Role of lipopolysaccharide on the structure and function of α-hemolysin from Escherichia coli

Chemistry and Physics of Lipids 135 (2005) 107–115

Role of lipopolysaccharide on the structure and function of ␣-hemolysin from Escherichia coli V. Herlax a , M.J.T. de Alaniz a , L. Bak´as a,b,∗ b

a Instituto de Investigaciones Bioqu´ımicas La Plata (INIBIOLP) Facultad de Ciencias M´ edicas, 60 y 120, 1900 La Plata, Argentina Departamento de Ciencias Biol´ogicas, Facultad de Ciencias Exactas, Universidad Nacional de La Plata, 47 y 115, 1900 La Plata, Argentina

Received 20 October 2004; received in revised form 1 February 2005; accepted 7 February 2005 Available online 28 March 2005

Abstract ␣-Hemolysin (HlyA) is a protein toxin (107 kDa) secreted by some pathogenic strains of E. coli. Several studies suggested the relationship between HlyA and lipopolysaccharide (LPS). We have studied experimentally the role of LPS on the stability and function of this toxin. The HlyA conformation in both, LPS-free and LPS-bound forms was investigated by tryptophan fluorescence. Studies about HlyA thermal and chemical denaturation indicated that its stability increased in the presence of LPS. On the other hand, the presence of negative and polar residues on the LPS reduced the tendency of HlyA to self-aggregation, and they may be the reservoir of calcium, cation essential for the lytic action of this toxin on red blood cells. These results suggest that HlyA and LPS are combined mainly via hydrophobic force to form an active toxin which stability is favored by the LPS. © 2005 Elsevier Ireland Ltd. All rights reserved. Keywords: Bacterial toxins; Lipopolysaccharide; Lipid–protein interaction; Protein fluorescence; Protein stability

1. Introduction Escherichia coli hemolysin is the prototype for the RTX exotoxin family produced by Gram-negative bacteria, it includes the leukotoxins of Pasteurella haemolytica and Actinobacillus actinomycetemcomitans and hemolysins from four Gram-negative genera. Abbreviations: HlyA, ␣-hemolysin; LPS, lipopolysaccharide; LUV, large unilamellar vesicle; Trp, tryptophan ∗ Corresponding author. Tel.: +54 221 482 4894; fax: +54 221 425 8988. E-mail address: [email protected] (L. Bak´as).

While the associated diseases and target cell specificities are diverse, they share several common structural and functional features, such as genetic organization, secretion by a leader-independent pathway and a tandemly-repeated nine-amino-acid sequence that is responsible for calcium binding (Welch, 2001). Escherichia coli hlyCABD operon encodes the polypeptide component (HlyA) of extracellular cytolytic toxin as well as proteins required for its acylation (HlyC) and sec-independent secretion (HlyB,D). HlyA, the mature form of the toxin, is a single polypeptide chain whose complex molecular organization was identified by different mutational and structure predic-

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tion studies. It comprises (a) amphiphilic ␣-helix at the N-terminal end (Erb et al., 1987), (b) a region rich in amphipatic stretches organized as ␣-helices (Soloaga et al., 1999), (c) a Ca2+ -binding domain containing 11–17 copies of a glycine-rich repeated nonapeptide (Ludwig et al., 1990) and (d) a C-terminal export sequence (Chevaux and Holland, 1996). The toxin is synthesized as an inactive precursor activated via a post-translational acylation at two internal lysine residues (Standley et al., 1996). Previous studies suggested the connection between HlyA and lipopolysaccharide (LPS). Several genes involved in the LPS biosynthetic pathway have been shown to affect either the expression or activity of HlyA, such as mutations in rfaC. Although transcription and secretion were decreased up to two-fold, specific hemolytic activity of toxin produced by the rfaC mutant strain was significantly reduced (Bauer and Welch, 1997). HlyA from the rfaC mutant strain compared to the protein produced by wild-type strain exhibited much slower kinetics of hemolysis, faster rate of decay of activity and greater formation of apparently inactive HlyA containing aggregates in culture supernatants. On the other hand, HlyA prepared in roughLPS mutant backgrounds is greatly reduced in lytic activity compared to the one from a smooth-LPS strain (Bauer and Welch, 1997; Standley et al., 1993; Wandersman and Letouffe, 1993). LPS might be required for the maximal production of some RTX toxins and might be a cofactor in several biological effects of RTX toxins (Czuprynski and Welch, 1995). It is a unique component of the outer membrane of all Gram-negative bacteria, and it is localized exclusively in the outer leaflet of this membrane, forming an impermeable barrier in addition to imparting strong hydrophilicity to the surface. The structure of E. coli LPS was reviewed by Schnaitman and Klena (Schnaitman and Klena, 1993). This molecule has amphipathic properties and consists of a hydrophobic fatty-acyl-containing lipid A; a highly charged and hydrophilic core containing 2-keto-3-deoxyoctosonic acid (KDO) linked to phosphate and ethanolamine; and a polar uncharged hydrophilic repeating polysaccharide containing an O-specific chain. LPS readily interacts with numerous biomolecules including phospholipids, membranes and serum proteins. Some of these interactions are non specific, and they probably involve nonsaturable binding, whereas

LPS binds stoichiometrically to certain proteins, suggesting a specific binding process (Ma et al., 2004; Augusto et al., 2003). The ability to bind LPS was also identified for other toxins, i.e. cholera toxins which is secreted solubly by Vibrio cholerae and heat labile enterotoxin which is retained on the surface of enterotoxigenic Escherichia coli (Horstman et al., 2004) although those studies were set out to elucidate which parts of the LPS molecules participate in toxin binding. Ostolaza et al. (1991) purified HlyA by size exclusion and ion exchange chromatography, demonstrating the presence of the LPS-associated 3hydroxytetradecanoic acid and KDO in active fractions of HlyA. Bohach and Snyder (Bohach and Snyder, 1985) showed that hemolytically active fractions affinity purified from HlyA producing culture supernatans with an anti-HlyA monoclonal antibodies (Mab) also reacted with an anti-LPS Mab, and they postulated that the toxin may exist as a complex. A model was designed for the physical interaction between LPS and HlyA in which LPS participates in forming or maintaining an active conformation of HlyA, and helps to protect it from aggregation or degradation (Bauer and Welch, 1997). However, the importance of bound LPS for HlyA structure and function has still to be experimentally resolved.

2. Materials and methods 2.1. Protein purification ␣-Hemolysin (HlyA) was purified from the culture filtrates of over producing strains of E. coli WAM 1824 (psF4000 in the JM15 background) kindly provided by Welch, RA. The cultures were grown to late log phase in LB to an optical density at 600 nm (OD600 = 0.8–1.0). Cells were pelleted and the supernatant was concentrated and partially purified by precipitation with 20% cold ethanol. Precipitate containing HlyA was collected by centrifugation (1 h, 10,000 rpm in a Sorvall centrifuge, rotor SSA 34), then resuspended in Tris 20 mM pH 7.0, NaCl 150 mM. This preparation showed on SDS–PAGE a main band at 107 kDa corresponding to more than 90% of total protein, which will be assigned as LPS boundHlyA. Other bands at low molecular weight corre-

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sponded to 10% of protein impurities were removed by dialysis (membrane cut off 30 kDa). The sample assigned as LPS-free HlyA was obtained by a modification of the method reported by Maheswaran et al. (1993) for large-scale preparation of the Pasteurella haemolytica LktA toxin, in which proteins were denatured and its interaction with LPS disrupted by boiling in the presence of SDS. The different molecules were separated by SDS gel electrophoresis, then the polypeptides were isolated from the gels by electroelution and renatured by removing the SDS with KCl (Sandri et al., 1993; Pellett and Welch, in preparation). HlyA samples were dialyzed against 20 mM Tris–HCl pH 7.0, NaCl 150 mM (TC buffer). In this way, the toxin preparations were almost free of detectable LPS by chemical assays and hemolysis activity was measured by a standard procedure to insure the correct folding of the protein. The protein can be stored at −70 ◦ C. 2.2. Gel electrophoresis Samples were resolved on a Biorad minislab gel SDS–PAGE apparatus. Samples were mixed with the sample diluter (2% SDS, 10% glycerol, w/v, 0.01% bromophenol blue, 10 mM Tris–HCl, pH 6.8). The samples were electrophoresed on 10% acrylamide gels in the presence of SDS according to Laemmli (1970). Protein bands were visualized by Coomassie Blue staining and LPS was visualized using a modified silver stain (Fomsgaard et al., 1990). 2.3. Immunoblotting analysis Samples from 10% SDS–polyacrylamide gels were transferred to nitrocellulose by the method of Towbin et al. (1979). Blots were blocked with 3% skim milk in TBS buffer (10 mM Tris–HCl, 150 mM NaCl, pH 7.5) 2 h at room temperature. They were then incubated with a solution containing a polyclonal rabbit antihemolysin antibody (1:1000) in 3% skim milk/TBS overnight at 4 ◦ C, washed with TBS buffer, and finally reacted with peroxidase-conjugated anti-rabbit Ig antibody (Sigma) (1:1000) in TBS buffer with 3% skim milk for 2 h at room temperature. After incubation and washing as above, the nitrocellulose was transferred to a peroxidase substrate solution containing 6 mg of 4-chloro-1-naphthol (Sigma) in 1 ml methanol,

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1 ml TBS, 3 ml H2 O and 8 ul H2 O2 for the detection of horseradish peroxidase-conjugate antibodies on the membrane. 2.4. Isolation of LPS A modified phenol–water procedure was used to isolate LPS from E. coli WAM 1824 (Westphal and Jann, 1965). The bacterial pellet (1 g) was resuspended in 10 ml PBS, and 10 ml of distilled water saturated with phenol were added and vortexed at 60–70 ◦ C for 10 min. After this time, it was placed on ice for 10 min and centrifuged at 7000 × g at 4 ◦ C for 20 min. Isolated LPS in the aqueous phase was subjected to a complete dialysis and lyophilization. 2.5. KDO assay 2-Keto-3-deoxyoctonate concentrations were determined by a colorimetric microassay (Karkhanis et al., 1978), in which 45 ␮l of samples were mixed with 5 ␮l of 0.2 N H2 SO4 and boiled for 20 min in sealed microcentrifuge tubes. Following boiling, the tubes were centrifuged at low speed to return all liquid to the bottom of the tubes, and 25 ␮l of periodate reagent (0.04 M NaIO4 in 0.125 N H2 SO4 ) was added. The mixtures were incubated in dark at room temperature for 20 min, and then 65 ␮l of arsenite reagent (2% NaAsO2 in 0.5 N HCl) was added with mixing. The samples were mixed until the yellowish color disappeared, then 250 ␮l of 0.3% thiobarbituric acid (Sigma Chemical Co.) was added, the reaction mixtures were vortexed and incubated in a boiling water bath for 10 min, 125 ␮l of dimethyl sulfoxide was added immediately to each sample while the contents were still hot. After the assay tubes had cooled, the OD550nm was measured. Buffer served as the blank, and a 0.1–2 ␮g of KDO standard range (Sigma Chemical Co.) was used to quantify the KDO present in unknowns. 2.6. Reconstitution of LPS–HlyA complex LPS (200 ␮g) isolated from WAM 1824 and LPSfree HlyA (10 ␮g) were mixed in TC buffer and incubated for 4 h at 4 ◦ C in the presence of deoxycholate (DOC) 0.6% and EDTA 0.1 mM. Then, the complex was subjected to dialysis (MWCO dyalisis membranes 12 kDa) against TC buffer.

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2.7. Measurements of intrinsic fluorescence Protein intrinsic fluorescence spectra were recorded on a SLM 4800 Aminco spectrofluorometer with a temperature controlled sample holder. The excitation wavelength was 295 nm in order to minimize tyrosine emission (Valpuesta et al., 1987). Slit width was 8 nm for both, excitation and emission. An appropriate amount of protein was added to 1-ml cuvettes containing buffer TC (20 mM Tris, 150 mM NaCl), under continuous stirring and the emission spectra were recorded in the 320–400 nm wavelength range. Fluorescence measurements were corrected for light scattering (Lackowicz, 1984). 2.8. Thermal and GdnHCl denaturation Fluorescence emission of Trp was used to monitor HlyA denaturation by temperature or by increasing concentrations of GdnCl. The thermal stability was measured after incubating LPS-free HlyA or LPS-bound HlyA (10 ␮g/ml) for 10 min at different temperatures ranging from 20 to 85 ◦ C for 10 min. The chemical stability was measured after incubating LPS-free HlyA or LPS-bound HlyA (10 ␮g/ml) at various concentration ranging from 0 to 5 M GdnHCl buffered with TC buffer at 25 ◦ C for 2 h. After the incubation, the Trp fluorescence spectra were recorded as above described. The ratio of the fluorescence intensity at 350 and 330 nm was used to quantify the thermal or chemical denaturation degree. 2.9. Light scattering measurements The effect of Triton X-100 on the size of protein aggregates for both LPS-free HlyA and LPS-bound HlyA was determined by right-angle light scattering measurements. The scattered light intensity was measured using an SLM 4800 spectrofluorometer with both, excitation and emission monochromators set at 500 nm. The excitation and emission band passes were 4 nm each. All the measurements were carried out at 24 ◦ C. 2.10. Hemolytic assays For the hemolytic assays, an aliquot of purified or HlyA–LPS complex was serially diluted in TC cold buffer containing 10 mM CaCl2 on a 96-well microtiter

plate. One hundred microliters of the diluted suspensions were mixed with 100 ␮l of standardized horse red blood cells, and the mixture was incubated at 37 ◦ C for 30 min. The absorbance of supernatants was read at 412 nm (Snyder and Zwadyk, 1969). In order to study the dependence of the activity on calcium concentration, proteins were first dialyzed against TC buffer with 0.5 mM EDTA to eliminate calcium ions that may interact with LPS. Then the percentage of hemolysis was assayed at different calcium concentrations (0–10 mM).

3. Results and discussion Purification of HlyA was found to be difficult and many of the contradictory published results about the action mechanism of this toxin are due to the different purification schemes. HlyA preparations contained significant amounts of LPS. Binding between LPS and HlyA was not broken during chromatography in nondenaturating conditions. Li and Clinkenbeard (Li and Clinkenbeard, 1999), reported that LPS could be removed from Pasteurella haemolytica leukotoxin, another member of the RTX toxin family by preparative SDS–PAGE. LPS-bound HlyA and LPS-free HlyA purified as described in Section 2 showed on a SDS–PAGE a main band at 107 kDa corresponding to more than 90% of total protein (Fig. 1A). HlyA samples were analyzed for their lipopolysaccharide contents by SDS–PAGE silver stain. Fig. 1B shows the absence of detectable LPS in the LPS-free HlyA, while LPS-bound HlyA shows typical bands of LPS extracted from E. coli WAM 1824. To confirm the presence of HlyA in both samples, immunoblotting assay was done as shown in Fig. 1C. On the other hand, we determined the presence of 2-keto3-deoxyoctonate, which is a marker of the occurrence of lipopolysaccharide associated with protein. Values of 35.5 and ≤0.5 ␮g KDO/mg protein were detected for LPS-bound HlyA and LPS-free HlyA samples, respectively. It is interesting to assess the interaction of HlyA with LPS by fluorescence measurements, as HlyA contains four Trp residues at positions 431, 480, 579 and 914 that can work as intrinsic fluorophores, and information about their environment can be obtained. For this reason, the Trp average fluorescence properties have of-

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Fig. 1. Analysis of LPS-free HlyA and LPS-bound HlyA by protein contents (A) LPS contents (B) and Immuno Blot using antiHly antibodies (C). Ten percent polyacrylamide gel was prepared and run according to the method of Laemmli. (A) Stained with Coomasie (lanes: A, molecular markers; B, LPS-bound HlyA; C, LPS-free HlyA. (B) Stained by silver stain (lanes: B, LPS-bound HlyA; C, LPS-free HlyA; D, LPS from Salmonella abortus equi included as standard; and E, LPS extracted from E. coli WAM 1824. (C) Immuno Blot (lanes: B, LPS-bound HlyA; C, LPS-free HlyA).

ten been used to monitor the unfolding, refolding and stability of proteins. This procedure is based on the knowledge that the spectral properties of these amino acids vary providing the residue is situated in a non polar or polar environment. Fig. 2 shows the normalized fluorescence emission spectra of both, LPS-free HlyA and LPS-bound HlyA. The wavelength corresponding to fluorescence intensity maximum, which reflects in average the polarity of the local environment of these residues is blue shifted for Trp residues in LPS-bound HlyA as compared with LPS-free HlyA, indicating a significantly less polar average environment for the Trp residues in the LPScomplex form. The reconstitution of HlyA with LPS was done in the presence of DOC 0.6% and EDTA 0.1 mM. LPS has self-aggregating properties which re-

sult in large polydisperse molecular forms, and it may be dispersed by detergents. LPS seems to be maximally disaggregated in 0.6% DOC (Shauds and Chun, 1980) and EDTA was added to remove divalent cations, increasing the LPS dispersion by charge–charge repulsion. Also, in mild denaturants as DOC, which might relax the protein conformation enabling it to interact with LPS, and the complex can be reconstituted. In those conditions LPS–HlyA reconstituted complex showed overlapped fluorescence spectra with protein before electrolution (LPS-bound HlyA). Further insight into structural changes that occur with LPS binding was obtained from thermal and chemical stability studies. Minor structural changes occurring in proteins at room temperature can sometimes be fully appreciated by subjecting the protein

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Fig. 2. Normalized fluorescence emission spectra of LPS-free HlyA (䊉), LPS-bound HlyA () and LPS-HlyA reconstituted complex (
) (excitation at 295 nm). The protein concentration was 10 ␮g/ml.

Fig. 3. Temperature denaturation of LPS-free HlyA (䊉) and LPSbound LPS () monitored by intrinsic tryptophan fluorescence and presented as ratios between intensities at 350/330 nm (excitation at 295 nm).

to thermal stress (Arrondo et al., 1994). For that purpose, the intrinsic fluorescence of HlyA was recorded in the 25–85 ◦ C interval. In general, fluorescence intensity of any fluorophore will decrease with increasing temperature, because collisional relaxation is favored. Since Trp residues in aqueous environments emit around 350 nm, while in nonpolar media they do so at 330 nm, the ratio of fluorescence intensities F350/F330 has been used to monitor changes in protein conformation (Blewit et al., 1984). Fig. 3 depicts the temperature dependence of Trp fluorescence for LPS-bound HlyA and LPS-free HlyA. At temperatures around 60 ◦ C an abrupt increase in F350/F330 ratio was detected in the absence of LPS, but at higher temperatures (above 70 ◦ C) it was found in the LPS–HlyA complex form. A discontinuity or break is most frequently interpreted in terms of thermally induced structural transition. Thus, these results showed a clear difference in the protein stability by LPS interaction. In Fig. 4, the fluorescence intensity at various concentrations of GnCl was plotted as the ratio between the fluorescence intensities at 350 nm and 330 nm (F350/F330). The measurement of fluorescence to follow the denaturation in LPS-bound HlyA and LPSfree HlyA at various GnCl concentrations reveals that, in both cases, the presence of denaturant induces a red

shift of the emission maximum and a decrease of intensity. Therefore, a plot of the ratio F350/F330 as a function of GnCl indicates cooperative unfolding events for both LPS-free or LPS-bound HlyA. As expected, LPS-

Fig. 4. GnHCl denaturation of LPS-free HlyA (䊉) and LPS-bound HlyA () monitored by intrinsic tryptophan fluorescence and presented as ratios between intensities at 350/330 nm (excitation at 295 nm).

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free HlyA denatures at lower GnCl concentration than LPS–HlyA complex. Moreover, this figure shows that, even in denatured state, the HlyA association with LPS takes place according to lower values of F350/F330. There is previous evidence about the tendency of HlyA to form aggregates in solution (Ostolaza et al., 1997; Gonzalez Carrero et al., 1985). The fact that HlyA purified according to Ostolaza et al. (1991) is excluded from Sephacryl S 200 column whose exclusion limit for globular proteins corresponds to a molecular weight of 500 kDa, clearly suggests that this preparation consists of multimeric aggregates. The tendency of HlyA to aggregate in aqueous media points towards the possibility of hydrophobic forces mediating the interaction. In order to test this hypothesis, both samples (LPS-bound HlyA and LPS-free HlyA) at 0.1 mg/ml concentration were treated with the nonionic surfactant Triton X-100, which is known to effectively disperse aggregate held together by hydrophobic forces. At this concentration, the equilibrium between monomer and aggregates is shifted toward aggregate side. As seen in Fig. 5, the scattering falls to about 25% the original value in LPS-free and about 50% in LPSbound HlyA, supporting the idea that LPS increases the stability of HlyA in solution. At high concentration employed in this experiments both, LPS-free and LPS

Fig. 5. The effect of 0.25% TX 100 on light-scattering from LPSfree HlyA and LPS-bound LPS suspension (100% is the scattering in the absence of detergent).

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bound HlyA produce a relatively high light scattering in buffer. At 0.25% TX-100 concentration is enough to break the association in LPS-bound HlyA but not in LPS-free state. The possible relationship between activity and degree of aggregation was mentioned previously (Gonzalez Carrero et al., 1985). Both HlyA and LPS have large regions of hydrophobicity, including attached fatty acids, which may allow complex formation between the two molecules. Charge interaction can also be present in the complex. HlyA does not exist in the aqueous medium as a monomer or with a fixed quaternary structure, but in the form of polydispersed aggregates. Such aggregation suggests a certain amphipatic character for the protein molecule, fact also suggested by its primary structure, rich in hydrophobic residues at the N-terminal end and with a large polar Ca2+ -binding domain near the C-terminal end. However, the HlyA–LPS association prevents the spontaneous irreversible inactivation of the protein that takes place in its absence. When addressing the questions dealing with the effect of LPS on the HlyA properties and its functional consequences, a series of tests were carried out in which the toxin was suspended in buffers containing different calcium concentrations. Previous results show that Ca2+ is essential for the membrane lytic activity of HlyA and the toxin must bind calcium prior to its attachment to the membrane for lysis occurrence (Ostolaza and Go˜ni, 1995). The calcium binding induces a competent state in the protein, allowing its insertion into the membrane. The binding of this cation does not lead to major changes in the secondary structure, judging from circular dichroism spectra. However, binding to Ca2+ exposes new hydrophobic residues at the surface protein, thus facilitating its irreversible insertion (Bak´as et al., 1998). HlyA activity was quantitatively tested in red blood cells, following a standard procedure. The results are shown in Fig. 6. The activation profiles are rather similar in both cases, LPS-free HlyA and LPS-bound HlyA, though 100% of hemolysis of LPS-bound HlyA was reached at lower calcium concentration. LPS molecule has negative charge that may act as reservoir of calcium, increasing the calcium concentration near the HlyA calcium-binding domain. As was previously suggested by Soloaga et al. (1998) the monomer is the active lytic form of this toxin. The absence of LPS bound to protein favors the

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membrane consistent with this phospholipid acting as a molecular chaperon (Bogdanov and Dowhan, 1999; De Cock et al., 2001). Our results support the hypothesis that LPS as a component of RTX polypeptide–LPS complexes, indirectly affects RTX toxins activities, being its main action to maintain the protein stability in solution. However, studies about the possible contribution of LPS to the insertion and oligomerization of HlyA, need to be done.

Acknowledgements Fig. 6. Effect of LPS on the calcium concentration dependence on the lytic activities of HlyA on red blood cells, in the assay medium. LPS-free HlyA (䊉) and LPS-bound HlyA (). The activity at 10 mM Ca2+ was taken to normalize as 100% hemolytic activity.

formation of irreversibly inactivated aggregates, and consequently the concentration of these active form decreases for LPS-free HlyA as was explained above it and it is shown in Fig. 5. If divalent cations act as counter ions for the phosphate groups present in lipid A and the phosphate and carboxyl groups in the polysaccharide core, neutralization of the negative charge of these groups would reduce repulsive charges between LPS subunit and allow LPS–protein complex to aggregate. However, Ca2+ binding has the effect modifying the tertiary structure of HlyA, and this leads to protein self-aggregation (in the absence of membrane) or the membrane insertion (in their presence). These results confirm that the monomeric species involved in aggregation are different according to the presence or absence of LPS associated with HlyA, as was previously observed in the effect of pH and ionic strength on the formation of reversible or irreversible aggregates (Ostolaza et al., 1997). In this way, several reports on the unfolding, refolding and assembly of proteins strongly suggest a molecular chaperon role for LPS. For instance, OmpA undergoes conformational changes upon interaction with LPS making the protein outer membrane compatible, but no lipid was found associated with any conformational variant of OmpA. Similarly, LPS appears to be required for folding outer membrane PhoE porin monomers of E. coli. The transient interactions of LPS with the PhoE monomer hydrophobic interfaces could prevent aggregation when passing through the inner

This work was partially supported by grants from UNLP, CICPBA and Ministerio de Salud de la Naci´onArgentina. LSB is member of the Carrera del Investigador, CICPBA, Argentina and VH is a fellow from Consejo Nacional de Investigaciones Cient´ıficas y Tecnol´ogicas, CONICET, Argentina.

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