JBC INTERFERin siRNA transfection reagent

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M312978200 on February 17, 2004

J. Biol. Chem., Vol. 279, Issue 17, 17834-17841, April 23, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/17/17834    most recent
M312978200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Hecht, O.
Right arrow Articles by Grötzinger, J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Hecht, O.
Right arrow Articles by Grötzinger, J.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Solution Structure of the Pore-forming Protein of Entamoeba histolytica*

Oliver Hecht{ddagger}, Nico A. van Nuland§, Karin Schleinkofer{ddagger}, Andrew J. Dingley||, Heike Bruhn**, Matthias Leippe{ddagger}{ddagger}, and Joachim Grötzinger{ddagger}§§

From the {ddagger}Biochemisches Institut der Christian-Albrechts-Universität Kiel, Olshausenstr. 40 24118 Kiel, Germany, the §Department of NMR Spectroscopy, Bijvoet Center for Biomolecular Research, Utrecht University, 3584 CH Utrecht, The Netherlands, the ||Department of Biochemistry and Molecular Biology, University College London, Gower Street, London, WC1E 6BT, United Kingdom, **Molekulare Parasitologie, Zentrum für Infektionsforschung, Röntgenring 11, 97070 Würzburg, Germany, and {ddagger}{ddagger}Zoologisches Institut der Christian-Albrechts-Universität Kiel, Olshausenstr. 40, 24118 Kiel, Germany

Received for publication, November 30, 2003 , and in revised form, February 9, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
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.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
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 {alpha}-helical (9), and this has also been predicted for amoebapore B and C (10). All amoebapores are able to form pores in membranes by oligomerization and thus affect the integrity of target cell membranes (10, 11).

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.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
 REFERENCES
 
Protein Purification—Frozen 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 Spectroscopy—Lyophilized 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 Spectropolarimetry—CD 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 Concentration—Protein 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 Activity—Pore-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 Calculation—Structure 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] ).


View this table:
[in this window]
[in a new window]
  		TABLE I
Structural statistics for the 20 conformers of amoebapore A

No NOE distance restraint was violated by more than 0.5 Å in any of the structures.

 
Size Exclusion Chromatography—Freeze 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-linking—For 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 Modification—Diethylpyrocarbonate (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.


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
REFERENCES
 
Structure of Amoebapore A—Since 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{alpha}N(i + 3) and d{beta}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").



View larger version (20K):
[in this window]
[in a new window]
  		FIG. 1.
Schematic representation of the short and medium range upper distance limits, classified as strong, middle, and weak NOEs, according to the height of the bars. Amide protons that are not exchanged with D2O after increasing the temperature up to 50 °C and an exposure of 24 h are indicated by filled circles.

 
The structure of amoebapore A (for statistics, see Table I) consists of five {alpha}-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 {Phi}/{psi}-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.



View larger version (43K):
[in this window]
[in a new window]
  		FIG. 2.
The structure of amoebapore A. A, a stereo view of 20 superimposed backbone traces representing the NMR structure. B, a ribbon representation of the energy-minimized average structure (same orientation as above). Disulfide bonds are depicted in blue, and the N and C terminus are labeled. Helices are numbered by roman numerals. Figures were created using GRASP (43) and Ribbons (44).

 



View larger version (42K):
[in this window]
[in a new window]
  		FIG. 3.
Ribbon (44) representations of amoebapore A (left), NK-lysin (middle), and granulysin (right) are shown in the upper panel. The C and N termini are depicted in black. All molecules were superimposed onto C{alpha} atoms of residues 30-60. Helices are numbered by roman numerals. The electrostatic potential of the molecular surface is shown in the lower panel in the same orientation as in the upper panel, indicating positive potential in blue and negative potential in red. The representation of the surface was generated using GRASP (43).

 
Amoebapore A Forms Stable Dimers—As 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.



View larger version (13K):
[in this window]
[in a new window]
  		FIG. 4.
pH-dependent behavior of amoebapore A. A, CD spectroscopy of amoebapore A. CD spectroscopy was performed at pH 3 (solid line), 4 (dotted line), 5 (dashed line), 6 (long dashed line), and 7 (dot-dashed line) respectively. B, the lower panel contains the elution profiles of size exclusion experiments showing the pH dependence of the amoebapore A oligomerization states. Experiments were performed at pH 3.5 (black), 5.2 (green), 8.0 (red), and 5.2 after DEPC treatment (blue). The protein elutes as a monomer at pH 3.5, as a dimer at pH 5.2, predominantly as a monomer at pH 8.0, and as a monomer at pH 5.2 after DEPC treatment. The varying hydrodynamic volumes are responsible for the various elution times of the monomer and dimer protein under different solvent conditions. Absorption (arbitrary units, AU) was measured at either 280 or 218 nm and scaled to comparable size. The upper panel shows the calibration curve as well as the elution times of amoebapore A under different pH values. Elution times of proteins used for calibration are indicated by circles, and elution times of amoebapore A under the different conditions are indicated by colored symbols according to the different pH values. C, the molecular organization of amoebapore A in solution was examined with and without cross-linking, which was performed using 1-ethyl-3-(dimethylaminopropyl)-carbodiimide (EDC) at pH 4.5 and dithiobis(succinimidylpropionate) (DSP) at pH 7.0, respectively.

 
Dimerization of Amoebapore A Is Mediated by a Histidine Residue—Although 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.



View larger version (11K):
[in this window]
[in a new window]
  		FIG. 5.
CD spectra of the DEPC treated (dashed line) and untreated (solid line) 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 {alpha}-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 {alpha}-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 Oligomerization—As 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.



View larger version (41K):
[in this window]
[in a new window]
  		FIG. 6.
Modeling the amoebapore A oligomerization. A, schematic representation of different possible monomer orientations within a dimer. The location of the single histidine is indicated. B, ribbon representation of the dimer. Residues responsible for electrostatic interactions are depicted. Helices 1 and 2 forming the hydrophobic surface are labeled (left). Ribbon representation was created using Ribbons (44). The electrostatic potential of the molecular surface of the modeled hexameric structure is shown, indicating positive potential in blue and negative potential in red (right). Figures of the electrostatic potential were generated using GRASP (43).

 
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 ~2 nm, in excellent agreement with the experimentally derived value. Fig. 6B shows the electrostatic potential of the molecular surface of the hexamer model, demonstrating the differences in charge distribution. In contrast to the exterior surface, the inside of the pore has a predominantly hydrophilic character.

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.


   FOOTNOTES
 
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. Back

Present address: European Molecular Biology Laboratory, Meyerhofstrasse 1, 69117 Heidelberg, Germany. Back

§§ To whom correspondence should be addressed. Tel.: 49-431-8801686; Fax: 49-431-8805007; E-mail: jgroetzinger{at}biochem.uni-kiel.de.

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. Back


   ACKNOWLEDGMENTS
 
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.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 

  1. Ravdin, J. I. (1995) Clin. Infect. Dis. 20, 1453-1464; quiz, 1465-1466[Medline] [Order article via Infotrieve]
  2. Stanley, S. L., Jr. (2003) Lancet 361, 1025-1034[CrossRef][Medline] [Order article via Infotrieve]
  3. Walsh, J. A. (1986) Rev. Infect. Dis. 8, 228-238[Medline] [Order article via Infotrieve]
  4. Petri, W. A., Jr., Haque, R., and Mann, B. J. (2002) Annu. Rev. Microbiol. 56, 39-64[Medline] [Order article via Infotrieve]
  5. Bruchhaus, I., Jacobs, T., Leippe, M., and Tannich, E. (1996) Mol. Microbiol. 22, 255-263[CrossRef][Medline] [Order article via Infotrieve]
  6. Leippe, M. (1997) Parasitol. Today 13, 178-183[CrossRef][Medline] [Order article via Infotrieve]
  7. Bracha, R., Nuchamowitz, Y., and Mirelman, D. (2003) Eukaryot. Cell 2, 295-305[Abstract/Free Full Text]
  8. Leippe, M., Ebel, S., Schoenberger, O. L., Horstmann, R. D., and Müller-Eberhard, H. J. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 7659-7663[Abstract/Free Full Text]
  9. Leippe, M., Tannich, E., Nickel, R., van der Goot, G., Pattus, F., Horstmann, R. D., and Müller-Eberhard, H. J. (1992) EMBO J. 11, 3501-3506[Medline] [Order article via Infotrieve]
  10. Leippe, M., Andrä, J., Nickel, R., Tannich, E., and Müller-Eberhard, H. J. (1994) Mol. Microbiol. 14, 895-904[Medline] [Order article via Infotrieve]
  11. Gutsmann, T., Rieckens, B., Bruhn, H., Wiese, A., Seydel, U., and Leippe, M. (2003) Biochemistry 42, 9804-9812[CrossRef][Medline] [Order article via Infotrieve]
  12. Munford, R. S., Sheppard, P. O., and O'Hara, P. J. (1995) J. Lipid Res. 36, 1653-1663[Medline] [Order article via Infotrieve]
  13. Andersson, M., Gunne, H., Agerberth, B., Boman, A., Bergman, T., Sillard, R., Jornvall, H., Mutt, V., Olsson, B., Wigzell, H., Dagerlind, A., Boman, H. G., and Gudmundsson, G. H. (1995) EMBO J. 14, 1615-1625[Medline] [Order article via Infotrieve]
  14. Pena, S. V., Hanson, D. A., Carr, B. A., Goralski, T. J., and Krensky, A. M. (1997) J. Immunol. 158, 2680-2688[Abstract]
  15. Bruhn, H., and Leippe, M. (1999) Biol. Chem. 380, 1001-1007[CrossRef][Medline] [Order article via Infotrieve]
  16. Miteva, M., Andersson, M., Karshikoff, A., and Otting, G. (1999) FEBS Lett. 462, 155-158[CrossRef][Medline] [Order article via Infotrieve]
  17. Ruysschaert, J. M., Goormaghtigh, E., Homble, F., Andersson, M., Liepinsh, E., and Otting, G. (1998) FEBS Lett. 425, 341-344[CrossRef][Medline] [Order article via Infotrieve]
  18. Anderson, D. H., Sawaya, M. R., Cascio, D., Ernst, W., Modlin, R., Krensky, A., and Eisenberg, D. (2003) J. Mol. Biol. 325, 355-365[CrossRef][Medline] [Order article via Infotrieve]
  19. Keller, F., Hanke, W., Trissl, D., and Bakker-Grunwald, T. (1989) Biochim. Biophys. Acta 982, 89-93[Medline] [Order article via Infotrieve]
  20. Liepinsh, E., Andersson, M., Ruysschaert, J. M., and Otting, G. (1997) Nat. Struct. Biol. 4, 793-795[CrossRef][Medline] [Order article via Infotrieve]
  21. Schägger, H., and von Jagow, G. (1987) Anal. Biochem. 166, 368-379[CrossRef][Medline] [Order article via Infotrieve]
  22. Delagio, F., Grzesiek, S., Vuister, G. W., Zhu, G., Pfeifer, J., and Bax, A. (1995) J. Biomol. NMR 6, 277-293[Medline] [Order article via Infotrieve]
  23. Johnson, B. A., and Blevins, R. A. (1994) J. Biomol. NMR 4, 603-614[CrossRef]
  24. Chen, G. C., and Yang, J. T. (1977) Anal. Lett. 10, 1195-1207
  25. Waxman, E., Rusinova, E., Hasselbacher, C. A., Schwartz, G. P., Laws, W. R., and Ross, J. B. (1993) Anal. Biochem. 210, 425-428[CrossRef][Medline] [Order article via Infotrieve]
  26. Guntert, P., Mumenthaler, C., and Wuthrich, K. (1997) J. Mol. Biol. 273, 283-298[CrossRef][Medline] [Order article via Infotrieve]
  27. Hoare, D. G., and Koshland, D. E., Jr. (1967) J. Biol. Chem. 242, 2447-2453[Abstract/Free Full Text]
  28. Carraway, K. L., and Koshland, D. E., Jr. (1968) Biochim. Biophys. Acta 160, 272-274[Medline] [Order article via Infotrieve]
  29. Lomant, A. J., and Fairbanks, G. (1976) J. Mol. Biol. 104, 243-261[CrossRef][Medline] [Order article via Infotrieve]
  30. Murzin, A. G., Brenner, S. E., Hubbard, T., and C. C. (1995) J. Mol. Biol. 247, 536-540[CrossRef][Medline] [Order article via Infotrieve]
  31. Bransburg-Zabary, S., Kessel, A., Gutman, M., and Ben-Tal, N. (2002) Biochemistry 41, 6946-6954[CrossRef][Medline] [Order article via Infotrieve]
  32. Andrä, J., and Leippe, M. (1994) FEBS Lett. 354, 97-102[CrossRef][Medline] [Order article via Infotrieve]
  33. Bruhn, H., Riekens, B., Berninghausen, O., and Leippe, M. (2003) Biochem. J. 375, 737-744[CrossRef][Medline] [Order article via Infotrieve]
  34. Hanson, D. A., Kaspar, A. A., Poulain, F. R., and Krensky, A. M. (1999) Mol. Immunol. 36, 413-422[CrossRef][Medline] [Order article via Infotrieve]
  35. Garland, W. J., and Buckley, J. T. (1988) Infect. Immun. 56, 1249-1253[Abstract/Free Full Text]
  36. Green, M. J., and Buckley, J. T. (1990) Biochemistry 29, 2177-2180[CrossRef][Medline] [Order article via Infotrieve]
  37. Jursch, R., Hildebrand, A., Hobom, G., Tranum-Jensen, J., Ward, R., Kehoe, M., and Bhakdi, S. (1994) Infect. Immun. 62, 2249-2256[Abstract/Free Full Text]
  38. Song, L., Hobaugh, M. R., Shustak, C., Cheley, S., Bayley, H., and Gouaux, J. E. (1996) Science 274, 1859-1866[Abstract/Free Full Text]
  39. Boheim, G. (1974) J. Membr. Biol. 19, 277-303[CrossRef][Medline] [Order article via Infotrieve]
  40. Andrä, J., Herbst, R., and Leippe, M. (2003) Dev. Comp. Immunol. 27, 291-304[CrossRef][Medline] [Order article via Infotrieve]
  41. Leippe, M., Andrä, J., and Müller-Eberhard, H. J. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 2602-2606[Abstract/Free Full Text]
  42. Berninghausen, O., and Leippe, M. (1997) Infect. Immun. 65, 3615-3621[Abstract]
  43. Nicholls, A., Sharp, K. A., and Honig, B. (1991) Proteins 11, 281-296[CrossRef][Medline] [Order article via Infotrieve]
  44. Carson, M. (1991) J. Appl. Crystallogr. 24, 946-950[CrossRef]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 Innate ImmunityHome page
J. Andra, T. Gutsmann, P. Garidel, and K. Brandenburg
Invited review: Mechanisms of endotoxin neutralization by synthetic cationic compounds
Innate Immunity, October 1, 2006; 12(5): 261 - 277.
[Abstract] [PDF]

Home page
 Protein Sci.Home page
V. E. Ahn, P. Leyko, J.-R. Alattia, L. Chen, and G. G. Prive
Crystal structures of saposins A and C
Protein Sci., August 1, 2006; 15(8): 1849 - 1857.
[Abstract] [Full Text] [PDF]

Home page
 J. Immunol.Home page
M. A. Ryan, H. T. Akinbi, A. G. Serrano, J. Perez-Gil, H. Wu, F. X. McCormack, and T. E. Weaver
Antimicrobial Activity of Native and Synthetic Surfactant Protein B Peptides
J. Immunol., January 1, 2006; 176(1): 416 - 425.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
T. Kolter, F. Winau, U. E. Schaible, M. Leippe, and K. Sandhoff
Lipid-binding Proteins in Membrane Digestion, Antigen Presentation, and Antimicrobial Defense
J. Biol. Chem., December 16, 2005; 280(50): 41125 - 41128.
[Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/17/17834    most recent
M312978200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Hecht, O.
Right arrow Articles by Grötzinger, J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Hecht, O.
Right arrow Articles by Grötzinger, J.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit