Institut für Molekulare Mikrobiologie und Biotechnologie, Westfälische Wilhelms-Universität Münster, Corrensstraße 3, 48149 Münster, Germany
Correspondence
Alexander Steinbüchel
steinbu{at}uni-muenster.de
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ABSTRACT |
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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.
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INTRODUCTION |
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Biosynthesis of the most abundant type of PHASCL, poly(3-hydroxybutyrate) [poly(3HB)], starts from acetyl-CoA and in R. eutropha is catalysed by -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).
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METHODS |
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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 7080 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 Km.
For inactivation of the phaP2 gene by insertion of a kanamycin resistance cassette (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
Km, which was recovered from SmaI-digested pSKsym
Km DNA (Overhage et al., 1999
). E. coli Top10 was transformed with the ligation mixture, and transformants harbouring the resulting pUCBM20 : :
phaP2
Km were obtained. To exchange the functional phaP2 with the inactivated gene,
phaP2
Km was isolated from pUCBM20 : :
phaP2
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 : :
phaP2
Km were obtained. Subsequently, pSUP202 : :
phaP2
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 : : 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 : :
phaP4, which was used as template to amplify
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 : :
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 : :
phaP3 or pJQ200mp18Tc : :
phaP4 were used to generate the corresponding phaP3 and phaP4 mutants R. eutropha
phaP3, R. eutropha
phaP4, R. eutropha
phaP123 and R. eutropha
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 His6PhaR (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 His6PhaR fusion protein (0·051·25 µg) in binding buffer (1 mM EDTA, 10 mM Tris/HCl, pH 7·0, 80 mM NaCl, 10 mM -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
).
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RESULTS |
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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 (P2,
P3 and
P4), with the exception of the
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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Cells of R. eutropha wild-type and the 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
phaP2 and
phaP3 mutants revealed granules of similar number and size as in the wild-type and the
phaP4 mutant, and were therefore not analysed by electron microscopy.
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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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DISCUSSION |
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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 3646 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 35 % 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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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.
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ACKNOWLEDGEMENTS |
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Received 9 September 2004;
revised 16 November 2004;
accepted 19 November 2004.
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