane 1, binding buffer alone (negative control); for lanes 2 to 5, crude lysates from recombinant E. coli BL21(DE3) expressing wild-type PhaR (5 µg) (lane 2), C{Delta}173-195 (5 µg) (lane 3), C{Delta}164-195 (5 µg) (lane 4), and C{Delta}73-195 (5 µg) (lane 5). (D) Binding of the deleted PhaR mutants to artificial P(3HB) granules. Proteins were separated in an SDS-18% polyacrylamide gel and stained with CBB. Lanes: 1, molecular mass standard proteins; lanes 2 to 12, proteins from recombinant E. coli BL21(DE3) expressing wild-type PhaR (lanes 2 to 6), C{Delta}173-195 (lanes 7 and 8), C{Delta}164-195 (lanes 9 and 10), and C{Delta}73-195 (lanes 11 and 12). Proteins were fractionated from crude cell extracts by centrifugation. S, soluble proteins; U0, P(3HB)-unbound fraction after mixing with P(3HB) suspension; U1, P(3HB)-unbound fraction after one wash of the U0 fraction; U2, P(3HB)-unbound fraction after two washes of the U0 fraction; B, P(3HB)-bound fraction after two washes of the U0 fraction. Open arrowheads denote the deleted PhaR proteins.

 
Functional mapping of PhaR amino acid residues responsible for DNA binding by a point mutation experiment. For a more refined functional mapping of amino acid residues related to the DNA binding of PhaR, we established a new plate assay system consisting of error-prone PCR-mediated random point mutagenesis and a GFP reporter-based in vivo monitoring assay to monitor the ability of PhaR that binds to the target DNA upon point mutation. To this end, we constructed the plasmid vector pTV-R to measure the mutational effects of PhaR on DNA-binding ability (see Materials and Methods). This pTV-R vector system was made by locating the expression elements in the following order from the upstream end: phaR, the T7 RNA polymerase promoter, the PhaR binding DNA region (containing the phaP promoter), and gfp, as shown in Fig. 3. In principle, if the wild-type phaR is expressed, its gene product would be able to bind to the target DNA and hinder gfp expression because of the repression of read-through by T7 RNA polymerase. We confirmed the feasibility of this system in a stepwise manner. First, fluorescence specific for gfp expression was detected for the vector pTV119N::gfp-harbored transformant E. coli. When pTV-R was used with the wild-type phaR, no fluorescence signal was detected, indicating the functional expression of phaR. Fluorescence was also detected using vector pTV-dR (negative control), in which the wild-type phaR was replaced by the mutant phaR with the deletion of C73-195. Therefore, it was expected that the change in DNA-binding ability of PhaR upon point mutation could be readily estimated on the plate by using the in vivo monitoring system and pTV-R. A mutant library of the phaR was prepared by forming colonies (hereafter termed "clones") of transformant cells of E. coli DH5{alpha}.

Out of about 12,000 clones, 650 clones that exhibited fluorescence much higher than that of the wild-type clone were selected as candidates for PhaR mutants for which the DNA-binding ability of PhaR would be reduced or lost. To eliminate clones in which decreased DNA-binding ability (increase in fluorescence) was caused by structural breakdown or aggregation of expressed PhaR, Western blot analysis was carried out. As a result, only six mutant proteins were expressed as soluble forms at almost same level as expression in the wild-type clone (data not shown). This suggested that these proteins had mutations associated with their DNA-binding abilities. Nucleotide sequencing revealed that all of the mutants had single mutations that were introduced into the locally restricted N-terminal region (positions 13, 15, 17, 18, and 21). At position 18, two amino acid substitutions (to Cys and His) occurred. Table 2 summarizes these results. No band shifts were observed for any of the mutants in the gel shift assay, while the wild-type phaR product resulted in a band shift (Fig. 4A). However, quantitative DNA-binding ability is not carried out by gel shift assay. Therefore, we attempted to adopt SPR analysis using purified protein samples. As a result, we actually detected DNA-binding abilities with various levels of decreases (down to dissociation constant values of 3.4% to 16.4% of that for the wild type) for three mutants, S17R, R18C, and R18H (Fig. 5 and Table 2). The dissociation constant for the wild-type PhaR to the target DNA was calculated to be 1.8 x 10–7 M. Recently, Kojima et al. reported that the dissociation constant was 1.2 x 10–7 M for the same combination of PhaR and the target DNA (16). Two distinct factors were found to influence PhaR-DNA binding by SPR. One consists of the DNA fragment size and target sequence recognized by the PhaR molecule (56 bp in this study and 18 bp for the fragment used previously [16]). The other is the His-tag modification of the PhaR protein. Previous studies used C-terminally-His-tagged PhaR, while the current study used N-terminally-His-tagged PhaR. Despite these differences, similar dissociation constants of the PhaR-DNA complex were obtained for each system. These studies differed in that the current study showed that P(3HB)-binding affinity was retained for all of the mutants (Fig. 4B).


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  		TABLE 2. DNA- and P(3HB)-binding abilities of PhaR mutants selected using an in vivo assay system

 

Figure 4
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  		FIG. 4. Binding abilities toward the target DNA and P(3HB) of PhaR mutants selected using in vivo assay system. (A) Gel shift assay with PhaR mutants. The target DNA fragment of about 200 bp includes the PhaR-binding site in the phaP promoter. The DNA fragments (2 µg) were mixed with the purified wild-type PhaR or each of the mutants in binding buffer in a total volume of 20 µl (lanes 2 to 8). Recombinant PhaR proteins were purified as previously described (47). Additives were as follows: for lane 1, binding buffer alone (negative control); for lanes 2 to 8, wild-type PhaR (150 ng) (lane 2), K13E (150 ng) (lane 3), Y15C (150 ng) (lane 4), S17R (150 ng) (lane 5), R18C (150 ng) (lane 6), R18H (150 ng) (lane 7), and Y21H (150 ng) (lane 8). (B) Binding of PhaR mutants to artificial P(3HB) granules. Proteins were separated in an SDS-13.5% polyacrylamide gel and stained with CBB. Lanes: 1, molecular mass standard proteins; lanes 2 to 9, proteins from recombinant E. coli BL21(DE3) expressing wild-type PhaR (lanes 2 and 3), K13E (lane 4), Y15C (lane 5), S17R (lane 6), R18C (lane 7), R18H (lane 8), and Y21H (lane 9). Proteins were fractionated from crude cell extracts by centrifugation. S, soluble proteins; B, P(3HB)-bound fraction after two washes.

 

Figure 5
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  		FIG. 5. Sensorgrams showing the interaction of purified PhaR with the target DNA by surface plasmon resonance. The 5'-biotinylated DNA fragment of 56 bp containing the promoter region of phaP was prepared by PCR. The target DNA fragment was immobilized on a sensor chip to be 400 resonance units (RU), and wild-type PhaR or mutants thereof were injected at 10 µg/ml. The measuring temperature was set at 25°C. Sample injection was done at a flow rate of 35 ml/min in 10 mM HEPES (pH 7.4), 150 mM NaCl, 3 mM EDTA, and 0.005% surfactant P-20.

 
Furthermore, to analyze the effects of these mutations on the secondary structure of PhaR, CD measurement was carried out. CD spectra showed that there were no differences between the wild-type PhaR and the mutants except for Y21H (Fig. 6). This suggested that the mutations causing the loss of DNA-binding ability were not associated with changes in the secondary structure but with subtle local structural changes.


Figure 6
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  		FIG. 6. Circular dichroism (CD) spectra of purified samples of PhaR. The spectra were taken at room temperature, and the PhaR (wild type or mutant) concentration was approximately 20 µg/ml. The measurements were performed in 600 µl of 50 mM Tris-HCl (pH 7.5).

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this study, for the first time, we present direct evidence that PhaR binds to the target DNA and P(3HB) at the same time by QCM analysis. This implies that PhaR possibly has two separate domains that bind to these two molecules. In our previous QCM analysis (47), PhaR was found to bind to the surface of P(3HB) in an apparently irreversible manner mainly by nonspecific hydrophobic interactions. This finding illustrates the high affinity and low specificity for binding of PhaR to P(3HB), which contrasts with the high DNA-binding specificity of PhaR. In the cellular environment, a complex of the three molecules, PhaR/P(3HB)/DNA, may arise, but its occurrence may be limited because the surface of the natural P(3HB) granule has a higher concentration of PhaP proteins than of PhaR proteins (23). It is more likely that the PhaR associated with the DNA is derived from free PhaR proteins rather than the P(3HB)-bound PhaR proteins. The localization of free PhaR to the P(3HB) surface (observationally irreversible) or to the target DNA (reversible) is dependent on the capacity of the surface area of P(3HB) granule. The access of PhaR to the granule is limited by the stage of maturation of the granule. QCM analysis allowed us to examine the DNA-binding affinities of the PhaR-P(3HB) complexes in the absence of PhaP, observations that could not be detected by gel shift assay or SPR. The presence of two binding domains of PhaR would be feasible, considering the derepression of the expressions of phaP and phaR through the dissociation of autoregulator PhaR from the complexes of PhaR and the target DNAs.

Generally, the binding of an effector substance to the corresponding protein is presumed to result in a conformational change that inhibits or prevents DNA binding, resulting in transcriptional derepression of the regulon (6). Unlike many other transcriptional regulators, the binding of PhaR to its target DNA sequence or to P(3HB) is not dictated by a simple "concentration equilibrium." In most cases observed for transcriptional regulation responses, an effector ligand which can bind to the transcription factor via reversible molecular interactions determines whether or not the transcription factor binds to its target DNA sequence. PhaR presents a rather unique case because it is initially sequestered by the P(3HB) granule after it is produced. An insignificant conformational change would be expected to occur, considering the temporary complex formation by PhaR, the target DNA, and P(3HB) observed during QCM analysis. It is of interest to quantitatively evaluate the presence or absence of conformational change of the DNA-bound PhaR upon binding of the effector P(3HB), and this is major thrust of our current and continuing studies. Based on the reversible binding of the target DNA to PhaR, the dissociation of the target DNA from the complex of PhaR/P(3HB)/DNA would be dependent on environmental conditions. Stringent conditions (electrophoresis for the gel shift assay and buffer substitution for QCM) favored the dissociation of the PhaR/P(3HB)/DNA complex to yield the PhaR/P(3HB) complex and dissociated target DNA.

The discovery of two separate binding domains within PhaR prompted us to further identify the amino acid residues responsible for DNA binding, and we used a GFP reporter-based in vivo monitoring system coupled with random point mutagenesis. By use of this screening system, six single soluble mutant proteins with decreased DNA-binding abilities were obtained. Interestingly, all effective mutations were introduced into the N-terminally limited region (positions 11 to 23), which is highly conserved among the PhaR homologs identified to date (Fig. 7). Overall structural similarities reported in functional studies conducted in vitro (23-25) and in vivo (30-32, 48, 49) suggest that the PhaR-mediated regulatory mechanism would be widespread among SCL PHA-producing bacteria. In this study, we have provided insight into structural features commonly shared by PhaR homologs, which will be of use for understanding the molecular mechanism of PhaR-mediated PHA biosynthesis.


Figure 7
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  		FIG. 7. Alignment of protein sequences homologous to the PhaR sequence. The amino acid sequence of PhaR was compared with sequences in GenBank using the NCBI BLAST program (1). Nine protein sequences exhibiting high homology to the PhaR sequence were aligned by using CLUSTAL W (41). PhaRPd, PhaR of P. denitrificans (25); AniARe, AniA of Rhizobium etli (44.0% identity) (5); AniASm, AniA of Sinorhizobium meliloti (33.9% identity) (42); ORF4Me, ORF4 of Methylobacterium extorquens AM1 (32.0% identity) (17); ORF4Tv, ORF4 of Thiocystis violacea (34.8% identity) (21); ORF4Cv, ORF4 of Chromatium vinosum (32.0% identity) (13); PhaRRe, PhaR of Ralstonia eutropha (36.0% identity) (36); PhaRBt, PhaR of Burkholderia thailandensis E264 (35.0% identity) (15); StdCCt, StdC of Comamonas testosteroni (30.9% identity) (3); PhbFAs, PhbF of Azotobacter sp. strain FA8 (37.0% identity) (27). The asterisks and dots indicate identical and similar residues (lower similarity for single dots and higher similarity for double dots), respectively. The highly conserved region is boxed. The amino acid residues responsible for DNA binding mapped in this study are shaded in the box. Based on the Pfam tertiary structure prediction, a putative DNA-binding domain was mapped in the N-terminal region (positions 10 to 73) and a putative P(3HB)-binding domain was found in the middle region (75 to 115) of PhaR. These predicted regions are marked by inverted arrows.

 
Using the Pfam tertiary structure prediction software (http://www.sanger.ac.uk/Software/Pfam/), a DNA-binding domain was mapped in the N-terminal region (positions 10 to 73). This prediction is consistent with the importance of the N-terminal region for the binding of PhaR to the target DNA, as revealed by functional mapping. In the deletion experiment, the most minimized construct (C{Delta}73-195), with a small N-terminal region containing the predicted helix-turn-helix motif, was expressed as the soluble form, suggesting that this region would be an independently foldable domain structure. It is generally thought that the DNA-binding motifs of most transcription factors possess basic amino acids to mediate electrostatic interactions with the phosphate backbone of DNA. In our study, mutational effects (Table 2) indicate that a Lys residue at position 13 and an Arg residue at position 18 possibly correspond to basic amino acids that interact with phosphates of the target DNA. In addition, a Tyr residue frequently occupied in DNA-binding motifs of many transcription factors contributes to stacking interactions with rings of DNA bases. A Tyr at position 15 of PhaR might take part in this stacking interaction upon the binding of PhaR to the target DNA.

The helix-turn-helix GntR family of bacterial regulators was presented as transcription factors that share similar N-terminal DNA-binding domains, but it was observed that divergence exists in the C-terminal effector-binding domains (33). Such a mosaic organization comprising two domains of different origins would be applicable to PhaR. A plausible binding motif for the effector P(3HB) was found in the middle region (75 to 115) of PhaR by use of the Pfam software. In contrast to the high sequence homology in the N-terminal regions among PhaR homologs, sequence similarity is partly distributed in the predicted P(3HB)-binding motif region shared by PhaR family members. The next project has been initiated to address the amino acid residues of PhaR responsible for binding to P(3HB) by two approaches, X-ray crystallographic analysis and further random mutagenesis analysis, as also employed for a member of the granule-associated proteins, P(3HB) depolymerase (10, 11).


   ACKNOWLEDGMENTS
 
We are very grateful to C. T. Nomura, State University of New York, College of Environmental Science and Forestry, for critical reading. We would like to also thank M. Yao, Y. Satoh, K. Tojima, and T. Aizawa, Hokkaido University, for useful technical discussions.

Our work described here was partly supported by a Grant-in-Aid for Scientific Research of Japan (no. 70216828) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan (to S. Taguchi) and the Industrial Technology Research Grant Program in 2003 from the New Energy and Industrial Technology Development Organization (NEDO).


   FOOTNOTES
 
* Corresponding author. Mailing address: Graduate School of Engineering, Hokkaido University, N13W8, Kita-ku, Sapporo, Hokkaido 060-8628, Japan. Phone and fax: 81-(0)11-706-6610. E-mail: staguchi{at}eng.hokudai.ac.jp. Back

{triangledown} Published ahead of print on 22 November 2006. Back


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 Materials and Methods
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 Discussion
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Journal of Bacteriology, February 2007, p. 1118-1127, Vol. 189, No. 3
0021-9193/07/$08.00+0     doi:10.1128/JB.01550-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.




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