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• W1994458920 abstract "The Sec-dependent protein translocation pathway promotes the transport of proteins into or across the bacterial plasma membrane. SecA ATPase has been shown to be a nanomotor that associates with its protein cargo as well as the SecYEG channel complex and to undergo ATP-driven cycles of membrane insertion and retraction that promote stepwise protein translocation. Previous studies have shown that both the 65-kDa N-domain and 30-kDa C-domain of SecA appear to undergo such membrane cycling. In the present study we performed in vivo sulfhydryl labeling of an extensive collection of monocysteine secA mutants under topologically specific conditions to identify regions of SecA that are accessible to the trans side of the membrane in its membrane-integrated state. Our results show that distinct regions of five of six SecA domains were labeled under these conditions, and such labeling clusters to a single face of the SecA structure. Our results demarcate an extensive face of SecA that interacts with SecYEG and is in fluid contact with the protein-conducting channel. The observed domain-specific labeling patterns should also provide important constraints on model building efforts in this dynamic system. The Sec-dependent protein translocation pathway promotes the transport of proteins into or across the bacterial plasma membrane. SecA ATPase has been shown to be a nanomotor that associates with its protein cargo as well as the SecYEG channel complex and to undergo ATP-driven cycles of membrane insertion and retraction that promote stepwise protein translocation. Previous studies have shown that both the 65-kDa N-domain and 30-kDa C-domain of SecA appear to undergo such membrane cycling. In the present study we performed in vivo sulfhydryl labeling of an extensive collection of monocysteine secA mutants under topologically specific conditions to identify regions of SecA that are accessible to the trans side of the membrane in its membrane-integrated state. Our results show that distinct regions of five of six SecA domains were labeled under these conditions, and such labeling clusters to a single face of the SecA structure. Our results demarcate an extensive face of SecA that interacts with SecYEG and is in fluid contact with the protein-conducting channel. The observed domain-specific labeling patterns should also provide important constraints on model building efforts in this dynamic system. The major Sec-dependent protein translocation pathway of Eubacteria, which is essential for both protein secretion and integral membrane protein insertion, has been subjected to intensive investigation for over two decades. Its core components consist of the SecA ATPase nanomotor along with SecYEG, the presumed protein-conducting channel (reviewed in Refs. 1.Veenendaal A. van der Does C. Driessen A.J.M. Biochim. Biophys. Acta. 2004; 1694: 81-95Crossref PubMed Scopus (102) Google Scholar and 2.Vrontou E. Economou A. Biochim. Biophys. Acta. 2004; 1694: 67-80Crossref PubMed Scopus (104) Google Scholar). Peripheral components such as the export-specific SecB chaperone, which maintains preproteins in a loosely folded export-competent state and transfers them to SecA, along with the integral membrane SecDF complex, which plays an as yet unknown role in the process, are important for rapid and efficient protein translocation (3.Randall L.L. Hardy S.J. Cell. Mol. Life Sci. 2002; 59: 1617-1623Crossref PubMed Scopus (120) Google Scholar, 4.Pogliano J.A. Beckwith J. EMBO J. 1994; 13: 554-561Crossref PubMed Scopus (171) Google Scholar, 5.Duong F. Wickner W. EMBO J. 1997; 16: 4871-4879Crossref PubMed Scopus (166) Google Scholar). The protein translocation reactions are organized by the SecA ATPase nanomotor, a complex protein that consists of a 65-kDa N-domain and a 30-kDa C-domain (6.Eichler J. Wickner W. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 5574-5581Crossref PubMed Scopus (73) Google Scholar, 7.Karamanou S. Vrontou E. Slanidis G. Baud C. Roos T. Kuhn A. Politou A.S. Economou A. Mol. Microbiol. 1999; 35: 1133-1145Crossref Scopus (118) Google Scholar). The N-domain consists of two nucleotide-binding domains, NBF-I and NBF-II, 2The abbreviations used are: NBF, nucleotide-binding domain; PPXD, preprotein-binding domain; HSD, helical scaffold domain; HWD, helical wing domain; CTR, carboxyl-terminal region; AMS, 4-acetamido-4′-maleimidyl-stilbene-2,2′-disulfonic acid; MPB, 3-(N-maleimido-propinyl)biocytin; P300, membrane fraction; RSO, right-side-out membrane vesicles; S300, soluble fraction; M/S, moderate to strong. which together form a high affinity nucleotide-binding cleft, whereas a protein substrate-binding domain (PPXD) is attached to NBF-I and is subject to its modulation (8.Hunt J.F. Weinkauf S. Henry L. Fak J.J. McNicholas P. Oliver D.B. Deisenhofer J. Science. 2002; 297: 2018-2026Crossref PubMed Scopus (241) Google Scholar, 9.Sharma V. Arockiasamy A. Ronning D.R. Savva C.G. Holzenburg A. Braunstein M. Jacobs W.R. Sacchettini J.C. Proc. Natl. Acad. Sci. U. S. A. 2003; 100: 2243-2248Crossref PubMed Scopus (155) Google Scholar, 10.van Voorst F. Vereyken I.J. de Kruijff B. FEBS Lett. 2000; 486: 57-62Crossref PubMed Scopus (14) Google Scholar). The C-domain consists of a central helical scaffold domain (HSD), an organizing center upon which the various other domains of SecA reside, a helical wing domain (HWD), and a carboxyl-terminal region (CTR) that contains both acidic phospholipid and SecB binding sites (11.Fekkes P. van der Does C. Driessen A.J. EMBO J. 1997; 16: 6105-6113Crossref PubMed Scopus (158) Google Scholar, 12.Breukink E. Nouwen N. van Raalte A. Mizushima S. Tommassen J. de Kruijff B. J. Biol. Chem. 1995; 270: 7902-7907Abstract Full Text Full Text PDF PubMed Scopus (132) Google Scholar). Soluble, peripheral membrane and integral membrane pools of SecA have been described previously (13.Cabelli R.J. Dolan K.M. Qian L. Oliver D.B. J. Biol. Chem. 1991; 266: 24420-24427Abstract Full Text PDF PubMed Google Scholar). Membrane-bound SecA consists of a pool bound via acidic phospholipids as well as one bound with nanomolar affinity to SecYEG, the SecA receptor (14.Hartl F.-U. Lecker S. Schiebel E. Hendrick J.P. Wickner W. Cell. 1990; 63: 269-279Abstract Full Text PDF PubMed Scopus (449) Google Scholar, 15.Eichler J. Rinard K. Wickner W. J. Biol. Chem. 1998; 273: 21675-21681Abstract Full Text Full Text PDF PubMed Scopus (33) Google Scholar). Both free and membrane-bound SecA have the capacity to interact directly with preproteins or to receive them via transfer from the SecB-preprotein complex (14.Hartl F.-U. Lecker S. Schiebel E. Hendrick J.P. Wickner W. Cell. 1990; 63: 269-279Abstract Full Text PDF PubMed Scopus (449) Google Scholar, 16.Karamyshev A. Johnson A. J. Biol. Chem. 2005; 280: 37930-37940Abstract Full Text Full Text PDF PubMed Scopus (45) Google Scholar, 17.Fekkes P. de Wit J.G. van der Wolk J.P. Kimsey H.H. Kumamoto C.A. Driessen A.J.M. Mol. Microbiol. 1998; 29: 1179-1190Crossref PubMed Scopus (102) Google Scholar). By simultaneously binding both protein substrates as well as SecYEG, SecA is able to initiate protein translocation. The dominant model for Sec-dependent protein translocation posits that a mobile region of SecA undergoes ATP-driven cycles of membrane insertion and retraction at SecYEG to promote the stepwise translocation of proteins (referred to as SecA membrane cycling) (18.Economou A. Wickner W. Cell. 1994; 78: 835-843Abstract Full Text PDF PubMed Scopus (483) Google Scholar). This model was originally based on the observation that both the N- and C-domains of SecA appeared to undergo membrane insertion in an ATP-, preprotein-, and SecYEG-dependent fashion based on their protease-resistant state as well as their accessibility to labeling reagents that specifically label only the exterior side of the membrane (6.Eichler J. Wickner W. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 5574-5581Crossref PubMed Scopus (73) Google Scholar, 18.Economou A. Wickner W. Cell. 1994; 78: 835-843Abstract Full Text PDF PubMed Scopus (483) Google Scholar, 19.Kim Y.J. Rajapandi T. Oliver D. Cell. 1994; 78: 845-853Abstract Full Text PDF PubMed Scopus (146) Google Scholar, 20.Ramamurthy V. Oliver D. J. Biol. Chem. 1997; 272: 23239-23246Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar, 21.Eichler J. Wickner W. J. Bacteriol. 1998; 180: 5776-5779Crossref PubMed Google Scholar). SecA has also been shown to undergo a default membrane insertion reaction in the presence of nonhydrolyzable ATP analogs (22.Economou A. Pogliano J.A. Beckwith J. Oliver D.B. Wickner W. Cell. 1995; 83: 1171-1181Abstract Full Text PDF PubMed Scopus (272) Google Scholar). However, the additional observation that the protease-resistant state of the C-domain could also be induced in the presence of micellar SecYEG and a nonhydrolyzable ATP analog, under conditions in which SecYEG is degraded to small peptides, has led to the alternative suggestion that these translocation ligands simply induce a stable SecA conformational state (23.van der Does C. Manting E.H. Kaufmann A. Lutz M. Driessen A.J.M. Biochemistry. 1998; 37: 201-210Crossref PubMed Scopus (94) Google Scholar). The structure of SecYEG protein, its conformational flexibility, and the dimensions and topology of the protein-conducting channel should provide important clues into the molecular basis of SecA membrane cycling. However, these topics have been the source of considerable controversy. Monomeric, dimeric, and tetrameric states for SecYEG have been detected in various biochemical and structural studies in which the presence of SecA, preprotein, and ATP stimulated the formation of dimeric and tetrameric forms of SecYEG (see Refs. 1.Veenendaal A. van der Does C. Driessen A.J.M. Biochim. Biophys. Acta. 2004; 1694: 81-95Crossref PubMed Scopus (102) Google Scholar and 24.Scheuring J. Braun N. Nothdurft L. Stumpf M. Veenendaal A. Kol S. van der Does C. Driessen A.J.M. Weinkauf S. J. Mol. Biol. 2005; 354: 258-271Crossref PubMed Scopus (46) Google Scholar and references contained therein). A small 5–8-Å protein-conducting channel has been proposed to lie within the SecYEG protomer, whereas a much larger channel of ∼20 Å would be formed at the interface of SecYEG oligomers in the ring-like structures that have been observed previously (25.van den Berg B. Clemons W.M. Collinson I. Modls Y. Hartmann E. Harrison S.C. Rapoport T.A. Nature. 2003; 427: 36-44Crossref PubMed Scopus (995) Google Scholar, 26.Meyer T. Menetret J.-F. Breitling R. Miller K. Akey C. Rapoport T. J. Mol. Biol. 1999; 285: 1789-1800Crossref PubMed Scopus (125) Google Scholar, 27.Breyton C. Haase W. Rapoport T.A. Kuhbrandt W. Collinson I. Nature. 2002; 418: 662-664Crossref PubMed Scopus (213) Google Scholar, 28.Manting E.H. van der Does C. Remigy H. Engel A. Driessen A.J.M. EMBO J. 2000; 19: 852-861Crossref PubMed Scopus (169) Google Scholar). Recent studies favor the former model. In one study utilizing cryoelectron microscopy reconstruction, a ribosome-nascent chain complex was captured associated with a SecYEG dimer, where one protomer formed the active protein-conducting channel, while the second protomer was in an inactive state (29.Mitra K. Schaffitzen C. Shaikh T. Tama F. Simon J. Brooks C. Ban N. Frank J. Nature. 2005; 438: 318-324Crossref PubMed Scopus (220) Google Scholar). In a second study utilizing cysteine-scanning mutagenesis and disulfide bond formation, the translocating polypeptide chain was located exclusively within the central region of SecY (30.Cannon K. Or E. Clemons W.M. Shibata Y. Rapoport T.A. J. Cell Biol. 2005; 169: 219-225Crossref PubMed Scopus (131) Google Scholar). Such a narrow channel, which begins with a 20–25-Å opening but narrows to 5–8 Å at the pore ring at the middle of the membrane, would significantly limit both the depth and extent of SecA membrane insertion at SecYEG. Clearly a high-resolution structure of SecA in its membrane-inserted state at SecYEG with or without a translocation intermediate is required to elucidate the structural dynamics of the translocon. However, given the difficulty in obtaining crystals of dynamic membrane proteins that diffract to atomic dimensions, this approach is likely to be difficult to achieve. The structure of the SecY complex from Methanocaldococcus jannaschii, obtained recently, is of limited use, because Archaea do not possess a SecA homolog but instead utilize a signal recognition particle-mediated pathway for protein secretion (25.van den Berg B. Clemons W.M. Collinson I. Modls Y. Hartmann E. Harrison S.C. Rapoport T.A. Nature. 2003; 427: 36-44Crossref PubMed Scopus (995) Google Scholar, 31.Pohlschroder M. Prinz W.A. Hartmann E. Beckwith J. Cell. 1997; 91: 563-566Abstract Full Text Full Text PDF PubMed Scopus (147) Google Scholar). Therefore, other approaches to obtain such structural information need to be sought. Cysteine-scanning mutagenesis, combined with either topologically specific sulfhydryl labeling or disulfide bond formation, has been shown to be a powerful method of assessing membrane protein structure and topology. For example, considerable information on the proximity of the different transmembrane helices and cytosolic or periplasmic domains of SecYEG protein has been obtained utilizing this approach (reviewed in Ref. 1.Veenendaal A. van der Does C. Driessen A.J.M. Biochim. Biophys. Acta. 2004; 1694: 81-95Crossref PubMed Scopus (102) Google Scholar). In addition, in vivo assessment of the periplasmic accessibility of engineered cysteine residues within an integral membrane protein to membrane-impermeable sulfhydryl reagents has been utilized to derive the topology of membrane transporters in a less invasive manner than through the utilization of more conventional in vitro approaches (reviewed in Ref. 32.van Geest M. Lolkema J. Microbiol. Mol. Biol. Rev. 2000; 64: 13-33Crossref PubMed Scopus (165) Google Scholar). Previously we utilized cysteine-scanning mutagenesis along with MPB labeling in RSO to demonstrate that at least three distinct regions within PPXD, NBF-II, and CTR of integral membrane SecA were periplasmically accessible (20.Ramamurthy V. Oliver D. J. Biol. Chem. 1997; 272: 23239-23246Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar). This approach is limited however by the laboriousness of the methodology as well as potential artifacts induced during sphero-plating and osmotic rupture, which is a particular concern given the highly dynamic nature of the Sec system. In the present study we have utilized in vivo rather than in vitro topology labeling to minimize system perturbation and have greatly expanded the number of cysteine mutants that have been examined. Our results provide the first detailed look at the membrane topology of integral membrane SecA protein in a more physiological manner than used previously. Strains, Plasmids, and Chemicals−Escherichia coli BL21.19 (secA13(Am) supF(Ts) trp(Am) zch::Tn10 recA::CAT clpA:: KAN) is derived from BL21(λDE3) (33.Studier W.F. Rosenberg A.H. Dunn J.J. Dubendorff J.W. Methods Enzymol. 1990; 185: 60-89Crossref PubMed Scopus (6006) Google Scholar) and was used as the host for all secA-containing plasmids. Plasmid pT7secA-Cys-0, a derivative of pT7secA2 that has all four cysteine codons within secA changed to serine, has been described previously (20.Ramamurthy V. Oliver D. J. Biol. Chem. 1997; 272: 23239-23246Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar); it was used to create the collection of monocysteine secA mutants described here utilizing a QuikChange site-directed mutagenesis kit (Stratagene) and appropriate oligonucleotides (Integrated DNA Technologies) as described by the manufacturer. All secA mutants were verified by DNA sequence analysis utilizing the DNA sequence facility at the University of Pennsylvania. The efficiency of plating of a given secA mutant, obtained by plating an appropriate dilution of an overnight culture of the strain on LB ampicillin (100 μg/ml) plates and incubating them overnight at either 42 or 30 °C, is defined as the ratio of the titer of colonies obtained at 42 °C divided by that obtained at 30 °C × 100%. MPB and AMS were purchased from Molecular Probes. Unless otherwise noted, most other chemicals were reagent grade or better and were obtained from Sigma or a comparable supplier. In Vivo MPB Labeling of Cells−Each monocysteine secA mutant was grown in LB medium supplemented with ampicillin (100 μg/ml) at 42 °C to an A600 of 0.65–0.7, after which the culture was chilled rapidly on ice for 10–20 min and then harvested by sedimentation at 7,000 × g for 10 min at 4 °C. The cell pellet was resuspended in 0.075 volume of buffer 1 (50 mm Hepes, pH 7.6, 250 mm sucrose, 5 mm EDTA). Where specified, the resuspended culture was incubated with 5 mm AMS at 4 °C for 90 min followed by sedimentation at 20,000 × g for 10 min at 4 °C, when the cell pellet was washed and resuspended in an equivalent volume of buffer 1 prior to MPB labeling. In other experiments the resuspended culture was incubated with 0.1% Triton X-100 at 0 °C for 15 min prior to MPB labeling. Biotinylation was performed by incubation with 75 μm MPB for 3 min at 0 °C. Labeling was quenched by the addition of 2-mercaptoethanol to 500 mm and incubation at 0 °C for 5 min followed by sedimentation of cells at 20,000 × g for 10 min at 4 °C. The cell pellet was resuspended in an equivalent volume of buffer 2 (50 mm Hepes, pH 7.6, 150 mm NaCl, 5 mm EDTA) supplemented with 200 mm 2-mercaptoethanol. In certain instances the MPB labeling pattern was analyzed directly on total cell protein by the addition of sample buffer (2% SDS, 125 mm Tris-HCl, pH 6.8, 5% 2-mercaptoethanol, 15% glycerol, 0.005% bromphenol blue) followed by SDS-PAGE and immunoblotting as described previously (34.Jilaveanu L.B. Zito C.R. Oliver D. Proc. Natl. Acad. Sci. U. S. A. 2005; 102: 7511-7516Crossref PubMed Scopus (85) Google Scholar). In other cases the MPB labeling pattern was analyzed on subcellular fractions. For this purpose cells were broken by two passages at 8,000 lb/in2 in a French pressure cell, and unbroken cells were removed by sedimentation at 13,000 × g for 10 min at 4 °C, giving rise to the total cleared lysate. Soluble (S300) and membrane (P300) fractions were prepared by sedimentation of the total cleared lysate at 320,000 × g for 30 min at 4 °C in a Sorvall RC M100 Micro-Ultracentrifuge. S300 was removed, and P300 was resuspended in one-sixth of the original volume of buffer 2. Following the addition of sample buffer and SDS-PAGE and immunoblotting, visualization of biotinylated proteins utilized streptavidin-conjugated horseradish peroxidase (Molecular Probes) and ECL (Pierce), whereas visualization of SecA content employed primary rabbit anti-SecA antisera and secondary goat anti-rabbit IgG-conjugated horseradish peroxidase (Pierce) and ECL. Construction of Monocysteine secA Mutants and In Vivo Labeling with MPB−Previous studies have demonstrated the feasibility of performing a topological analysis of membrane proteins utilizing a cysteine-scanning approach combined with in vivo labeling with sulfhydryl-reactive reagents (reviewed in Ref. 32.van Geest M. Lolkema J. Microbiol. Mol. Biol. Rev. 2000; 64: 13-33Crossref PubMed Scopus (165) Google Scholar). To undertake a similar approach with SecA ATPase, we created an extensive collection of monocysteine secA mutants. The mutations were targeted according to four principles. (i) We sought a mutation density that would provide us with detailed topological information on the different SecA domains, resulting in a collection of 63 monocysteine mutants for the 901-amino acid-residue SecA protein. (ii) By inspection of the homologous Bacillus subtilis SecA structure (8.Hunt J.F. Weinkauf S. Henry L. Fak J.J. McNicholas P. Oliver D.B. Deisenhofer J. Science. 2002; 297: 2018-2026Crossref PubMed Scopus (241) Google Scholar) (the two proteins have 50% identity at the amino acid sequence level (35.Schmidt M.G. Rollo E.E. Grodberg J. Oliver D.B. J. Bacteriol. 1988; 170: 3404-3414Crossref PubMed Scopus (134) Google Scholar, 36.Sadaie Y. Takamatsu H. Nakamura K. Yamane K. Gene. 1991; 98: 101-105Crossref PubMed Scopus (94) Google Scholar)), we attempted to confine our selection of monocysteine substitutions to either surface-accessible residues or residues that are located at domain-domain interfaces; the latter residues could readily become surface-accessible by a change in SecA conformation during its membrane insertion. (iii) To avoid nonfunctional secA mutants that would be difficult to grow in a haploid state and that could give rise to spurious results, we utilized an alignment of existing eubacterial SecA protein sequences to avoid highly conserved amino acid residues, and where feasible, we placed the substitution at a site where a naturally occurring cysteine residue was located in one of the SecA homologs. (iv) Finally, we tried to choose amino acid residues that were structurally/chemically similar to cysteine for substitution. The monocysteine substitutions were made on a plasmid-borne functional copy of the secA gene in which its four naturally occurring cysteine codons were substituted with serine (see “Experimental Procedures”). secA function was assessed in BL21.19, where chromosomal secA expression can be shut off by growth at 42 °C because of the presence of a secA amber mutation and a temperature-sensitive amber suppressor (37.Mitchell C. Oliver D.B. Mol. Microbiol. 1993; 10: 483-497Crossref PubMed Scopus (188) Google Scholar). Nearly all of our monocysteine secA mutants were functional in vivo as assessed by the ability of the appropriate plasmid-borne secA allele to complement the secA amber defect at 42 °C, and they gave rise to plating efficiencies of 25% or greater in general (defined under “Experimental Procedures”). The few monocysteine secA mutants that were nonfunctional were not subjected to further analysis in this study. To label regions of SecA in a topologically specific manner, we grew our strains under conditions in which only the monocysteine-containing secA gene copy was expressed at a moderate level (at 42 °C and without isopropyl-1-thio-β-d-galactopyranoside induction), and we employed the readily detectable sulfhydryl-labeling reagent, MPB. Previous studies have shown that when MPB is utilized at low concentrations, it is impermeable to the plasma membrane and can be used to map periplasmically accessible portions of membrane proteins (Refs. 38.Bayer E. Zalis M. Wilchek M. Anal. Biochem. 1985; 149: 529-536Crossref PubMed Scopus (55) Google Scholar and 39.Loo T. Clarke D. J. Biol. Chem. 1995; 270: 843-848Abstract Full Text Full Text PDF PubMed Scopus (262) Google Scholar; also shown below). The topological specificity of MPB labeling of SecA in RSO has also been demonstrated previously (20.Ramamurthy V. Oliver D. J. Biol. Chem. 1997; 272: 23239-23246Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar). In piloting our experiments we found that analysis of membrane fractions of in vivo labeled strains gave better clarity and sensitivity in our Western blots, although it was possible to analyze unfractionated cells directly as well. These procedures allowed us to directly compare the SecA labeling intensities of our different mutants without the use of immunoprecipitation or affinity purification, which complicate comparisons because of the high degree of variability in sample recovery. In addition, we found that by running the SDS-polyacrylamide gels for a longer time, better separation of high molecular weight membrane proteins in the range of SecA was achieved. By utilizing an MPB concentration and labeling time similar to those used in our previous study, we were able to achieve topologically specific labeling of SecA in vivo based on four criteria. (i) Cys-530, which has been shown previously to label strongly with MPB in RSO (20.Ramamurthy V. Oliver D. J. Biol. Chem. 1997; 272: 23239-23246Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar), was strongly labeled under our new regimen when the relevant P300 fraction was examined. By contrast, Cys-0, which lacked any cysteine residues, was not labeled even though both proteins were present at comparable levels (Fig. 1, compare panels A and B). (ii) Cys-530 labeling was prevented by pretreatment of cells with AMS (Fig. 1A), which has been utilized extensively to demonstrate topologically specific labeling by sulfhydryl-reactive reagents because of its membrane impermeability (39.Loo T. Clarke D. J. Biol. Chem. 1995; 270: 843-848Abstract Full Text Full Text PDF PubMed Scopus (262) Google Scholar). (iii) Cys-530, Cys-350, and Cys-470, which have been shown previously to label strongly, moderately, and weakly, respectively, in RSO (20.Ramamurthy V. Oliver D. J. Biol. Chem. 1997; 272: 23239-23246Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar), gave a similar in vivo labeling pattern; furthermore, strong labeling was observed in these latter two cases if the integrity of the plasma membrane was breached by Triton X-100 treatment prior to labeling (Fig. 1A). (iv) Finally, only membrane-associated SecA and not soluble SecA was labeled in vivo unless the plasma membrane was permeabilized by Triton X-100 treatment prior to labeling (Fig. 1, compare panels A and C, which contain the P300 and S300 fractions, respectively). The single, prominent, MPB-labeled, soluble protein of unknown identity in S300 fractions was presumably periplasmic in origin and was unrelated to SecA. We noted, as previous authors have done, that there was a relatively small number of MPB-labeled membrane proteins in our P300 (Fig. 1A) due to the fact that most naturally occurring cysteine residues that are accessible to the trans side of the membrane often participate in disulfide bond formation (40.Stewart J. Hermodson M. J. Bacteriol. 2003; 185: 5234-5239Crossref PubMed Scopus (13) Google Scholar). This circumstance makes in vivo labeling with sulfhydryl-reactive reagents ideal when combined with a cysteine-scanning mutagenesis approach, provided that expression levels of the test membrane protein are sufficient for ready detection. To demonstrate that our procedure labeled SecYEG-bound SecA protein, we investigated the SecYEG dependence of MBP labeling. For this purpose we compared the MPB labeling efficiency of BL21.19 (pBBsecA-his), which contains secA on a low copy number plasmid (34.Jilaveanu L.B. Zito C.R. Oliver D. Proc. Natl. Acad. Sci. U. S. A. 2005; 102: 7511-7516Crossref PubMed Scopus (85) Google Scholar), with that of BL21.19 (pBBsecA-his, pET610). Plasmid pET610 overproduces SecYEG protein, utilizing the powerful Trc promoter (41.van der Does C. de Keyzer J. van der Laan M. Driessen A. Methods Enzymol. 2003; 372: 86-98Crossref PubMed Scopus (44) Google Scholar). The results of this analysis showed that the specific activity of MPB labeling of SecA was increased 2.6-fold by SecYEG overproduction after accounting for a 34% lower SecA level in BL21.19 (pBBsecA-his, pET610) compared with BL21.19 (pBBsecA-his) (supplemental Fig. S1). Although the observed increase in specific activity of SecA labeling appears to be lower than the increase in SecYEG overproduction, we have shown previously that only a fraction of overproduced SecYEG protein properly assembles in the membrane where it gives rise to an increase in SecA high affinity binding sites and SecA-dependent translocation ATPase activity (42.Zito C.R. Oliver D. J. Biol. Chem. 2003; 278: 40640-40646Abstract Full Text Full Text PDF PubMed Scopus (22) Google Scholar). However, we cannot rule out the possibility that some of the MPB-labeled residues of SecA arose from periplasmic exposure of a phospholipid-bound pool of SecA, although such speculation is inconsistent with the extractability of this pool of SecA by reagents that classically remove only peripherally bound membrane proteins (15.Eichler J. Rinard K. Wickner W. J. Biol. Chem. 1998; 273: 21675-21681Abstract Full Text Full Text PDF PubMed Scopus (33) Google Scholar). In addition, this latter concern is inconsistent with our data given below. Data Analysis−Analysis of the MPB labeling pattern of 63 monocysteine secA mutants is given in Table 1. To ensure that our data were consistent, we performed two or more independent experiments (with separate Western blots) for each mutant, and positive and negative controls (Cys-530 and Cys-0, respectively) were included with each experiment. Given a modest degree of variability in labeling intensity and a continuum in labeling strength between strongly and moderately labeled mutants, we divided the labeling patterns into three groups: moderately to strongly labeled, weakly labeled, and unlabeled (M/S, W, and U, respectively, in Table 1). This scoring system provided a fairer representation of our data and eliminated a potentially more subjective bias. A large proportion (43%) of the SecA monocysteine residues are contained within the M/S group, a similar proportion (46%) are contained within the unlabeled (U) group, and the remainder (11%) resides within the weakly labeled (W) group.TABLE 1In vivo MPB labeling of monocysteine secA mutants Two different types of controls were used to ensure the quality of the entire data set. First, to confirm the topological specificity of labeling of the M/S group of mutants, we compared their labeling pattern in the absence and presence of AMS. Inclusion of AMS prevented MPB labeling of SecA and other proteins in all cases (supplemental Fig. S2). Second, to better understand the accessibility properties of the unlabeled group of mutants, we compared their labeling pattern in the absence and presence of 0.1% Triton X-100. This experiment indicated that these mutants could be divided into two subgroups. Cys-47, Cys-142, Cys-190, Cys-213, Cys-287," @default.
• W1994458920 created "2016-06-24" @default.
• W1994458920 creator A5045021053 @default.
• W1994458920 creator A5067591137 @default.
• W1994458920 date "2007-02-01" @default.
• W1994458920 modified "2023-09-30" @default.
• W1994458920 title "In Vivo Membrane Topology of Escherichia coli SecA ATPase Reveals Extensive Periplasmic Exposure of Multiple Functionally Important Domains Clustering on One Face of SecA" @default.
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