A rivet model for channel formation by aerolysin-like pore-forming toxins
2006; Springer Nature; Volume: 25; Issue: 3 Linguagem: Inglês
10.1038/sj.emboj.7600959
ISSN1460-2075
AutoresIoan Iacovache, Patrick Paumard, Holger Scheib, Claire Lesieur, Naomi Sakai, Stefan Matile, Michael W. Parker, Gijs R. van den Brink,
Tópico(s)Fluid Dynamics Simulations and Interactions
ResumoArticle19 January 2006free access A rivet model for channel formation by aerolysin-like pore-forming toxins Ioan Iacovache Ioan Iacovache Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland Search for more papers by this author Patrick Paumard Patrick Paumard Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland Search for more papers by this author Holger Scheib Holger Scheib Department of Structural Biology, University of Geneva, Geneva, Switzerland Swiss Institute of Bioinformatics, University of Geneva, Geneva, Switzerland SBC Lab AG, Winkel, Switzerland Search for more papers by this author Claire Lesieur Claire Lesieur Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland Search for more papers by this author Naomi Sakai Naomi Sakai Department of Organic Chemistry, University of Geneva, Geneva, Switzerland Search for more papers by this author Stefan Matile Stefan Matile Department of Organic Chemistry, University of Geneva, Geneva, Switzerland Search for more papers by this author Michael W Parker Michael W Parker Biota Structural Biology Laboratory, St Vincent's Institute of Medical Research, Fitzroy, Victoria, Australia Search for more papers by this author F Gisou van der Goot Corresponding Author F Gisou van der Goot Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland Search for more papers by this author Ioan Iacovache Ioan Iacovache Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland Search for more papers by this author Patrick Paumard Patrick Paumard Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland Search for more papers by this author Holger Scheib Holger Scheib Department of Structural Biology, University of Geneva, Geneva, Switzerland Swiss Institute of Bioinformatics, University of Geneva, Geneva, Switzerland SBC Lab AG, Winkel, Switzerland Search for more papers by this author Claire Lesieur Claire Lesieur Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland Search for more papers by this author Naomi Sakai Naomi Sakai Department of Organic Chemistry, University of Geneva, Geneva, Switzerland Search for more papers by this author Stefan Matile Stefan Matile Department of Organic Chemistry, University of Geneva, Geneva, Switzerland Search for more papers by this author Michael W Parker Michael W Parker Biota Structural Biology Laboratory, St Vincent's Institute of Medical Research, Fitzroy, Victoria, Australia Search for more papers by this author F Gisou van der Goot Corresponding Author F Gisou van der Goot Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland Search for more papers by this author Author Information Ioan Iacovache1,‡, Patrick Paumard1,‡, Holger Scheib2,3,4, Claire Lesieur1, Naomi Sakai5, Stefan Matile5, Michael W Parker6 and F Gisou van der Goot 1 1Department of Microbiology and Molecular Medicine, University of Geneva, Geneva, Switzerland 2Department of Structural Biology, University of Geneva, Geneva, Switzerland 3Swiss Institute of Bioinformatics, University of Geneva, Geneva, Switzerland 4SBC Lab AG, Winkel, Switzerland 5Department of Organic Chemistry, University of Geneva, Geneva, Switzerland 6Biota Structural Biology Laboratory, St Vincent's Institute of Medical Research, Fitzroy, Victoria, Australia ‡These authors contributed equally to this work *Corresponding author. Department of Genetics & Microbiology, CMU, University of Geneva, 30 quai Ernest Ansermet, 1211 Geneva 4, Switzerland. Tel.: +41 22 379 5652; Fax: +41 22 379 5896; E-mail: [email protected] The EMBO Journal (2006)25:457-466https://doi.org/10.1038/sj.emboj.7600959 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The bacterial toxin aerolysin kills cells by forming heptameric channels, of unknown structure, in the plasma membrane. Using disulfide trapping and cysteine scanning mutagenesis coupled to thiol-specific labeling on lipid bilayers, we identify a loop that lines the channel. This loop has an alternating pattern of charged and uncharged residues, suggesting that the transmembrane region has a β-barrel configuration, as observed for Staphylococcal α-toxin. Surprisingly, we found that the turn of the β-hairpin is composed of a stretch of five hydrophobic residues. We show that this hydrophobic turn drives membrane insertion of the developing channel and propose that, once the lipid bilayer has been crossed, it folds back parallel to the plane of the membrane in a rivet-like fashion. This rivet-like conformation was modeled and sequence alignments suggest that such channel riveting may operate for many other pore-forming toxins. Introduction Membrane proteins constitute some 30% of the human proteome, yet relatively little is still known about the structure of these proteins and how they fold and insert into the membrane. These important issues are difficult to address because the proteins of interest must first be isolated in detergents or chaotropic agents and then studied in the presence of membranes. Studies have therefore been confined mainly to two model proteins, bacteriorhodopsin and outer membrane protein A (OmpA) of Escherichia coli. A unique type of system to address folding and membrane insertion is however provided by pore-forming toxins (PFT) produced by a wide range of pathogenic bacteria. These include aerolysin from Aeromonas hydrophila (Abrami et al, 2000), α-toxin from Staphylococcus aureus (Menestrina et al, 2003) and the cholesterol-dependent toxins produced by a variety of Gram-positive bacteria (Tweten et al, 2001). These toxins are extremely intriguing because they are produced as water-soluble proteins but acquire their active state only once they have undergone a major conformational change that leads to their insertion into a lipid bilayer. A key element in this conformational change is the association into multimeric structures: either heptamers, as in the case of aerolysin and Staphylococcus α-toxin, or higher order structures (consisting of up to approximately 50 protomers) as for the cholesterol-dependent toxins. Subsequently, part of the oligomers inserts into the lipid bilayer and forms a transmembrane water-filled channel. The mechanisms that govern folding of the transmembrane region and membrane insertion are not understood at the molecular and atomic levels. Here we have focused on channel formation by aerolysin. Aerolysin is secreted as a soluble precursor named proaerolysin, which can exist as a dimer or a monomer, and requires the removal of a C-terminal peptide by proteolysis to gain activity (Abrami et al, 2000). Upon high-affinity interaction with glycosyl-phosphatidylinositol (GPI)-anchored proteins on the plasma membrane of the target cell (Abrami et al, 2000), the protein is concentrated (Abrami and van der Goot, 1999) thereby promoting the formation of stable amphipathic heptamers that can insert into the membrane and form channels (Abrami et al, 2000). Channel formation induces ion fluxes leading to membrane depolarization and ultimately cell death (Abrami et al, 2000). Despite the availability of the X-ray structure of the dimeric form of proaerolysin (Parker et al, 1994), the precise roles of each of its four domains (Figure 1A) have not been fully established and the mechanisms that lead to membrane insertion and channel formation remain poorly understood. Based on electron microscopy and docking experiments, models of the aerolysin channel have been proposed that suggest that domain 4 is the transmembrane region and domain 3 constitutes the mouth of the channel (Figure 1A) (Parker et al, 1994; Tsitrin et al, 2002). We were however intrigued by the presence of a loop in domain 3 (corresponding to amino acids 239–264, further called the DIII-loop) that has a striking pattern of alternating charged and none charged amino acids that could form a transmembrane β-hairpin. Alternating hydrophobic–hydrophilic residues are found in the transmembrane region of Staphylococcal α-toxin (Song et al, 1996), the proposed transmembrane regions of the protective antigen of anthrax toxin (Nassi et al, 2002), perfringolysin O (Shatursky et al, 1999), Clostridium septicum α-toxin (Melton et al, 2004) and bacterial porins (Schulz, 2002). We have previously shown that the DIII-loop in aerolysin is important because hemolytic activity is lost when this loop is covalently linked to the core of domain 3 by engineering a disulfide bridge between residues 253 and 300 (Rossjohn et al, 1998b). Recently, it has been shown that for the α-toxin of C. septicum, a toxin that shares 28% identity and 45% similarity with aerolysin but lacks domain 1 (Ballard et al, 1995), every other residue within the corresponding DIII-loop contacts a hydrophobic environment, compatible with a transmembrane β-strand configuration (Melton et al, 2004). Figure 1.Proaerolysin structure. (A) Ribbon diagram of the proaerolysin X-ray structure (Parker et al, 1994). The DIII-loop in domain 3 (residues K238–A261) is indicated in black. (B) Schematic representation of the DIII-loop. Acidic residues are shown in red, basic residues in blue, polar uncharged residues in yellow and hydrophobic residues in gray. The alternating pattern of charged/uncharged (hydrophobic) residues is broken by a patch of five consecutive hydrophobic residues (W247–G251). Underlined residues represent residues mutated in the K244C-A261C and K246C-E258C double cysteine mutants. Figures were generated using DeepView (Guex and Peitsch, 1997) and rendered using Pov-Ray (http://www.povray.org/). Download figure Download PowerPoint Here we have studied the topology of the DIII-loop and its functional role in channel formation. By restraining size and/or flexibility of the DIII-loop through the introduction of disulfide bridges at different positions, we first show that the DIII-loop is not required for heptamerization, but is crucial for proper membrane insertion and is directly involved in channel formation. Next, using cysteine scanning mutagenesis coupled to channel analysis using planar lipid bilayers and cysteine-specific labeling, we identify the lumen-lining residues and position the tip of the transmembrane hairpin. The hydrophobicity of this tip leads us to propose a 'rivet'-like model for membrane anchoring of aerolysin, which also seems relevant for a number of other PFT based on sequence alignments. Results Generation of single and double cysteine mutants To address the role of the DIII-loop in channel formation, we first engineered disulfide bonds within the loop to restrain its size and flexibility. Based on the structure of proaerolysin, the following double mutants were designed and generated: K244C-A261C, K246C-E258C (residues underlined in Figure 1B) and W247C-I257C. In addition, single cysteine mutants were generated for all residues from T240 to E258. C-terminally histidine-tagged variants of all the mutants were produced in E. coli and purified on nickel columns. Their activity was tested by measuring the ability to lyse erythrocytes. As shown in Table I, the double cysteine mutants K244C-A261C and K246C-E258C were non-hemolytic unless reduced with β-mercaptoethanol (βMeOH), indicating that the mutant proteins were properly folded but that disulfide bridge formation occurred and that it impaired activity. The W247C-I257C mutant was always inactive irrespective of its reducing state—and therefore not further studied—most likely because the W247C mutation in itself leads to loss of activity (Table I). All other single cysteine mutants showed significant hemolytic activity (Table I). Table 1. Hemolytic activity, single-channel conductance and sensitivity to thiol-specific labeling of DIII-loop single cysteine mutants Mutant Hemolytic activity Single-channel conductance (pS) Percentage of current decrease MTSEA-biotin MTSEA-X-biotin WT 11 449±24 K244C-A261C 0 No channels K244C-A261C+βMeOH 9 374±35 22±4 34±14 K246C-E258C 0 No channels K246C-E258C+βMeOH 9 454±80 84±2 96±1 T240C 10 378±23 31±13 16±10 T241C 10 388±32 0±0 1±1 K242C 9 395±26 26±9 27±13 N243C 8 431±31 0±2 K244C 10 443±32 15±5 16±9 F245C 9 423±40 0±0 K246C 9 422±29 0±1 0±1 W247C 0 452±27 2±1 P248C 9 446±35 2±3 1±1 L249C 12 409±13 2±2 0± V250C 8 367±13 0±1 0±1 G251C 8 389±51 0±1 2±2 E252C 8 400±35 1±2 1±1 T253C 10 423±29 2±2 1±1 E254C 9 426±33 26±10 31±11 L255C 12 416±37 2±1 S256C 10 431±23 35±6 24±5 I257C ND ND ND E258C 9 428±41 28±9 34±11 ΔLV 0 ND ΔPLV 0 ND V250D 9 378±3 L249D 9 383±4 L249C-V250D 7 No channels The hemolytic activity of mutant aerolysins in comparison to WT was assayed as described in Materials and methods and expressed in the number of wells lysed after 60 min. The single-channel conductance (measured in 1 M NaCl-containing buffer at ±25 mV) for each single cysteine mutant as well as for K246C-E258C, K246C-E258C and WT aerolysin was calculated from three different experiments as a mean value of 10 single-channel opening events. The percentage of channel closure upon addition of MTSEA-biotin and MTSEA-X-biotin was calculated as (Imax−IMTS) × 100/Imax (see Figure 4A). ND: not determined. Binding, heptamerization and membrane insertion of the disulfide-containing mutants To investigate which step in the mode of action of aerolysin was affected by the introduced disulfide bonds, we tested whether the two double cysteine mutants were able to bind to their GPI-anchored receptors. As shown in Figure 2A, in a toxin overlay assay, both mutants recognized N-CAM and semaphorin 7, two GPI-anchored proteins present in BHK cells (Fivaz et al, 2002). Consistently, these mutants were able to bind to BHK cells (Figure 2B) and oligomerize (Figure 2C), as witnessed by Western blot analysis of extracts from toxin-treated cells. Both mutants were also able to form heptamers in solution (Figure 2D, Coomassie blue-stained gel), although oligomerization was significantly slower for the K244C-A261C mutant. Figure 2.Receptor binding, heptamerization and membrane association of the disulfide-containing mutants. (A) The disulfide-containing mutants were tested in an aerolysin overlay assay on BHK cell extracts (40 μg protein per lane). Aerolysin was revealed using anti-aerolysin antibodies followed by an HRP-labeled secondary antibody. Binding to two GPI-anchored proteins, N-CAM and semaphorin 7 (sema. 7), is illustrated. (B) BHK cells were incubated with trypsin-activated WT or disulfide-containing mutant aerolysins (400 ng/ml) at 4°C for 1 h. Cell extracts (40 μg per lane), from which the nuclei were cleared by centrifugation, were analyzed by Western blotting using an anti-toxin antibody. (C) BHK cells, treated with trypsin-activated WT or mutant aerolysin as in panel B, were washed and further incubated at 37°C for 45 min. The presence of heptamers was revealed by Western blotting against aerolysin. (D) The ability of the disulfide-containing mutants to oligomerize in vitro was assayed by monitoring the appearance of the heptamer as a function of time after trypsin activation. SDS gels were revealed by Coomassie blue staining. (E) WT, Y221G and disulfide-containing mutant proaerolysins were activated with trypsin and allowed to heptamerize in vitro and then submitted to a Triton X-114 partitioning assay. Aliquots from the total sample (Tot), the aqueous (Aq) and detergent (Det) phases were analyzed by SDS–PAGE and Coomassie blue staining. (F) WT, Y221G, K246C-E258C and reduced K246C-E258C heptamer were reconstituted into proteoliposomes and submitted to separation by sucrose density flotation gradients. The six fractions of the gradient, from top to bottom, were analyzed by SDS–PAGE and Coomassie blue staining. Download figure Download PowerPoint The ability of the disulfide-containing mutants to interact specifically with GPI-anchored proteins and to heptamerize suggests that the overall structure of the toxin was not grossly altered by the introduced disulfide bonds. In the normal course of events, concomitant to heptamerization, hydrophobic patches become exposed, leading to membrane insertion of aerolysin and channel formation. Owing to its amphipathic nature, the heptamer partitions into the detergent phase after solubilization in Triton X-114 (Bordier, 1981), as we have previously shown (Tsitrin et al, 2002). When similar experiments were performed on the disulfide-containing mutants, both mutant heptamers were found in the detergent phase, indicating the presence of hydrophobic surfaces (Figure 2E, Coomassie blue-stained gel). A previously characterized single point aerolysin mutant, Y221G, was used as a control, as this mutant forms heptamers that are soluble (Tsitrin et al, 2002). Focusing on the K246C-E258C mutant, we next tested whether the heptamers could be reconstituted into proteoliposomes, as can be performed for the wild-type (WT) channel (Cabiaux et al, 1997). Heptamers were generated in solution and mixed with liposomes and detergent. The detergent was subsequently removed using Bio-beads and the liposomes were separated from free toxin by floatation on density gradients. As a negative control, we again used the Y221G heptamers, which indeed did not associate with the liposomes and remained at the bottom of the gradient—where the sample was initially loaded—in marked contrast to WT heptamers, which were used as a positive control (Figure 2F). A significant fraction of the K246C-E258C heptamers clearly floated in the gradient, demonstrating that they were bound/inserted to liposomes. However, they were found throughout the gradient, indicating that they associated with lipids less efficiently than WT heptamers, and were thus partially released during the separation procedure. Membrane association was significantly increased when the K246C-E258C mutant was reduced by βMeOH before the generation of heptamers. Channel formation by the double cysteine mutants requires disulfide reduction As the disulfide-containing mutants were able to bind to their receptors, heptamerize and expose hydrophobic patches enabling them to interact, albeit less efficiently, with lipids, the simplest explanation for their complete lack of hemolytic activity is that they are unable to form channels. We therefore analyzed their behavior using planar lipid bilayers, an extremely sensitive technique that permits the analysis of single-channel events. For all experiments, the toxin was preactivated with trypsin and added to the cis side of the bilayer setup. When approximately 10 channel events were detected, the cis chamber was perfused (with 5–10 times the chamber volume) to remove the toxin remaining in the solution. Whereas addition of WT aerolysin led to a stepwise increase in membrane current (not shown), as previously observed (Wilmsen et al, 1990), addition of either of the disulfide mutants did not, even after 1 h (not shown). Subsequently adding a reducing agent to the cis chamber was insufficient to trigger channel formation. However, when the mutants were preincubated with βMeOH before the addition to the bilayer chamber, then channels readily formed (Figure 3), and had a single-channel conductance similar to that of WT channels (Table I). These observations suggest that the disulfide bond was accessible to reducing agents in the monomer/dimer form but not in the membrane-associated heptameric form, and more importantly that, once the bond was broken, WT-like channels were formed. Figure 3.Channel analysis of the disulfide-containing mutants. The K244C-A261C (A) and K246C-E258C (B) mutants were incubated with βMeOH, activated with trypsin and added to the cis chamber of the bilayer setup, after having generated an EggPC:DOPE (1:1) membrane between the two chambers. After 5–10 channels had formed, the cis chamber was perfused as indicated. At the time indicated by an arrow, MTSEA-X-biotin was added to the trans chamber at 200 nM. For both K244C-A261C (A) and K246C-E258C (B), a decrease in current was observed, an effect that could be reversed by the addition of βMeOH as shown for K246C-E258C mutant (B). Download figure Download PowerPoint We next investigated whether the free sulfhydryl groups resulting from disulfide bridge reduction were accessible to labeling with thiol-reactive probes and whether this labeling would affect the channel properties. Addition of the thiol-specific reagent MTSEA-biotin to the trans chamber led to a small decrease (variable from 30 to 50% from one experiment to another) in current for the K244C-A261C mutant (Figure 3A) and a drastic decrease (93%) for the K246C-E258C mutant (Figure 3B), indicating that for both mutants at least one of the cysteines was accessible to labeling and located in the lumen of the channel. The inhibitory effect of MTSEA-biotin could be reversed by the addition of βMeOH, as illustrated for K246C-E258C (Figure 3B). For both mutants, the transmembrane current was reduced, irrespective of whether the MTSEA-biotin was added to the trans or the cis side of the bilayer. Also, current decrease showed little sensitivity toward the spacer length between MTSEA and the biotin moiety (using MTSEA-biotin, MTSEA-X-biotin or MSTEA-XX-biotin) as well as toward the concentration of the MTS reagent (not shown). The negatively charged MTSES in contrast did not lead to a current decrease, for either of the mutants. Taking into account these observations, all subsequent labeling experiments were performed with MTSEA-biotin and/or MTSEA-X-biotin, at a single MTS concentration. Cysteine scanning analysis of the channel structure We next analyzed the ability of all the single cysteine mutants to form channels. As described above for the disulfide-containing mutants, single cysteine mutants were preactivated with trypsin, treated with βMeOH (pretreatment with a reducing agent often increases the reactivity of cysteines) and added to the cis chamber of the bilayer setup. All mutants formed channels with a single conductance similar to that of WT channels (Table I). After about 5–10 channels had formed, the toxin remaining in solution was perfused out of the chamber. MSTEA-(X)i-biotin was then added to the trans chamber and the reduction in current was monitored, as illustrated for the E258C mutant in Figure 4A and for other mutants in Supplementary Figure S1. Changes in current upon the addition of MTS probes (MSTEA-biotin as well as MSTEA-X-biotin) were observed for cysteines at positions 240, 242 and 244 and then 254, 256 and 258 (Table I and Figure 4B). In contrast, no significant changes in current were observed for cysteines at the alternating positions, that is, 241, 243 and 245, and then 253 and 255 (Table I and Figure 4B), consistent with their orientation toward the lipid bilayer, which would render them inaccessible to labeling. Figure 4.Effect of MTSEA labeling on the single cysteine mutants. (A) The E258C single cysteine mutant was incubated with βMeOH and activated with trypsin and added to the cis chamber of the bilayer setup. After formation of some channels, free toxin was removed by perfusion of the cis chamber, as indicated. At the time indicated by an arrow, MTSEA-X-biotin was added to the trans chamber at 200 nM. (B) Schematic diagram illustrating the positions in the DIII-loop where cysteine residues were sensitive to thiol labeling (red) or insensitive (black). On the left panel, the loop is shown with the conformation it has in the crystal structure of the proaerolysin dimer (Parker et al, 1994). On the right panel, the loop is shown in the β-hairpin configuration as modeled in the current work (see Figure 6A and B). Download figure Download PowerPoint When cysteine mutants corresponding to the central continuous stretch 246–252 were analyzed, there was no alternating behavior: the conductance of all mutant channels was insensitive to MTSEA-biotin addition (Table I and Figure 4B). The central position of this region combined with the break in the alternating behavior toward MTSEA-biotin labeling suggests that it is not part of the transmembrane β-hairpin but rather forms the loop between the two β-strands (Figure 4B). This hairpin loop is expected to be on the trans side of the membrane and thus accessible to labeling. The lack of effect of MTSEA-biotin on channel conductance could be due to the fact that the biotin molecules when added to the hairpin loops are too small to obstruct the channel. With the aim of increasing the size of the labels, we removed excess MSTEA-(X)i-biotin from the bilayer chambers by perfusion and then added avidin or streptavidin, which are large molecules that bind biotin with extremely high affinity. We however did not observe any effect of avidin/streptavidin for any of the mutants with cysteines at positions 246–252, strongly suggesting that these residues were in fact inaccessible to labeling with MSTEA-(X)i-biotin. Modification of the WPLVG sequence The above cysteine scanning mutagenesis experiments identify the WPLVG sequence as the center of the transmembrane hairpin. This sequence, which is absolutely conserved among aerolysins of all Aeromonas species (Gurcel et al, 2006), is very hydrophobic, raising the possibility that it could drive membrane insertion, especially considering that bringing seven sequences together in the heptamer would generate a 35-residue patch. To address the importance of the hydrophobic WPLVG stretch in membrane insertion, we first generated a WPLVG deletion mutant. This mutant failed to fold, in hindsight not surprisingly considering the numerous contacts this sequence has with the rest of the proaerolysin molecule: Trp-247 is involved in an aromatic ring stacking interaction with Phe-404 and its indole nitrogen is involved in a hydrogen bond with Tyr-304; Pro-248 is involved in van der Waals interactions with Phe-184; finally, Val-250 forms van der Waals contacts with Phe-184, His-186, Val-189 and Tyr-304. The inability of the ΔWPLVG mutant to fold suggests that exposure of the core of domain 3, which is normally covered by the loop (Figure 1A), leads to instability in solution. We next analyzed the effects of introducing a charged residue in the middle of the WPLVG sequence. Leu-249 and Val-250 were changed to aspartic acid in single point mutants, leading to a 75% loss in hemolytic activity (corresponding to three wells difference in Table I), despite the fact that binding to erythrocytes (Figure 5A) and oligomerization (Figure 5B) were not significantly altered. The mutants were also able to expose hydrophobic patches upon heptamerization, as illustrated by their ability to partition into the detergent phase of Triton X-114 in the oligomeric form (shown for V250D in Figure 5C). Channels could be observed in lipid bilayers, with normal conductances (Table I), albeit only when adding 10-fold higher concentrations of toxin to the bilayer chamber. Altogether, these observations suggest that the introduction of a charged residue (times 7 in the heptamer) impairs crossing the lipid bilayer, that is, the step that separates heptamerization from the final transmembrane channel, but that once formed the channel has a normal conductance. Figure 5.Mutagenesis of the hairpin LV tip does not prevent aerolysin binding and heptamerization. WT or mutant (L249D, V250D, L249C-V250D and DLV) proaerolyins were incubated for 10 min at room temperature with erythrocyte ghosts and analyzed by Western blotting with anti-PA antibodies (A). (B) Following proaerolysin binding, erythrocytes were treated with trypsin for 10 min at room temperature to convert proaerolysin to aerolysin, and then with a 10-fold excess of trypsin inhibitor. Oligomerization was allowed to proceed for 60 min and samples were analyzed by Western blotting against the toxin. (C) WT and mutants were activated with trypsin for 20 min at room temperature followed by addition of trypsin inhibitor, and allowed to oligomerize for 60 min. Samples were submitted to Triton X-114 partitioning. Aqueous (Aq) and detergent (Det) phases were analyzed by SDS–PAGE and Coomassie blue staining. Download figure Download PowerPoint When both Leu-249 and Val-250 were modified, to cysteine and aspartic acid respectively, channel formation in lipid bilayers was completely abrogated (Table I) and hemolytic activity was reduced by 94%, despite normal binding and oligomerization at the surface of the erythrocytes (Figure 5A and B), altogether again indicating that membrane insertion was impaired. Deleting these two residues was similarly deleterious because the ΔLV mutant was hemolytic inactive, despite normal binding to erythrocytes (Figure 5A) and the exposure of some hydrophobic surface (Figure 5C). This mutant could only be produced in small amounts, most of it coming out of solution during the production in E. coli as for the full WPLVG deletion. This altered stability might also explain why the ΔLV mutant shows an oligomerization defect (Figure 5B). This can however not account for the loss in hemolytic activity and channel formation if one compares the ΔLV mutant to the K244C-A261C mutant, which also oligomerizes more slowly (Table I and Figure 2). Modeling of the aerolysin β-barrel The pattern observed by the cysteine scanning mutagenesis in the aerolysin transmembrane hairpin tip differs from the pattern seen in
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