Institut für Mikrobiologie der Westfälischen Wilhelms-Universität Münster, Corrensstrasse 3, D-48149 Münster, Germany1
Institut für Mikrobiologie und Genetik der Georg-August-Universität Göttingen, Grisebachstrasse 8, D-37077 Göttingen, Germany2
Author for correspondence: Alexander Steinbüchel. Tel: +49 251 8339821. Fax: +49 251 8338388. e-mail: steinbu{at}uni-muenster.de
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ABSTRACT |
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Keywords: phaR, phaP regulation, repressor, inclusion bodies, autoregulation of phaR
Abbreviations: GARG, goat-anti-rabbit IgGgold; His6, hexahistidine; MM, mineral salts medium; PHA, polyhydroxyalkanoate; PHB, polyhydroxybutyrate; poly(3HB), poly(3-hydroxybutyrate)
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INTRODUCTION |
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R. eutropha is regarded as a model organism, along with Allochromatium vinosum, for studying short-carbon-chain-length PHA biosynthesis in bacteria (Steinbüchel & Schlegel, 1991 ). The genes responsible for PHA biosynthesis in R. eutropha have been cloned and characterized (Slater et al., 1988
; Schubert et al., 1988
; Peoples & Sinskey, 1989a
, b
) and comprise a ß-ketothiolase (phaA), an acetoacetyl-CoA reductase (phaB) and the PHA synthase (phaC). ß-Ketothiolase (EC 2 . 3.1.9) catalyses the first step of poly(3HB) biosynthesis, i.e. the condensation of two molecules of acetyl-CoA to acetoacetyl-CoA. Acetoacetyl-CoA is then reduced to R-(-)-3-hydroxybutyryl-CoA by an acetoacetyl-CoA reductase (EC 1.1.1.36). In the last step of the reaction, R-(-)-3-hydroxybutyryl-CoA is polymerized to poly(3HB) by PHA synthase, the key enzyme of PHA biosynthesis. The resulting PHA granules are coated with a layer of phospholipids and proteins, with phasins as the predominant compound. Phasins are a class of proteins of between 14 and 28 kDa in size that form a layer at the surface of the hydrophobic core of PHA granules; they also influence the number and size of PHA granules (Wieczorek et al., 1995
; Pieper-Fürst et al., 1995
; Steinbüchel et al., 1995
). The phasin of R. eutropha is encoded by phaP, and the formation of PhaP is dependent on PHA biosynthesis and accumulation (Wieczorek et al., 1995
). In R. eutropha, the amount of phasin present in cells has been shown to parallel the amount of PHA present in cells (York et al., 2001a
, b
).
Adjacent to the phaCAB gene cluster, an ORF (designated phaR) has been detected in R. eutropha (Slater et al., 1998 ). In this bacterium, PhaR seems to be involved in the regulation of phasin and PHA biosynthesis (York et al., 2002
). Other genes homologous to phaR have been found in several other PHA-accumulating bacteria (Rehm & Steinbüchel, 1999
), such as Sinorhizobium meliloti (Tombolini et al., 1995
; Povolo & Casella, 2000
). Recently, a phaR homologue (encoding PhaR) was identified in Paracoccus denitrificans which was capable of binding to the intergenic regions of phaCphaP and phaPphaR, and it was proposed that PhaR functions as a negative regulator of phasin synthesis (Maehara et al., 1999
, 2001
). Whereas much knowledge of PhaR has been gained by studying P. denitrificans, less is known about PhaR in the model organism R. eutropha. York et al. (2002)
suggested that PhaR promotes poly(3HB) synthesis in R. eutropha by regulating the expression of PhaP and by possibly regulating the expression of additional proteins. The aim of this study was to reveal the molecular mechanisms by which PhaR couples the synthesis of PhaP to the presence of poly(3HB) in R. eutropha. To achieve this, the phaR gene product of R. eutropha was analysed by electron microscopy investigations and DNA-binding experiments and by detailed analyses of a knock-out mutant. In combination with the results of a study that has been published recently (York et al., 2002
), a comprehensive model for the regulation of PHA granule formation in R. eutropha is suggested and discussed in this work.
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METHODS |
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Transfer of DNA.
Competent cells of E. coli were prepared and transformed by using the CaCl2 procedure described by Hanahan (1983) .
DNA sequencing.
This procedure was done by using the Sequi Therm EXCEL TM II Long Read Cycle Sequencing Kit (Epicentre Technologies), IRD800-labelled oligonucleotides (MWG-Biotech) and a Li-Cor 4000L (Li-Cor Biosciences) automated sequencer (MWG-Biotech).
PCR amplifications.
All PCR amplifications of DNA were carried out as described by Sambrook et al. (1989) , employing Pfx-DNA-polymerase (Gibco-BRL) and an Omnigene HBTR3CM DNA Thermal Cycler (Hybaid).
Inactivation of phaR in R. eutropha H16 by insertion of Km.
To amplify phaR from the genomic DNA of R. eutropha H16, oligonucleotides phaR_up (5'-AAAACGATCGCAGCACCATGTTCGTGCAGC-3') and phaR_rp (5'-AAAACGATCGATTCATTGCCTACGCACTCG-3') were designed (PvuI restriction sites are shown in bold). The 987 bp PCR product was cloned into pSK Bluescript-, to create pSKphaR. This plasmid was then digested with StuI. The linearized plasmid (pSKphaR) was ligated with Km that had been recovered from SmaI-digested pSKsym
Km DNA, whose construction has been described by Overhage et al. (1999)
. E. coli XL-1 Blue was transformed with the ligation mixture and transformants harbouring the resulting hybrid pSKphaR
Km, conferring kanamycin resistance, were obtained. To exchange the functional phaR gene for the inactivated one, phaR
Km was cloned into the suicide vector pSUP202 (Simon et al., 1983a
). To achieve this, phaR
Km was isolated from pSKphaR
Km after its digestion with PvuI and the fragment was ligated with PvuI-digested pSUP202. E. coli S17-1 was transformed with the ligation mixture and transformants harbouring the hybrid pSUP202phaR
Km, conferring resistance to tetracycline and kanamycin, were obtained. Subsequently, pSUP202phaR
Km was transferred to R. eutropha H16 by conjugation. Homogenotes resulting from a double crossover exhibiting a kanamycin-resistant phenotype were selected and distinguished from heterogenotes resulting from only a single crossover and exhibiting a tetracycline- plus kanamycin-resistant phenotype on MM agar plates containing the respective antibiotics. The genotype of homogenotes was controlled by PCR and DNA sequencing.
Cloning of phaR and purification of recombinant hexahistidine(His6)-tagged PhaR from recombinant E. coli.
For the cloning of phaR into E. coli, PCR was done using phaR_His_5 (5' - AAAAAA CATATG CATCAC CAC CAC CAC CACATGGCCACGACCAAAAAAGGCGC-3') as the sense primer and phaR_His_3 (5'-AAAAAATCTAGACAGCGTGCGGGATATGCG-3') as the anti-sense primer. These primers were deduced from the upstream and downstream regions, respectively, of the phaR gene of R. eutropha H16 (Slater et al., 1998 ). The phaRHis6 PCR product obtained was purified and ligated into pMa/c5-915 (Table 1
), which harbours the cI857ts gene encoding the temperature-sensitive
repressor. The recombinant His6PhaR (N-terminal fusion) was purified from E. coli TOP10 harbouring pMa/c5-914phaRHis6. Protein purification under native and denaturing conditions was conducted with Ni-NTA Spin Columns (Qiagen), as described by the manufacturer.
PAGE and Western immunoblotting.
Protein samples were resuspended in gel loading buffer [0·6% (w/v) SDS, 1·25% (w/v) ß-mercaptoethanol, 0·25 mM EDTA, 10% (v/v) glycerol, 0·001% (w/v) bromophenol blue, 12·5 mM Tris/HCl; pH 6·8] and separated in SDS 12·5% (w/v) polyacrylamide gels, as described by Laemmli (1970) . Proteins were stained with Coomasie brilliant blue R-250 (Weber & Osborn, 1969
) or with silver (Heukeshofen & Dernick, 1985
). Immunological detection of the PhaR protein blotted from the SDS-polyacrylamide gel onto PVDF membranes was done exactly as described by Towbin et al. (1979)
.
Preparation and purification of antibodies.
Approximately 850 µg of the PhaRHis6 protein was dissolved in 500 µl Tris/HCl (pH 7·0). This was then separated by SDS-PAGE and blotted onto a PVDF membrane. The membrane was sent to Eurogentec (Herstal, Belgium) for the production of polyclonal anti-PhaR antiserum in rabbits. The IgG fraction of the serum was purified on a protein A-Sepharose CL-4B affinity column (Hjelm et al., 1972 ). To obtain highly monospecific antibodies against PhaR, the antiserum was subjected to an affinity purification according to the method of Olmsted (1981)
with the modifications of Pieper-Fürst et al. (1994)
. Antibodies against PhaP were available from previous studies (Wieczorek et al., 1995
).
Primer extension.
The oligonucleotide primer PE_phaR (5'-ATCGCTGTTTGCGCGCTGCTGCACC-3') was 5'-labelled with IRD800. Total RNA (1 µg) extracted from R. eutropha H16 cells, which were grown under the storage conditions described above, was purified by using the RNeasy Mini Kit (Qiagen). The extension reaction was carried out at 50 °C for 50 min in a total volume of 50 µl in the presence of 200 U of Superscript II reverse transcriptase, as described by the manufacturer (Invitrogen). After its digestion with RNase A for 30 min at 37 °C, the sample was precipitated with ethanol, dissolved in H2O and then analysed on a 6% (w/v) polyacrylamide sequencing gel containing 8 M urea, employing the Li-Cor 4000L automated DNA sequencer. A sequencing reaction using the same labelled primer was run alongside the sample to determine the size of the primer-extension product. The phaR PCR product described in the gel-mobility-shift assay was used as template for the sequencing reaction.
Gel-mobility-shift assay.
To generate fragments comprising the upstream and downstream regions of phaP and phaR that could be employed in this assay, PCR was done using the genomic DNA of R. eutropha H16 and the appropriate primers. Amplification using primers USP_phaP_shift (5'-GCAATCGCGCATCGTTGAACACGCA-3') and DSP_phaP_shift (5'-GCCGAGGATCCTGTGCGCATCGGAG-3') with subsequent digestion of the PCR products with PstI and SmaI produced fragments of 126, 232, 518 and 775 bp. PCR products generated using primers USP_phaR_shift (5'-CTGCAGGCCGCCACG-3') and DSP_phaR_shift (5'-CCAGGATGATCTGCAGCAAGA-3') were digested with PstI to give fragments of 330 and 685 bp. These DNA fragments (1·5 µg) were mixed with purified PhaRHis6 (102000 ng) in binding buffer [1 mM EDTA, 10 mM Tris/HCl (pH 7·0), 80 mM NaCl, 10 mM ß-mercaptoethanol, 5% (w/v) glycerol] in a total volume of 20 µl. Incubation was carried out for 40 min at room temperature. After this incubation, 10xloading dye [250 mM Tris/HCl (pH 7·5), 0·2% (w/v) bromophenol blue, 0·2% (w/v) xylene cyanol, 40% (w/v) glycerol] was added to the samples. DNAprotein complexes were separated from unbound DNA fragments in 8% (w/v) native polyacrylamide gels, using 2xTris/borate/EDTA buffer (Sambrook et al., 1989 ). After electrophoresis, the gels were stained with ethidium bromide and the bands were visualized with a UV light.
DNaseI footprinting.
DNaseI footprinting was performed by using non-radioactive probes containing the IRD800 label in combination with a Li-Cor sequencer. For this purpose, DNA probes were prepared as follows. The PCR products described in the gel-mobility-shift assay were used as template DNA in PCR reactions with an IRD800-labelled primer (footprint III, 5'-CCCGGAGTGGCGTCACAGCCGCTCCC-3') in combination with DSP_phaP_shift (5'-GCCGAGGATCCTGTGCGCATCGGAG-3'), and USP_phaR_shift (5'-CTGCAGGCCGCCACG-3') in combination with PE_phaR (5'-ATCGCTGTTTGCGCGCTGCTGCACC-3'). Ten nanograms of the IRD800-labelled fragment was used per reaction. The binding reaction conditions for DNaseI footprinting were identical to the conditions used in the gel-mobility-shift assay. DNaseI cleavage was done by adding 20 µl of a solution containing 5 mM CaCl2, 10 mM MgCl2 and 2·5 mU of DNaseI (Gibco-BRL) to the PCR products. After 1 min, the DNaseI reaction was stopped by the addition of 20 µl of 4 M ammonium acetate and 30 mM EDTA. The DNA was extracted with 60 µl of phenol, precipitated with 96% ethanol in the presence of 40 µl of 50% glycogen and washed with 70% (v/v) ethanol. The pellet was dissolved in 1 µl of formamide loading buffer, heated at 95 °C for 5 min and then chilled on ice. Subsequently, 0·8 µl of the sample was analysed on a Li-Cor 4000L sequencer using a 6% denaturating sequence gel with 0·2 mm spacers and settings at 2000 V, 25 mA, 50 W and 45 °C.
Electron microscopy studies.
Cells were washed and suspended in 50 mM potassium phosphate buffer (pH 6·8), fixed in the presence of a mixture of 0·2% (v/v) glutaraldehyde plus 0·3% (w/v) paraformaldehyde and embedded in Spurrs low-viscosity resin (Spurr, 1969 ), as described by Walther-Mauruschat et al. (1977)
. For the post-embedding and immunogold labelling of PhaR and PhaP, cells were embedded in Lowicryl K4M (Lowri) as described by Roth et al. (1981)
, except that methanol was used instead of ethanol for dehydration. Immunological detection of PhaR and PhaP in ultra-thin sections, employing the primary antibodies and goat-anti-rabbit IgGgold (GARG) complex (Dakopatts), was done exactly as described by Pieper-Fürst et al. (1994)
. The specificity of the labelling was demonstrated by a control experiment using only the GARG complexes and mAbs against the Cap proteins of Sendai virus. Micrographs were taken with a Philips EM301 electron microscope at an acceleration voltage of 80 kV. Magnifications were calibrated with a cross-lined grating replica (Balzers).
Analysis of the PHA.
The samples [i.e. their poly(3HB)] were subjected to methanolysis in the presence of 15% (v/v) sulfuric acid. The resulting methyl esters of 3-hydroxybutyric acid were analysed by gas chromatography, as described by Brandl et al. (1988) and Timm & Steinbüchel (1990)
.
Preparation of PHA granules.
Granules consisting of poly(3HB) were isolated from R. eutropha H16 cells that had been grown in sodium gluconate, by employing the hypochlorite treatment described previously (Jendrossek et al., 1993 ). PHA granules were also isolated by centrifugation in sucrose gradients according to the method of Preusting et al. (1993)
with the modifications of Wieczorek et al. (1995)
.
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RESULTS |
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Localization of PhaR by Western immunoblotting
Because the in vitro binding studies shown above demonstrated the high affinity of His6PhaR for artificial poly(3HB) granules, it was very likely that the native protein is one of the granule-associated proteins occurring in R. eutropha. Various methods were employed to study the in vivo localization of PhaR. The cellular localization of native PhaR was analysed by Western immunoblotting using monospecific polyclonal anti-PhaR antibodies to probe the native PHB granules fraction, the membrane fraction and the soluble protein fraction (see Methods for details of the fractionation procedure). In addition, the crude cellular extract was subjected to Western immunoblotting. Fig. 4(a) clearly shows that PhaR was detected in the crude extract (lane 1), in the insoluble fraction (lane 3) and in the poly(3HB) granule-associated protein fraction (lane 4), whereas no PhaR could be detected in the soluble protein fraction (lane 2). The experiments showed that the purified polyclonal antibodies against PhaR were highly specific for this protein. Only one band was obtained in Western blots prepared from crude extracts of E. coli TOP10 harbouring pMa/cphaRHis6 or of the poly(3HB) granule-associated protein fraction (Fig. 4a
, lanes 5 and 4, respectively). To determine the quantities of the monospecific polyclonal anti-PhaR antibodies present, different concentrations (14, 28, 56, 112 and 224 µg) of the soluble protein fraction were subjected to Western immunoblotting and probing with the monospecific polyclonal anti-PhaR antibodies. The results clearly showed that PhaR was detected in the soluble protein fraction as a faint band only when 224 µg soluble protein was applied (data not shown).
Immunoelectron microscopic localization of PhaR
The monospecific polyclonal anti-PhaR antibodies were also used to localize PhaR in the cells at the ultrastructural level, by using immunoelectron microscopy. Cells of R. eutropha H16 and of the phaR knock-out mutant were embedded and ultra-thin sections were subjected to immunogold labelling. The labelling was mainly confined to only the periphery of the PHA granules in cells of the wild-type and there was no evidence that PhaR occurred within the core of the granules (Fig. 8). PhaR was not detected in the mutant.
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DISCUSSION |
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As the conclusion of their physiological study, York et al. (2002) proposed a rather simple and straightforward mechanism for the regulatory action of PhaR in R. eutropha. According to these authors, PhaR is a repressor that binds to a regulatory sequence upstream of phaP; however, DNA-binding studies were not done by this group. Upon the onset of PHB synthesis, PhaR was postulated to be removed from the binding region, thus allowing the formation of PhaP; however, the titrating factor was not identified. In the current study, we also found that PhaR binds upstream of phaP, as revealed by gel-mobility-shift assays (Fig. 5
). Moreover, the exact location of the binding site was identified by DNaseI footprinting experiments. The latter experiments showed that PhaR protects two DNA regions: one region is located directly upstream of the -35 region of the promoter that was identified previously (Steinbüchel et al., 1996
) and the second region maps from position -12 to +21. The protected regions comprise 14 and 33 bp and exhibit G+C ratios of 35 and 40 mol%, respectively. When the 14 and 33 bp binding sequences were analysed for sequence motifs that could serve as targets for DNA-binding proteins (e.g. palindromes, inverted or direct repeats), a 12 bp repeat sequence GCAMMAAWTMMD was identified on the sense and anti-sense strands, with the -10 and -35 regions of phaP located in between. These data clearly demonstrate (by the application of various independent methods) that PhaR binds to the promoter region of phaP. Maehara et al. (2001)
have also shown in gel-mobility-shift assays that the PhaR protein of P. denitrificans specifically binds to the intergenic phaCphaP and phaPphaR regions.
This study also revealed that the PHA granules are a very plausible candidate for the titrating factor of PhaR. The capability of PhaR to bind to PHA granules was demonstrated by in vitro experiments employing artificial granules as well as by in vivo cell fractionation and an immunoelectron microscopic approach employing specific anti-PhaR antibodies. Therefore, it seems plausible that upon the onset of PHA biosynthesis (i.e. when the nascent granules lack PhaP) PhaR is bound to the granules. Because of this, repression is removed from the phaP upstream binding region and phaP can be transcribed. The surface of the PHA granules is sufficiently large to bind the relatively small amounts of PhaR protein. However, the phasin PhaP most probably possesses a higher affinity for PHA granules than PhaR does. This could explain why in the later stages of PHA accumulation excess PhaR protein can again bind to the phaP upstream region, thus preventing the formation of more PhaP than can be bound to the granules.
By use of gel-mobility-shift assays, this study also clearly showed that PhaR binds not only to the phaP upstream region and to the PHA granules, but it also binds to the phaR upstream region. DNaseI footprinting experiments mapped the PhaR binding site to being approximately 86 bp upstream of the translational start site of phaR, which was also identified in this study. The transcriptional start site was identified as being 8 bp upstream of the putative -12 region. Normally, the distance from the transcriptional start to the -12 region is 48 bp (Hawley & McClure, 1983 ; Rosenberg & Court, 1979
). Since the number of copies of the phaR upstream region is limited by the number of copies of chromosomes in the cell, this region has a lower capacity to titrate significant quantities of PhaR than the surface of the PHA granules. However, this binding may allow the efficient autoregulation of PhaR expression and, thus, may prevent the synthesis of more PhaR than is required for the repression of PhaP expression. Steinbüchel et al. (1996)
have already suggested an autoregulation mechanism for the expression of PhaP; however, we have found that phaR (the repressor gene) is itself autoregulated and that phaP is not. Fig. 10
shows a model for the regulatory action of PhaR and its interactions with the phaP and phaR upstream regions and the PHA granules. This model is based on the results of this study and is also consistent with the results of previous studies (Wieczorek et al., 1995
; Steinbüchel et al., 1996
; Maehara et al., 2001
; York et al., 2002
).
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In conclusion, the interactions of the autoregulated repressor PhaR with the PHA granules and the phaP and phaR upstream promoter regions allow R. eutropha to regulate the expression of the phasin PhaP very efficiently. On the one hand, this regulation guarantees that the required large amounts (35% of total cellular protein) of PhaP can be synthesized if PHAs are synthesized. On the other hand, it ensures that PhaP is not synthesized in excess, thereby decreasing the burden imposed on the cells.
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ACKNOWLEDGEMENTS |
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Received 18 March 2002;
revised 8 April 2002;
accepted 29 April 2002.