Influence of homologous phasins (PhaP) on PHA accumulation and regulation of their expression by the transcriptional repressor PhaR in Ralstonia eutropha H16

Markus Pötter, Helena Müller and Alexander Steinbüchel

Institut für Molekulare Mikrobiologie und Biotechnologie, Westfälische Wilhelms-Universität Münster, Corrensstraße 3, 48149 Münster, Germany

Correspondence
Alexander Steinbüchel
steinbu{at}uni-muenster.de


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Phasins play an important role in the formation of poly(3-hydroxybutyrate) [poly(3HB)] granules and affect their size. Recently, three homologues of the phasin protein PhaP1 were identified in Ralstonia eutropha strain H16. The functions of PhaP2, PhaP3 and PhaP4 were examined by analysis of R. eutropha H16 deletion strains ({Delta}phaP1, {Delta}phaP2, {Delta}phaP3, {Delta}phaP4, {Delta}phaP12, {Delta}phaP123 and {Delta}phaP1234). When cells were grown under conditions permissive for poly(3HB) accumulation, the wild-type strain and all single-phasin negative mutants ({Delta}phaP2, {Delta}phaP3 and {Delta}phaP4), with the exception of {Delta}phaP1, showed similar growth and poly(3HB) accumulation behaviour, and also the size and number of the granules were identical. The single {Delta}phaP1 mutant and the {Delta}phaP12, {Delta}phaP123 and {Delta}phaP1234 mutants showed an almost identical growth behaviour; however, they accumulated poly(3HB) at a significantly lower level than wild-type and the single {Delta}phaP2, {Delta}phaP3 or {Delta}phaP4 mutants. Gel-mobility-shift assays and DNaseI footprinting experiments demonstrated the capability of the transcriptional repressor PhaR to bind to a DNA region +36 to +46 bp downstream of the phaP3 start codon. The protected sequence exhibited high similarity to the binding sites of PhaR upstream of phaP1, which were identified recently. In contrast, PhaR did not bind to the upstream or intergenic regions of phaP2 and phaP4, thus indicating that the expression of these two phasins is regulated in a different way. Our current model for the regulation of phasins in R. eutropha strain H16 was extended and confirmed.


Abbreviations: CDW, cell dry weight; His6, hexahistidine; PHA, polyhydroxyalkanoate; PHASCL, short carbon-chain-length PHA; poly(3HB), poly(3-hydroxybutyrate)

A list of oligonucleotides used in this study is given in Supplementary Table S1, the appearance of confluently grown colonies of the various Ralstonia eutropha mutants in comparison to the wild-type is shown in Supplementary Fig. S1, and the results of gel-mobility-shift assays of PhaR binding to DNA fragments comprising up- and downstream regions of phaP2, phaP3 and phaP4, and footprinting are shown in Supplementary Fig. S2 with the online version of this paper at http://mic.sgmjournals.org.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Polyhydroxyalkanoates (PHAs) are polyoxoesters that are synthesized and accumulated as cytoplasmatic inclusions by diverse bacteria. Today, more than 140 different PHA constituents have been identified, and PHAs represent a versatile class of microbial polymers (Steinbüchel & Valentin, 1995). The major role of PHAs is that of a useful reserve material for carbon and energy, considering the ability to store large quantities of reduced carbon without significant effects on the osmotic pressure of the cell (Anderson & Dawes, 1990). Referring to beneficial properties, such as biodegradability and origin from renewable resources, PHAs have attracted much interest in academia and industry, and many technical and medical applications of PHAs have been developed (Anderson & Dawes, 1990; Hocking & Marchessault, 1994; Asrar & Gruys, 2002; Williams & Martin, 2002). Special emphasis has been given to transgenic plants that produce Biopol (Poirier & Gruys, 2002). The Gram-negative facultative chemolithoautotrophic hydrogen-oxidizing bacterium Ralstonia eutropha is regarded as a model organism, along with Allochromatium vinosum, for studying the biosynthesis of short carbon-chain-length PHA (PHASCL). Therefore, it is necessary to investigate in detail the morphogenesis and granule structure of PHASCL, as well as synthesizing and mobilizing enzymes of PHASCL in R. eutropha.

Biosynthesis of the most abundant type of PHASCL, poly(3-hydroxybutyrate) [poly(3HB)], starts from acetyl-CoA and in R. eutropha is catalysed by {beta}-ketothiolase (PhaA), acetoacetyl-CoA reductase (PhaB) and the key enzyme of PHA biosynthesis, PHA synthase (PhaC): all three are encoded by the phaCAB operon (Oeding & Schlegel, 1973; Haywood et al., 1988a, b; Slater et al., 1988; Schubert et al., 1988; Peoples & Sinskey, 1989). The resulting PHA granules are surrounded by a layer of phospholipids and proteins, with phasins as the predominant compound. Phasins are a class of low-molecular-mass amphipathic proteins that form a layer at the surface of the lipophilic poly(3HB) granule (Steinbüchel et al., 1995). They occur in any PHASCL-accumulating bacterium, and are analogues of oleosins, which are bound to the surface of the oleosome in plants (Wieczorek et al., 1995; Pieper-Fürst et al., 1995; Huang, 1992; Steinbüchel et al., 1995).

It was recently shown that R. eutropha strain H16 expresses, in addition to PhaP1, three homologous phasins (PhaP2, PhaP3, PhaP4), which are also located at the surface of PHA granules or which possess the capability to bind to PHA granules (Pötter et al., 2004; Srinivasan et al., 2002; Schwartz et al., 2003). Absence of phasin PhaP1 influences the size and number of PHA granules in bacteria, and the amount of PhaP1 parallels the quantity of PHA present in the cells (Wieczorek et al., 1995; York et al., 2001a, b). The expression of phaP1 is regulated by the transcriptional repressor PhaR (Pötter et al., 2002; York et al., 2002).

The occurrence of four phasin proteins in R. eutropha suggests that the three additional homologous phasins might also have an influence on PHA homeostasis and on the amount of PHA accumulated in the cells. To understand the functions of PhaP2, PhaP3 and PhaP4, knock-out mutants of the various phasin genes were generated. The effects of the various phaP deletions on poly(3HB) accumulation were monitored and discussed. Furthermore, DNA-binding experiments were performed to reveal whether or not the expression of the phasin homologues is also regulated by PhaR (Pötter et al., 2002).


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Bacterial strains and growth conditions.
Bacterial strains used in this study are listed in Table 1. Cells of R. eutropha were grown in 1 l Erlenmeyer flasks equipped with baffles at 30 °C in 200 ml mineral salts medium (MSM) supplemented with 1·5 % (w/v) sodium gluconate (Schlegel et al., 1961). To promote accumulation of PHA, the concentration of NH4Cl was reduced to 0·02 % (storage conditions). Tetracycline and kanamycin were used at final concentrations of 25 and 160 µg ml–1, respectively. Cells of Escherichia coli were cultivated at 37 °C in Luria–Bertani (LB) medium (Sambrook et al., 1989). Solid media contained 1·5 % (w/v) agar-agar. Growth was monitored photometrically with a Klett–Summerson photometer (Manostat) using filter no. 54 (520–580 nm). Mating of E. coli S17-1 with R. eutropha was performed at 30 °C on LB agar plates. The sacB gene selection was done on LB agar plates supplemented with 5 % sucrose and 0·2 % fructose at 30 °C.


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Table 1. Bacterial strains and plasmids used in this study

 
Viable-colony staining.
To visualize poly(3HB) in cells, 0·5 µg Nile red (ml medium)–1 was added (Spiekermann et al., 1999). The agar plates were exposed to UV light (312 nm) after appropriate cultivation periods to detect accumulation of PHB.

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 Spurr's low-viscosity resin (Spurr, 1969; Walther-Mauruschat et al., 1977). Sections with a thickness of 70–80 nm were made with a diamond knife (Ultracut, Leica) and placed on a 200 mesh copper grid. Imaging was performed with an H-500 TEM (Hitachi) in the bright-field mode at 80 kV acceleration voltage and at room temperature.

Isolation and manipulation of DNA.
Chromosomal DNA of R. eutropha H16 was isolated by the method of Marmur (1961). Plasmid DNA was isolated by the protocol of Birnboim & Doly (1979). DNA restriction fragments were purified with the Nucleotrap kit (Machery-Nagel) and restriction enzymes, ligases and other DNA-manipulating enzymes were used according to the manufacturer's instructions.

Transfer of DNA.
Competent cells of E. coli were prepared and transformed by the CaCl2 procedure, as described by Hanahan (1983).

PCR amplification.
All PCR amplifications of DNA were carried out as described by Sambrook et al. (1989), employing Pfx-DNA-polymerase (Invitrogen), an Omnigene HBTR3CM DNA thermal cycler (Hybaid) and the primers listed in Supplementary Table SI (available as supplementary data with the online version of this paper at http://mic.sgmjournals.org).

DNA sequencing.
Sequencing was done by using the Sequi Therm EXCEL TM II long read cycle sequencing kit (Epicentre Technologies) and IRD 800-labelled oligonucleotides (MWG-Biotech) in a Li-Cor 4000L (Li-Cor Biosciences) automatic sequencing apparatus (MWG-Biotech).

Inactivation of phaP2 in R. eutropha by insertion of the omega element {Omega}Km.
For inactivation of the phaP2 gene by insertion of a kanamycin resistance cassette ({Omega}Km), hybrid plasmid pUCBM20 : : phaP2 was constructed. For this, two oligonucleotides (phaP2_XbaI_EcoRI_fw and phaP2_EcoRI_rv) were designed to amplify phaP2 and its adjacent regions. The 2214 bp PCR product was cloned into pUCBM20 to create pUCBM20 : : phaP2, which was then digested with HincII. The linearized plasmid was ligated with {Omega}Km, which was recovered from SmaI-digested pSKsym{Omega}Km DNA (Overhage et al., 1999). E. coli Top10 was transformed with the ligation mixture, and transformants harbouring the resulting pUCBM20 : : {Delta}phaP2{Omega}Km were obtained. To exchange the functional phaP2 with the inactivated gene, {Delta}phaP2{Omega}Km was isolated from pUCBM20 : : {Delta}phaP2{Omega}Km after digestion with EcoRI and ligated with EcoRI-digested pSUP202 DNA (Simon et al., 1983a). E. coli S17-1 was transformed with the ligation mixture, and transformants harbouring pSUP202 : : {Delta}phaP2{Omega}Km were obtained. Subsequently, pSUP202 : : {Delta}phaP2{Omega}Km was transferred to R. eutropha H16 by conjugation. The genotype of homogenotes was controlled by PCR and DNA sequencing.

Modification of the suicide vector pJQ200mp18.
Plasmid pJQ200mp18 was modified to use tetracyline resistance as selectable marker. A 1304 bp PCR fragment using the oligonucleotides Tc_BglII_fw and Tc_BglII_rv encoding tetracyline resistance was excised from pBR322 (Bolivar et al., 1977) and cloned into the BglII site of pJQ200mp18 to yield pJQ200mp18Tc.

Construction of phaP3 and phaP4 precise deletion gene replacement plasmids.
All oligonucleotides used for PCR are listed in Supplementary Table S1 (available as supplementary data with the online version of this paper at http://mic.sgmjournals.org). The 919 bp and 777 bp fragments upstream and downstream of phaP3 were amplified employing phaP3_fw and phaP3_EcoRI_rv or phaP3_EcoRI_fw and phaP3_rv, respectively. The resulting fragments were EcoRI digested and ligated to yield a 1696 bp fragment. This fragment was amplified using phaP3_BamHI_fw and phaP3_BamHI_rv, and the resulting PCR product was cloned into the SmaI site of pJQ200mp18Tc to yield pJQ200mp18Tc : : {Delta}phaP3. Similarly, the 999 bp and 584 bp fragments upstream and downstream of phaP4 were amplified employing phaP4_BamHI_fw and phaP4_XbaI_rv or phaP4_XbaI_fw and phaP4_BamHI_rv, respectively. The 999 bp fragment was digested with BamHI and XbaI and cloned into pUCBM20 to obtain pUCBM20 : : phaP4A. The 584 bp fragment was cloned into XbaI- and EcoRV-digested pUCBM20 : : phaP4A to obtain pUCBM20 : : {Delta}phaP4, which was used as template to amplify {Delta}phaP4 by PCR employing phaP4_BamHI_fw and phaP4_BamHI_rv. The resulting 1583 bp fragment was cloned into SmaI-digested pJQ200mp18Tc to yield pJQ200mp18Tc : : {Delta}phaP4.

Construction of phaP gene replacement strain using the sacB system.
Gene replacement was accomplished by adaptation of standard protocols (Slater et al., 1998; Quandt & Hynes, 1993). Plasmids pJQ200mp18Tc : : {Delta}phaP3 or pJQ200mp18Tc : : {Delta}phaP4 were used to generate the corresponding phaP3 and phaP4 mutants R. eutropha {Delta}phaP3, R. eutropha {Delta}phaP4, R. eutropha {Delta}phaP123 and R. eutropha {Delta}phaP1234. The plasmids were mobilized from E. coli donor strain S17-1 to the respective R. eutropha recipient strains by the spot agar mating technique (Hogrefe et al., 1981). Successful gene replacement strains were identified and confirmed by PCR analyses and DNA sequencing.

Expression and purification of recombinant hexahistidine (His6)-tagged PhaR from E. coli.
The recombinant His6–PhaR (N-terminal fusion) was expressed in E. coli Top10 (Invitrogen) harbouring pMa/c5-914 : : phaRHis6 and purified using a Ni-NTA-Superflow column (Qiagen), as described by Pötter et al. (2002).

Gel-mobility-shift experiments.
Fragments comprising the upstream and downstream regions of phaP2, phaP3 and phaP4 or the respective structural genes, which were to be employed in gel-mobility-shift experiments, were obtained by PCR using genomic DNA of R. eutropha H16 and the following primers, plus subsequent treatment of the PCR products with restriction endonucleases: phaP2_gelshift_fw and phaP2_gelshift_rv with PvuII to give 162, 220, 362, 632 and 695 bp fragments; phaP3_gelshift_fw and phaP3_gelshift_rv with AvaI to give 367, 438, 608 and 713 bp fragments; phaP4_gelshift_fw and phaP4_gelshift_rv with StuI to give 254, 468, 666 and 823 bp fragments. These DNA fragments (1·5 µg) were mixed with purified His6–PhaR fusion protein (0·05–1·25 µg) in binding buffer (1 mM EDTA, 10 mM Tris/HCl, pH 7·0, 80 mM NaCl, 10 mM {beta}-mercaptoethanol, 5 %, w/v, glycerol) in a total volume of 20 µl. Incubation and separation were performed exactly as described by Pötter et al. (2002). After electrophoresis, the gels were stained with ethidium bromide and the DNA bands were visualized with UV light.

DNaseI footprinting.
DNaseI footprinting experiments were performed using non-radioactive probes containing the IRD800 label with a Li-Cor sequencer. The PCR products described above in the gel-mobility-shift experiments were used as template DNA in PCR reactions with an IRD800-labelled primer (footprint_phaP3). Ten nanograms of this IRD800-labelled fragment was used for each reaction. Binding reactions for DNaseI footprinting were identical to the binding reaction conditions in gel-mobility-shift experiments (see above). For DNaseI cleavage, 20 µl of a solution containing 5 mM CaCl2, 10 mM MgCl2 and 2·5 mU of DNaseI (Gibco) was added; the reaction was stopped after 1 min by addition of 20 µl 4 M ammonium acetate and 30 mM EDTA. DNA was extracted with 60 µl phenol, precipitated with 96 % (v/v) ethanol in the presence of 40 µl 50 % glycogen, and washed with 70 % (v/v) ethanol. The pellet was dissolved in 1 µl formamide loading buffer, heated at 95 °C for 5 min, and chilled on ice. Subsequently, 0·8 µl was analysed on the Li-Cor sequencer using a 6 % denaturating sequence gel with 0·2 mm spacers and the following settings: 2000 V, 25 mA, 50 W and 45 °C. The protected nucleotide sequences of phaP1, phaP3 and phaR were aligned using CLUSTALW, created by Thompson et al. (1994).

PHA quantification.
Samples were subjected to methanolysis in the presence of 15 % (w/v) sulfuric acid and the methyl esters of 3-hydroxybutyric acid were analysed by gas chromatography (Brandl et al., 1988; Timm & Steinbüchel, 1990).


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Generation of phasin mutants
In addition to an already existing phaP1 mutant, two sets of mutants were generated. The first set comprised single mutants defective in phaP2 ({Delta}P2), phaP3 ({Delta}P3) or phaP4 ({Delta}P4). The phaP2 mutant was obtained by inserting a kanamycin-resistance cassette into phaP2 as described in Methods. From phaP3 and phaP4 precise deletion mutants were generated by gene replacement employing suicide plasmids that comprise the upstream and downstream regions of these genes only. The second set comprised multiple mutants that were all derived from the {Delta}phaP1 mutant. The sacB system from Bacillus subtilis was applied to R. eutropha to generate deletion mutants of the respective phasin genes to avoid additional antibiotic resistances. This sacB system is inducible by sucrose, and is lethal when expressed in Gram-negative bacteria (Quandt & Hynes, 1993). Using the suicide vector pJQmp18Tc, which was constructed in this study, tetracycline resistance can now also be used as a convenient antibiotic-resistance marker for selection of homogenotes. Applying this system, a double mutant defective in phaP1 and phaP2 ({Delta}P12), a triple mutant defective in phaP1, phaP2 and phaP3 ({Delta}P123), and a quadruple mutant defective in all four phasin genes ({Delta}P1234) were generated as described in Methods.

Phenotypic characterization of the phasin mutants
To determine the growth behaviour and the capability to synthesize and accumulate poly(3HB) of the various phaP mutants of R. eutropha H16, the wild-type and mutant strains were cultivated under conditions permissive for poly(3HB) accumulation in liquid MSM containing 1·0 % (w/v) sodium gluconate and 0·02 % (w/v) NH4Cl.

The wild-type strain and all single-phasin negative mutants ({Delta}P2, {Delta}P3 and {Delta}P4), with the exception of the {Delta}P1 strain, showed initially a similar growth behaviour according to the optical density of the cultures (Fig. 1a). However, the increase of the optical density became slightly slower after about 12 h cultivation in the single phaP2, phaP3 and phaP4 mutants. After about 18 h cultivation, all cultures of these mutants had reached the same density of about 950 Klett Units. These studies indicated that growth is almost unaffected in these phaP mutants and that none of the phasin genes is essential for growth or PHA accumulation in R. eutropha. The latter was also true for the phaP1 mutant; however, the increase of optical density and final density were significantly slower and less, respectively, than those of the wild-type or other phaP mutants (see below). The phaP2, phaP3 and phaP4 mutants, like the wild-type, reached their maximum poly(3HB) amount after 28 h and accumulated poly(3HB) to 80 % (w/w) of the cell dry weight (CDW) in the stationary growth phase (Fig. 1c). This was also visible by inspecting colonies on MSM gluconate agar plates (Supplementary Fig. S1, available as supplementary data with the online version of this paper at http://mic.sgmjournals.org). The single phaP2, phaP3 and phaP4 mutants, in contrast to the phaP1 mutant, did not exhibit any significant deviation from that of the wild-type in colony size or opacity, or Nile red-mediated fluorescence due to poly(3HB), which is consistent with the other observations described above.



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Fig. 1. Growth behaviour (a, b) and poly(3HB) accumulation (c, d) of R. eutropha strain H16 and derivatives in MSM. Composition of the medium and cultivation conditions were as described in Methods. After 8, 16, 24 and 28 h, samples were withdrawn and the poly(3HB) contents of the cells were analysed by gas chromatography. (a) Growth of wild-type (x) and the mutants {Delta}P1 ({blacksquare}), {Delta}P2 ({bullet}), {Delta}P3 ({blacktriangleup}) and {Delta}P4 ({blacklozenge}). (b) Growth of wild-type (x) and the mutants {Delta}P1 ({blacksquare}), {Delta}P12 ({bullet}), {Delta}P123 ({blacktriangleup}) and {Delta}P1234 ({blacklozenge}). (c) Poly(3HB) contents of cells of wild-type (black shading) and of the mutants {Delta}P2 (large dots), {Delta}P3 (diagonal hatching), {Delta}P4 (small dots) and {Delta}P1 (grey shading). (d) Poly(3HB) contents of cells of wild-type (black shading) and of the mutants {Delta}P1 (grey shading); {Delta}P12 (large dots), {Delta}P123 (diagonal hatching) and {Delta}P1234 (small dots). Each experiment was done twice. Error bars are shown.

 
In contrast, the {Delta}P1, {Delta}P12, {Delta}P123 and {Delta}P1234 mutants exhibited very similar but significantly slower growth compared to the wild-type and after 24 h, maximum optical densities of only 800–830 Klett Units were obtained (Fig. 1d). Each of these strains was also examined for poly(3HB) accumulation after cultivation on MSM medium under storage conditions (Fig. 1d). The single {Delta}P1 mutant as well as the double ({Delta}P12), triple ({Delta}P123) and quadruple ({Delta}P1234) mutants contained after 28 h incubation a significantly lower amount of poly(3HB) than the wild-type strain (Fig. 1d). The poly(3HB) amounts in cells of the double (38 %, w/w, of CDW), triple (36 %) and quadruple (30 %) mutants were significantly lower than in cells of the single phaP1 mutant (50 %) (Fig. 1d). As a consequence, the final optical density in these mutants was lower (about 800–830 Klett Units, Fig. 1b) than in mutants of the first set harbouring intact phaP1 (about 900–950 Klett Units, Fig. 1a). This was also clearly visible on solid media and by the visible-colony staining method (Supplementary Fig. S1). The {Delta}phaP1 mutant, the double mutant {Delta}phaP12, the triple mutant {Delta}phaP123 and the quadruple mutant {Delta}phaP1234 exhibited a much weaker fluorescence and also a significantly lower opacity than the wild-type.

Cells of R. eutropha wild-type and the {Delta}phaP4 mutant were also analysed by electron microscopy. In contrast to the phaP1 mutant, which harboured only one large poly(3HB) granule (Wieczorek et al., 1995), both strains exhibited a large number of closely packaged poly(3HB) granules of medium size in the cytoplasm (Fig. 2). The mean size and number of poly(3HB) granules was almost identical to that of the wild-type when a large number of electron microscopic pictures of thin sections of cells of the phaP4 mutant were statistically analysed. Light microscopic studies of the {Delta}phaP2 and {Delta}phaP3 mutants revealed granules of similar number and size as in the wild-type and the {Delta}phaP4 mutant, and were therefore not analysed by electron microscopy.



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Fig. 2. Morphology of cells of the wild-type R. eutropha H16 (a), the {Delta}phaP1 mutant (b) and the {Delta}phaP4 mutant (c). Cells were cultivated in MSM containing 0·02 % (w/v) NH4Cl plus 1·5 % (w/v) sodium gluconate and harvested in the stationary growth phase. Thin sections were prepared and electron micrographs were obtained as described in Methods. Bars, 0·5 µm.

 
Binding of the regulator PhaR to phaP2, phaP3 and phaP4
To investigate the interaction of PhaR with phaP2, phaP3 and phaP4, gel-mobility-shift assays were performed. Fragments derived from the phaP2 and phaP4 regions exhibited no shift in migration while the concentration of the His6–PhaR fusion protein was increased (Supplementary Fig. S2a, c, respectively, available as supplementary data with the online version of this paper at http://mic.sgmjournals.org). In contrast, the 713 bp AvaI–AvaI fragment containing phaP3 DNA clearly shifted (Supplementary Fig. S2b), whereas migration of the other fragments (i.e. 376 bp, 438 bp and 608 bp) obtained by AvaI restriction was not retarded. These gel-mobility-shift assays clearly demonstrated binding of PhaR to an intragenic region of phaP3.

Determination of the PhaR binding site by DNaseI footprinting
To identify the exact binding site of the transcriptional repressor PhaR (Pötter et al., 2002; York et al., 2002) within phaP3, DNaseI footprinting experiments were performed. For this, purified PhaR protein and the PCR product harbouring phaP3 were incubated as described in Methods. Addition of PhaR to the sample resulted in a distinct DNaseI protection of a DNA region +36 to +46 bp downstream of the phaP3 start codon. The sequence of this region is shown in Supplementary Fig. S2d. Pötter et al. (2004) identified three 12 bp sequences as PhaR binding regions upstream of phaP1; in addition, a binding site of PhaR was identified upstream of phaR. These regions were aligned with the region protected in phaP3, which is also shown in Fig. 3. Comparison of these sequences revealed several conserved nucleotides. Five nucleotides of the phaP3 binding region were identical to the binding sites of PhaR to phaP1, and two nucleotides of the phaR binding region were identical to the binding sites of phaP1 and phaP3.



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Fig. 3. Comparison of PhaR DNA binding sites. Multiple sequence alignment of the DNA sequence of phaP1 (–49 to –36 bp relative to the phaP1 transcriptional start site), phaP1' (+10 to +21 bp), phaP1'' (+6 to –7 bp), phaR (+11 to +18 bp) and the protected sequence of phaP3. Cons', consensus sequence.

 

   DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Recent studies and the analysis of the R. eutropha strain H16 genome sequence have shown that the metabolism of the storage compound poly(3HB) in this bacterium is much more complex than previously assumed. For example, in addition to phaP1, three other phasin homologous genes (phaP2, phaP3 and phaP4) have been annotated, and it has been shown that all four phasin proteins are expressed in R. eutropha (Schwartz et al., 2003; Pötter et al., 2004). To gain more information about the function of these homologous phasins, two sets of knock-out mutants were generated and the influence of the phasins on growth and poly(3HB) accumulation was investigated. Cultivation experiments with the wild-type and the {Delta}phaP2, {Delta}phaP3 and {Delta}phaP4 single mutants of R. eutropha revealed only small differences between these strains with regard to growth and PHA accumulation. These results clearly showed that in single mutants lacking expression of PhaP2, PhaP3 or PhaP4, the amounts of poly(3HB) accumulated by the cells were not significantly affected and their phenotypes were identical. Only the phaP1 mutant exhibited a clear poly(3HB)-leaky phenotype and was clearly distinguished from the wild-type and the other single-phasin mutants. Mutants of the second set, which were all defective in phaP1 and which were in addition also defective in phaP2, or phaP2 and phaP3, or phaP2, phaP3 and phaP4, behaved clearly differently and exhibited a phenotype. Growth of these mutants was only slightly affected (Fig. 1b) and cells of mutants with an increasing number of defective phasin genes accumulated smaller amounts of poly(3HB) than the wild-type (Fig. 1d). In addition, the size and number of the poly(3HB) granules in the single-phasin mutants {Delta}phaP2, {Delta}phaP3 and {Delta}phaP4 exhibited no differences from the wild-type strain (Fig. 2). In contrast, the {Delta}phaP1 mutant formed only one, large poly(3HB) granule (Wieczorek et al., 1995). All these data and the results of previous studies clearly demonstrated that only PhaP1 has a major influence on poly(3HB) accumulation and that it is the major phasin protein in R. eutropha H16.

In addition to the high sequence similarities of PhaP1 and PhaP3 (Pötter et al., 2004), further common features of these two phasins were found. Gel-mobility-shift experiments showed that the transcriptional repressor PhaR binds to phaP3, whereas it does not bind to phaP2 or phaP4, respectively (Supplementary Fig. S2). This indicates that the expression of PhaP1 and PhaP3 is regulated by a common mechanism. It may also indicate that both phasins have a similar function. This assumption is supported by the recent finding that PhaP3 occurred at high levels in cells of the phaP1 deletion mutant (Pötter et al., 2004). DNaseI footprinting analysis (Supplementary Fig. S2d) showed that PhaR binds to an intragenic region of phaP3 located 36–46 bp downstream of the translational start site. This makes binding of PhaR to phaP3 different from the binding of this protein to phaP1 and phaR, where binding occurs upstream of the structural genes. In addition, an alignment of the DNA sequences of the PhaR binding sites upstream of phaP1 and phaP3 clearly indicated high similarities, whereas the PhaR binding site upstream of its own structural gene exhibited a lower similarity. Therefore, different binding intensities of PhaR to the phaP1 and phaP3 binding sites in comparison to the phaR binding site can be expected.

Therefore, our previous model of the regulation of PhaP1 formation was extended and PhaP3 was included (Fig. 4). If the cells are cultivated under conditions non-permissive for PHA biosynthesis, PhaR cannot bind to poly(3HB) granules because they are absent from the cells. The cytoplasmic concentration of PhaR is sufficiently high to repress transcription of phaP1 and phaP3. If physiological conditions permissive for poly(3HB) biosynthesis occur, the constitutively expressed PHA synthase starts to synthesize poly(3HB) molecules, which remain covalently linked to this enzyme. Initially, small micelles are formed, which become larger and constitute the nascent poly(3HB) granules. Proteins such as PhaR, with a binding capacity to the hydrophic surface, bind to the granules. This lowers the cytoplasmic concentration of PhaR. From a certain point, the cytoplasmic concentration of PhaR becomes too low to sufficiently repress transcription of phaP1 and phaP3 any longer. PhaP1 and also PhaP3 are therefore synthesized and subsequently bind to the poly(3HB) granules. The granules become larger and reach their maximum size. Therefore, the PhaP1 protein is being continuously synthesized in sufficient amounts. In addition, small amounts of PhaP3 are also synthesized. The reasons for formation of less PhaP3 than PhaP1 (Pötter et al., 2004) may be many. One reason may be a stronger repression of the transcription of the phaP3 gene by PhaR in comparison to the phaP1 gene. When the poly(3HB) granules have reached the maximum size possible, which may be due to the limited space in the cytoplasm or to the depletion of the carbon source, most of the poly(3HB) granule surface will be covered with PhaP1 protein, which contributes about 3–5 % of the total cellular protein, in addition to lower amounts of PhaP3 and PhaR. In this situation, no more space will be available at the PHA granule surface for binding additional PhaR, or PhaR may even be displaced by PhaP1 (and PhaP3). Consequently, the cytoplasmic concentration of PhaR will increase and exceed the threshold concentration required again to repress transcription of phaP1 and phaP3. PhaP1 and PhaP3 proteins are, as a consequence, no longer synthesized, and these phasins are therefore not produced in higher amounts than required to cover the surface of poly(3HB) granules. In addition, the binding capacity of PhaR to the promoter region of its own gene prevents overexpression of this repressor protein, which is therefore under autocontrol. Where no PhaP1 is produced in a phaP1 deletion mutant, we have found higher concentrations of PhaP3 protein in the cells (Pötter et al., 2004). This is consistent with the extended model described above, because PhaR no longer has to compete with the major phasin protein PhaP1 for binding at the PHA granule surface. The cytoplasmic concentration of PhaR may therefore be lower in cells of a phaP1 mutant, and repression of phaP3 transcription may be further diminished.



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Fig. 4. Summary of interactions between PhaR, PhaP1, PhaP3, PHA granules, phaR, phaP1 and phaP3. The PHA synthase and PHA depolymerases are not shown for reasons of simplification.

 
In conclusion, the homologous phasins have different influences on PHA accumulation behaviour. PhaP1 can be assigned to the major phasins, and the absence of PhaP1 dramatically influences the amounts of poly(3HB) accumulated in the cells. The influence of each of the PhaP1 homologues on poly(3HB) accumulation is less severe. However, the absence of the major phasin PhaP1 in combination with a deletion of phaP2, or phaP2 and phaP3, or phaP2, phaP3 and phaP4, affects poly(3HB) accumulation more severely.

The functions of PhaP2 and PhaP4 are not understood and should be revealed in further studies. Both proteins are expressed at much lower levels than PhaP1 and even than PhaP3, and transcription of both genes is not repressed by PhaR. In addition, PhaP2 seems in vivo not to be bound to the granules under the conditions which were tested, although it is capable of binding to artificial poly(3HB) granules (Pötter et al., 2004). PhaP2 and PhaP4 cannot therefore be considered as phasins sensu strictu and may have a different function for which only low concentrations of these proteins are required. In the future, we will investigate whether the functions of these two proteins are related to the mobilization of poly(3HB) and whether they interact with one of the five PHA depolymerases of R. eutropha.


   ACKNOWLEDGEMENTS
 
This study was supported by a grant provided by the Deutsche Forschungsgemeinschaft (STE 386/6-1 and STE 386/6-2). The project was also carried out within the framework of the Competence Network Göttingen ‘Genome research on bacteria’ (GenoMik) financed by the German Federal Ministry and Research (BMBF). We thank Ursula Malkaus and Rudolf Reichelt for expert electron microscopic preparation and for taking the TEM micrographs. The authors are indebted to A. J. Sinskey (MIT, Cambridge, MA, USA) for providing R. eutropha strain Re1052. We thank Sandra Kohaus for technical assistance.


   REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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Received 9 September 2004; revised 16 November 2004; accepted 19 November 2004.



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