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1 Institut für Molekulare Mikrobiologie und Biotechnologie, Westfälische Wilhelms-Universität, Corrensstraße 3, D-48149 Münster, Germany
2 Department of Cell Biology and Ultrastructural Research, Leibniz Institute for Arteriosclerosis Research, University of Münster, Germany
Correspondence
Alexander Steinbüchel
steinbu{at}uni-muenster.de
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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-galactosidase (LacZ) were expressed separately in the triacylglycerol (TAG)-accumulating actinomycetes Rhodococcus opacus PD630 and Mycobacterium smegmatis mc2155, employing the M. smegmatis acetamidase (ace) promoter of the Escherichia–Mycobacterium/Rhodococcus shuttle plasmid pJAM2. PhaP1 and the PhaP1 fusion proteins were expressed stably in the recombinant strains. Western blot analysis of cell fractions of Rh. opacus revealed that PhaP1 and the PhaP1–eGFP fusion protein were associated with the TAG inclusions, whereas no phasin or phasin fusion protein was detected in the soluble and membrane fractions. Additional electron microscopy/immunocytochemistry studies demonstrated that PhaP1 was mainly located on the surface of intracellular TAG inclusions; in addition, some PhaP1 also occurred at the plasma membrane. Fluorescence microscopic investigations of the subcellular distribution of the PhaP1–eGFP fusion protein in vivo and on isolated TAG inclusions revealed that the fusion protein was bound to TAG inclusions at all stages of their formation, and to some extent at the cytoplasmic membrane. The PhaP1–LacZ fusion protein also bound to the TAG inclusions, and could be separated together with the inclusions from Rh. opacus crude extracts, thus demonstrating the immobilization of -galactosidase activity on the inclusions. This is believed to be the first report demonstrating the ability of PhaP1 to bind to lipid inclusions in addition to PHA inclusions. Furthermore, it was demonstrated that this non-specificity of PhaP1 can be utilized to anchor enzymically active fusion proteins to a matrix of bacterial TAG inclusions.
These authors contributed equally to this work.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Wältermann & Steinbüchel, 2006). In prokaryotes, polyhydroxyalkanoates (PHAs) are the most common type of such storage compounds (Steinbüchel & Valentin, 1995). As the most prominent example, Ralstonia eutropha H16 is able to accumulate poly(3-hydroxybutyrate) (PHB) at up to 90 % of its cell dry weight (Steinbüchel, 1991). In contrast, inclusions composed of triacylglycerols (TAGs) or wax esters (WEs) occur less frequently in prokaryotes. The latter seem to be restricted to nocardioform actinomycetes, streptomycetes and some Gram-negative strains (Alvarez & Steinbüchel, 2002; Wältermann & Steinbüchel, 2005).
Until recently, the molecular architecture of hydrophobic inclusions was not well understood. They are generally composed of the respective storage lipid or polyester representing the hydrophobic core of the inclusion, and are surrounded by a shell consisting of a monolayer of phospholipids and proteins attached to the surface. This amphiphilic shell separates the core from the cytoplasm, thereby preventing the inclusions from coalescing, or other proteins from binding non-specifically to the inclusions (Wältermann et al., 2005; Jurassek & Marchessault, 2004). However, besides having this in common, large differences have been revealed regarding the protein composition and formation between PHA inclusions on the one hand, and WE or TAG inclusions on the other. Whereas no specific proteins are known to be abundantly associated with bacterial TAG and WE inclusions, PHA inclusions are coated by phasins, which represent a unique class of proteins (Pötter & Steinbüchel, 2005; Wältermann & Steinbüchel, 2005; Steinbüchel et al., 1995). PhaP1, which represents the major phasin on the surface of PHA inclusions in Ra. eutropha H16, plays an important role in the formation and structure of these inclusions, because its presence or absence affects the number and size of the inclusions, and the amount of PHB in the cells (Wieczorek et al., 1995, Pötter et al., 2002, 2004; York et al., 2002). According to the most accepted model, PHA inclusions are formed from soluble PHA synthases that polymerize the 3-hydroxybutyrate (3HB) of 3HB-CoA to PHB, with concomitant release of CoA. Since PHA synthases remain covalently linked to the growing PHB chain, an amphiphilic complex composed of the hydrophilic synthase and the elongating polymer chain is formed (Gerngross et al., 1993). These complexes are thought to aggregate to form micelle-like structures, which enlarge to PHA granules due to subsequent extension of the PHA chains. During granule growth, phasins and phospholipids are thought to migrate into the exposed hydrophobic surface of the polymer core, thereby generating an interphase between the hydrophobic core and the cytoplasm (Stubbe & Tian, 2003). However, how phasin molecules are targeted to the hydrophobic surface of growing PHA granules, and whether this recognition is specific for PHA granules or also occurs with other types of hydrophobic inclusions, is unknown (Pieper-Fürst et al., 1995).
In contrast to the formation of PHA granules, TAG and WE inclusions are formed at the cytoplasmic surface of the plasma membrane by WE synthase/acyl-CoA : diacylglycerol acyltransferase (WS/DGAT). The latter is the key enzyme for biosynthesis of these lipids in bacteria, and is bound to lipid droplets (Kalscheuer & Steinbüchel, 2003; Wältermann et al., 2005). These small droplets coalesce to larger structures which are then released into the cytoplasm, and appear finally as large lipid inclusions (Wältermann et al., 2005).
In this study, we demonstrate the heterologous expression of Ra. eutropha H16 phaP1 in TAG-accumulating cells of Rhodococcus opacus and Mycobacterium smegmatis, the distribution of the phasin protein in the cells, and its binding to intracellular TAG inclusions. In addition, we demonstrate the binding of fusion proteins consisting of PhaP1 and the enhanced green fluorescent protein (eGFP) or -galactosidase to lipid inclusions, and that this binding is mediated by the phasin.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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) were routinely cultivated in Luria–Bertani medium (Sambrook et al., 1989). Cells of Rh. opacus PD630 (DSM 44193; Alvarez et al., 1996) and M. smegmatis mc2155 (ATCC 700084; Snapper et al., 1990) were cultivated in Standard I (Std1) medium (Merck). To promote biosynthesis of TAGs and formation of inclusions, cells were transferred to mineral salt medium (MSM) containing 0.1 g NH4Cl l–1, and cultivated for 24, 48, 72 and 96 h (Schlegel et al., 1961). After 72 h, cells achieved their maximum TAG content, which was routinely assessed by TLC, according to a previously described method (Wältermann et al., 2000), and by microscopic examination. In addition, M. smegmatis was also cultivated in Sauton's medium (SM) (Darzins, 1958). To promote TAG accumulation in SM, the potassium phosphate concentration was reduced to 0.05 g l–1. Carbon was supplied in MSM and SM as sodium gluconate or glucose (10 g l–1) for Rh. opacus or M. smegmatis, respectively. To maintain plasmid pJAM2 and its derivatives, kanamycin was used at a final concentration of 50 µg ml–1, following Sambrook et al. (1989). Induction of the acetamidase (ace) promoter of pJAM2 and its derivatives was routinely achieved by addition of 0.5 % (w/v) acetamide to the respective cultures, if not otherwise mentioned (Triccas et al., 1998). All liquid cultures were incubated in Erlenmeyer flasks equipped with baffles at 37 °C for E. coli or at 30 °C for Rh. opacus and M. smegmatis. Solid media were prepared by the addition of 18 g agar l–1.
Cloning of phaP1 downstream of the ace promoter of pJAM2.
Standard molecular biology protocols were used (Sambrook et al., 1989). All PCR products were first cloned into a TA vector (pGEM-T Easy; Promega). Ligation products were first checked by DNA sequencing, and then released by digestion with appropriate restriction enzymes before they were cloned into the expression vector pJAM2, which represents an E. coli–Mycobacterium/Rhodococcus shuttle vector containing the 1.5 kb ace promoter region (Triccas et al., 1998). For subcloning, restriction enzyme recognition sites (as underlined below) were incorporated into the oligonucleotide sequences. The coding region of PhaP1 was amplified without its native start and stop codons (582 bp) by PCR from Ra. eutropha H16 genomic DNA, using the oligonucleotides phaP1-5' (5'-AAAGGATCCATCCTCACCCCGGAACAAGTT-3') and phaP1-3' (5'-AAAGGATCCCGATATGCTTTGCCAACGGAC-3'). Subsequently, the PCR product was cloned downstream of the ace promoter into the BamHI site of pJAM2. In this way, a functional in-frame fusion with the first six codons of the amiE gene was generated, yielding pJAM2 : : phaP1. The phaP1 gene in the constructed fusion lacked its own stop codon, but contained a stop codon after the His6-tag linker sequence of pJAM2. Therefore, the amino acids SRHHHHHH occurred at the C-terminal region of the protein.
Construction of the phaP1–egfp and phaP1–lacZ fusion-expressing plasmids.
A 720 bp fragment representing the complete eGFP gene was amplified, without the start codon, from pEGFP-N3 (BD Bioscience Clontech) using PCR primers egfp-5' (5'-AAATCTAGAGTGAGCAAGGGCGAGGAGCTG-3') and egfp-3' (5'-AAATCTAGATTACTTGTACAGCTCGTCCATG-3'), harbouring the native stop codon (bold type). The PCR product was then cloned downstream of the ace promoter and phaP1 into the XbaI site of pJAM2 : : phaP1, yielding pJAM2 : : phaP1–egfp. To investigate the expression and distribution of unfused eGFP in control experiments, the phaP1 portion of pJAM2 : : phaP1–egfp was released from the expression plasmid by BamHI restriction and religation, yielding pJAM2 : : egfp. For construction of pJAM2 : : phaP1–lacZ, the 3075 bp coding region of lacZ was amplified from genomic DNA of E. coli S17-1 without its native start codon, using PCR primers lacZ-5' (5'-AAATCTAGAACCATGATTACGGATTCACTGG-3') and lacZ-3' (5'-AAATCTAGATTATTTTTGACACCAGACCAACTG-3') harbouring the native stop codon (bold type). The PCR product was then cloned downstream of the ace promoter and phaP1 into the XbaI site of pJAM2 : : phaP1.
Preparations of the electrocompetent cells and transfer of the constructed plasmids to M. smegmatis and Rh. opacus were carried out according to Snapper et al. (1990) and Kalscheuer et al. (1999), respectively, by electroporation in a model 2550 electroporator (Eppendorf–Netheler–Hinz).
Preparation of crude cell extracts, soluble fractions, membrane fractions and TAG inclusions.
Cells of Rh. opacus and M. smegmatis were grown in MSM with reduced ammonium concentration, as described above, harvested by centrifugation (20 min, 6000 g, 4 °C), and resuspended in two volumes of 0.1 M sodium phosphate buffer (pH 7.5). After threefold passage through a French pressure cell (1000 MPa), crude extracts were obtained. To obtain soluble fractions, cell debris was removed from crude extracts by centrifugation for 30 min at 16 000 g at 4 °C, followed by 90 min at 100 000 g at 4 °C in a Sorvall Discovery 90SE ultracentrifuge. Membrane fragments were pelleted by the 100 000 g ultracentrifugation step, and subsequently resuspended in 0.1 mM sodium phosphate buffer (pH 7.5), after washing in the same buffer. TAG inclusions were prepared by loading 1–2 ml crude extracts onto the top of a discontinuous glycerol gradient consisting of 3 ml each of 22, 44 and 88 % (v/v) glycerol in 0.1 M sodium phosphate buffer (pH 7.5). The gradient was centrifuged for 1 h at 170 000 g at 4 °C. The TAG inclusions were withdrawn and subsequently washed twice in 0.1 M sodium phosphate buffer (pH 7.5), and used for further analyses.
SDS-PAGE and immunoblot analysis.
Samples from crude cell extracts were analysed for their protein content according to the method of Bradford (1976), using BSA as a standard. Samples of crude extracts representing a quantity of 50 µg protein, and samples of soluble fractions, membrane fragments and TAG inclusions obtained from corresponding amounts of the extracts, were separated on SDS-polyacrylamide gels. Proteins in the gels were visualized using Coomassie brilliant blue R250 according to the method of Weber & Osbourne (1969). For immunological detection of eGFP, and of PhaP1 and its fusions, the proteins in the gels were transferred onto a PVDF membrane, according to the method of Towbin et al. (1979). Proteins on the membrane were stained with Ponceau S and analysed immunologically employing 0.5 µg polyclonal rabbit anti-PhaP1 IgGs ml–1 (Wieczoreck et al., 1995) or 0.5 µg polyclonal rabbit anti-eGFP IgGs ml–1 (BD Bioscience). IgGs were visualized on immunoblots using goat anti-rabbit IgG–alkaline phosphatase conjugates (diluted 1 : 10 000 in 10 mM Tris/HCl, pH 7.5), converting 5-bromo-4-chloro-3-indolyl-phosphate dipotassium/nitrotetrazolium blue chloride (Sigma) into an insoluble dark product.
Preparation of light microscopy samples.
Nile Red-labelled cells and isolated TAG inclusions were prepared by incubating samples for 30 min at 4 °C in 0.1 M sodium phosphate buffer (pH 7.5) containing 0.5 µg Nile Red ml–1 (stock solution 0.5 mg ml–1 in DMSO). After labelling, cells and inclusions were sedimented by centrifugation at 16 000 g at 4 °C, and resuspended in 0.1 M sodium phosphate buffer (pH 7.5). Cells and TAG inclusions were attached to glass slides via electrostatic interaction, employing slides that were positively charged through adsorption of poly(
-L-lysine) (PL) hydrobromide. In order to coat a glass surface with PL hydrobromide, cleaned glass slides were rinsed thoroughly with tap water, dipped in methanol, and rinsed again with demineralized water. Afterwards, a drop of 0.01 % (w/v) PL hydrobromide solution was added. After air-drying, slides were rinsed with demineralized water, and a drop of a cell suspension or TAG inclusions was added. After 15 min, the coated slides were rinsed with demineralized water to remove loosely attached bacteria or TAG inclusions, and analysed by fluorescence microscopy.
Microscopy and image processing.
Slides were examined on a Zeiss Axio Imager M1 upright wide field fluorescence microscope fitted with a x100/1.4 NA oil-immersion Plan-Apochromat objective lens, and x4 or x2.5 auxiliary tube lenses in phase contrast (PH) or differential interference contrast (DIC) mode. Images were collected by using a Peltier-cooled AxioCam MRm 16 bit digital monochrome charge-coupled device (CCD) camera. The two-thirds of an inch-sized CCD chip consisted of 1388 (horizontal)x1040 (vertical) pixels, each 6.45x6.45 nm in size. Nile Red and eGFP fluorescence were excited using a Zeiss HBO 103 W/2 high-pressure mercury arc lamp. Recording of single and multichannel fluorescence images was performed by using emission bandpass filters at EX/EM 470±40/525±50 nm for eGFP and EX/EM 550±25/605±70 nm for Nile Red. Image stacks consisting of 45–96 planes of optical sections covering the entire z axis were generated by collecting images at focal positions differing in increments of 0.275 µm, by employing a high-precision motorized xyz stage. Depending on samples and fluorescence channels, the exposure times varied between 50 and 1000 ms to obtain sufficiently saturated images suitable for deconvolution. To reduce photobleaching, illumination was controlled by a Zeiss high-speed shutter device. Care was taken to avoid exposing the field to be recorded to the fluorescence light source until recording had begun and the camera had been adjusted to provide the optimum image. Images were stored in zvi data format for subsequent image data processing. All images were acquired using the Zeiss Axiovision 4.5 software. Where indicated, constrained iterative deconvolution of acquired images was performed using the Zeiss AxioVision 3D deconvolution module. All image processing was performed on a Siemens 2.8 GHz Line Celsius R630 workstation.
Cryosectioning and immunogold labelling.
For cryosectioning, cell suspensions of Rh. opacus were prefixed for 5 min by adding an equal volume of 4 % (w/v) paraformaldehyde in PBS (pH 7.4). Cells were washed briefly in the same buffer, and fixed further in 4 % (w/v) paraformaldehyde for 1 h, followed by 1 h incubation in 4 % (w/v) paraformaldehyde with 0.9 M sucrose and 90 % (w/v) polyvinylpyrrolidone 25 buffered with 50 mM sodium carbonate (pH 7.0) as a cryoprotectant. The cells were concentrated by centrifugation at 16 000 g for 20 min at 4 °C, placed on pins in a small volume of cryoprotectant, and frozen in liquid nitrogen. Ultrathin sections were performed as described by Tokuyasu (1980). For immunogold labelling of cryosections, 5 µg polyclonal rabbit anti-PhaP1 IgGs ml–1 (Wieczoreck et al., 1995) was used, followed by gold-conjugated (12 nm gold) goat anti-rabbit IgGs (Jackson Immunoresearch), diluted 1 : 20 from the commercially obtained stock solution with 10 mM Tris/HCl (pH 7.5). Each set of three samples of the replicas and control specimens, which were prepared without the first antibody, were analysed by electron microscopy.
Determination of -galactosidase activity on isolated TAG inclusions.
TAG inclusions isolated from cells of Rh. opacus grown for 72 h in ammonium-reduced MSM harbouring pJAM2 : : phaP1–lacZ or pJAM2 : : phaP1 as a control were prepared as described above. Inclusions (10 mg wet weight) were suspended in 100 µl 0.1 M sodium phosphate buffer (pH 7.5), followed by addition of 650 µl reaction solution consisting of 17 ml 0.1 M sodium phosphate buffer (pH 7.5), 3 ml ONPG solution (8 %, w/v), 1 mM magnesium chloride, 45 mM -mercaptoethanol and 4 µl SDS solution (20 %, w/v). The assay mixture was incubated for 30 min at 37 °C. To stop the reaction, 400 µl 1 M disodium carbonate were added. Subsequently, TAG inclusions were eliminated from the assay by filtration through a 0.22 µm syringe filter, and the absorbance of the filtrate was examined at 405 nm to analyse the amount of cleaved ONPG. For calculation of enzyme activities an 
405 of 4.6 mM–1 cm–1 was used for the product o-nitrophenol (ONP). Measured -galactosidase activity was essentially associated with TAG inclusions, since further cleavage of ONPG did not occur in assays after the inclusions were removed. -Galactosidase activity assays were done in triplicate in three separate experiments.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). The intracellular stability of PhaP1 in recombinant M. smegmatis was also demonstrated by extending the expression time to 96 h (Fig. 1B).
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). This might have been due to cleavage of the inducer by the chromosomally encoded acetamidase, thus providing the cells with sufficient ammonium for growth. Attempts to achieve a sufficient accumulation of TAG inclusions under phosphate limitation in SM, as described in Methods, failed due to poor growth and little lipid accumulation (data not shown).
To circumvent this obstacle, all constructed plasmids were subsequently introduced into Rh. opacus. In contrast to M. smegmatis, SDS-PAGE electropherograms of crude extracts of the corresponding recombinant strains of Rh. opacus revealed no additional visible protein bands, in comparison to those obtained from the wild-type, when grown for 72 h in ammonium-reduced MSM, even when the cells were induced with 0.5 % (w/v) acetamide, indicating that expression of genes controlled by the M. smegmatis ace promoter was significantly lower in Rh. opacus. However, according to the results obtained in recombinant M. smegmatis, the anti-PhaP1 IgGs recognized a 25 kDa protein in Western blots obtained from crude extracts of cells harbouring pJAM2 : : phaP1, although immunological recognition of the phasin was significantly weaker than that in recombinant M. smegmatis. As in M. smegmatis, no degradation products of the phasin were detected in Rh. opacus. Similarly, eGFP and the PhaP1–eGFP fusion protein were easily recognized on Western blots of crude cell extracts harbouring pJAM2 : : egfp and pJAM2 : : phaP1-egfp, respectively, as was demonstrated by employing the anti-eGFP IgGs (Fig. 2). In contrast to M. smegmatis, addition of 0.5 % (w/v) acetamide to the cultures did not affect TAG accumulation in Rh. opacus (not shown). To investigate the cellular distribution of PhaP1, eGFP and the PhaP1–eGFP fusion protein in Rh. opacus, crude extracts of induced cells were fractionated into soluble fractions, membrane fractions and fractions representing the TAG inclusions. On Western blots of the respective fractions of Rh. opacus pJAM2 : : phaP1, the phasin was recognized by the anti-PhaP1 IgGs in the fraction representing the TAG inclusions, whereas no signal occurred in the soluble fraction. This indicates that PhaP1 was associated with the TAG inclusions in recombinant Rh. opacus. The result obtained for the distribution of PhaP1 in cell fractions of Rh. opacus was also confirmed by the localization of the PhaP1–eGFP fusion protein by employing anti-eGFP IgGs on Western blots of the strain harbouring pJAM2 : : phaP1-egfp. In this recombinant strain, the fusion protein also occurred only in the fraction representing the TAG inclusions. We also tried to localize PhaP1 and its eGFP fusion in electropherograms of total membrane fractions of the recombinant strains, but failed (data not shown). As expected, the unfused eGFP was only localized in the soluble fraction of the control strain harbouring pJAM2 : : egfp (Fig. 2).
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). As revealed by constrained iterative deconvolution of images obtained from Std1-grown cells, fluorescence also occurred to some extent in areas of the cell wall and plasma membrane. However, fluorescence at these sites was much weaker than that of intracellular TAG inclusions (see deconvolved image in Fig. 3A). After 72 h in ammonium-reduced MSM, cells were fully packed with brightly fluorescent TAG inclusions. In fact, after deconvolution, large TAG inclusions in these cells often exhibited a ring of fluorescence, indicating a localization of the fusion protein at the surface of the inclusions (Fig. 3D). Fluorescence of the fusion protein was thoroughly distuinguishable from Nile Red fluorescence at all stages of TAG accumulation, which, in addition to the TAG inclusions, also clearly labelled the cellular envelope (Fig. 3A–D).
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). In contrast to this, TAG inclusions from cells expressing unfused eGFP exhibited no fluorescence when observed without Nile Red labelling (not shown). Cells expressing unfused egfp, which served as a negative control, exhibited a diffuse green fluorescence throughout the cytoplasm, whereas intracellular TAG inclusions were easily detectable by their Nile Red fluorescence (Fig. 3F).
Immunogold labelling of cryosections
To investigate whether PhaP1 is targeted exclusively to the surface of TAG inclusions in Rh. opacus, or also to other components of the cells, the fusion protein was localized on cryosections by postembedding immunogold labelling. Ultrathin cryosections were prepared from recombinant cells harbouring pJAM2 : : phaP1-egfp grown under ammonium-reduced conditions for 72 h. For immunogold labelling of cryosections, rabbit anti-PhaP1 IgGs were used in combination with goat anti-rabbit IgG gold conjugates. In cryosectioned Rh. opacus cells, TAG inclusions appeared as nearly spherical, electron-translucent areas with little internal structure. Strong labelling of PhaP1 was found at the surface of the inclusions, whereas almost no label was observed in the cytoplasm. Label was also detected at the plasma membrane, thus confirming the fluorescence-microscopic localization of the PhaP1–eGFP fusion protein at this site in the cells. However, although immunogold labelling of sections is not a suitable method for the quantification of proteins, the concentration of PhaP1 label at the periphery of the cells was lower than that at the surface of the TAG inclusions (Fig. 4A). Control specimens, prepared without the primary antibody, were essentially free from gold particles (Fig. 4B).
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; Wältermann & Steinbüchel, 2005). The specific -galactosidase activity [9.5±0.5 U (mg wet weight)–1] was significantly higher in samples containing TAG inclusions which were isolated from Rh. opacus PD630 harbouring pJAM2 : : phaP1-lacZ. When TAG inclusions were removed from the assay, specific -galactosidase activity was only 0.3±0.3 U (mg wet weight)–1. For control experiments, cells harbouring pJAM2 : : phaP1 were analysed in the same way. Since Rh. opacus also expresses a chromosomally encoded -galactosidase, low enzyme activity was also expected to occur in the control samples. In the controls, i.e. in samples containing TAG inclusions, which were isolated from Rh. opacus harbouring pJAM2 : : phaP1, specific -galactosidase activity was only 1.8±0.8 or 0.3±0.3 U (mg wet weight)–1 before and after removal of the TAG inclusions, respectively. The data represent the mean±SD of three independent experiments. These data demonstrate a stable immobilization of LacZ to bacterial TAG inclusions mediated by PhaP1 as an anchor. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). The motif in PhaP1 which targets the phasin to PHB granules in Ra. eutropha H16 has not yet been identified. However, PHB granules, as well as TAG inclusions, possess a hydrophobic core of the polyester or lipid, respectively, which is thought to be surrounded by a monolayer of phospholipids (de Koning & Maxwell, 1993; Hocking & Marchessault, 1994; Mayer & Hoppert, 1997; Wältermann et al., 2005). This common structure allows the targeting of PhaP1 to PHB granules and also TAG inclusions, as demonstrated in this study. Therefore, our data indicate that targeting of PhaP1 to PHB granules in Ra. eutropha H16 is most probably not mediated by a direct mutual recognition of the phasin and the polymer in the granules. Our results indicate that PhaP1 has the ability to bind to any type of hydrophobic inclusion, irrespective of whether a PHA or a different hydrophobic compound is present in the core of the inclusions. Furthermore, it is also unlikely that an additional as-yet-unidentified component involved in PHA metabolism mediates targeting of PhaP1 to the inclusions, since such components would have been absent in the strains used in our study. Most probably, binding of PhaP1 to the inclusions is mediated only by the presence of the amphiphilic interphase consisting of the monolayer membrane between the inclusions and the surrounding cytoplasm, by the hydrophobic surface of the core, or by a combination of both.
Combined electron microscopy and postembedding immunocytochemistry revealed that PhaP1 is distributed mostly on the amphiphilic surface of the TAG inclusions. However, in contrast to its exclusive distribution on the surface of PHB granules in Ra. eutropha H16, it was demonstrated that some PhaP1 was also present at the plasma membrane and cell wall regions in Rh. opacus. This distribution has also been reported by Pieper-Fürst et al. (1994) while investigating the cellular distribution of the 14 kDa phasin in Rhodococcus ruber, which is able to synthesize equal amounts of TAGs and the copolymer poly(3-hydroxybutyrate-co-3-hydroxyvalerate) [poly(3HB-co-3HV)]. Although, in R. ruber, it is not known whether TAGs and poly(3HB-co-3HV) occur separately or simultaneouly in the inclusions, it has been demonstrated that the phasin occurs on the surface of any inclusion in the cells, and also at the cytoplasmic surface of the plasma membrane. According to our recently proposed model, TAGs are synthesized by WS/DGAT localized at the cytoplasmic surface of the plasma membrane as small lipid droplets, which coalesce to form larger inclusions by prolonged TAG synthesis, and are then released into the cytoplasm (Wältermann et al., 2005). Thus, a binding of phasins to nascent TAG inclusions at their site of synthesis is the most probable explanation for this distribution, rather than a direct interaction of the phasin with the cytoplasmic membrane. This was also confirmed by our unsuccessful experiments, which aimed at localizing the phasin in the total membrane fractions of Rh. opacus. Most probably, this was caused by the disintegration of the plasma membrane–lipid droplet association during cell disruption.
In Ra. eutropha H16, the amount of PHB and the number of granules is directly influenced by the amount of phasin molecules in the cells (Wieczorek et al., 1995; Pötter et al., 2002). The presence of the phasin neither altered the amount of TAGs in Rh. opacus, nor influenced the size or number of the TAG inclusions. As revealed by our expression analysis, the total amount of PhaP1 in Rh. opacus was very low, since expression of the protein was limited by the ace promoter of pJAM2. Whether the presence of a high amount of phasins can influence TAG metabolism in the cells remains to be elucidated. This may be clarified by inactivation of the chromosomally encoded acetamidase in M. smegmatis, through the use of an alternative gene expression system that does not influence cellular TAG metabolism. However, a more suitable expression system or an acetamidase-negative M. smegmatis strain are as yet unavailable.
We demonstrated that TAG inclusions tagged with a PhaP1–LacZ fusion protein exhibited -galactosidase acitivity in vitro. The immobilization of enzymes and other surface proteins or defined particles offers interesting applications. One example could be the synthesis of functionalized nanoparticles, e.g. to carry antibodies for analytic purposes, or hormones and other therapeutic agents. Such nanoparticles can be purified easily from crude cell extracts. Moldes et al. (2004) have created a system for the synthesis and purification of enzymes using PHA granules as a matrix, and the N terminus of the phasin PhaF from Pseudomonas putida as a linker. Furthermore, PHB granules in recombinant E. coli have been successfully demonstrated as a matrix for the purification of target proteins by fusions with phasins and self-cleaving affinity tags based on protein-splicing elements known as inteins (Banki et al., 2005). Also, TAG inclusions have been utilized as a matrix for purification of enzymes (Moloney, 1998, 2002). The latter author created an analogous system based on plant cells, by attaching target enzymes to oil bodies via oleosins. Both purification systems described above are patented and commercially available (Prieto et al., 2004; Moloney, 1997; Moloney et al., 2005). In addition, anchoring of enzymes and other proteins to TAG inclusions by a PhaP1 tag offers an interesting possibility to establish alternative, bioengineered pathways on the monolayer surface of intracellular TAG inclusions.
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ACKNOWLEDGEMENTS |
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Received 3 March 2006;
revised 25 July 2006;
accepted 25 July 2006.
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