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• W2007142983 abstract "Cholesterol-dependent cytolysins are a family of poreforming proteins that have been shown to be virulence factors for a large number of pathogenic bacteria. The mechanism of pore formation for these toxins involves a complex series of events that are known to include binding, oligomerization, and insertion of a transmembrane β-barrel. Several features of this mechanism remain poorly understood and controversial. Whereas a prepore mechanism has been proposed for perfringolysin O, a very different mechanism has been proposed for the homologous member of the family, streptolysin O. To distinguish between the two models, a novel approach that directly measures the dimension of transmembranes pores was used. Pore formation itself was examined for both cytolysins by encapsulating fluorescein-labeled peptides and proteins of different sizes into liposomes. When these liposomes were re-suspended in a solution containing anti-fluorescein antibodies, toxin-mediated pore formation was monitored directly by the quenching of fluorescein emission as the encapsulated molecules were released, and the dyes were bound by the antibodies. The analysis of pore formation determined using this approach reveals that only large pores are produced by perfringolysin O and streptolysin O during insertion (and not small pores that grow in size). These results are consistent only with the formation of a prepore complex intermediate prior to insertion of the transmembrane β-barrel into the bilayer. Fluorescence quenching experiments also revealed that PFO in the prepore complex contacts the membrane via domain 4, and that the individual transmembrane β-hairpins in domain 3 are not exposed to the nonpolar core of the bilayer at this intermediate stage. Cholesterol-dependent cytolysins are a family of poreforming proteins that have been shown to be virulence factors for a large number of pathogenic bacteria. The mechanism of pore formation for these toxins involves a complex series of events that are known to include binding, oligomerization, and insertion of a transmembrane β-barrel. Several features of this mechanism remain poorly understood and controversial. Whereas a prepore mechanism has been proposed for perfringolysin O, a very different mechanism has been proposed for the homologous member of the family, streptolysin O. To distinguish between the two models, a novel approach that directly measures the dimension of transmembranes pores was used. Pore formation itself was examined for both cytolysins by encapsulating fluorescein-labeled peptides and proteins of different sizes into liposomes. When these liposomes were re-suspended in a solution containing anti-fluorescein antibodies, toxin-mediated pore formation was monitored directly by the quenching of fluorescein emission as the encapsulated molecules were released, and the dyes were bound by the antibodies. The analysis of pore formation determined using this approach reveals that only large pores are produced by perfringolysin O and streptolysin O during insertion (and not small pores that grow in size). These results are consistent only with the formation of a prepore complex intermediate prior to insertion of the transmembrane β-barrel into the bilayer. Fluorescence quenching experiments also revealed that PFO in the prepore complex contacts the membrane via domain 4, and that the individual transmembrane β-hairpins in domain 3 are not exposed to the nonpolar core of the bilayer at this intermediate stage. Cholesterol-dependent cytolysins (CDCs) 1The abbreviations used are: CDC, cholesterol-dependent cytolysin; PFO, perfringolysin O; SLO, streptolysin O; αHL, S. aureus α-hemolysin; TMH, transmembrane β-hairpin; NBD, 7-nitrobenz-2-oxa-1,3-diazole; FITC, fluorescein isothiocyanate; CA, carbonic anhydrase; Amy, β-amylase; Thy, thyroglobulin; GSH, reduced glutathione; Fl, fluorescein; POPC, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine; x-doxyl-PC, 1-palmitoyl-2-stearoyl-(x-doxyl)-sn-glycero-3-phosphocholine; DPA, 2,6-pyridinedicarboxylic acid or dipicolinic acid.1The abbreviations used are: CDC, cholesterol-dependent cytolysin; PFO, perfringolysin O; SLO, streptolysin O; αHL, S. aureus α-hemolysin; TMH, transmembrane β-hairpin; NBD, 7-nitrobenz-2-oxa-1,3-diazole; FITC, fluorescein isothiocyanate; CA, carbonic anhydrase; Amy, β-amylase; Thy, thyroglobulin; GSH, reduced glutathione; Fl, fluorescein; POPC, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine; x-doxyl-PC, 1-palmitoyl-2-stearoyl-(x-doxyl)-sn-glycero-3-phosphocholine; DPA, 2,6-pyridinedicarboxylic acid or dipicolinic acid. are produced by a variety of pathogenic Gram-positive bacteria (reviewed in Refs. 1Tweten R.K. Parker M.W. Johnson A.E. Curr. Top. Microbiol. Immunol. 2001; 257: 15-33Crossref PubMed Google Scholar and 2Palmer M. Toxicon. 2001; 39: 1681-1689Crossref PubMed Scopus (155) Google Scholar). The monomeric forms of the CDCs are highly water soluble, but the proteins bind to cholesterol-containing membranes and then spontaneously self-associate to form large aqueous pores in the bilayer. These oligomeric complexes vary in size and may contain up to 50 individual monomers (3Bhakdi S. Tranum-Jensen J. Sziegoleit A. Infect. Immun. 1985; 47: 52-60Crossref PubMed Google Scholar, 4Olofsson A. Hebert H. Thelestam M. FEBS Lett. 1993; 319: 125-127Crossref PubMed Scopus (78) Google Scholar, 5Morgan P.J. Hyman S.C. Rowe A.J. Mitchell T.J. Andrew P.W. Saibil H.R. FEBS Lett. 1995; 371: 77-80Crossref PubMed Scopus (50) Google Scholar). The only crystal structure of a water soluble, monomeric form of a CDC was solved by Rossjohn et al. (6Rossjohn J. Feil S.C. Mckinstry W.J. Tweten R.K. Parker M.W. Cell. 1997; 89: 685-692Abstract Full Text Full Text PDF PubMed Scopus (401) Google Scholar) for perfringolysin O (PFO) from Clostridium perfringens, and their data revealed that PFO is comprised of four domains. The crystal structure of a membrane-inserted oligomer of a CDC is not presently available. However, several fluorescence-based studies have identified the regions of PFO that form a transmembrane β-barrel and have also provided other structural information about the membrane-inserted oligomer. Domain 3 of PFO contains two stretches of amino acids (190–217 and 288–311) that interact with the membrane during pore formation and create an amphipathic β-sheet that serves as an aqueous-lipid interface after insertion into the membrane (7Shepard L.A. Heuck A.P. Hamman B.D. Rossjohn J. Parker M.W. Ryan K.R. Johnson A.E. Tweten R.K. Biochemistry. 1998; 37: 14563-14574Crossref PubMed Scopus (268) Google Scholar, 8Shatursky O. Heuck A.P. Shepard L.A. Rossjohn J. Parker M.W. Johnson A.E. Tweten R.K. Cell. 1999; 99: 293-299Abstract Full Text Full Text PDF PubMed Scopus (309) Google Scholar). Domain 4 (residues 391–500) is involved in membrane recognition and binding, and remains close to the membrane surface in the membrane-inserted oligomer where it contacts, but is not deeply embedded in, the bilayer (9Heuck A.P. Hotze E.M. Tweten R.K. Johnson A.E. Mol. Cell. 2000; 6: 1233-1242Abstract Full Text Full Text PDF PubMed Scopus (150) Google Scholar, 10Ramachandran R. Heuck A.P. Tweten R.K. Johnson A.E. Nat. Struc. Biol. 2002; 9: 823-827PubMed Google Scholar). The mechanism of action of the CDCs involves a complex series of events that are known to include the binding and stable association with cholesterol-containing membranes by the toxin monomers (11Oberley T.D. Duncan J.L. Infect. Immun. 1971; 4: 683-687Crossref PubMed Google Scholar, 12Ohno-Iwashita Y. Iwamoto M. Mitsui K. Ando S. Nagai Y. Eur. J. Biochem. 1988; 176: 95-101Crossref PubMed Scopus (59) Google Scholar), the lateral diffusion on the bilayer with concomitant formation of cytolysin oligomers (13Harris R.W. Sims P.J. Tweten R.K. J. Biol. Chem. 1991; 266: 6936-6941Abstract Full Text PDF PubMed Google Scholar, 14Palmer M. Valeva A. Kehoe M. Bhakdi S. Eur. J. Biochem. 1995; 231: 388-395Crossref PubMed Scopus (45) Google Scholar, 15Palmer M. Harris R. Freytag C. Kehoe M. Tranum-Jensen J. Bhakdi S. EMBO J. 1998; 17: 1598-1605Crossref PubMed Scopus (156) Google Scholar, 16Shepard L.A. Shatursky O. Johnson A.E. Tweten R.K. Biochemistry. 2000; 39: 10284-10293Crossref PubMed Scopus (173) Google Scholar, 17Hotze E.M. Wilson-Kubalek E.M. Rossjohn J. Parker M.W. Johnson A.E. Tweten R.K. J. Biol. Chem. 2001; 276: 8261-8268Abstract Full Text Full Text PDF PubMed Scopus (114) Google Scholar), and ordered and coupled conformational changes that result in pore formation (9Heuck A.P. Hotze E.M. Tweten R.K. Johnson A.E. Mol. Cell. 2000; 6: 1233-1242Abstract Full Text Full Text PDF PubMed Scopus (150) Google Scholar, 17Hotze E.M. Wilson-Kubalek E.M. Rossjohn J. Parker M.W. Johnson A.E. Tweten R.K. J. Biol. Chem. 2001; 276: 8261-8268Abstract Full Text Full Text PDF PubMed Scopus (114) Google Scholar, 18Abdel-Ghani E.M. Weis S. Walev I. Kehoe M. Bhakdi S. Palmer M. Biochemistry. 1999; 38: 15204-15211Crossref PubMed Scopus (43) Google Scholar, 19Hotze E.M. Heuck A.P. Czajkowsky D.M. Shao Z. Johnson A.E. Tweten R.K. J. Biol. Chem. 2002; 277: 11597-11605Abstract Full Text Full Text PDF PubMed Scopus (110) Google Scholar). The relative timing of oligomer formation and of the insertion of individual transmembrane β-hairpins (TMHs) has been a subject of controversy (1Tweten R.K. Parker M.W. Johnson A.E. Curr. Top. Microbiol. Immunol. 2001; 257: 15-33Crossref PubMed Google Scholar, 2Palmer M. Toxicon. 2001; 39: 1681-1689Crossref PubMed Scopus (155) Google Scholar, 20Bayley H. Curr. Biol. 1997; 7: R763-R767Abstract Full Text Full Text PDF PubMed Google Scholar, 21Gilbert R.J.C. Cell. Mol. Life Sci. 2002; 59: 832-844Crossref PubMed Scopus (158) Google Scholar). Two different models have been proposed to explain the insertion of the CDC β-sheet into the membrane, one based on the prepore mechanism that has been shown to mediate the formation of small pores by several toxins (Fig. 1A; Refs. 22Walker B. Krishnasastry M. Zorn L. Bayley H. J. Biol. Chem. 1992; 267: 21782-21786Abstract Full Text PDF PubMed Google Scholar, 23van der Goot F.G. Pattus F. Wong K.R. Buckley J.T. Biochemistry. 1993; 32: 2636-2642Crossref PubMed Scopus (84) Google Scholar, 24Song L. Hobaugh M.R. Shustak C. Cheley S. Bayley H. Gouaux J.E. Science. 1996; 274: 1859-1866Crossref PubMed Scopus (1937) Google Scholar, 25Sellman B.R. Kagan B.L. Tweten R.K. Mol. Microbiol. 1997; 23: 551-558Crossref PubMed Scopus (68) Google Scholar), and the other on the gradual enlargement of a small oligomer and pore into a large oligomer and pore by the sequential addition of monomers to an initial inserted complex (Fig. 1B; Ref. 15Palmer M. Harris R. Freytag C. Kehoe M. Tranum-Jensen J. Bhakdi S. EMBO J. 1998; 17: 1598-1605Crossref PubMed Scopus (156) Google Scholar). Shepard et al. (16Shepard L.A. Shatursky O. Johnson A.E. Tweten R.K. Biochemistry. 2000; 39: 10284-10293Crossref PubMed Scopus (173) Google Scholar) showed that a PFO oligomeric complex is formed on liposomes at both 4 and 37 °C, and also showed by SDS-agarose electrophoresis that this oligomer is large and relatively uniform in size. At low temperatures, these authors were able to identify the formation of a prepore complex formed in the absence of significant insertion of the TMHs. Furthermore, they showed that PFO was found to increase the ion conductivity through a planar bilayer by large and discrete stepwise changes in conductance that are consistent with the insertion of a preassembled pore complex into the bilayer. In contrast, Palmer et al. (15Palmer M. Harris R. Freytag C. Kehoe M. Tranum-Jensen J. Bhakdi S. EMBO J. 1998; 17: 1598-1605Crossref PubMed Scopus (156) Google Scholar) proposed that individual SLO monomers are inserted and added to the pre-existing oligomers on erythrocyte membranes to produce a pore that grows in size continuously until it reaches its final state. To establish unambiguously whether PFO and SLO only form pores of a discrete, large size or, alternatively, pores whose diameter is continually increasing, we have analyzed the mechanism of pore formation using fluorophores of different sizes to monitor pore dimensions. Several fluorescein-labeled peptides and proteins with different hydrodynamic radii were encapsulated into cholesterol-containing liposomes, and the kinetics of pore formation was detected by the rate of fluorophore exposure to quenchers (fluorescein-specific antibodies) located in the external medium. If a prepore complex is an obligatory intermediate for the insertion of the transmembrane β-barrel, all sizes of trapped molecules should be released at the same time from the liposomes. On the contrary, if the pore starts small and grows continuously, the smaller trapped molecules should be released faster than the larger ones. As we have pointed out (10Ramachandran R. Heuck A.P. Tweten R.K. Johnson A.E. Nat. Struc. Biol. 2002; 9: 823-827PubMed Google Scholar), major conformational changes in the elongated monomeric PFO structure (6Rossjohn J. Feil S.C. Mckinstry W.J. Tweten R.K. Parker M.W. Cell. 1997; 89: 685-692Abstract Full Text Full Text PDF PubMed Scopus (401) Google Scholar) are required to bring the TMHs to the membrane surface while leaving only the tip of domain 4 exposed to the bilayer. Thus, an important question is to what extent the conformational changes have occurred by the time the prepore complex is formed. In other words, what structural changes have occurred at the stage where the PFO prepore complex is poised to insert its TMHs into the bilayer, but cannot because of a lack of energy (16Shepard L.A. Shatursky O. Johnson A.E. Tweten R.K. Biochemistry. 2000; 39: 10284-10293Crossref PubMed Scopus (173) Google Scholar), a mutation (19Hotze E.M. Heuck A.P. Czajkowsky D.M. Shao Z. Johnson A.E. Tweten R.K. J. Biol. Chem. 2002; 277: 11597-11605Abstract Full Text Full Text PDF PubMed Scopus (110) Google Scholar), or a disulfide bond (9Heuck A.P. Hotze E.M. Tweten R.K. Johnson A.E. Mol. Cell. 2000; 6: 1233-1242Abstract Full Text Full Text PDF PubMed Scopus (150) Google Scholar, 17Hotze E.M. Wilson-Kubalek E.M. Rossjohn J. Parker M.W. Johnson A.E. Tweten R.K. J. Biol. Chem. 2001; 276: 8261-8268Abstract Full Text Full Text PDF PubMed Scopus (114) Google Scholar). Are its TMHs lying on the surface of the membrane prior to the cooperative insertion of the TMHs to form the β-barrel (19Hotze E.M. Heuck A.P. Czajkowsky D.M. Shao Z. Johnson A.E. Tweten R.K. J. Biol. Chem. 2002; 277: 11597-11605Abstract Full Text Full Text PDF PubMed Scopus (110) Google Scholar)? If so, have the nonpolar sides of the hairpins already been exposed to the hydrophobic core of the bilayer? Does domain 4 sit on the membrane surface with the same orientation that it has in the membrane-inserted oligomer? Because so little is known about the prepore structure, we have also addressed the above important questions. Fluorescence quenching analyses of the mechanism of pore formation for a functional PFO molecule reveal the following. (i) The insertion of the oligomeric transmembrane β-barrel for PFO requires the formation of a prepore complex. (ii) Neither PFO nor SLO are able to form small pores on cholesterol-containing membranes, and (iii) the interaction of PFO with the bilayer during binding and prepore assembly does not result in small molecule leakage through the membrane. In addition, the topographical examination of the prepore complex reveals that only the tip of domain 4 is embedded in the membrane bilayer, a topography similar to that observed in the final membrane-inserted oligomer. In contrast, the domain 3 TMHs are not exposed to the nonpolar interior of the membrane in the prepore complex. Preparation of Toxin Derivatives—The gene for PFOC459A, the cysteine-less derivative of PFO in which Cys-459 was replaced by alanine, was cloned in pTrcHisA (Invitrogen, Carlsbad, CA) as described previously (7Shepard L.A. Heuck A.P. Hamman B.D. Rossjohn J. Parker M.W. Ryan K.R. Johnson A.E. Tweten R.K. Biochemistry. 1998; 37: 14563-14574Crossref PubMed Scopus (268) Google Scholar). This plasmid (named pRT20) was used as the template for all cysteine-substitution mutagenesis. The generation of cysteine-substituted derivatives of PFOC459A, their expression, and their purification have been described previously (7Shepard L.A. Heuck A.P. Hamman B.D. Rossjohn J. Parker M.W. Ryan K.R. Johnson A.E. Tweten R.K. Biochemistry. 1998; 37: 14563-14574Crossref PubMed Scopus (268) Google Scholar). Single cysteine mutants of PFO were 90–100% labeled with NBD and purified as before (7Shepard L.A. Heuck A.P. Hamman B.D. Rossjohn J. Parker M.W. Ryan K.R. Johnson A.E. Tweten R.K. Biochemistry. 1998; 37: 14563-14574Crossref PubMed Scopus (268) Google Scholar). The toxin concentration was calculated using a molar absorptivity (ϵ) at 280 nm of 84,000 cm–1m–1 for PFO and of 71,300 cm–1m–1 for SLO (26Pace C.N. Vajdos F. Fee L. Grimsley G. Gray T. Prot. Sci. 1995; 4: 2411-2423Crossref PubMed Scopus (3436) Google Scholar). Peptide and Protein Labeling with FITC—The labeling of carbonic anhydrase from bovine erythrocytes (CA), β-amylase from sweet potato (Amy), and thyroglobulin from bovine thyroid (Thy) (all from Sigma) with fluorescein isothiocyanate isomer I (FITC, Molecular Probes, Eugene, OR) was performed as described by Heuck and Wolosiuk (27Heuck A.P. Wolosiuk R.A. J. Biochem. Biophys. Methods. 1997; 34: 213-225Crossref PubMed Scopus (16) Google Scholar). Briefly, the proteins (10 mg/ml) were incubated with FITC (molar ratio 1:2, protein/FITC) in 50 mm NaHCO3 (pH 9.0) for 12 h at room temperature with gentle mixing. Unreacted FITC was removed by gel filtration (Sephadex G-25 or G-50, 1.5 cm inner diameter × 20 cm) equilibrated with 50 mm HEPES (pH 7.5), 100 mm NaCl (buffer A), and the homogeneity of the sample was determined by FPLC using a Superdex-200 column (Amersham Biosciences) equilibrated in the same buffer. Fractions corresponding to a single symmetric peak of the expected molecular weight were pooled and storaged at –20 °C until use. Reduced l-glutathione (Sigma) was incubated with FITC (molar ratio 1:1) in buffer A for 18 h and stored at –20 °C until use. Liposome Preparation—All phospholipids were obtained from Avanti Polar Lipids (Alabaster, AL). Cholesterol was obtained from Steraloids (Newport, RI). Large unilamellar liposomes were generated using an Avestin Inc. (Ottawa, Canada) Liposofast extruder. A mixture of cholesterol and 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) (55:45 mole%, respectively) in chloroform was dried at 37 °C under N2 and kept under vacuum for at least 3 h. To hydrate the lipid mixture, 0.55 ml of buffer A was added to the dried phospholipid/sterol mixture (final total lipid concentration 10 mm), and the sample was incubated for 30 min at 37 °C. The lipids were then resuspended by vortexing. The suspended phospholipid/sterol mixture was frozen in liquid N2 and thawed at 37 °C a total of five times to reduce the number of multilamellar liposomes and to enhance the trapped volumes of the vesicles (28Nayar R. Hope M.J. Cullis P.R. Biochim. Biophys. Acta. 1989; 986: 200-206Crossref Scopus (180) Google Scholar). Then the sample was passed 21 times through the extruder, equipped with both a 100-nm and a 200-nm pore size polycarbonate filter, at room temperature (20–25 °C). Whereas the 100-nm filter dictates the final size of the vesicles (mean diameter ∼110 nm; 28), the 200 nm filter faces the initial incoming solution and reduces the chances of contamination with larger particles or foreign material. The resulting liposomes were stored at 4 °C and used within 2 weeks of production. Liposomes used in lipophilic quenching experiments were prepared using POPC and cholesterol in the same way, except that 10 mole% of the POPC (i.e. 4.5% of the total lipids) was replaced by a nitroxide-labeled phospholipid, 1-palmitoyl-2-stearoyl-(x-doxyl)-sn-glycero-3-phosphocholine (x-doxyl-PC, where x is equal to 5, 7, or 12 and indicates the location of the quencher in the acyl chain of the phospholipid). The composition of these liposomes was therefore 55:40.5:4.5 mole% for cholesterol, POPC, and doxyl-PC, respectively. Liposomes containing Tb(DPA)33- and the fluorescein-labeled peptides or proteins were prepared as above, except that buffer A included 3 mm TbCl3 (Alfa Aesar, Ward Hill, MA), 9 mm 2,6-pyridinedicarboxylic acid (DPA, Sigma, neutralized to pH 7), and 20 μm fluorescein (attached to the indicated protein), and was added to the lipid film to yield a final total lipid concentration of 30 mm (final volume, 0.35 ml). The resulting liposomes were separated from non-encapsulated Tb(DPA)33- and fluorescein-labeled proteins by gel filtration (Sepharose CL-6B-200, 0.7 cm inner diameter × 50 cm) in buffer A. Traces of l-3-phosphatidylcholine-1,2-di[14C]oleoyl (Amersham Biosciences) were added to the lipid mixture in order to quantify the lipid concentration and recovery after gel filtration. The Tb(DPA)33- complex was included in each preparation to allow simultaneous analysis of the formation of small and large pores, as well as an internal control for differences in the leakage properties of each preparation. When these liposome preparations were exposed to PFOC459A, the rate of quenching of the Tb(DPA)33- complex by 5 mm EDTA in the extravesicular medium was identical in each case (not shown). Steady State Fluorescence Spectroscopy—Intensity measurements were performed using the same instrumentation described earlier (7Shepard L.A. Heuck A.P. Hamman B.D. Rossjohn J. Parker M.W. Ryan K.R. Johnson A.E. Tweten R.K. Biochemistry. 1998; 37: 14563-14574Crossref PubMed Scopus (268) Google Scholar). The excitation wavelength and bandpass, and the emission wavelength and bandpass, were, respectively: 470, 4, 530, and 4 nm for NBD; 295, 2, 348, and 4 nm for Trp; 495, 4, 520, and 4 nm for fluorescein; and 278, 2, 544, 4 nm for Tb(DPA)33- . For Tb(DPA)33- measurements, an Oriel 5215 cutoff filter (0% transmittance below 350 nm) was placed in the emission light path to block any second-order excitation light. When measurements were performed at low temperature, the cuvette chamber was continuously flushed with N2 to prevent condensation of water on the cuvettes. Kinetics measurements were done using 1 × 1 cm quartz cells, and the samples were continuously stirred using a magnetic stirring bar (1.5 × 8 mm). End-point measurements were done in 4 × 4 mm quartz microcells that were coated with POPC vesicles to minimize protein adsorption (29Ye J. Esmon N.L. Esmon C.T. Johnson A.E. J. Biol. Chem. 1991; 266: 23016-23021Abstract Full Text PDF PubMed Google Scholar). When additions were made to micro-cells, the contents were mixed thoroughly with a 2 × 2 mm magnetic stirring bar as described previously (30Dell V.A. Miller D.L. Johnson A.E. Biochemistry. 1990; 29: 1757-1763Crossref PubMed Scopus (63) Google Scholar). Time-dependent Detection of PFO NBD or Trp Emission—A sample containing the water-soluble PFO monomer (final concentration 25–100 nm) in buffer A was placed in the temperature-controlled cuvette chamber and measurements were taken until a stable signal was obtained (F 0). The kinetic analysis was started by the addition of liposomes (final concentration 50–100 μm in buffer A, final volume 1.6 ml), and data acquisition was initiated 15 s later, after complete mixing of the sample. Emission intensities were recorded at the appropriate intervals and at least sixty measurements were taken for each experiment. Blank measurements were made using an otherwise identical sample that lacked the fluorophore. The blank data were subtracted from the corresponding sample data, and the net F 0 value was then dilution-corrected. For experiments using doxyl quenchers, the emission intensities of 50 nm aliquots of the indicated monomeric NBD-labeled PFO sample in buffer A were measured at the indicated temperature as explained above. Time-dependent data acquisition was started after the addition of liposomes (final concentration 50 μm) that either contained or lacked 5-, 7-, or 12-doxyl-PC. Detection of Pore Formation—(a) End point measurements: liposomes (100 μm total lipids) were suspended in 280 μl of buffer A containing 5 mm EDTA and 4 μl of a 1:10-diluted (in buffer A) solution of anti-fluorescein antibody (0.5 μl of this rabbit polyclonal IgG fraction quenches ∼95% of the emission intensity of 0.8 fmol of GSH-Fl in buffer A; lot 84B1, Molecular Probes). The net initial emission intensity (F 0) was determined after equilibration of the sample at 25 °C for 5 min. Aliquots of 20 μl containing the amount of toxin that gives the indicated final concentration were added and the samples were incubated 30 min at 37 °C. After re-equilibration at 25 °C, the final net emission intensity (F f) of the sample was determined (i.e. after blank subtraction and dilution correction) and the fraction of marker quenched was estimated using (F 0 – F f)/(F 0 – F T), where F T is the net emission intensity obtained when the same liposomes are treated with an excess of toxin (i.e. under conditions of maximal release of the fluorophore). Typical values of F T/F 0 for Tb(DPA)33- , GSH-Fl, and Amy-Fl are 0.07, 0.12, and 0.23, respectively. (b) Kinetics measurements: liposomes (50 μm total lipids) loaded with Tb(DPA)33- and the fluorescein-labeled protein were suspended in buffer A supplemented with 5 mm EDTA and 20 μl of a 1:10-diluted (in buffer A) solution of anti-fluorescein antibody. After equilibration of the sample at 25 °C, the net initial emission intensity was determined and PFOC459A was added to a final concentration of 25 nm (final volume of 1.6 ml). Unless indicated, data collection and analysis were performed as described above. Determining the Size of Pores in Liposomal Membranes—A detailed analysis of the mechanism of PFO pore formation requires an assay that can directly measure and efficiently discriminate between transmembrane pores having different diameters. To achieve this, we have used a spectroscopic approach that detects the release of liposome-encapsulated fluorophores following toxin addition. By surrounding the liposomes with quenchers of the fluorophores, any release of fluorophore into the medium is detected by a reduction in emission intensity as a quencher contacts a fluorophore. Such an approach monitors the co-localization of fluorophore and quencher because no quenching will be observed if the quencher cannot access and interact with the fluorophore (e.g. Refs. 31Hamman B.D. Hendershot L.M. Johnson A.E. Cell. 1998; 92: 747-758Abstract Full Text Full Text PDF PubMed Scopus (349) Google Scholar, 32Sharpe J.C. London E. J. Membrane Biol. 1999; 171: 209-221Crossref PubMed Scopus (66) Google Scholar, 33Heuck A.P. Johnson A.E. Cell Biochem. Biophys. 2002; 36: 89-102Crossref PubMed Scopus (35) Google Scholar). However, as soon as a pore is formed in the liposomal membrane, a fluorophore can diffuse out of the liposome and its release will be detected by the quenching of the emission intensity. The size of the pore can then be estimated by using fluorophores of different sizes within the liposomes because a fluorophore will only move through pores that are larger than the fluorophore (32Sharpe J.C. London E. J. Membrane Biol. 1999; 171: 209-221Crossref PubMed Scopus (66) Google Scholar). Staphylococcus aureus α-hemolysin (αHL) is a β-barrel poreforming toxin that forms a transmembrane pore that is ∼16 Å in diameter (24Song L. Hobaugh M.R. Shustak C. Cheley S. Bayley H. Gouaux J.E. Science. 1996; 274: 1859-1866Crossref PubMed Scopus (1937) Google Scholar). When liposomes encapsulating fluorescent molecules of different sizes were incubated with αHL, only the small fluorophores with a diameter of about 10 Å were able to pass through the αHL pore and be quenched (Fig. 2A, Table I). A large molecule like Amy-Fl (∼100 Å in diameter) was unable to pass through the αHL transmembrane pore.Table IMolecular dimensions of the fluorophores and quenchers employed to determine the size of transmembrane poresMoleculeMWDiameterDaÅTb(DPA)33-650∼10Glutathione-FI600∼10Carbonic Anhydrase-FI29,000∼45β-Amylase-FI200,000∼100Thyroglobulin-FI669,000∼140EDTA4-290∼10Anti-FI antibody150,00060 × 120 × 150 Open table in a new tab To confirm that the observed fluorescence quenching was dictated by the size of the pore, and not by the nature of the fluorophore-quencher pair used in the assay, liposomes containing both a small fluorophore (Tb(DPA)33-) and a large fluorescent molecule (Amy-Fl) were incubated with αHL (Fig. 2B). The fast quenching of the emission intensity of Tb(DPA)33- after addition of αHL clearly shows that the integrity of the membrane has been disrupted by the formation of pores that are at least 10 Å in diameter (the diameter of the pore required for the mixing of encapsulated Tb(DPA)33- and extraliposomal EDTA). In contrast, Amy-Fl was not quenched because the fluorescent molecule and its quencher (the anti-fluorescein antibody) are too big to pass through the αHL pore. Thus, the emission intensity of fluorescein was unaffected by αHL addition. However, when PFO (pore diameter ∼250 Å) was added to the same sample, efficient quenching of Amy-Fl intensity was observed. This fluorescence approach therefore not only allows direct detection of the formation of a transmembrane pore, but it also permits a direct estimation of the diameter of the newly formed pore. Hence, this spectroscopic assay is both efficient and informative. PFO Pore Formation: a Growing Pore or a Prepore Complex Assembly?—Using the fluorescence approach described above, we can analyze pore-formation directly using a functional toxin and discriminate between the two models of pore formation proposed for the CDCs. If, as suggested by Palmer et al. (15Palmer M. Harris R." @default.
• W2007142983 created "2016-06-24" @default.
• W2007142983 creator A5001106575 @default.
• W2007142983 creator A5044088516 @default.
• W2007142983 creator A5056897795 @default.
• W2007142983 date "2003-08-01" @default.
• W2007142983 modified "2023-10-15" @default.
• W2007142983 title "Assembly and Topography of the Prepore Complex in Cholesterol-dependent Cytolysins" @default.
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