The coat protein of Pseudomonas aeruginosa phage Pf3 is transiently inserted into the bacterial inner membrane with a single transmembrane anchor sequence in the NoutCin orientation. The N‐terminal sequence immediately flanking the membrane anchor contains one negatively charged residue, whereas the C‐terminal hydrophilic segment has two positively charged residues. To investigate how the orientation of this protein is achieved, the three flanking charged amino acid residues were altered. Membrane insertion was analyzed in vivo using the accessibility to externally added protease and in vitro by testing the insertion into inverted Escherichia coli membrane vesicles. In both systems, the orientation of the protein was completely reversed for the oppositely charged mutant coat protein (RD mutant). In addition, we show in vivo that the electrochemical membrane potential is necessary for the translocation of both the wild‐type and the mutant Pf3 coat proteins, suggesting that membrane insertion is driven by electrophoretic forces.
The 44‐amino‐acid, single membrane‐spanning (type III) coat protein of the Pseudomonas aeruginosa phage Pf3 has been studied as a model protein for Sec‐independent membrane translocation (Kuhn et al., 1990; Rohrer and Kuhn, 1990; Lee et al., 1992; Cao and Dalbey, 1994). When expressed from a plasmid in Escherichia coli, it is inserted into the inner membrane with an NoutCin orientation. Due to its small size and simple membrane topology, the Pf3 coat is a very attractive system for the study of the fundamental mechanisms of membrane insertion and also for determining the structural properties which are important for this process. Recently, it was demonstrated that the Pf3 protein adopts a conformation in lipid vesicles that is 75% α‐helical and that in aqueous solution it shows a 40% α‐helical structure (Thiaudiere et al., 1993). In organic solvent, Pf3 coat protein exists as a trimer with a 90% α‐helical conformation, whereas in planar lipid bilayers it can form voltage‐gated channels, probably as tetrameric units (Pawlak et al., 1994). This suggests that the Pf3 protein undergoes considerable structural changes upon the insertion process. However, it remains unknown which structural changes occur during the membrane insertion process and whether the charged amino acid residues are transported through the lipid bilayer in an ionic state with a hydration shell or whether they are neutralized by protonation or deprotonation.
Most prokaryotic and eukaryotic membrane proteins have an asymmetric distribution of charged amino acid residues flanking the transmembrane segments of the protein (von Heijne and Gavel, 1988; Hartmann et al., 1989). In many cases, insertion of proteins into the membrane requires the Sec machinery. Spiess (1995) has suggested that the membrane topology of eukaryotic proteins may be controlled by the translocase or the associated TRAM component of the endoplasmic reticulum which recognizes the charge distribution of the protein regions. Similarly, in E.coli, the interaction of the translocated proteins with the components of the Sec machinery may very well be important for correctly orienting the transmembrane segments of larger protein domains (Kimura et al., 1991; Gafvelin and von Heijne, 1994). Short periplasmic tails or loops, which typically contain few charged residues, are thought to be translocated in a sec‐independent manner, whereas longer translocated segments can contain several charged residues and their membrane transport is strongly sec dependent. Andersson and von Heijne (1994) have recently shown that, besides the size of a translocated region, positively charged residues influence the Sec dependency of these segments. Hence, an interaction with the Sec machinery is often decisive for the topology of loops (Kuhn, 1988; Andersson and von Heijne, 1993).
In contrast, long N‐terminal tails are translocated independently of Sec by a still unknown mechanism (Dalbey et al., 1995). Interestingly, while the N‐terminal tails of such proteins typically have negatively charged residues, they contain only very few positively charged amino acid side chains. The E.coli inner‐membrane protein ProW has an extraordinarily long, highly negatively charged periplasmic tail (Whitley et al., 1994), whereas the simpler Pf3 coat protein has an 18‐amino‐acid‐residue N‐terminal tail with two negatively charged aspartyl residues. We demonstrate here that the orientation of the Pf3 coat protein is strongly determined by the distribution of charged amino acid residues directly flanking the membrane anchor. This is shown in vivo by protease mapping experiments as well as in an in vitro translation system supplemented with isolated E.coli inner‐membrane vesicles (INVs). In addition, our results suggest that the electrochemical membrane potential is the driving force for the translocation of negatively charged residues.
The translocation of the N‐terminal tail is affected by mutations in the C‐terminal tail
The Pf3 coat protein is 44 residues long with one transmembrane hydrophobic segment, an N‐terminal tail of 18 amino acid residues in length and a short C‐terminal region of eight residues. The N‐terminus is exposed to the periplasm, whereas the C‐terminus remains on the cytoplasmic side of the membrane. Since the N‐terminal region is sensitive to protease but the C‐terminal region is resistant, the membrane insertion of this protein can be analyzed on intact spheroplasts (Rohrer and Kuhn, 1990). To study how the transmembrane orientation of the Pf3 coat protein is achieved, we generated a set of mutants with alterations in the charged amino acid residues that directly border the transmembrane segment. The primary sequences of these various Pf3 mutants are shown in Figure 1. The aspartyl residue at position 7 remained unchanged in all constructs. It is unlikely that this residue is decisive for the topology because it is too far from the membrane anchor. The translocation of the Pf3 coat proteins across the membrane was investigated in vivo and in vitro with protease protection experiments using [35S]methionine‐labeled proteins in spheroplasts and INVs, respectively. For the in vivo experiments, the Pf3 proteins were expressed from a pT7‐7 plasmid in transformed BL21 cells. The cells were induced, labeled with [35S]methionine for 3 min, immediately converted to spheroplasts and treated with proteinase K. The samples were immunoprecipitated with antibodies to Pf3 coat and analyzed by SDS–PAGE, and the bands were quantified on a phosphorimager. The intactness of the spheroplasts was verified by immunoprecipitation using antibodies against the cytoplasmic GroE protein. The protease activity was verified by the accessibility of the outer‐membrane protein OmpA (data not shown). Whereas the wild‐type protein and the mutant NN were fully accessible to the protease added to the outside of the spheroplasts, indicating that the N‐terminal region of the protein had translocated, the mutants RR, RS, RN and RD were not digested and only degraded when the spheroplasts were disrupted by detergent (Figure 2A). The mutants ND, DN and DD were partially degraded, indicating that their membrane insertion was less efficient.
For the in vitro translocation experiments, the Pf3 proteins were synthesized in an E.coli translation system (S‐135; see Materials and methods) in the presence of [35S]methionine and analyzed by SDS–PAGE. For each case, the Pf3 coat protein appears as one single band of the expected size. Isolated E.coli inverted inner‐membrane vesicles were added cotranslationally. Protease digestion was performed (0.5 mg/ml proteinase K) for 30 min at 25°C. Samples were then analyzed by SDS–PAGE. Translocation rates were calculated from the ratio of totally synthesized protein and the material protected from protease digestion. Interestingly, the Pf3 coat protein and the mutants NN, DN and ND were translocated with an efficiency of ∼40%, which corresponds to the translocation rates observed for the wild‐type protein. In contrast, the N‐terminal mutants RR, RS, RN and RD did not show protection from the protease (Figure 2B). The mutant with two negatively charged residues at each end (DD) was substantially reduced in its translocation efficiency. These results show that mutations in both regions flanking the membrane anchor can affect the translocation of the N‐terminal tail, both in vivo and in vitro.
Specific labeling can discriminate the N‐terminal tail and the C‐terminal tail of the Pf3 coat protein
As the only methionyl residue in the Pf3 coat protein is the starting residue at position one, only the proteolytic fragments with the N‐terminus intact can be detected. Protein fragments that lost the N‐terminal methionine were not detectable and therefore could not be analyzed for their cellular localization. To visualize these proteolytic fragments, the Pf3 coat proteins were labeled with [3H]phenylalanine instead of [35S]methionine. The only phenylalanyl residues in the Pf3 coat protein are located at positions 43 and 44 and therefore the C‐terminus can be specifically labeled. Protease digestion experiments with the 3H‐labeled wild‐type Pf3 coat protein showed a protected fragment of a reduced size, suggesting that the protein was cleaved in the N‐terminal region. The C‐terminal segment with the radioactive phenylalanine was not accessible by proteinase K in the presence or absence of membrane vesicles. The inaccessibility of the C‐terminal segment was verified by digestion with a variety of different proteases. By using microsequencing and mass spectroscopy, only two proteinase K cleavage sites were found in the Pf3 coat protein, which were located both in the N‐terminal region between the threonyl residue at position 13 and the alanyl residue at position 14, and between the glutamyl residue at position 11 and the leucyl residue at position 12 (M.Soekarjo, 1995).
Extension of the C‐terminal tail of the Pf3 coat protein by short uncharged epitopes
To make the C‐terminal tail more accessible to protease, we engineered a spacer epitope between residues 42 and 44 of the Pf3 coat protein (Figure 3). The epitope sequences were derived from antigenic peptides originating from the HIV nef protein. Three different epitope tags with lengths of six, seven and eight uncharged amino acid residues, respectively, were selected to create the modified Pf3 proteins. The primary sequences of the three tagged versions of the Pf3 protein are shown in Figure 3B. When expressed in vitro from a pT7‐7 plasmid in the presence of [3H]phenylalanine, a translation product of the expected size was visible in each case (Figure 3A, lanes 1, 3, 5 and 7). When the control sample with the non‐tagged Pf3 protein (lanes 1 and 2) was treated with proteinase K (lane 2), the Pf3 coat protein was cleaved, presumably between the residues 13 and 14, and an 3H‐labeled C‐terminal fragment was detected. In contrast, the two tagged versions Pf3tag1 (lanes 3 and 4) and Pf3tag3 (lanes 7 and 8) did not result in a visible protease‐resistant fragment (lanes 4 and 8), indicating an additional proteolytic cleavage site in the C‐terminal region. In the case of Pf3tag2 (lanes 5 and 6), a fragment was detectable after protease treatment (lane 6). Since the tag2 sequence contains an additional phenylalanyl residue at its N‐terminus (Figure 3B), we suspect that the cleavage by the protease is most probably after this phenylalanyl residue, leaving a visible Pf3 fragment behind. For each of the subsequent experiments we used the tag1 version for studying the orientation of the Pf3 proteins by specific labeling, since the location of the two termini can be determined by labeling either the unique methionyl residue at position 1 or the unique phenylalanyl residue at position 50. We consequently engineered the tag1 epitope into the various Pf3 coat protein mutants that are schematically shown in Figure 4.
The orientation of the Pf3 coat protein is controlled by the charged residues in the tail regions and depends on the membrane potential
Cells expressing Pf3 coat proteins with tag1 were pulse‐labeled with [35S]methionine for 3 min, converted to spheroplasts and analyzed by protease mapping experiments (Figure 5). The Pf3tag1 wild‐type protein (panel A) was almost completely digested after treatment for 60 min with proteinase K (lane 3), whereas the mutants RStag1 and RNtag1 were not accessible to protease or were accessible to a very minor extent, respectively (lanes 3 and 7 of panel C). This suggests that neither the N‐terminal tail nor the C‐terminal tail was translocated. About half of the mutants RDtag1 (panel B, lanes 1–3) and DDtag1 (panel C, lanes 9–11) were digested to a smaller fragment, suggesting a cleavage in the C‐terminal tail. Because these fragments were radioactively labeled at position 1 with [35S]methionine and recognized by the Pf3 antibody that is specific for the N‐terminal region (data not shown), these Pf3 coat proteins are most likely inserted into the membrane in an inverted orientation with the C‐terminus located in the periplasmic space.
To investigate whether the translocation of the wild‐type Pf3tag1 and RDtag1 proteins required the electrochemical membrane potential, the cells were treated with CCCP, a protonophore, 45 s prior the addition of the [35S]methionine. The protease mapping experiments show that the wild type (panel A, lanes 5–8) and the RD mutant (panel B, lanes 5–8) were resistant to the externally added protease. This demonstrates that the insertion of the Pf3 proteins, no matter which membrane orientation they adopt, is strongly dependent on an intact membrane potential. The membrane insertion of the wild‐type Pf3tag1 and mutant RDtag1 was also studied in cells where the ATPase activity of the SecA protein was inhibited by 2 mM NaN3 2 min prior to the addition of [35S]methionine. Whereas cleavage of proOmpA was inhibited in both cases (data not shown), membrane insertion of the wild‐type and RD mutant was quite efficient (panels A and B, lanes 9–12).
We then investigated the membrane orientation of the Pf3tag1 proteins in E.coli INVs (Figure 6). Specific labeling of either the N‐ or the C‐terminal tail was performed with radiolabeled methionine or phenylalanine, respectively. The Pf3tag1 proteins were synthesized in vitro in the presence of E.coli INV plus either [35S]methionine (Figure 6A) or [3H]phenylalanine (Figure 6B), respectively. When the wild‐type Pf3tag1 protein was radiolabeled at the N‐terminus (panel A, lanes 1 and 2), a protease resistant fragment was detected which was slightly smaller than the original Pf3tag1 protein (lane 2), suggesting that the C‐terminal tail is accessible to the protease, leading to the smaller protected fragment. In contrast, no such fragment was detectable when the C‐terminal phenylalanyl residue was labeled (panel B, lane 2). This is consistent with the label staying at the cis‐side of the membrane and the protease cleaving off the C‐terminal region. The membrane topology of the Pf3tag1 protein is illustrated in Figure 6C, emphasizing that the radioactive label was translocated to the inside of the membrane vesicles (protease protected) in the case of [35S]methionine‐labeled wild‐type Pf3tag1 protein (left scheme) and outside the vesicles (protease accessible) in the case of a [3H]phenylalanine‐labeled protein (right scheme). These results clearly demonstrate that the N‐terminal tail of the Pf3tag1 wild‐type protein was translocated, whereas the C‐terminus stayed at the cytoplasmic side of the membrane.
To define further the features of the protein that are crucial for its membrane orientation, we examined in parallel the Pf3 coat protein mutants NNtag1, DDtag1, RStag1, RNtag1 and RDtag1 (Figure 6). The mutant NNtag1 (lanes 3 and 4) was inserted in the wild‐type orientation, although the insertion was less efficient. The mutant DDtag1 (lanes 5 and 6) showed protease resistant fragments for the N‐terminally labeled (panel A) as well as for the C‐terminally labeled (panel B) protein. When no membrane vesicles were added, no protease resistant fragments were observed (data not shown). This suggests that about equal amounts of the DDtag1 protein were inserted in an Nout or Cout orientation. No protease resistant fragments were seen for the RStag1 and the RNtag1 mutants (lanes 8 and 10 in both panels), demonstrating that neither the N‐terminus nor the C‐terminus of these two mutants was translocated and that the proteins did not adopt a transmembrane orientation at all. The RDtag1 protein (lanes 11 and 12), in contrast, clearly showed a protease resistant fragment if the C‐terminus was labeled (panel B, lane 12), but no protease resistant material was detected if the N‐terminus had been labeled (panel A, lane 12). This result is exactly the opposite to what is observed for the Pf3tag1 wild‐type protein (lanes 1 and 2 in both panels) and thus clearly demonstrates that all of the RDtag1 mutant protein had the reverse membrane orientation compared with that of the wild‐type Pf3tag1 protein. Figure 6C illustrates this reversed orientation of the RDtag1 protein with a protease protected 3H‐labeled C‐terminus (inside the vesicle) and a protease accessible 35S‐labeled N‐terminus (outside the vesicle). For all the cases analyzed, control experiments without vesicles were performed, resulting in the complete digestion of the coat proteins. Similarly, when the added vesicles were lyzed by detergent no undigested Pf3 coat protein was detected (data not shown). Taken together, our results show that the orientation of the Pf3 coat protein is strongly determined by the charged amino acid residues directly flanking the hydrophobic membrane anchor. In particular, protein regions with positively charged residues inhibit translocation, whereas negatively charged regions promote translocation. Strikingly, the DD mutant, with both hydrophilic segments being negatively charged, is the only Pf3 variant that adopts both possible transmembrane orientations. In contrast, if both flanking regions are positively charged (RStag1), membrane insertion is completely inhibited. In no case are positively charged residues translocated across the membrane.
We have studied how proteins insert into the membrane in E.coli using the small, single‐spanning Pf3 coat protein which has an N‐terminal hydrophilic tail of 18 and an extended C‐terminal tail of 14 amino acid residues. We show here that the orientation of the Pf3 coat protein in biological membranes is entirely controlled by the distribution of the charged residues present in the tails flanking the transmembrane segment. By reversing the charge distribution (mutant RDtag1) of Pf3 coat, we found that it inserts into E.coli INVs with the inverted orientation of that of the wild‐type protein. In contrast, preliminary experiments with small unilamellar phospholipid vesicles showed that only one orientation of the protein was found, independent of the charge distribution (D.Kiefer, unpublished results). On the basis of these data we believe that a component in the E.coli membrane which is distinct from the phospholipids decides which tail is translocated. Our hypothesis is that in addition to the distribution of charged amino acid residues, the electrochemical membrane potential might be the key determinant of the orientation of the translocating protein. We show that the negatively charged residues in the Pf3 coat protein respond directly to the potential and most likely play an active role in the translocation process. Positively charged tails were not translocated and supposedly were passively hindered in their translocation across the membrane bilayer possibly by electrostatic interaction with the negatively charged phospholipid headgroups on the cis‐side of the membrane. Indeed, when the membrane potential of the cell was destroyed, neither the wild‐type nor the inverted mutant inserted into the membrane at all. Since the membrane potential is such that the periplasmic side is positively charged it may therefore induce the translocation of negatively charged regions according to an electrophoretic mechanism. In this context it is interesting to note that neutral tails (C‐terminal segments of mutants NN and RN) did not efficiently translocate. When both tails were negatively charged (in mutant DD) the Pf3 protein was inserted in both orientations. Taken together, these experiments show that the negatively charged residues in the Pf3 coat protein respond directly to the potential and play an active role. Consequently, this suggests that in vivo the ionic state of the charged amino acyl side chains is the translocated form and might also control which domain of the protein is translocated across the membrane.
The results reported here for the Pf3 coat protein differ from previous studies of the M13 procoat protein, in which we had shown that a strongly positively charged loop region is translocated across the membrane in the presence or absence of a membrane potential, demonstrating that electrophoresis is not the primary driving force for insertion of this protein (Kuhn et al., 1990; Cao et al., 1995). However, the insertion mechanism of the M13 procoat protein substantially differs from that of the Pf3 coat protein, since the procoat has two membrane‐spanning regions, which both contribute to the translocation process (Kuhn et al., 1986; Kuhn, 1987). Thermodynamic studies corroborate that the wild‐type M13 procoat is inserted mainly due to hydrophobic interactions (Soekarjo et al., 1996). Electrostatic and electrophoretic forces might therefore only modulate the efficiency of membrane insertion of the M13 procoat protein.
Electrophoretic effects of translocating N‐terminal tails have been observed with mutant leader peptidase proteins (Cao and Dalbey, 1994). Whereas positively charged residues in the N‐terminal tail hindered translocation, negatively charged residues promoted the translocation. The involvement of charged amino acid residues, particularly positively charged ones, has been thoroughly studied with the leader peptidase of E.coli. The membrane orientation of the protein was inverted when the strongly positively charged cytoplasmic loop of 25 amino acid residues that links the two membrane‐spanning regions was reduced in charge and four additional lysines were added to the N‐terminus (von Heijne, 1989). Mutants with a deletion in the cytoplasmic loop which leaves two positively and three negatively charged amino acid residues (of the normal ten positively and five negatively charged residues) were membrane‐inserted in the wild‐type orientation, whereas the addition of positively charged residues to the N‐terminus increased the percentage of the inverted orientation of that mutant (Andersson et al., 1992). Similar effects were obtained by varying the charge of the loop region; increasing the number of negatively charged residues favored the inverted orientation (Andersson and von Heijne, 1994). Since the membrane insertion of the inverted leader peptidase is Sec independent, in contrast to the wild‐type protein, components of the Sec machinery might respond to the charge of the substrate and indirectly influence the orientation. This might also explain why most of the leader peptidase mutants have a mixture of both orientations.
Although the endoplasmic reticulum shows only a very marginal membrane potential, charged residues also play a role in the membrane orientation of eukaryotic proteins. This was examined in detail for three different proteins. First, for the cytochrome P‐450, the addition of positively charged residues into the short N‐terminal region preceding the transmembrane segment of the type III protein led to an almost complete inversion of the topology (Sato et al., 1990). Second, the receptor domain of the asialoglycoprotein receptor H1 was found partially to adopt a reversed orientation when four charged residues flanking the signal peptide were inverted (Beltzer et al., 1991). Third, 75% of the paramyxovirus hemagglutinin neuraminidase inverted its orientation after the introduction of charged residues into the membrane anchor flanking regions (Parks and Lamb, 1991). It has been suggested that for these proteins the translocase complex or the TRAM protein recognize the polarity and align the orientation of the translocating protein (Spiess, 1995). Membrane topology may also be influenced by the folding of a hydrophilic tail. A fusion protein of the asialoglycoprotein receptor H1 and dihydrofolate reductase (DHFR) showed that the orientation can be inverted by folding the DHFR domain in the presence of methotrexate (Denzer et al., 1995).
To investigate the role of the charged residues in orienting a membrane protein we therefore chose a small Sec‐independent protein with two comparable tails that can translocate. We observed that charged residues on either side of the membrane anchor control the orientation of the protein. We found that both orientations require the membrane potential and that the protein remains in the cytoplasm in the absence of the potential. Hence, the membrane potential supports the movement of a negatively charged side chain of the translocating protein region. The active role of negatively charged residues is also demonstrated in a mutant Pf3 coat protein where both tails are negatively charged and the protein (DD) assumes both orientations. Similarly, if one tail is uncharged and the other tail still negatively charged (NN) the negatively charged tail is translocated normally across. These two mutant proteins do not contain any positively charged residues, yet they insert into the membrane very efficiently. Uncharged tails as present in mutants RN and NN are impaired for translocation, which underscores the importance of the role that negatively charged residues play in the process.
Materials and methods
The pT7‐7 plasmid (Tabor and Richardson, 1985) with the Pf3 insert coding for the major coat protein was constructed by ligation into the restriction sites NdeI and EcoRI. By site‐directed mutagenesis (Kunkel, 1985), an NdeI restriction site was introduced at the start codon of the Pf3 coat gene. The EcoRI site was 33 bp downstream of the stop codon of the coat gene of Pf3 (Rohrer and Kuhn, 1990). Similarly, the various Pf3 coat protein mutants were obtained by oligonucleotide‐directed mutagenesis using as template DNA the M13mp19 with the Pf3 coat gene in the XbaI site. The mutants were then subcloned into the pT7‐7 vector using NdeI and EcoRI. The epitope tags within the Pf3 coat proteins were constructed by first introducing a MunI restriction site into the penultimate codon of the Pf3 coat gene. This was performed by PCR mutagenesis of the respective plasmids using the oligonucleotides 5′CGGATCAAAACAATTGCGC and 5′TAATACGACTCACTATAGGG to generate a 230 bp fragment which was ligated into pMOSBlue (Amersham) and subsequently subcloned into pT7‐7 using NdeI and EcoRI. For each of the Pf3 mutants the MunI site was introduced by this method. Into the opened MunI sites, two short, complementary linker oligonucleotides with AATT overhangs were ligated; these complementary oligonucleotides code for the respective epitope tags.
In vitro expression system
The Pf3 coat proteins were expressed by programing an E.coli translation system with the respective mRNAs transcribed in vitro using the T7 transcription Genescribe kit from United States Biochemicals. The cell‐free translation system consists of a high‐speed supernatant (S‐135) prepared according to Müller and Blobel (1984) from an E.coli wild‐type strain lysed by two cycles in a French press cell. The reactions were performed in volumes of 25 μl, containing 4 μl S‐135 and 4 μl transcription mix. Buffer and ion conditions in the translation reactions were 10 mM HEPES pH 8.0, 140 mM potassium acetate, 10 mM magnesium acetate, 20 mM ammonium acetate, 0.8 mM spermidine, 0.1 mM EDTA, 3% PEG, 40 μM 19 amino acids, 2 mM DTT, 2.5 mM ATP, 0.5 mM GTP, 12 mM phosphoenolpyruvate, 8 mM creatine phosphate, 40 μM creatine kinase and 5 μCi [35S]methionine or [3H]phenylalanine. Reactions were performed at 37°C for 30 min and stopped by the addition of 5% TCA on ice. Samples were subsequently prepared for SDS–PAGE. Bands were quantified using a Fuji BAS 1000 bio‐imaging analyzer.
Preparation of inverted membrane vesicles
Inverted inner‐membrane vesicles were prepared from exponentially growing E.coli cells essentially as described in Müller and Blobel (1984). The cells were harvested and resuspended in 1 ml/g 50 mM HEPES buffer pH 7.5, 250 mM sucrose, 1 mM EDTA and 1 mM DTT. Cell lysis was performed by two passages through a French pressure cell. The resulting supernatant was loaded on a three‐step sucrose gradient with 2.02 M, 1.4 M and 0.7 M sucrose and centrifuged in a TST 28.38‐17 Kontron swing out rotor for 16 h. The visible band between the two upper steps was collected by a syringe and concentrated by ultracentrifugation. The resulting pellet was homogenized by douncing in 50 mM HEPES pH 7.5, 250 mM sucrose and 1 mM DTT and the suspension was adjusted to an absorption at 280 nm of ∼20.
In vivo protease mapping
Five hundred microliter cultures of BL21(DE3)pLysS (Studier et al., 1990) with the respective pT7‐7 plasmids were grown to mid‐log phase in M9 minimal medium (Miller, 1972) with 0.5% fructose and 20 μg/ml of each amino acid but methionine. IPTG was added to induce synthesis of the plasmid‐borne proteins. After 10 min the cells were labeled with 50 μCi [35S]methionine for 3 min, chilled on ice and centrifuged for 1 min at 4°C. The pellet was resuspended in 250 μl 0.1 M Tris–acetate, pH 8.2, 0.5 M sucrose, 5 mM EDTA, and was treated with 80 μg/ml lysozyme (Serva) and diluted with 250 μl of water. Eight min later, 50 μl of 200 mM MgSO4 was added to stabilize the spheroplasts and collected by centrifugation. The spheroplasts were resuspended in 400 μl of 50 mM Tris–acetate, pH 8.2, 250 mM sucrose, 10 mM MgSO4. An aliquot of 100 μl was removed and precipitated with 25% TCA. To the remaining sample 1 mg/ml proteinase K (Sigma) was added and incubated on ice. One aliquot of 100 μl was removed, treated with 2.5% Triton X‐100 and incubated with proteinase K for 30 min. After quenching the protease with 5 mM PMSF, the samples were precipitated with TCA and analyzed by SDS–PAGE.
In vitro protease mapping
Insertion into isolated E.coli INVs was performed cotranslationally by adding 3 μl vesicles to the translation mixtures. After in vitro protein synthesis, 0.5 mg/ml proteinase K was added and proteolytic digestion was performed at 25°C for 30 min. Proteolysis was stopped by the addition of 2 mM PMSF and 5% TCA on ice. Samples were then prepared for SDS–PAGE.
The technical assistance of Sabine Bünger is greatly acknowledged. This work was supported by the grant Ku 749/1 from the DFG to A.K. and by the National Science Foundation grant MCB‐93166891 to R.D.
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