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J. Biol. Chem., Vol. 279, Issue 17, 17834-17841, April 23, 2004
Solution Structure of the Pore-forming Protein of Entamoeba histolytica*![]() ![]() ¶![]() ![]() ![]() ![]() ![]()
From the
Received for publication, November 30, 2003 , and in revised form, February 9, 2004.
Amoebapore A is a 77-residue protein from the protozoan parasite and human pathogen Entamoeba histolytica. Amoebapores lyse both bacteria and eukaryotic cells by pore formation and play a pivotal role in the destruction of host tissues during amoebiasis, one of the most life-threatening parasitic diseases. Amoebapore A belongs to the superfamily of saposin-like proteins that are characterized by a conserved disulfide bond pattern and a fold consisting of five helices. Membrane-permeabilizing effector molecules of mammalian lymphocytes such as porcine NK-lysin and the human granulysin share these structural attributes. Several mechanisms have been proposed to explain how saposin-like proteins form membrane pores. All mechanisms indicate that the surface charge distribution of these proteins is the basis of their membrane binding capacity and pore formation. Here, we have solved the structure of amoebapore A by NMR spectroscopy. We demonstrate that the specific activation step of amoebapore A depends on a pH-dependent dimerization event and is modulated by a surface-exposed histidine residue. Thus, histidine-mediated dimerization is the molecular switch for pore formation and reveals a novel activation mechanism of pore-forming toxins.
The protozoan parasite, Entamoeba histolytica, inhabits the colon of infected humans. It is the causative agent of human amoebiasis that often leads to tissue damage, colitis, and extraintestinal abscesses (1). Amoebiasis is the second leading cause of death from parasitic diseases worldwide (2). About 50 million people suffer from invasive amoebiasis, of whom up to 100,000 die annually (3).
In the amoebic trophozoite, several factors have been identified that are involved in pathogenesis. In addition to a galactose-/N-acetylgalactosamine-specific lectin on the amoebic surface that mediates adhesion to colonic mucus and host cells (4) and secreted cysteine proteinases that disintegrate tissues by cleaving extracellular matrix proteins (5), a family of membrane-active polypeptides have been discovered (6). These polypeptides exist as three isoforms and are named amoebapore A, B, and C, respectively. They are capable of lysing a broad spectrum of target cells, including human host cells and bacteria. It has been recently shown that trophozoites of E. histolytica lacking amoebapore A, due to transcriptional silencing of the encoding gene, became avirulent (7), demonstrating that this protein is a key pathogenicity factor of the parasite. All three amoebapore isoforms have been isolated and biochemically characterized, and their primary structure has been elucidated by molecular cloning of the genes of their precursors (8-10). The mature proteins consist of 77 amino acid residues each and are localized within cytoplasmic granules. The overall sequence identity between the three amoebapores is between 35 and 57% (10). Despite the substantial sequence divergence, they possess a characteristic disulfide bond pattern and a single conserved C-terminal histidine residue. The secondary structure of the major isoform A has been determined to be exclusively By sequence similarity, amoebapores have been grouped into the family of saposin-like proteins (SAPLIP).1 Although the members of the SAPLIP family have different biological functions, they are all able to interact with lipids. With one exception, all of them possess six conserved cysteine residues that are involved in forming the disulfide bond pattern characteristic of this protein family (12). Despite the enormous evolutionary distance, amoebapores reveal a substantial sequence similarity with membrane-permeabilizing effector molecules of mammalian lymphocytes such as porcine NK-lysin and human granulysin. Like their amoebic counterparts, the mammalian proteins reside in intracellular granules and are able to lyse bacteria and eukaryotic cells (10, 13, 14). Based on structural information and biochemical data, several mechanisms have been suggested to explain how saposin-like proteins may form membrane pores (15, 16). For both NK-lysin and granulysin, it has been suggested that binding of the monomeric proteins to the membrane via their charged epitopes results in destabilization of the lipid bilayer (17, 18), whereas amoebapore A has been proposed to act via the barrel stave mechanism (19).
Here, we present the high resolution three-dimensional structure of monomeric amoebapore A solved by NMR spectroscopy. The structure reveals that the charge distribution on the surface of amoebapore A differs markedly from that of other members of the saposin-like family of proteins, namely the porcine NK-lysin (20) and human granulysin (18). In contrast to NK-lysin and granulysin with their positively charged epitopes that are suggested to be responsible for membrane interaction, amoebapore A shows an extended hydrophobic surface area. Moreover, we report that dimerization of amoebapore A is a prerequisite for membrane binding and pore formation. This dimerization mechanism is a pH-dependent activation step of cell lysis by amoebapore A, which is mediated by a key histidine residue located at the dimer interface.
Protein PurificationFrozen trophozoites of E. histolytica HM-1: IMSS were extracted twice in 10 volumes of 1 N HCl, 5% acetic acid, 1% trifluoroacetic acid, 1% NaCl overnight at 4 °C under constant shaking. The acid extracts were centrifuged at 150,000 x g at 4 °C for 1 h. The supernatants were combined and passed through a tC18 12-cc (2 g) Sep-Pak cartridge (Waters). The cartridge was washed with 12 ml of 0.1% trifluoroacetic acid, 35% acetonitrile, and elution of adsorbed material was achieved with 15 ml of 0.1% trifluoroacetic acid, 80% acetonitrile. The eluate was lyophilized, redissolved in 4 ml of 0.1% trifluoroacetic acid, and subsequently applied in portions to reversed-phase HPLC using an Aquapore Butyl 300 column (2.1 x 220 mm; Brownlee Laboratories, San Jose, CA) connected with a 130 A separation system (Applied Biosystems). Peptides were eluted with a nonlinear gradient of 0% (3 min), 40-50% (10 min), 50-70% (22 min), and 70% (10 min) acetonitrile in 0.1% trifluoroacetic acid at 50 °C and at a flow rate of 0.2 ml/min. All fractions were immediately tested for pore-forming activity. Active fractions were lyophilized, resuspended in 0.1% trifluoroacetic acid, and subjected to rechromatography using the same column and a linear gradient of 0-70% acetonitrile in 0.1% trifluoroacetic acid and otherwise the same conditions. Homogeneity of purified amoebapore A, which elutes at 55% acetonitrile, has been verified by N-terminal sequencing and mass spectrometry and was routinely confirmed by analytical reversed-phase HPLC and Tricine-SDS-PAGE using 13% separation gels (21). Amoebapore A was repeatedly lyophilized and redissolved in 10 mM HCl to evaporate residual trifluoroacetic acid bound to the peptide and stored at -20 °C. NMR SpectroscopyLyophilized protein was dissolved in 93% H2O, 7% D2O, and the pH was adjusted to 3.5 The protein concentration was 1.5 mM. Sets of two-dimensional homonuclear 1H-NMR total correlation spectroscopy and NOESY spectra were recorded at 600 or 750 MHz and 24 °C on either a Varian INOVA 750 or Bruker Avance 600/750 spectrometers. The mixing times for the six NOESY spectra were 60, 100, 140, 180, 250, and 300 ms, respectively. The spectral width was 8200 Hz in both dimensions, and the water signal was suppressed by a weak radio frequency pulse during the relaxation delay. For each spectrum, 400 t1 increments were acquired, each with 2048 complex points using the time-proportional phase incrementation scheme. Prior to Fourier transformation, a 60° shifted sine bell window function was applied in both dimensions, and the spectra were zero-filled in t1 so that 1024 x 1024 data points were obtained. Finally, the spectra were baseline-corrected with a polynomial function. To verify the amide proton resonance assignment, we recorded a natural abundance 1H-15N-heteronuclear single quantum coherence spectrum at 750 MHz with spectral widths of 9000 and 2500 Hz for 1H and 15N, respectively. For the spectrum, 180 t1 increments were acquired each with 1600 complex points. For the H/D exchange experiments, the protein was lyophilized again and dissolved in 100% D2O. The pH and concentration were the same as in the above mentioned experiments. NOESY spectra were recorded with a mixing time of 150 ms at 500 MHz on a Bruker AVANCE 500. The spectral width was 6009 Hz in both dimensions, and the water signal was suppressed by a weak radio frequency pulse during the relaxation delay. Because of the slowly exchanging amide protons, partly due to the low pH, the temperature was increased from 24 °C in the first recordings to 50 °C in the last recording. All data processing was performed on an SGI Indigo work station using the program nmrDraw (22). All subsequent procedures, such as spectral assignment, cross-peak integration, and distance determination, were performed by using the NMRview program (23). CD SpectropolarimetryCD measurements were carried out with a Jasco J-720 spectropolarimeter (Japan Spectroscopic Co., Ltd., Tokyo, Japan), calibrated according to Chen and Yang (24). The spectral bandwidth was 1 nm. The measurements were carried out at 24 °C. Protein ConcentrationProtein concentrations were calculated from absorption spectra in the range of 240-320 nm using the method of Waxman et al. (25). Assay for Pore-forming ActivityPore-forming activity was determined by monitoring the dissipation of a valinomycin-induced diffusion potential in liposomes as described previously (8). Briefly, liposomes prepared from crude soybean phosphatidylcholine IIS (Sigma) (40 mg/ml) in 50 mM K2SO4, 0.5 mM EDTA, 50 mM Tris maleate, pH 5.2, were diluted 1: 4000 in a similar buffer containing Na+ instead of K+ and adjusted to pH 5.2. The addition of 1 nM valinomycin induced a diffusion potential that was monitored by the fluorescence quenching of 3,3'-diethylthiodicarbocyanine iodide dye (1 µM; Eastman Kodak Co.) using a fluorescence spectrophotometer (LS50B; PerkinElmer Life Sciences) with excitation and emission wavelengths of 620 and 670 nm, respectively. Pore-forming activity was measured as the initial change of fluorescence over time after the addition of sample. One unit is defined as a fluorescence increase to 5% of the prevalinomycin level in 1 min at 25 °C. Structure CalculationStructure calculations were performed using the software program DYANA (26). The structure calculation was based on a set of 1418 NOE distance restraints. In addition to the NOE constraints, 2 x 58 hydrogen bond distance restraints, resulting from the H/D exchange experiments, and 3 x 12 distances, representing the spaces within a disulfide bond, were introduced. NOE cross-peak intensities were classified as strong, medium, and weak and assigned to restraints of 1.8-2.8, 1.8-3.4, and 1.8-5.0 Å., respectively, with appropriate pseudo-atom corrections. Hydrogen bond restraints were 1.8-2.4 and 2.8-3.4 Å for H/O and N/O distances, respectively. Disulfide bond restraints were 2.03-2.15, 3.03-3.13, and 2.97-4.49 Å for SG/SG, SG/CB, and SG/CA distances, respectively. In the final structure calculation, all 1570 distance restraints were used to calculate 75 structures. Of these structures, 20 structures with the lowest target functions were selected. The structural statistics of these structures are listed in Table I. The coordinates have been deposited in the Protein Data Bank (accession code 1OF9 [PDB] ).
Size Exclusion ChromatographyFreeze dried amoebapore A was solved in 50 mM sodium citrate buffer (pH 3.5), 50 mM sodium phosphate buffer (pH 5.2), and 50 mM Tris/HCl buffer (pH 8.0), respectively, incubated for 1-2 h at room temperature, and then subjected to size exclusion chromatography with an equilibrated Superdex 75 (16/60) column (Amersham Biosciences) at a constant flow rate of 1 ml/min at 4 °C. The column was calibrated using a mixture of four proteins of known molecular mass, i.e. albumin (67 kDa), ovalbumin (43 kDa), chymotrypsinogen A (25 kDa), and ribonuclease A (13.7 kDa). The column was equilibrated with 50 mM sodium citrate buffer (pH 3.5), 50 mM sodium phosphate buffer (pH 5.2), and 50 mM Tris/HCl buffer (pH 8.0), respectively, and loaded with 1.0 ml of protein solution. Fractions of 3 ml were collected. Cross-linkingFor cross-linking in solution, 0.5 µg of amoebapore A were incubated with 100 µM 1-ethyl-3-(dimethylaminopropyl)-carbodiimide in 1 M glycine methylester (pH 4.5) at 20 °C for 2 h (27, 28). Alternatively, dithiobis(succinimidyl propionate) was added from a freshly prepared stock solution in dimethyl sulfoxide to the same amount of amoebapore A in 50 mM sodium phosphate (pH 7.0) to reach a final concentration of 100 µM (29). After an incubation period of 30 min at 20 °C, the reaction was stopped by adding an excess of Tris. The samples were subjected to Tricine-SDS-PAGE using 13% gels (21). DEPC ModificationDiethylpyrocarbonate (DEPC) modification was performed according to Andrä and Leippe (see Ref. 32). Briefly, 0.58 mg of amoebapore A were dissolved in 4.0 ml of a 50 mM sodium phosphate (pH 6.0) buffer. After incubation with a 200-fold excess of DEPC for 1.5 h at 4 °C and another 1 h at room temperature, the solution was subjected to size exclusion chromatography and eluted with a 50 mM sodium phosphate buffer (pH 5.2) at 4 °C.
Structure of Amoebapore ASince the protein could not be recombinantly expressed in Escherichia coli and was purified from its natural source E. histolytica, structure elucidation of the amoebapore A relied on conventional two-dimensional 1H-NMR experiments. The helical secondary structure elements in amoebapore A were identified by the presence of characteristic d N(i + 3) and d N(i + 3) sequential connectivities in the two-dimensional NOESY spectra of this protein. Fig. 1 shows these patterns of connectivities derived from the NOESY spectra. This specific pattern is missing in the loop regions and incomplete for helix 1 due to partial spectral overlap. Additional information about the secondary structure elements was derived from H/D exchange experiments. Even after increasing the temperature to 50 °C for 24 h, several amide protons were still observable (Fig. 1) and therefore included in the structure calculation as hydrogen bonds (see "Experimental Procedures").
The structure of amoebapore A (for statistics, see Table I) consists of five -helices (Fig. 2, A and B), which comprise residues 4-16 (helix I), 25-35 (helix II), 42-52 (helix III), 55-63 (helix IV), and 67-73 (helix V). A single disulfide bridge connects helices 2 and 3, whereas two disulfide bridges connect helices 1 and 5. According to structural classification of proteins (SCOP) (30), the structure can be described as a folded leaf in which helices 1 and 2 are one-half of the leaf and are folded against helices 3, 4, and 5 as the second half. Helices 1 and 2 are connected by a loop consisting of eight residues (17-24), whereas helices 3 and 4 virtually merge and are separated by only two residues (53 and 54). Residues 64-66 link helices 4 and 5. The two parts of the folded leaf are connected by a loop between helices 2 and 3 (residues 36-41). The superposition of 20 independently calculated structures demonstrates the coherence of the generated ensemble, which is shown as a schematic representation in Fig. 2A. In particular, within the helical regions, the root mean square deviations for the backbone atoms are 0.25 Å. Approximately 75% of the backbone / -dihedral angle pairs were in the most favorable region of a Ramachandran plot, and none were observed in disallowed regions. On the basis of this ensemble, an average structure of amoebapore A was calculated and resembles the typical fold of the saposin-like proteins (Fig. 2B). A comparison of this structure with the structures of porcine NK-lysin and human granulysin is shown in Fig. 3. Although the three structures show a similar global fold, they differ substantially in the spatial arrangement of the helices. The orientations of helices 2, 3, and 4 relative to each other are similar in the three structures. However, the orientation of helices 1 and 5 relative to 2, 3, and 4 diverges considerably in amoebapore A when compared with NK-lysin and granulysin (Fig. 3). In amoebapore A, helix 1 runs almost parallel to helices 2 and 3, whereas in NK-lysin and granulysin, helix 1 is kinked by 90°. Remarkably, the molecular surface character of amoebapore A constituted by residues from helices 1 and 2 is predominantly hydrophobic (Fig. 3). The length of this hydrophobic area of about 36 Å is sufficient to span a lipid bilayer (31). In contrast, both mammalian molecules, NK-lysin and granulysin, have clusters of positively charged amino acids, which are proposed to be responsible for the initial contact with the membrane and ultimately lead to its destruction (16, 18). As such cationic clusters are not present on the surface of amoebapore A (Fig. 3), a different mode of interaction with the lipid bilayer must exist.
Amoebapore A Forms Stable DimersAs amoebapore A has been shown to be most active at pH 5.2 (32), initial NMR experiments were performed at this pH. Unfortunately, the proton T2 values measured under these conditions were indicative of protein oligomerization. Stepwise lowering of the pH until 3.5 resulted in proton T2 values corresponding to a monomeric protein species (data not shown). Apparently, the oligomerization of amoebapore A, which pI has been calculated to 5.9, is driven by electrostatic interactions that are disrupted by protonation of acidic residues upon lowering the pH. To examine whether the global conformation of amoebapore A changes with pH, CD spectra of amoebapore A were recorded at pH values of 3, 4, 5, 6, and 7. The identical CD spectra observed in Fig. 4A clearly indicate that the overall conformation of amoebapore A is not influenced by shifting the pH between 3 and 7. Although CD spectroscopy is a very sensitive method to observe changes in secondary structure, we cannot rule out changes in tertiary structure. As the conformation of amoebapore A did not change over the pH range tested, NMR experiments were performed at a pH value (i.e. 3.5) at which the protein does not self-associate. The observed pH-dependent oligomerization of amoebapore A raised the question about the stoichiometry of the functional protein complexes. To determine the stoichiometry and the stability of the oligomers, we performed size exclusion chromatography at different pH values. In accordance with the T2 measurements, chromatography at pH 3.5 revealed that the protein exists as a stable monomer (Fig. 4B). At pH 5.2, the protein eluted from the column as a single peak at a retention time corresponding to the molecular weight of the dimeric species (Fig. 4B). No higher order oligomers were detectable. The fact that amoebapore A is most active at this pH (32) raised the possibility that dimerization is a prerequisite for pore formation. Fractions containing the amoebapore A dimer were examined for pore-forming activity by monitoring the dissipation of a valinomycin-induced diffusion potential in liposomes. All of the pore-forming activity (54,375 units) of the material loaded onto the column was recovered in fractions containing the dimer. To further examine the relationship between pH-dependent activity and the oligomerization state, we performed size exclusion chromatography experiments at pH 8, at which amoebapore A has been described to be inactive (32). Under these conditions, the protein elutes predominantly as a monomer (Fig. 4B). Although some entity was detected near the position of the dimer, it is known that amoebapore does not exert pore-forming activity at this pH (32). Notably, under all the conditions examined, proteins eluted as distinct species from the column, indicating that no monomer/dimer equilibria exist during the time course of the experiment (3 h). To further confirm this result, we performed cross-linking experiments. Results from the cross-linking experiments at pH 4.5, at which we anticipate that acidic residue side chains are deprotonated and histidine side chains are protonated, indicated that only dimers were present (Fig. 4C). The same experiment performed at pH 7.0 revealed that amoebapore A existed solely in the monomeric state (Fig. 4C). From the fact that amoebapore A is monomeric at pH 7.0 and 8.0, respectively, hydrophobic interactions can be excluded to be the driving forces for dimerization.
Dimerization of Amoebapore A Is Mediated by a Histidine ResidueAlthough NK-lysin is biologically active at a broad range of pH (33), pore formation by amoebapore A is restricted to a very narrow pH range (32). No pore-forming activity has been observed between pH values 10 and 6.5, whereas activity increased dramatically at pH 6.0 and reached a maximum at pH 5.2 (32). In contrast, granulysin is most active at pH 7.4 (34). Together with the structural analysis, these results suggest that the protonation state of a histidine side chain may be responsible for the pH-dependent activity observed. Interestingly, the single histidine present in amoebapore A is solvent-exposed, and no long range NOEs were observed in the NOESY spectra recorded. Consequently, the histidine residue shows a high root mean square deviation value of 1.87 Å when compared with the average value of 1.20 Å. The histidine side chain therefore displays a high degree of unrestricted motion. The importance of this amino acid residue for the activity of amoebapore A was further supported by reversible chemical modification of the histidine residue. Treatment with DEPC, leading to a non-ionizable N-carbethoxyhistidyl derivative, resulted in an almost complete loss of pore-forming activity (32). To examine the oligomerization status of the DEPC-treated protein, we performed size exclusion chromatography (Fig. 4B). At pH 5.2, the chemically modified amoebapore A elutes as a monomer in contrast to the native molecule. Furthermore, cross-linking experiments with amoebapore A have shown that oligomers, i.e. dimers up to hexamers, exist when the protein was bound to liposomes (8). This oligomerization was no longer observed when the histidine-modified protein was used in such a cross-linking experiment, although the liposome binding capacity was hardly affected (32). To exclude the possibility that chemical modification induced a conformational change, we recorded CD spectra of the DEPC treated and untreated protein (Fig. 5). Both spectra are identical, demonstrating that derivatization of the histidine, resulting in a non-ionizable side chain, does not influence the overall conformation of amoebapore A. We therefore conclude that protonation of the sole histidine residue is indispensable for the dimerization of amoebapore A.
The participation of histidines in membrane-associated oligomerization processes has been suggested for other pore-forming toxins, although these molecules belong to a structurally unrelated family of proteins. For aerolysin (35, 36) from Aeromonas hydrophyla and for the -toxin (37) of Staphylococcus aureus, it has been reported that histidine modification with DEPC or side-directed mutagenesis blocks aggregation and pore formation. The crystal structure of the -toxin revealed that a histidine is involved in oligomerization by promoting electrostatic interactions between two monomers (38). In contrast to Amoebapore A, no distinct oligomers have been observed for these molecules in solution. A Model of Amoebapore A OligomerizationAs dimers of amoebapore A were observed to be the functionally important species, we concluded that dimerization had to occur in a head-to-head orientation (Fig. 6A) since a head-to-tail interaction would lead to higher order oligomers, which were not observed. As a consequence, the two histidines from the two monomers must be located at the dimer interface. Two different dimer models can be envisaged. The first involves a parallel orientation of the monomers, and the second involves an antiparallel orientation (Fig. 6A). In the parallel orientation, the hydrophobic areas of the two molecules point in opposite directions, whereas in the antiparallel orientation, they point in the same. These two possibilities result in two different dimerization interfaces. A comparison revealed that a parallel orientation would bring similarly charged epitopes together, resulting in a repulsive interaction (not shown). In contrast, a more favorable conformation would be the antiparallel orientation, as this would lead to an attractive electrostatic interaction between two monomers. As no conformational change has been observed during dimerization, the solution structure of the amoebapore A monomer was used to build a model of the biologically active dimer.
Although a description of the dimerization interface on an atomic level is not possible by such a model, it allows the identification of certain amino acid side chains responsible for dimerization. Since we have shown that the interaction is dominated by charged amino acid side chains, we first looked for potential ion pairs located at the dimer interface. In Fig. 6B, a model of the dimer is shown, and two ion pairs identified within the interface are depicted. Due to the antiparallel orientation of the two monomers, all interactions occur twice. Histidine 75 of monomer 1 is in close contact to aspartic acid 63 of monomer 2, and the second ion pair is formed by glutamic acid 2 and lysine 64. No long range NOEs have been observed for these four residues, indicating their possible involvement in such an interaction. The interface shows neither overlapping van der Waals radii of amino acid side chains nor extended cavities, demonstrating that the shape of the involved molecular surfaces are complementary to each other. A striking consequence of this dimerization mode is that the two hydrophobic areas formed by helices 1 and 2 in each molecule are very close and thus augment each other (Fig. 6B). Since amoebapore A is a pore-forming toxin, this extended hydrophobic epitope might be involved in the interaction with the hydrophobic environment of the surrounding membrane. According to early biophysical experiments, it has been proposed that amoebapore A acts via the barrel stave model (19). This model describes the pore as an assembly of proteins inserted into a membrane such that the staves are oriented by their hydrophobic areas toward the lipids of the membrane, whereas the hydrophilic areas constitute the inside of the pore (39). The suggested barrel stave model is in excellent agreement with the properties of the amoebapore A dimer. In contrast, other pore-forming molecules of the SAPLIP family, NK-lysin and granulysin, have been described to interact with the membrane only superficially, thereby causing primarily a membrane perturbance (17, 18). Upon insertion into membranes, amoebapore A appears to form hexamers as determined by cross-linking experiments using lipid vesicles (8). Like dimerization in solution, the formation of homooligomers, presumably hexamers, within the membrane is not accomplished by a conformational change as evidenced by CD spectropolarimetry (9).
Therefore, as in the case of the dimer, the solution structure of the amoebapore A monomer is a suitable module for the construction of a hexamer model. As shown in Fig. 6B, three dimers can be readily arranged into a circular configuration such that the hydrophobic surface is extended to form a belt surrounding the hexamer (Fig. 6B). As with the monomer/monomer interface, the dimer/dimer interface could be constructed without overlap of van der Waals radii from the involved side chains and without the introduction of cavities in the interaction area. It has been recently shown that amoebapore A forms pores with a diameter between 1.3 and 2.2 nm (11). The diameter of the modeled pore is The pH-dependent conversion into the permeabilization-competent dimer allows the storage of amoebapore A as a mature inactive form in cytoplasmic granules inside the amoebae. As amoebapore B and C proteins act in a pH-dependent manner and have the same key histidine residue, our conclusion from the structural data for amoebapore A presumably holds true for all three isoforms. The consensus view is that the primary function of amoebapores is to combat the intracellular growth of phagocytosed bacteria inside the amoeba. The protozoon discharges its effector proteins in the acidic environment of phagolysosomes in which engulfed bacteria are situated (40). Persuasive support for the involvement of amoebapores in extracellular killing of host cells comes from the finding that amoebapores are cytolytic toward eukaryotic cells (41, 42) and that amoebae lacking the major form amoebapore A are no longer capable of killing mammalian cells or destroying host tissues in vitro (7). For the scenario of the cytolytic reaction, it may be envisaged that after amoeba-target cell contact has been established, amoebapores are discharged from cytoplasmic granules into the confined environment of the contact zone in which a high protein concentration and a relatively low pH may be reached. Such an environment probably leads to the formation of dimers and ultimately the formation of pore-forming hexamers of amoebapore A. The presented structure of amoebapore A is the first structure of a toxin derived from an eukaryotic parasite. The structural information and the derived implications for the mode of action of this key factor in pathogenicity should lead to a better understanding of the enigma of invasive amoebiasis.
The atomic coordinates and structure factors (code 1OF9 [PDB] ) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
* This work was supported by grants from the Deutsche Forschungsgemeinschaft (Grant GR 1623/1-1, LE 1075/2-3, and the SFB 617) and the European UNION (Grant HPRI-CT-2001-00172). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ Present address: European Molecular Biology Laboratory, Meyerhofstrasse 1, 69117 Heidelberg, Germany.
1 The abbreviations used are: SAPLIP, saposin-like proteins; HPLC, high pressure liquid chromatography; NOE, nuclear Overhauser effect; NOESY, NOE spectroscopy; DEPC, diethylpyrocarbonate; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine.
We thank C. Ott for technical assistance during amoebapore purification, B. Riekens for performing the chemical cross-linking, and Dr. L. Shaw for critical reading of the manuscript. The support of the Bernhard Nocht Institute for Tropical Medicine at which the amoebae used for the study were cultured is also gratefully acknowledged.
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