Laboratoire de Microbiologie, INSERM U-411, Faculté de Médecine Necker-Enfants Malades, 156 rue de Vaugirard, 75730 Paris Cedex 15, France1
Author for correspondence: Alain Charbit. Tel: +33 1 40 61 53 76. Fax: +33 1 40 61 55 92. e-mail: charbit{at}necker.fr
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ABSTRACT |
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Keywords: linker insertions, truncated proteins, domain interactions, LLO mutants
Abbreviations: HRBC, horse red blood cells; LLO, listeriolysin O; PFO, perfringolysin
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INTRODUCTION |
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LLO belongs to the family of thiol-activated cytolysins that are secreted by a large number of pathogenic Gram-positive bacteria. These pore-forming toxins comprise more than 20 members to date (Bayley, 1997 ; Alouf, 2000
), including the extensively studied perfringolysin (PFO) from Clostridium perfringens, streptolysin O from Streptococcus pyogenes and pneumolysin from Streptococcus pneumoniae. LLO is composed of 529 residues and possesses at its N terminus a 25 residue typical signal sequence (Mengaud et al., 1988
). The protein is secreted into the culture supernatant as a monomer (Geoffroy et al., 1989
). The three-dimensional structure of LLO is currently unknown but that of monomeric PFO has been determined at 2·7
by X-ray crystallography (Rossjohn et al., 1997
). The molecule, which comprises 500 residues, is elongated and is composed of four domains that are rich in ß-sheet structures. Domain 2 is connected to domain 4 through a glycine linker (at residue 392). The autonomous domain 4 folds into a compact ß sandwich. Electron microscopy data showed that the PFO monomer is L shaped with four domains of equal size with one end, domain 4 (d4), flexibly linked to the three others.
A classical approach to understanding how the different regions of a polypeptide interact to stabilize the secondary, tertiary and quaternary structures of the native conformation consists of expressing these regions independently. Fragment complementation has been performed either with fragments produced by limited proteolysis or chemical cleavage, or with incomplete polypeptide chains expressed independently by genetic manipulations. The in vivo assembly of functional proteins from complementing fragments has been demonstrated for several proteins of Gram-negative bacteria, including integral membrane proteins (Bibi & Kaback, 1990 ) and soluble cytoplasmic, periplasmic or secreted proteins (Betton & Hofnung, 1994
; Diep et al., 1998
; Shiba & Schimmel, 1992
).
Taking advantage of the sequence similarities between LLO and PFO, we elaborated a theoretical 3D model of LLO folding, and engineered truncated and modified LLO proteins and expressed them in L. monocytogenes. We found that, when secreted simultaneously, two truncated proteins comprising the proximal (LLO-d123) and distal (LLO-d4) portions of LLO could reassemble to form an active molecule. In contrast, in-frame insertions in the region connecting these two domains drastically altered protein stability and abolished cytolytic activity.
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METHODS |
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Constructions.
PCR-amplified fragments were first cloned into pCR (Invitrogen). All the recombinant genes were finally subcloned into the high-copy-number Gram-negative/Gram-positive shuttle plasmid pAT28 (Trieu-Cuot et al., 1990 ). The wild-type hly gene preceded by its upstream promoter region, cloned into pAT28, has been described previously (Dubail et al., 2000
).
LLO-d123.
The truncated hly gene encoding the first 416 residues of LLO and its upstream promoter region were amplified by PCR using primers 1 (5'-CCGGATCCCTTAAAGTGACTTTTATGTTGAGGCA-3') and 2 (5'-CCCTGCAGTTAATCTGTATAAGCTTTTGAAGTTGTTTC-3'). The amplified product was first cloned into pTCV-lac and then subcloned into pAT28, yielding plasmid pAT28-LLO-d123. The corresponding protein was named LLO-d123.
LLO-d4.
To allow secretion of the predicted d4 domain of LLO into the culture medium, we fused residues 415529 to the proximal portion of the protein (residues 135) which contains the putative signal sequence. Two pairs of oligonucleotides were used for PCR amplification. The first pair of primers was 5'-GCTCTAGATCTCTTAAAGTGACTTTTATGTT- GAGGCA-3' (primer 1) and 5'-GGGAATTCCATATGTGAATTTTCTTTATTGAATGCAGATGCATCCTTTGCTTCAGTTTG 3' (primer 2). An NdeI site (underlined) was created at the 3' end directly after the triplet encoding residue 35. The second pair of primers was 5'-GGCCATATGACAGATGGAAAAATTAACATCGATCACTCT-3' (primer 1) and 5'-CCCTGCAGACAATTATTCGATTGGATTATCTAC-3' (primer 2). The two amplified fragments were cloned into pCR. An XbaIKpnI fragment, comprising the proximal part of hly, was first subcloned into the XbaIKpnI sites of pAT28 (yielding pAT28-LLO:135). Then, an NdeIKpnI fragment comprising the distal part of hly was inserted into the NdeIKpnI sites of pAT28-LLO:135, yielding plasmid pAT28-LLO-d4. The resulting protein was named LLO-d4.
LLO-lL.
A DNA fragment containing the promoter region of hly and the portion encoding domains d123 (residues 1 to 416) followed by a linker encoding the hexapeptide GGSGGS and an NdeI site was amplified by PCR using the primers 5'-GCTCTAGATCTCTTAAAGTGACTTTTATGTTGAGGCA-3' (primer 1) and 5'-GGCATATGGGATCCTCCGGATCCTCCATCTGTATAAGCTTTTGAAGTTGTTTCAATATATTCTGAG-3' (primer 2). The NdeI site was designed to be in-frame with the NdeI site present in construction pAT28-LLO-d4 (see above). The amplified fragment was first cloned into the pCR cloning vector. Then the XbaINdeI fragment encoding LLO-d123 was substituted for the XbaINdeI fragment of plasmid pAT28-LLO-d4. The sequence of the modified hinge region thus corresponds to the insertion of the nonapeptide GGSGGSHMTD flanked on both sides by the predicted original G417 linker.
LLO-lS.
A DNA fragment containing the promoter region of hly and the portion encoding residues 1 to 414 of d123, followed by a NdeI restriction site was first amplified by PCR with the primers 5'-GCTCTAGATCTCTTAAAGTGACTTTTATGTTGAGGCA-3' (primer 1) and 5'-CCCATATGATAAGCTTTTGAAGTTGTTTCAATATA-3' (primer 2). The amplified fragment was first cloned into the pCR cloning vector. Then the XbaINdeI fragment encoding LLO-d123 was substituted for the XbaINdeI fragment of plasmid pAT28-LLO-d4. The resulting modified protein thus corresponds to the insertion of the dipeptide HM between residues 414 and 415 of LLO.
Simultaneous expression of LLO-d123 and LLO-d4.
First, the gene encoding LLO-d123 was integrated into the chromosome of EGDhly. For that, we used the integrative vector pAT113 (Trieu-Cuot et al., 1991
; Autret et al., 2001
). Integration of pAT113 in the chromosome of Gram-positive bacteria requires the presence of the transposon-encoded integrase in the recipient. Therefore EGD
hly was transformed with plasmid pAT145, carrying the transposon-encoded integrase Int-Tn. The BamHISalI fragment of plasmid pAT28-LLO-d123 was cloned into the BamHISalI sites of vector pAT113 and the resulting plasmid (pAT113-LLO-d123) was transferred into EGD
hly/pAT145 by electroporation. One transformant, corresponding to a single chromosomal insertion (and showing normal growth capacities; not shown) was reserved. Finally, plasmid pAT28-LLO-d4, encoding LLO-d4, was introduced into this strain by electroporation.
Protein preparation and analysis
Protein preparation.
The LLO proteins were prepared from supernatants of EGDhly transformed with the different pAT28 derivatives. For each mutant, 25 ml of an 8 h culture at 37 °C in BHI-spectinomycin broth were added to 500 ml RPMI 1640 minimal medium containing glucose (3% final). The suspension was grown overnight with agitation at 37 °C (under these conditions, the cultures corresponded to
2x108 bacteria ml-1). After centrifugation, cell-free supernatants were filtered through a 0·22 mm pore size Millipore filter. The filtered supernatants were first concentrated to 15 ml by tangential flux through miniplate YM30 (Millipore) with a cut-off of 30 kDa (except for LLO-d4 and LLO-d123+LLO-d4 expressed simultaneously, and the negative control EGD
hly, for which a cut-off of 10 kDa was used). Supernatants were further concentrated to a final volume of 1 to 1·5 ml by centrifugation through ultrafree Biomax units. The total protein concentration of the preparations, determined by the Bradford colorimetric method, was
1 mg ml-1.
Western-blot analysis.
Ten microlitres of each concentrated supernatant were loaded per well onto SDS13% polyacrylamide gels. SDS-PAGE and Western-blot analyses were performed as described previously (Charbit et al., 2000 ) with polyclonal and monoclonal antibodies (see below).
Quantification.
The amounts of LLO present in the concentrated supernatants were determined by dot-blot assays. Serial twofold dilutions of each preparation were coated onto nitrocellulose sheets and detection was carried out with monoclonal (mAb SE2) or polyclonal anti-LLO antibodies. Serial twofold dilutions of purified LLOwt were used as standards in the assay (starting from 1 µg protein). The amounts of LLO detected in each spot were quantified by densitometry scanning of the nitrocellulose sheet using the NIH image software version 1.61. They varied between 1% and 5% of total proteins. On this basis (i.e. 10 to 50 µg LLO for 1011 bacteria), the number of molecules of LLO produced under our growth conditions corresponds to 1035x103 per bacterium, which is in agreement with previously reported values (Geoffroy et al., 1989
; Villanueva et al., 1995
). The final concentration of LLO in each protein preparation was finally adjusted to 0·5 µg µl-1.
Antibodies.
A polyclonal anti-LLO serum, raised in rabbits against denatured LLO (Geoffroy et al., 1989 ), was used in Western blots and dot blots at a final dilution of 1/1000 and in immunofluorescence at a final dilution of 1/500. A monoclonal anti-LLO antibody, mAb SE2 (kindly provided by Dr A. J. Ainsworth, Veterinary Medical Research, College of Veterinary Medicine, Mississippi State University, MS, USA), raised in mice after injection of concentrated L. monocytogenes extracellular proteins (Erdenlig et al., 1999
), was used in Western blots and dot blots at a final dilution of 1/1000 and in immunofluorescence at a final dilution of 1/100.
A monoclonal anti-pneumolysin antibody, mAb PLY-5 (kindly provided by Dr J. R. de los Toyos, Area de Immunologia, Facultad de Medicina, Universidad de Oviedo, Spain), was used in the membrane-binding assay to inhibit binding of LLO-d4 to erythrocyte membranes. This mAb was previously shown to recognize a peptide within the conserved undecapeptide at the tip of the d4 domains of thiol-activated cytolysins, including LLO (Jacobs et al., 1999 ).
Immunoprecipitation.
Concentrated supernatants were first incubated with mAb SE2 (25 µl antibody in 250 µl final) and then immunoprecipitated with 50 µl protein Aagarose (25 µl bed volume from a suspension at 3 mg ml-1; Boehringer Mannheim). For LLO-d123+LLO-d4 expressed simultaneously (denoted 123+4) or separately (denoted 123+4rec), 2 µg LLO were used (1 µg LLO-d123+1 µg LLO-d4). Protein preparations of either LLO-d123 or LLO-d4 alone were used as negative controls (1 µg). After electrophoresis, immunoprecipitated proteins were transferred onto nitrocellulose membranes. The LLO proteins were finally revealed using the rabbit polyclonal anti-LLO serum (1/1000 final). The assay was repeated twice.
Haemolysis.
Haemolytic phenotypes were visualized by spreading bacteria onto horse-blood agar plates (bioMérieux). Haemolytic activity was measured at pH 6·6 as described previously (Geoffroy et al., 1989 ; Portnoy et al., 1988
). Serial twofold dilutions of filtered supernatants from BHI-grown bacteria (starting from 20 µl supernatant) were incubated with 50 µl horse red blood cells (HRBC) at OD541 0·2, in a final volume of 100 µl. The haemolytic activity was estimated as the reciprocal of the dilution giving 50% haemolysis.
The haemolytic activity of the concentrated supernatants was also measured at pH 6·6. Serial twofold dilutions of each LLO preparation, starting from 1 µg LLO in the first well, were tested at different concentrations of HRBC (OD541 0·2, 0·1 or 0·05).
Membrane-binding assays
Binding to erythrocyte membranes.
Horse erythrocytes (50 ml) were lysed by sonication (on ice) with 20 mM MgCl2. After sonication, unbroken cells were removed by low-speed centrifugation (1000 g for 20 min). The supernatant was dialysed overnight at 4 °C in PBS/20 mM MgCl2. Dialysed membranes were concentrated by ultracentrifugation at 23500 g for 1 h and finally resuspended in PBS/20 mM MgCl2 at a concentration of 0·4 g l-1 (final).
Each concentrated supernatant, containing 5 µg LLO, was incubated with 80 µg erythrocyte membranes (unless otherwise stated) in a final volume of 1 ml for 30 min at room temperature. The mixture was then centrifuged at 23500 g for 1 h at 4 °C. The pellets containing the membrane were solubilized in SDS-PAGE loading buffer. Toxins from the supernatant were recovered by TCA precipitation. Each fraction (pellet or supernatant) was finally concentrated by centrifugation at 23500 g for 1 h at 4 °C. After electrophoresis, proteins were transferred onto nitrocellulose membranes. The LLO proteins were finally revealed using the rabbit polyclonal anti-LLO serum (1/1000 final). The percentage of LLO in the bound (pellet) and unbound (supernatant) fractions was determined by densitometry scanning of the nitrocellulose sheet.
Inhibition of LLO-d4 binding was performed after a pre-incubation with 1 µl mAb PLY-5 for 30 min at 37 °C (5 µg LLO in a final volume of 100 µl).
Binding to intact cells.
We used the epithelial cell line HeclB, a human endometrial adenocarcinoma cell line (obtained from the American Type Culture Collection, Manassas, VA, USA). Cells were cultured in Dulbeccos Modified Eagles Medium (DMEM; Gibco), containing 10% foetal bovine serum at 37 °C under 5% CO2. Cells were seeded at 8x104 cells cm-2 onto 12 mm diameter glass coverslips in 24-well plates. Monolayers were used 24 h after seeding.
Purified LLO-d123 and LLO-d4 were used in this assay. Binding of LLO-d123 and LLO-d4 to the surface of intact cells was detected by confocal microscopy, using either anti-LLO mAb SE2 for LLO-d123 or the polyclonal serum for LLO-d4. HeclB cells were incubated with 1 µg LLO d123 or LLO-d4 for 30 min at 37 °C. After three washes with PBS, cells were fixed with 3% (w/v) paraformaldehyde (in PBS) for 30 min and washed three times with PBS. Cells were then processed for fluorescence labelling. Cells were incubated sequentially with either mAb SE2 diluted 1/100 and then with CY3-labelled anti-mouse IgG (Jackson Immunoresearch Laboratories) diluted 1/200, or with the polyclonal serum diluted 1/500 and then with anti-rabbit IgG coupled to Alexa 546 diluted 1/200. Incubations were carried out for 30 min at room temperature and followed by three washings in PBS. Coverslips were mounted on slides and examined by fluorescence microscopy with a Leica TCS4D confocal laser scanning microscope. Each assay was repeated several times.
Sequence searches, alignments and graphics.
Similarity searches were done via the internet with BLAST software (Altschul et al., 1997 ). The theoretical 3D model of LLO folding was produced using the Automated Comparative Protein Modelling Server (available at http://www.expasy.ch/swissmod/SWISS-MODEL.html) (Guex et al., 1999
and references therein) and the RasMac Program (Version RasMol v2.6, 1994. Available from Roger Sayle, Biomolecular Structure, Glaxo Research and Development, Greenford, UK), using the X-ray coordinates of PFO.
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RESULTS |
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Expression of the LLO mutant proteins in L. monocytogenes
Two truncated LLO proteins corresponding to domains 1, 2 and 3 (LLO-d123) or domain 4 (LLO-d4), and two linker-insertion mutants corresponding to in-frame insertion of two (LLO-lS) or ten residues (LLO-lL) at amino acid site 416, were constructed (Fig. 1b). The recombinant proteins were expressed in EGD
hly from plasmid-borne genes. EGD
hly expressing LLOwt was used as a positive control (Dubail et al., 2000
).
The four LLO mutant proteins were efficiently secreted into the culture supernatant of L. monocytogenes. As shown in the Western blot of Fig. 2 (a), LLO-d123 and LLO-d4 were detected by the polyclonal anti-LLO antibody in concentrated culture supernatants. Strikingly, for the two linker mutants (LLO-lS and LLO-lL), two major degradation species were recognized. The apparent migration of the two species detected (
50 and 15 kDa, respectively) is compatible with a cleavage of the mutant proteins in the vicinity of the linker insertion that would generate peptides corresponding to d123 and d4 domains. The monoclonal anti-LLO antibody (mAb SE2) recognized LLOwt and LLO-d123 but failed to recognize LLO-d4, indicating that its recognition site lies within the proximal 2/3 portion of the molecule (Fig. 2b
). mAb SE2 could also recognize the mature form of the two linker-insertion mutants as well as a species of
50 kDa. The fact that the 15 kDa species, detected with the polyclonal serum, was not detected by the mAb (not shown) further supports the idea that this peptide might correspond to the d4 domain of LLO.
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Binding of LLO-d123 and LLO-d4 to eukaryotic membranes
Wild-type LLO is known to bind to cholesterol-containing membranes (Jacobs et al., 1998 ). The ability of the mutant proteins to bind to eukaryotic cell membranes was tested on erythrocyte membranes and on intact eukaryotic cells.
Binding to erythrocyte membranes. This was assayed essentially as described by Jacobs et al. (1998) on concentrated supernatants. Each protein preparation, containing 5 µg LLO, was incubated with erythrocyte membranes (80 µg ml-1 final). After 30 min incubation at room temperature, the membranes were collected by centrifugation and subjected to SDS-PAGE (see Methods). The LLO fraction bound to the membranes was revealed by Western blotting with anti-LLO polyclonal serum. As shown in Fig. 4(a)
), the four LLO mutant proteins (including LLO-d123 and LLO-d4 expressed simultaneously) bound to erythrocyte membranes. Strikingly, with both linker-insertion mutants, the two major proteolytic species were still detected. For LLO-d123,
50% of the protein was detected in the membrane fraction and 50% in the supernatant: a value comparable to that obtained with LLOwt. In contrast, for LLO-d4, the bound fraction corresponded to only 1520% (as determined by densitometry scanning of the nitrocellulose sheet using the NIH image software version 1.61).
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Two sets of controls were performed to check the specificity of the binding. i) The assay was carried out without membranes. Under these conditions, the two truncated and the two-linker insertion mutant proteins were essentially found in the supernatant. ii) The assay was performed with a heterologous protein, i.e. the maltose-binding protein (MBP) of Escherichia coli K-12 (Betton & Hofnung, 1994 ). MBP was essentially found in the supernatant (not shown). These two assays demonstrated that the presence of the LLO mutant proteins in the membrane fractions (pellets) was not due to the formation of non-specific protein aggregates and confirmed that the binding was specific.
Preincubation of LLO-d4 with mAb PLY-5 (directed against the d4 domain) almost totally inhibited the binding of LLO-d4 to erythrocyte membranes (Fig. 4c). This result is in agreement with previous data suggesting that binding of LLO involved directly its conserved motif, located in the d4 domain. However, it cannot be excluded that inhibition might be due to steric hindrance by the mAb and that membrane binding occurs via another portion of the LLO-d4 polypeptide.
Binding to intact cells. Binding of LLO-d123 and LLO-d4 to the surface of intact eukaryotic cells was then evaluated by confocal microscopy on human epithelial cells, using mAb SE2 (directed against this proximal portion of the molecule). One microgram of purified protein was used (Fig. 3b). As shown in Fig. 5(a)
, a strong and uniformly distributed binding of LLO-d123 protein was observed on all the cells. As a negative control, cells were preincubated with a supernatant from EGD
hly alone and we did not observe any non-specific binding of the monoclonal antibody to cell membranes (Fig. 5b
). Under the conditions of the assay, using mAb SE2, only a very weak labelling was observed when cells were preincubated with the LLO linker derivatives (not shown). The binding of LLO-d4 to intact cells was tested using the polyclonal anti-LLO serum. As shown in Fig. 5(c)
, binding of LLO-d4 was also uniformly distributed on the surface of all the cells. As a negative control, the polyclonal serum was tested on cells that were not preincubated with LLO: only a very weak background signal was recorded under these conditions (Fig. 5d
).
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Co-immunoprecipitation of LLO-d4 with LLO-d123. A concentrated supernatant from EGDhly expressing simultaneously the two polypeptides was incubated with mAb SE2 (specific for LLO-d123). The antigen-antibody complexes were immunoprecipitated with protein Aagarose and subjected to SDS-PAGE. After electrophoresis, proteins were transferred onto nitrocellulose membranes and the LLO proteins were revealed with rabbit polyclonal anti-LLO (recognizing both LLO-d123 and LLO-d4). As expected, LLO-d123 was efficiently immunoprecipitated by mAb SE2 and revealed by the polyclonal serum (Fig. 6a
). There was not any non-specific reaction with LLO-d4. In the concentrated supernatant containing both polypeptides (123+4, simultaneous expression), LLO-d4 was co-immunoprecipitated with LLO-d123, demonstrating that the proteins were able to interact physically. In contrast, when LLO-d123 and LLO-d4 produced separately were mixed (123+4rec), only LLO-d123 was immunodetected.
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These data, which demonstrated that LLO-d123 and LLO-d4 proteins could reassemble to form a haemolytically active LLO when expressed simultaneously, suggest that functional assembly of the two domains might occur during, or immediately after, the co-secretion of the two proteins.
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DISCUSSION |
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LLO-d123 and LLO-d4 bind to cell membranes and can reassemble to form an active molecule
Earlier studies have shown that a C-terminal proteolytic fragment of PFO (residues 304500) could bind erythrocytes like intact toxin (Tweten et al., 1991 ), and biophysical studies clearly established that domain 4 of PFO interacted with the bilayer (Nakamura et al., 1998
). Further biophysical analyses revealed that each PFO monomer contained a second region (domain 3), involved in pore formation (Shatursky et al., 1999
; Shepard et al., 1998
). A structural linkage between domains 3 and 4 of PFO was very recently demonstrated (Heuck et al., 2000
). That study showed that the two domains interacted sequentially with cholesterol-containing membranes: domain 4 interacted first, eliciting a conformational change in the proximal part of the molecule, allowing subsequent insertion of domain 3.
In agreement with these data, the present work revealed that LLO-d4 and LLO-d123, expressed individually, could bind to erythrocyte membranes and to intact eukaryotic cells. The binding capacity of the LLO-d123 alone might be due to a direct accessibility of the membrane-binding domain at the surface of the molecule. Further analyses will be required to determine the exact mode of interaction of this truncated protein with the membrane and of its oligomeric state in solution and upon binding to cell membranes.
LLO-d123 and LLO-d4, when secreted simultaneously by the same bacterium, could reassemble to form a haemolytically active molecule. Physical interactions between the two portions of LLO were confirmed by showing that the d4 domain could be immunoprecipitated with an antibody specifically directed against d123.
We also tested whether the expression of LLO-d123 could interfere with the activity of the chromosomally encoded wild-type protein (negative dominance). For that, we transformed EGDwt with the multicopy plasmid pAT28-LLO-d123 encoding LLO-d123. The haemolytic activity recorded in the culture supernatant of the recombinant strain was identical to that of the wild-type strain, indicating that the expression of the truncated polypeptide did not prevent pore formation (not shown).
Most of the mutations in LLO (or in PFO) affecting haemolytic activity identified so far were found in the d4 domain, within or in close vicinity to the conserved undecapeptide thought to be involved in cell binding (Michel et al., 1990 ; Jones et al., 1996
). We have shown here that modifications in the region connecting d123 and d4 were also critical for the haemolytic activity of LLO, by increasing the susceptibility of the protein to proteolytic degradation. These data are in agreement with the currently accepted notion that d123 and d4 interact together to form an active cytolysin molecule (Palmer et al., 1998
; Shepard et al., 1998
, 2000
; Ghani et al., 1999
; Heuck et al., 2000
). One hypothesis would be that the hinge region might be involved in this interaction. Biochemical and biophysical analyses of the truncated proteins should provide interesting information on the ability of the LLO protein domains to fold autonomously.
Finally, these data showed that the hinge region of LLO is not a permissive region of the molecule. In this respect, it is worth mentioning that a region close to the N terminus of LLO was very recently found to be critical for bacterial virulence (Decatur & Portnoy, 2000 ; Lety et al., 2001
). This region is involved in phagosomal escape and might also regulate the intra-cytosolic half life of LLO. Strikingly, this proximal region of LLO allowed local sequence alterations without any detectable effect on protein secretion and haemolytic activity. At this stage, it is tempting to speculate that the hinge region between d123 and d4 of LLO could also be a target of proteolytic degradation in infected cells.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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---|
Altschul, S. F., Madden, T. L., Schaffer, A. A., Zhang, J., Zhang, Z., Miller, W. & Lipman, D. J. (1997). Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25, 3389-3402.
Autret, N., Dubail, I., Trieu-Cuot, P., Berche, P. & Charbit, A. (2001). Identification of new genes involved in the virulence of Listeria monocytogenes by signature-tagged transposon mutagenesis. Infect Immun 69, 2054-2065.
Bayley, H. (1997). Toxin structure: part of a hole? Curr Biol 7, R763-R767.[Medline]
Berche, P., Gaillard, J. L. & Sansonetti, P. J. (1987). Intracellular growth of Listeria monocytogenes as a prerequisite for in vivo induction of T cell-mediated immunity. J Immunol 138, 2266-2271.
Betton, J. M. & Hofnung, M. (1994). In vivo assembly of active maltose binding protein from independently exported protein fragments. EMBO J 13, 1226-1234.[Abstract]
Bibi, E. & Kaback, H. R. (1990). In vivo expression of the lacY gene in two segments leads to functional lac permease. Proc Natl Acad Sci USA 87, 4325-4329.[Abstract]
Chakraborty, T., Leimeister-Wachter, M., Domann, E., Hartl, M., Goebel, W., Nichterlein, T. & Notermans, S. (1992). Coordinate regulation of virulence genes in Listeria monocytogenes requires the product of the prfA gene. J Bacteriol 174, 568-574.[Abstract]
Charbit, A., Andersen, C., Wang, J., Schiffler, B., Michel, V., Benz, R. & Hofnung, M. (2000). In vivo and in vitro studies of transmembrane beta-strand deletion, insertion or substitution mutants of the Escherichia coli K-12 maltoporin. Mol Microbiol 35, 777-790.[Medline]
Decatur, A. L. & Portnoy, D. A. (2000). A PEST-like sequence in listeriolysin O essential for Listeria monocytogenes pathogenicity. Science 290, 992-995.
Diep, D. B., Lawrence, T. S., Ausio, J., Howard, S. P. & Buckley, J. T. (1998). Secretion and properties of the large and small lobes of the channel-forming toxin aerolysin. Mol Microbiol 30, 341-352.[Medline]
Dramsi, S., Biswas, I., Maguin, E., Braun, L., Mastroeni, P. & Cossart, P. (1995). Entry of Listeria monocytogenes into hepatocytes requires expression of InIB, a surface protein of the internalin multigene family. Mol Microbiol 16, 251-261.[Medline]
Drevets, D. A., Sawyer, R. T., Potter, T. A. & Campbell, P. A. (1995). Listeria monocytogenes infects human endothelial cells by two distinct mechanisms. Infect Immun 63, 4268-4276.[Abstract]
Dubail, I., Berche, P., The European Listeria Genome Consortium & Charbit, A. (2000). Listeriolysin O as a reporter to identify constitutive and in vivo-inducible promoters in the pathogen Listeria monocytogenes. Infect Immun 68, 32423250.
Erdenlig, S., Ainsworth, A. J. & Austin, F. W. (1999). Production of monoclonal antibodies to Listeria monocytogenes and their application to determine the virulence of isolates from channel catfish. Appl Environ Microbiol 65, 2827-2832.
Gaillard, J. L., Berche, P. & Sansonetti, P. (1986). Transposon mutagenesis as a tool to study the role of hemolysin in the virulence of Listeria monocytogenes. Infect Immun 52, 50-55.[Medline]
Gaillard, J. L., Berche, P., Mounier, J., Richard, S. & Sansonetti, P. (1987). In vitro model of penetration and intracellular growth of Listeria monocytogenes in the human enterocyte-like cell line Caco-2. Infect Immun 55, 2822-2829.[Medline]
Gaillard, J. L., Jaubert, F. & Berche, P. (1996). The inlAB locus mediates the entry of Listeria monocytogenes into hepatocytes in vivo. J Exp Med 183, 359-369.[Abstract]
Geoffroy, C., Gaillard, J. L., Alouf, J. E. & Berche, P. (1989). Production of thiol-dependent haemolysins by Listeria monocytogenes and related species. J Gen Microbiol 135, 481-487.[Medline]
Ghani, E., Weis, S., Walev, I., Kehoe, M., Bhakdi, S. & Palmer, M. (1999). Streptolysin O: inhibition of the conformational change during membrane binding of the monomer prevents oligomerization and pore formation. Biochemistry 38, 15204-15211.[Medline]
Guex, N., Diemand, A. & Peitsch, M. C. (1999). Protein modelling for all. Trends Biochem Sci 24, 364-367.[Medline]
Guzman, C. A., Rohde, M., Chakraborty, T., Domann, E., Hudel, M., Wehland, J. & Timmis, K. N. (1995). Interaction of Listeria monocytogenes with mouse dendritic cells. Infect Immun 63, 3665-3673.[Abstract]
Heuck, A. P., Hotze, E. M., Tweten, R. K. & Johnson, A. E. (2000). Mechanism of membrane insertion of a multimeric ß-barrel protein: perfringolysin O creates a pore using ordered and coupled conformational changes. Mol Cell 6, 1233-1242.[Medline]
Jacobs, T., Darji, A., Frahm, N., Rohde, M., Wehland, J., Chakraborty, T. & Weiss, S. (1998). Listeriolysin O: cholesterol inhibits cytolysis but not binding to cellular membranes. Mol Microbiol 28, 1081-1089.[Medline]
Jacobs, T., Cima-Cabal, M. D., Darji, A. & 7 other authors (1999). The conserved undecapeptide shared by thiol-activated cytolysins is involved in membrane binding. FEBS Lett 459, 463466.[Medline]
Jones, S., Preiter, K. & Portnoy, D. A. (1996). Conversion of an extracellular cytolysin into a phagosome-specific lysin which supports the growth of an intracellular pathogen. Mol Microbiol 21, 1219-1225.[Medline]
Kathariou, S., Metz, P., Hof, H. & Goebel, W. (1987). Tn916-induced mutations in the hemolysin determinant affecting virulence of Listeria monocytogenes. J Bacteriol 169, 1291-1297.[Medline]
Kuhn, M. & Goebel, W. (1989). Identification of an extracellular protein of Listeria monocytogenes possibly involved in intracellular uptake by mammalian cells. Infect Immun 57, 55-61.[Medline]
Leimeister-Wachter, M., Haffner, C., Domann, E., Goebel, W. & Chakraborty, T. (1990). Identification of a gene that positively regulates expression of listeriolysin, the major virulence factor of Listeria monocytogenes. Proc Natl Acad Sci USA 87, 8336-8340.[Abstract]
Lety, M. A., Frehel, C., Dubail, I., Beretti, J. L., Kayal, S., Berche, P. & Charbit, A. (2001). Identification of a PEST-like motif in listeriolysin O required for phagosomal escape and for virulence of Listeria monocytogenes. Mol Microbiol 39, 1124-1140.[Medline]
Mackaness, G. B. (1962). Cellular resistance to infection. J Exp Med 116, 381-406.
Mengaud, J., Vicente, M. F., Chenevert, J., Pereira, J. M., Geoffroy, C., Gicquel-Sanzey, B., Baquero, F., Perez-Diaz, J. C. & Cossart, P. (1988). Expression in Escherichia coli and sequence analysis of the listeriolysin O determinant of Listeria monocytogenes. Infect Immun 56, 766-772.[Medline]
Michel, E., Reich, K. A., Favier, R., Berche, P. & Cossart, P. (1990). Attenuated mutants of the intracellular bacterium Listeria monocytogenes obtained by single amino acid substitutions in listeriolysin O. Mol Microbiol 4, 2167-2178.[Medline]
Nakamura, M., Sekino-Suzuki, N., Mitsui, K. I. & Ohno-Iwashita, Y. (1998). Contribution of tryptophan residues to the structural changes in perfringolysin O during interaction with liposomal membranes. J Biochem 123, 1145-1155.[Abstract]
Palmer, M., Harris, R., Freytag, C., Kehoe, M., Tranum-Jensen, J. & Bhakdi, S. (1998). Assembly mechanism of the oligomeric streptolysin O pore: the early membrane lesion is lined by a free edge of the lipid membrane and is extended gradually during oligomerization. EMBO J 17, 1598-1605.
Park, S. F. & Stewart, G. S. (1990). High efficiency transformation of Listeria monocytogenes by electroporation of penicillin-treated cells. Gene 94, 129-132.[Medline]
Portnoy, D. A., Jacks, P. S. & Hinrichs, D. J. (1988). Role of hemolysin for the intracellular growth of Listeria monocytogenes. J Exp Med 167, 1459-1471.[Abstract]
Renzoni, A., Cossart, P. & Dramsi, S. (1999). PrfA, the transcriptional activator of virulence genes, is upregulated during interaction of Listeria monocytogenes with mammalian cells and in eukaryotic cell extracts. Mol Microbiol 34, 552-561.[Medline]
Rossjohn, J., Fell, S. C., McKinstry, W. J., Tweten, R. K. & Parker, M. W. (1997). Structure of a cholesterol-binding, thiol-activated cytolysin and a model of its membrane form. Cell 89, 685-692.[Medline]
Shatursky, O., Heuck, A. P., Shepard, L. A., Rossjohn, J., Parker, M. W., Johnson, A. E. & Tweten, R. K. (1999). The mechanism of membrane insertion for a cholesterol-dependent cytolysin: a novel paradigm for pore-forming toxins. Cell 99, 293-299.[Medline]
Sheehan, B., Kocks, C., Dramsi, S., Gouin, E., Klarsfeld, A. D., Mengaud, J. & Cossart, P. (1994). Molecular and genetic determinants of the Listeria monocytogenes infectious process. Curr Top Microbiol Immunol 192, 187-216.[Medline]
Shepard, L. A., Heuck, A. P., Hamman, B. D., Rossjohn, J., Parker, M. W., Ryan, K. R., Johnson, A. E. & Tweten, R. K. (1998). Identification of a membrane-spanning domain of the thiol-activated pore-forming toxin Clostridium perfringens perfringolysin O: an alpha-helical to beta-sheet transition identified by fluorescence spectroscopy. Biochemistry 37, 14563-14574.[Medline]
Shepard, L. A., Shatursky, O., Johnson, A. E. & Tweten, R. K. (2000). The mechanism of pore assembly for a cholesterol-dependent cytolysin: formation of a large prepore complex precedes the insertion of the transmembrane beta-hairpins. Biochemistry 39, 10284-10293.[Medline]
Shiba, K. & Schimmel, P. (1992). Functional assembly of a randomly cleaved protein. Proc Natl Acad Sci USA 89, 1880-1884.[Abstract]
Trieu-Cuot, P., Carlier, C., Poyart-Salmeron, C. & Courvalin, P. (1990). A pair of mobilizable shuttle vectors conferring resistance to spectinomycin for molecular cloning in Escherichia coli and in Gram-positive bacteria. Nucleic Acids Res 18, 4296.[Medline]
Trieu-Cuot, P., Carlier, C., Poyart-Salmeron, C. & Courvalin, P. (1991). An integrative vector exploiting the transposition properties of Tn1545 for insertional mutagenesis and cloning of genes from gram-positive bacteria. Gene 106, 21-27.[Medline]
Tweten, R. K., Harris, R. W. & Sims, P. J. (1991). Isolation of a tryptic fragment from Clostridium perfringens theta-toxin that contains sites for membrane binding and self-aggregation. J Biol Chem 266, 12449-12454.
Villanueva, M. S., Sijts, A. J. & Pamer, E. G. (1995). Listeriolysin is processed efficiently into an MHC class I-associated epitope in Listeria monocytogenes-infected cells. J Immunol 155, 5227-5233.[Abstract]
Received 2 April 2001;
revised 4 June 2001;
accepted 13 June 2001.