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A new heat-shock gene, ppiD, encodes a peptidyl–prolyl isomerase required for folding of outer membrane proteins in Escherichia coli

1998; Springer Nature; Volume: 17; Issue: 14 Linguagem: Inglês

10.1093/emboj/17.14.3968

1460-2075

Claire Dartigalongue, Satish Raina,

Peptidase Inhibition and Analysis

Article15 July 1998free access A new heat-shock gene, ppiD, encodes a peptidyl–prolyl isomerase required for folding of outer membrane proteins in Escherichia coli Claire Dartigalongue Claire Dartigalongue Département de Biochimie Médicale, Centre Médical Universitaire, 1 rue Michel-Servet, 1211 Genève 4, Switzerland Search for more papers by this author Satish Raina Corresponding Author Satish Raina Département de Biochimie Médicale, Centre Médical Universitaire, 1 rue Michel-Servet, 1211 Genève 4, Switzerland Search for more papers by this author Claire Dartigalongue Claire Dartigalongue Département de Biochimie Médicale, Centre Médical Universitaire, 1 rue Michel-Servet, 1211 Genève 4, Switzerland Search for more papers by this author Satish Raina Corresponding Author Satish Raina Département de Biochimie Médicale, Centre Médical Universitaire, 1 rue Michel-Servet, 1211 Genève 4, Switzerland Search for more papers by this author Author Information Claire Dartigalongue1 and Satish Raina 1 1Département de Biochimie Médicale, Centre Médical Universitaire, 1 rue Michel-Servet, 1211 Genève 4, Switzerland *Corresponding author. E-mail: [email protected] The EMBO Journal (1998)17:3968-3980https://doi.org/10.1093/emboj/17.14.3968 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info We have identified a new folding catalyst, PpiD, in the periplasm of Escherichia coli. The gene encoding PpiD was isolated as a multicopy suppressor of surA, a mutation which severely impairs the folding of outer membrane proteins (OMPs). The ppiD gene was also identified based on its ability to be transcribed by the two-component system CpxR–CpxA. PpiD was purified to homogeneity and shown to have peptidyl–prolyl isomerase (PPIase) activity in vitro. The protein is anchored to the inner membrane via a single transmembrane segment, and its catalytic domain faces the periplasm. In addition, we have identified by site-directed mutagenesis some of the residues essential for its PPIase activity. A null mutation in ppiD leads to an overall reduction in the level and folding of OMPs and to the induction of the periplasmic stress response. The combination of ppiD and surA null mutations is lethal. This is the first time two periplasmic folding catalysts have been shown to be essential. Another unique aspect of PpiD is that its gene is regulated by both the Cpx two-component system and the σ32 heat shock factor, known to regulate the expression of cytoplasmic chaperones. Introduction Bacteria have evolved many adaptive systems in order to cope with various environmental stresses. Among these, heat shock produces a highly conserved response which in Escherichia coli is regulated by two alternative sigma factors, σ32 and σE. These two factors govern the transcription of two heat-shock regulons which, respectively, are specialized in coping with protein misfolding in the cytoplasm and the extra-cytoplasm, i.e. periplasm and outer membrane. Protein misfolding in the extra-cytoplasm induces an additional stress response (other than the σE-dependent stress response), which is regulated by the two component system CpxR–CpxA. The periplasmic protease HtrA, involved in the degradation of misfolded polypeptides, is encoded by a gene which is transcribed both by the EσE polymerase and the CpxR activator (Danese et al., 1995; Raina et al., 1995). However, fkpA which encodes a peptidyl–prolyl isomerase of the FKBP family in the periplasm (Missiakas et al., 1996), belongs to the σE regulon (Danese and Silhavy, 1997), whereas dsbA and ppiA, which encode periplasmic thiol:disulfide oxido-reductase and peptidyl–prolyl isomerase respectively, are under the control of the Cpx system (Danese and Silhavy, 1997; Pogliano et al., 1997). Unlike the cytoplasm which contains many ATP-dependent chaperones with wide substrate specificity, the periplasm appears to contain two defined types of folding catalysts. Folding of most translocated proteins encounters two types of rate-limiting steps, which are overcome by two classes of catalysts: protein disulfide isomerase (PDI) and peptidyl–prolyl cis–trans isomerase (PPIase). In E.coli, there are at least six known Dsb proteins (reviewed by Missiakas and Raina, 1997a). These proteins are involved in the oxidation of disulfide bonds or the rearrangement of wrongly paired disulfides (Bardwell et al., 1991; Missiakas et al., 1994; Zapun et al., 1995; Rietsch et al., 1996). PPIases are found in both the cytoplasm and the periplasm. They catalyse the rapid interconversion between the cis and trans forms of the peptide bond Xaa–Pro. Three distinct families of PPIases have been identified so far; these include the cyclophilins (PpiA in the periplasm), FKBP-like proteins (FkpA in the periplasm) and the newly discovered parvulin family. The first member of the parvulin family was identified as a cytosolic E.coli protein (Rahfeld et al., 1994), whose gene was later designated ppiC. Interestingly, the yeast PpiC homologue, ESS1, is the only essential PPIase for yeast. Surprisingly, yeast mutants lacking all the other 12 PPIases are viable, although some of them are heat-shock inducible (Dolinski et al., 1997). In E.coli, a second member of the parvulin family, known as SurA, was identified in the periplasm. SurA is involved in the correct folding of outer membrane protein (OMP) monomers. We isolated this gene as a multicopy suppressor of htrM (Missiakas et al., 1996). htrM mutants synthesize altered lipopolysaccharide (LPS), and as a consequence lack most of the OMPs in their outer membrane. This suppression effect is due specifically to correction of the proper outer membrane profile. Also, the accumulation of misfolded OMPs was shown to induce the σE-dependent response to protein misfolding in the extra-cytoplasm. Overexpression of surA was shown to dampen this response in many cases where misfolded polypeptides accumulate, suggesting a general chaperone-like function for SurA. We and others have shown that mutations in surA confer severe defects in the amounts or maturation of OMPs (Lazar and Kolter, 1996; Missiakas et al., 1996; Rouvière and Gross, 1996). Bacteria lacking surA turn on the σE stress response constitutively (Missiakas et al., 1996; Rouvière and Gross, 1996). Finally, surA mutants are extremely sensitive to detergents and hydrophobic drugs such as novobiocin, phenotypes reminiscent to the leakiness of the outer membrane. In the present study, we took advantage of the hypersensitivity of surA mutants to novobiocin and selected for multicopy suppressors. We identified a new gene and based on its sequence homology to members of the parvulin family, we have designated this gene ppiD. ppiD was also identified in an independent genetic screen aimed at identifying new members of the CpxR–CpxA regulon. We show here that ppiD encodes a membrane-anchored polypeptide of 623 amino acids, of which the last C-terminal 589 residues are located in the periplasm. Transcriptional analysis of ppiD revealed that the gene belongs to two stress regulons: the CpxR–CpxA regulon and the σ32 regulon. This is the first member of a periplasmic folding catalyst to be regulated by the classical heat-shock sigma factor σ32. Results ppiD is a multicopy suppressor of surA We have shown previously that a null mutation in surA leads to a highly pleiotropic phenotype. One of the defects is hypersensitivity to antibiotics such as novobiocin, due to a higher permeability of the outer membrane. surA mutant bacteria do not form colonies in the presence of novobiocin concentrations >10 μg/ml, while isogenic wild-type bacteria grow in the presence of 40 μg/ml of novobiocin (Missiakas et al., 1996). We constructed a chromosomal DNA library lacking surA and used it to transform the surA null mutant. Candidates resistant to 30 μg/ml of novobiocin were selected. Plasmid DNA from seven such candidates was isolated and shown to breed true by retransforming a surA null mutant. The DNA was used to probe the ordered E.coli DNA library (Kohara et al., 1987). All seven plasmids hybridized to bacteriophage λ 148(3B6) and λ 149(7E2) clones. This corresponds to a genetic map position in the 9.5–10 minute region on the E.coli chromosome. This area includes the known serine protease, Lon, and the DNA-binding protein, HupB. DNA from one such plasmid, pSR3239, was used to construct a minimal subclone 2.4 kbp BsaBI–SmaI DNA fragment in the p15A-based vector, pOK12. This subclone, pCD51, was found to be sufficient to restore novobiocin resistance to the surA null mutant (Figure 1; Table I). Since our initial clone pSR3239 also carries the intact hupB gene, we subcloned the 1 kbp EcoNI–KpnI DNA fragment which carries the intact hupB gene (pCD56), and found that it does not complement a surA null mutant (Figure 1). Sequence analyses of pCD51 revealed the presence of a single complete open reading frame (ORF) which encodes a polypeptide of 623 amino acids. Sequence examination also revealed homology to PPIase. Hence, this gene was designated ppiD. It carries a putative parvulin-like domain located between amino acids 227 and 357. This predicted amino acid sequence shares 34% identity with an ORF of unknown function from Haemophilus influenza (SwissProt accession No. P44092). The next closest homologue is a very recently sequenced ORF of unknown function from Acinetobacter (PID entry e1173385), with a sequence identity of 29% over its entire length. PpiD also has sequence similarity to the SurA protein. More precisely, residues 86–121 of PpiD align with residues 65–100 of SurA (44% identity). A second region lying between residues 334 and 357 of PpiD seems to correspond to the parvulin-like domain of SurA (residues 251–274). It is known that SurA has two very similar parvulin-like domains at its C-terminus. These residues include the motif VGFHIL and are highly conserved among the members of the PpiC (parvulin) family. However, it should be pointed out that the highly conserved residues H and L are replaced by L and V, respectively, in the PpiD amino acid sequence. Also, the predicted amino acid sequence of PpiD shows the presence of a single parvulin-like domain, whereas SurA carries two such domains. Figure 1.Restriction map of the ppiD gene and surrounding DNA sequences. The different plasmids constructed with either wild-type or the disrupted ppiD gene are also shown. Columns on the right indicate the ability of the different clones to confer novobiocin resistance to the surA mutant. Download figure Download PowerPoint Table 1. Complementation of surA and comparative resistance to various agents reflecting membrane defects of the ppiD mutant as compared with other folding agents LBA Nov SDS 10 15 20 0.2% 1% 2% MC4100 109 109 109 109 109 109 109 MC4100 ppiD::ΩKan 8×107 7×105 5×103 0 8×106 5×03 0 MC4100 skp− 2×109 7×108 6×105 3×105 109 7×107 6×105 MC4100 ppiD::ΩKan skp− 5×108 5×107 2×102 0 3×108 2×102 0 htrM− 2×107 0 0 0 0 0 0 htrM− (pCD275 ppiD+) 3×109 5×108 5×107 2×106 5×109 2×108 2×109 surA::ΩKan 3×108 102 0 0 5×103 4×103 0 surA::ΩKan (pCD275ppiD+) 109 9×108 6×106 5×105 8×107 8×107 5×105 surA::ΩKan (pCD273 A350) 5×108 102 0 0 9×103 5×103 0 Note that htrM mutants do not form colonies on MacConkey agar but htrM::Tn5 transformed with ppiD+ (pCD275) fully restored the growth on MacConkey agar plates, particularly at 37°C, and suppressed the mucoid phenotype. Identification of ppiD as a new member of the CpxR–CpxA regulon In our effort to understand the nature of the Cpx-dependent stress response, we have constructed a library of transcriptional fusions using the single copy promoter probe vector pFZY (see Materials and methods). We have shown previously that overexpression of the prpA gene leads to the constitutive induction of the Cpx pathway (Missiakas and Raina, 1997b). Hence, we looked for promoter fusions, which are highly induced (deeper blue colonies) on X-gal-containing plates, in the presence of a plasmid carrying the prpA gene under control of the inducible lac promoter (pDM1574). We isolated ∼70 such clones and transduced a genetically well characterized cpxR null allele (Danese et al., 1995) into them. Only those clones which showed a decline in β-galactosidase activity were retained (Figure 2). Fifteen such isolates were retained and plasmid DNA prepared from them was sequenced. Among them, 12 clones were found to contain sequences from the promoter region of the ppiD gene. Figure 2.The transcription of the ppiD gene is under the control of the CpxR–CpxA two-component system. The β-galactosidase activity was measured in the wild-type ppiD–lacZ fusion or when transduced with a null mutation of cpxR. Induction of htrA transcription serves as an internal control. The shaded bars show the β-galactosidase activity from the ppiD–lacZ fusion deleted for the region containing the three potential CpxR-binding boxes. Download figure Download PowerPoint ppiD is also a bona fide heat shock gene under the control of σ32 To analyse the transcriptional activity and promoter usage of ppiD, we constructed a single copy ppiD–lacZ fusion using the promoter region of ppiD and the λRS45 and pRS550 vectors (Simons et al., 1987). Using this fusion, we observed that ppiD transcription is heat-shock inducible (Figure 3), which is reminiscent of classical heat-shock gene induction. The single copy ppiD–lacZ-carrying strain (CD212) was transformed with a vector carrying either the rpoH gene under the control of an inducible promoter (pSR1332; Missiakas et al., 1993a) or the rpoE gene (pSR1628; Raina et al., 1995). As shown in Figure 3, induction of the rpoH gene product from pSR1332 produces a 10-fold increase in β-galactosidase activity using the ppiD–lacZ fusion, whereas no induction is observed with the rpoE+ plasmid (pSR1628). Furthermore, no reduction is observed in the presence of the rseA+-carrying plasmid (pSR2661; Missiakas et al., 1997) which encodes the specific anti-σE factor, which in high dosage represses the transcription from σE-dependent promoters (Table II). Figure 3.The ppiD is also a heat-shock gene under the control of σ32. Cultures of isogenic bacteria carrying single copy fusions to either the promoter of ppiD or the lon gene were tested for β-galactosidase activity at different temperatures. These fusions were transformed with rpoH+ plasmid with the inducible lac promoter (pSR1332) grown in minimal medium supplemented with 0.2% glucose up to an OD of 0.2; aliquots were treated with or without the lac inducer IPTG (1 mM) for 20 min. The β-galactosidase activity was measured as described above. The last lane represents, as a further control, the β-galactosidase activity from a ppiD–lacZ fusion in a dnaK103 background. Each sample was assayed for β-galactosidase activity four times, and the data presented are the average of three independent experiments. Download figure Download PowerPoint Table 2. ppiD transcription is not affected by overexpression of either rpoE or its specific anti-sigma factor rseA β-galactosidase activity (Miller units) ppiD–lacZ 515 ± 25 ppiD–lacZ (prpoE+) 595 ± 33 ppiD–lacZ (prseA+) 505 ± 23 htrA–lacZ 123 ± 10 htrA–lacZ (prpoE+) 945 ± 72 htrA–lacZ (prseA+) 23 ± 7 It is known that mutation in dnaK, dnaJ, grpE or htrC leads to a constitutive elevated induction of the σ32 heat-shock response. The ppiD–lacZ-carrying strain was transduced into the dnaK103 mutant. As can be seen in Figure 3, the β-galactosidase expression is highly induced. Taken together, the data clearly show that ppiD has another promoter in addition to its CpxR–CpxA-dependent promoter, and that this promoter is under σ32 control. This is the first example so far of a periplasmic folding catalyst placed under the control of σ32. Identification of the heat-shock protein PpiD on two-dimensional gels The PpiD protein has not up to now been identified as a heat-shock protein on two-dimensional gel isoelectric focusing gels. The reasons could be that its molecular weight coincides with that of HtpG and DnaK (all are ∼70 kDa in size). In addition, the predicted pI of HtpG is 5.09 and that of PpiD is 4.95. PpiD, being an inner membrane protein, may migrate differently somehow, perhaps overlapping HtpG on these gels. In fact, in many of our earlier two-dimensional heat-shock gels, we did observe a protein induced by a 50°C heat shock which runs close to HtpG and DnaK. Hence, we constructed two isogenic strains (ppiD::ΩTet htpG+ and ppiD::ΩTet ΔhtpG) and compared the global heat-shock profiles of cultures of these strains after a sudden 50°C heat shock for 5 min. Samples were analysed by two-dimensional gel electrophoresis. As can be seen in Figure 4, there is a clear induction of an ∼70 kDa polypeptide in the ppiD+ htpG+ sample (Figure 4A) but not in that of ppiD::ΩTet ΔhtpG (Figure 4C). As a control for the spot identified as PpiD, we also added [35S]methionine-labelled PpiD to heat-shock extracts from ppiD::ΩTet ΔhtpG (Figure 4B. This clearly indicates that PpiD is a heat-shock protein and accumulates at a temperature of 50°C. The positions of other heat-shock proteins such as GroEL and DnaK are also shown in Figure 4B. Figure 4.Identification of the PpiD protein as a heat-shock protein by two-dimensional electrophoresis. Cultures of isogenic strains wild-type (A), ppiD::ΩTet htpG− + PpiD (B) and ppiD::ΩTet ΔhtpG (C) were grown at 30°C, shifted to 50°C for 5 min, then labelled for another 5 min with [35S]methionine (50 μCi/ml). The proteins were resolved in the first dimension on 1.6% (pH 5.0–7.0) and 0.4% (pH 3.5–10.0) ampholines. For the second dimension, samples were electrophoresed on a 12.5% SDS gel. Autoradiograms of the relevant portions of dried gels are shown. The arrow marked D point to the position of PpiD, and K and EL indicate the positions of 70 kDa DnaK and 63 kDa GroEL heat-shock proteins. Download figure Download PowerPoint Acetyl phosphate and the cpxA* mutations influence the transcription of ppiD We further examined the CpxR–CpxA transcriptional dependence of the ppiD gene. It is known that the level of acetyl phosphate modulates the activation or repression of two-component systems such as CpxR–CpxA. Thus we determined the β-galactosidase activity of ppiD–lacZ under different growth conditions which affect the level of acetyl phosphate in vivo. It is known that the use of pyruvate as a carbon source results in the maximum level of intracellular acetyl phosphate. Indeed, a 3-fold activation of the ppiD promoter can be obtained when cultures are shifted from glycerol to pyruvate medium (Figure 5). Figure 5.Critical role of the CpxR-binding domain and the levels of acetyl phosphate in the transcription of the ppiD gene. Cultures of isogenic strains CD212, carrying the wild-type full-length ppiD–lacZ fusion, and CD221, carrying ppiD–lacZ fusion with deletion of the three putative CpxR boxes, were used for these experiments. Cultures were grown in regular LB medium up to an OD of 0.5 at 595 nm. Cultures were spun, washed, diluted 1:100 and transferred to minimal medium supplemented with different carbon sources such as glycerol, glucose or pyruvate and allowed to reach an OD of 0.2 at 595 nm. Download figure Download PowerPoint We have also used the SR3570 strain carrying a cpxA* mutation in which T252 is changed to R252 (Missiakas and Raina, 1997b). This chromosomal allele leads to a gain of function of CpxA, presumably a hyperkinase activity (Missiakas and Raina, 1997b). This mutation is similar to a gain-of-function mutation in envZ12 (Aiba et al., 1989). We transduced the ppiD–lacZ fusion into cpxA* mutant SR3570, resulting in CD220. This strain again showed a 2- to 3-fold induction of ppiD–lacZ activity (Table III). In control experiments we transduced a htrA–lacZ fusion into the same cpxA* mutant (Table III). The increased ppiD–lacZ expression is in fact more than that observed for the htrA–lacZ fusion. These results further confirm that the transcription of ppiD is in part regulated by the two-component system, CpxR–CpxA. Table 3. CpxR–CpxA-dependent regulation of the ppiD genea β-galactosidase activity (Miller units) CD212 = MC4100 ppiD–lacZ 510 ± 23 CD215 = CD212cpxA* 1420 ± 72 SR1458 = MC4100 phtrA–lacZ 110 ± 7 CD220= phtrA–lacZ cpxA* 177 ± 12 a The hyperkinase CpxA induces its transcription. All measurements were performed in triplicate and the averages are presented. Mapping of ppiD transcriptional start sites In order to understand the transcriptional regulation of ppiD, we determined the transcriptional initiation site(s) under different growth conditions and in different genetic backgrounds. RNA was extracted from bacterial cultures grown either at 30°C or following a brief shift to 50°C. Using the primer extension technique, the transcriptional start sites were mapped. As can be seen in Figure 6, lane 2, using RNA from those cultures subjected to the 50°C shift, two transcriptional start sites designated as Phs1 and P* could be mapped 75 and 83 nucleotides, respectively, upstream of the translational initiation start site. No corresponding signal is seen from RNA extracted at 30°C, which clearly indicates that transcription from the start site designated as Phs1 is heat-shock inducible. The −10 and −35 regions upstream of this start site show a perfect match with the σ32-regulated promoters (for the −10 box CCCC and for the −35 box CTTGTG, to be compared with the consensus CTTGAA). In addition, the spacing between the two canonical boxes is 15 nucleotides, a feature common to most of the σ32-regulated promoters. We do not know at present which sigma factor is responsible for the transcription initiating at nucleotide 83 (P*). The two additional start sites located at nucleotides 85 and 93 appear only in the RNA extracted at 30°C (lane 3) and not at 50°C. Thus, they are not heat-shock induced and may correspond to σ70-coupled CpxR–CpxA-dependent transcription. Examination of the sequence shows at least three conserved CpxR boxes at positions −261 (GGTAAAGAG), −221 (GGTAAGC) and −209 (GGTAACT) (Figure 7) upstream of the translational initiation codon. Figure 6.Mapping of ppiD transcriptional start sites. Primer extension reactions of total cellular RNA hybridized to a 32P end-labelled oligonucleotide probe, complementary to nucleotides −5 to 17 of the ppiD sense strand. The annealed primer was extended by AMV reverse transcriptase. RNA was extracted from wild-type MC4100 bacteria grown at 30°C (lane 3), or shifted to 50°C for 10 min (lane 2). Lane 1 represents a control annealed RNA plus primer without addition of AMV reverse transcriptase. Lanes labelled G, A, T and C correspond to the dideoxy sequencing reactions carried out using the same oligonucleotide as the primer. Download figure Download PowerPoint Figure 7.Nucleotide sequence of the promoter region of the ppiD gene. The initiation start designated as PHS corresponds to the promoter under σ32 control. The corresponding −10 and −35 regions are underlined. The other initiation site designated P1 corresponds to one of the start sites resulting from primer extension of RNA isolated at 30°C. The three putative CpxR-binding boxes are highlighted in bold and underlined. Download figure Download PowerPoint Deletion of the putative CpxR boxes leads to a decline in ppiD transcription As mentioned above, there are three putative CpxR-binding boxes (Figure 7) similar to what has been described for dsbA (Pogliano et al., 1997). To assay their role in vivo, we removed the first 147 nucleotides from plasmid pCD189 by digestion with BamHI and FokI, resulting in plasmid pCD191 (Table IV). This removes all of the three putative CpxR-binding boxes. This promoter fusion still carries the σ32-regulated promoter Phs1, as well as 43 nucleotides upstream of it. This promoter fusion was analysed and shows an overall decline in promoter strength (Figure 2). It no longer responded to changes in the level of acetyl phosphate (compare ppiD–lacZ and ppiD–lacZ CpxR boxes− in Figure 2). Also, introduction of the cpxR null allele did not change the ppiD promoter activity. Furthermore, this promoter fusion is not induced by overexpression of the prpA gene product, but is still subject to regulation by σ32, like the original fusion construct CD212. This further confirms that ppiD transcription is in part regulated by the two-component system CpxR–CpxA, and that the whole nucleotide region deleted contains the CpxR-binding boxes which are important in its regulation. Table 4. Bacterial strains and plasmids Relevant characteristics Reference or source Strains CA8000 HfrH thi our collection JB23 ΔhtpG Bardwell and Craig (1988) MC4100 F− araD139 Δ(argF-lac) U169 our collection SR21 CA8000 htrM :: Tn5 Raina and Georgopoulos (1991) SR3205 MC4100 surA::ΩKanr Missiakas et al. (1996) SR3570 MC4100 cpxA*(T252R) Missiakas and Raina (1997b) CD63 SR3205 (pCD51, ppiD+) this study CD162 MC4100 ppiD::ΩTet at the KpnI site this study CD212 MC4100 φ(ppiD-lacZ) this study CD213 CD212 transduced into SR1458 this study CD214 CD212 transduced into SR2195 this study CD215 MC4100 φ(ppiD-lacZ) cpxA* this study CD220 SR1458 cpxA* this study CD221 CD212 deleted for the three CpxR boxes this study CD247 MC4100 ppiD::ΩKan at the KpnI site this study CD249 MC4100 ppiD:ΩKan at the PstI site this study CD259 CD247 transduced into SR1421 this study CD261 CD247 transduced into SR1359 this study CD269 SR3205 surA::ΩKanr (pDM154 surA+) this study CD270 CD162 ppiD::ΩTet (pCD275) this study Plasmids pKO3 allelic replacement vector Link et al. (1997) pFZY1 promoter probe vector Koop et al. (1987) pDM1554 pSK surA+ Missiakas et al. (1996) pSR1232 pTTQ rpoH+ (1.3 kbp EcoRV) Missiakas et al. (1993a) pSR1574 pSE420 prpA+ (721 bp NdeI–BamHI) Missiakas and Raina (1997b) pSR1628 pOK12 rpoE+ (1.4 kbp Sau3A) Raina et al. (1995) pSR1865 pOK12 dsbA+ (1.8 kpb Sau3A) Missiakas et al. (1993b) pSR2661 pKO12 rseA+ (1210 bp HindIII–MscI) Missiakas et al. (1997) pSR3239 pOK12 carrying a 4 kbp ppiD+DNA fragment this study pCD50 pOK12 carrying the 2.8 kbp EcoNI–SmaI ppiD+ this study pCD51 pOK12 carrying the 2.5 kbp BsaBI–SmaI ppiD+ this study pCD52 pET24-a carrying the minimal ppiD+coding sequence this study (1.9 kbp NdeI–EcoRI) pCD56 pOK12 carrying the 1.1 kbp EcoNI–KpnI hupB+ this study pCD57 pAED-4 carrying the minimal ppiD+coding sequence this study pCD160 pCD57 with an ΩTet inserted at the KpnI site this study pCD174 pCD57 with an ΩKan inserted at the KpnI site this study pCD175 pKO3 with an ΩKan inserted at the PstI site this study pCD176 pKO3 with an ΩTet inserted at theKpnI site this study pCD189 pRS550 (pppiDlacZ) carrying 360 nucleotides of the ppiD promoter this study pCD191 pCD189 first 147 nucleotides containing the CpxR boxes deleted this study pCD271 pCD57 carrying mutation G347 to A this study pCD273 pCD57 carrying mutation I350 to A this study pCD275 pSE420 carrying the minimal ppiD+coding sequence this study Identification, purification and localization of the PpiD protein We constructed a minimal subclone of ppiD, pCD52, by PCR in the pET24a expression vector with a T7 promoter. This plasmid was first sequenced and then shown to complement ppiD fully and to suppress surA mutant bacteria. Expression of the protein was induced with isopropyl-β-D-thiogalactopyranoside (IPTG) and, consistent with the predicted mass of the deduced amino acid sequence, was shown to be a protein of ∼69 kDa (Figure 8). Examination of the PpiD amino acid sequence predicted it to be an inner membrane protein, with a single transmembrane domain spanning amino acid residues 16–34 with its N-terminus in the cytoplasmic membrane. The rest of the polypeptide (amino acids 35–623) is predicted to be in the periplasm. The overexpressed PpiD protein was indeed found to localize to the inner membrane (Figure 8A) and could be released either upon Triton X-100 (1%) or sarkosyl (0.2%) treatment. The solubilized membrane fraction containing PpiD was first purified by gel filtration followed by an anion exchange chromatography. The purified protein was eluted with a linear gradient of KCl (0.05–1 M) containing 0.2% NP-40. Figure 8.(A) Purification of PpiD. Cultures of strains carrying plasmid pCD51 in which the ppiD gene is under the transcriptional control of the T7 polymerase promoter were grown at 37°C in M9 minimal medium. Expression of the T7 RNA polymerase was induced by addition of IPTG (1 mM) for 2 h. Lane 1 is the total extract from induced bacteria carrying plasmid pCD51, lane 2 represents the soluble fraction, lane 3 represents the inner membrane fraction after extraction with sarkosyl, lane 4 contains the aggregated and outer membrane proteins not released by sarkosyl, and lane 5 contains purified PpiD protein after gel filtration and further purification on MonoQ (FPLC). Proteins were resolved by SDS–PAGE (12.5% acrylamide), and a Coommasie Blue-stained dried gel is shown. (B) For the immunoprecipitation experiments, a 10 ml culture of wild-type bacteria was labelled with [35S]methionine (100 μCi/ml) for 10 min and fractionated as above. Lane 1 is PpiD immunoprecipitated from total cell extracts, lane 2 represents sol