ECP (eosinophil cationic protein) is an eosinophil secretion protein with antipathogen activities involved in the host immune defence system. The bactericidal capacity of ECP relies on its action on both the plasma membrane and the bacterial wall. In a search for the structural determinants of ECP antimicrobial activity, we have identified an N-terminal domain (residues 1–45) that retains most of ECP's membrane-destabilizing and antimicrobial activities. Two sections of this domain, ECP-(1–19) and ECP-(24–45), have also been evaluated. All three peptides bind and partially insert into lipid bilayers, inducing aggregation of lipid vesicles and leakage of their aqueous content. In such an environment, the peptides undergo conformational change, significantly increasing their α-helix content. The bactericidal activity of the three peptides against Escherichia coli and Staphylococcus aureus has been assessed at both the cytoplasmic membrane and the bacterial envelope levels. ECP-(1–45) and ECP-(24–45) partially retain the native proteins ability to bind LPS (lipopolysaccharides), and electron microscopy reveals cell damage by both peptides. Interestingly, in the E. coli cells agglutination activity of ECP is only retained by the longest segment ECP-(1–45). Comparative results suggest a task distribution, whereby residues 1–19 would contribute to membrane association and destabilization, while the 24–45 region would be essential for bactericidal action. Results also indicate that ECP cytotoxicity is not uniquely dependant on its membrane disruption capacity, and that specific interactions at the bacteria wall are also involved.
- antimicrobial peptide
- electron microscopy
- host defence
Host defence proteins and peptides of the innate immune system are potential candidates for chemotherapeutic development. Among those, AMPs (antimicrobial peptides) are characterized by a variety of primary and secondary structures  and by a broad spectrum of activity. Their fast non-specific mechanism of action , mostly at the plasma membrane and cell surface level, tends to mean the development of resistant strains is rather unlikely . On the basis of these features, several native AMPs, or their synthetic analogues, have been proposed as alternatives to conventional antibiotics and some of them are currently in clinical trials, mostly for topical applications .
Some AMPs are derived from host defence proteins by limited proteolysis that releases in vivo active fragments, often corresponding to the N- or C-terminus . Proteolytic processing is very frequent in immunological cascade events, where local cleavage can release the active peptides in the area of infection or inflammation. There are also multiple examples of synthetic peptides corresponding to a protein N- or C-terminus that display antimicrobial activity . Thus lactoferricin, a naturally occurring bactericidal peptide derived from the N-terminus of lactoferrin, is very effective against some antibiotic-resistant strains . Likewise, BPI (bactericidal permeability-increasing protein) is expressed in neutrophils and displays cytotoxicity against Gram-negative bacteria. A recombinant fragment corresponding to the BPI N-terminus is currently in phase III clinical trials . Other examples of AMPs resulting from proteolysis include cathelicidins from neutrophils  and cryptdins, α-defensins secreted by intestinal Paneth cells, whose bactericidal activity requires activation of a precursor by a metalloprotease .
We are currently working on ECP (eosinophil cationic protein) as a model of the potential involvement of mammalian RNases in the host defence system. ECP is a secretory ribonuclease (also known as RNase 3) found in the eosinophilic leucocyte and potentially involved in innate immunity. ECP can also be expressed at lower levels by activated neutrophils. Although the role of eosinophils during infection is controversial, eosinophils are activated, and selectively release their content, at the inflammation areas . Some authors have reported antimicrobial activities for eosinophils and have proposed a complementary action, together with neutrophils, during bacterial infections . Besides local protein secretion, which can harm the host tissues, eosinophils may also participate in immunoregulation and tissue remodelling processes [12,13]. Although a wealth of structural and functional data have been reported, the physiological role of ECP remains elusive . Its cytotoxic activity is effective against a wide range of pathogens, suggesting a relatively non-specific mechanism of action. Although there is no evidence of direct in vivo involvement of ECP in the host response to bacterial infection, ECP kills both Gram-negative and Gram-positive strains at a low micromolar range and its activity depends on its action both at the bacterial cell wall and cytoplasmic membrane levels [15–18]. ECP belongs to the vertebrate RNase A superfamily, of which several members are endowed with antimicrobial properties (see Supplementary Figure S1 available at http://www.BiochemJ.org/bj/421/bj4210425add.htm) [19–21]; the family that may have started off playing a physiological role in the host immune system [22–24]. Interestingly, ECP antibacterial activity is not shared by EDN (eosinophil-derived neurotoxin), a closely related eosinophil ribonuclease . Antimicrobial RNases, as innate immune proteins with anti-infective and immunomodulatory properties, present substantial therapeutic potential in the drug development industry, both in the search for alternative antibiotics and for the treatment of inflammatory disorders.
In order to identify the structural determinants of ECP's cytotoxic mechanism of action, we have searched the ECP sequence for potential active domains using a theoretical approach to spot key antimicrobial regions (M. Torrent, V. Nogués and E. Boix, unpublished work), and have identified a potentially active region at the N-terminus. To validate our hypothesis, we have synthesized three peptides representative of this region (Figure 1) and assessed their properties both on a synthetic membrane model and on bacterial cell cultures.
DOPC (dioleoyl phosphatidylcholine), DOPG (dioleoyl phosphatidylglycerol), (6,7)-Br2PC, (9,10)-Br2PC, and 11,12-Br2PC [1-palmitoyl-2-stearoyl-(6,7, 9,10 and 11,12)-dibromo-sn-glycero-3-phosphocholine respectively] were from Avanti Polar Lipids. ANTS (8-aminonaphthalene-1,3,6-trisulfonic acid disodium salt) and DPX (p-xylenebispyridinium bromide) were from Invitrogen. Bovine pancreatic ribonuclease A, type XII-A, polymyxin B sulfate, LT (lipoteichoic acids) from Staphylococcus aureus, and LPS (lipopolysaccharides) from Eschericha coli serotype 0111:B4 were purchased from Sigma–Aldrich. BC (BODIPY® TR cadaverine; where BODIPY® is boron dipyrromethane (4,4-difluoro-4-bora-3a,4a-diaza-s-indacene) and DiSC3(5) (3,3 dipropylthiacarbocyanine) were purchased from Invitrogen. PD-10 desalting columns with Sephadex G-25 were from Amersham Pharmacia Biotech. E. coli BL21DE3 (Novagen) and S. aureus 502 A (ATCC) strains were used.
Peptide ECP-(1–19) was obtained from NeoMPS. ECP-(24–45) and ECP-(1–45), with cysteine residues at positions 23 and 37 replaced by serine, were prepared by Fmoc (fluoren-9-ylmethoxycarbonyl) solid-phase peptide synthesis methods . All peptides were purified by HPLC to ∼95% homogeneity and were satisfactorily characterized by MALDI–TOF-MS (matrix-assisted laser-desorption ionization–time-of-flight MS). Further experimental details can be found in the Supplementary Experimental section and Figures S6–S8 available at http://www.BiochemJ.org/bj/421/bj4210425add.htm.
Expression and purification of ECP
Wild-type ECP was obtained from a human ECP synthetic gene . Protein expression in the E. coli BL21DE3 strain, folding of the protein from inclusion bodies and the purification steps were carried out as described in .
Antimicrobial activity was expressed as the IC50. The IC50 of each peptide was determined from two independent experiments performed in triplicate for each peptide concentration. Peptides were dissolved in 10 mM sodium phosphate (Na2HPO4/NaH2PO4) buffer, pH 7.5, and serially diluted from 10 μM to 0.2 μM. Bacteria were incubated at 37 °C overnight in LB (Luria–Bertani) broth and diluted to give approximately 5×105 CFU (colony-forming units/ml. In each assay peptide solutions were added to each bacteria dilution, incubated for 4 h, and samples were plated on Petri dishes and incubated at 37 °C overnight. The CFU in each Petri dish was counted, the average and the S.E.M. calculated and represented in a semi-logarithmic plot.
MIC (minimal inhibitory concentration) determination
Antimicrobial activity was expressed as the MIC, which is defined as the lowest concentration of peptides that completely inhibits microbial growth. MIC of each peptide was determined from two independent experiments performed in triplicate for each peptide concentration. Peptides were dissolved in 10 mM sodium phosphate buffer, pH 7.5 and serially diluted from 100 μM to 0.2 μM. Bacteria were incubated at 37 °C overnight in LB broth and diluted to give approximately 5×105 CFU/ml. In each assay peptide solutions were added to each bacteria dilution, incubated for 4 h and samples were plated on to Petri dishes and incubated at 37 °C overnight.
LUVs (large unilamellar vesicles) of a defined size (approx. 200 nm) were prepared from a vacuum-drying lipids chloroform solution by extrusion through 800, 400 and 200 nm polycarbonate membranes as described in . The lipid suspension was frozen and thawed several times prior to extrusion. Liposomes containing DOPC/DOPG (3:2 molar ratio) were obtained. A 1 mM stock solution of liposome suspension in 10 mM Tris/HCl, pH 7.4, and 0.1 M NaCl was prepared.
Tryptophan fluorescence emission spectra were recorded using a 280 nm excitation wavelength. Slits were set at 2 nm for excitation and 5–10 nm for emission. Emission spectra were recorded from 300–400 nm at a scan rate of 60 nm/min, in a 10 mm×10 mm cuvette with stirring immediately after sample mixing. Protein and peptide spectra at 0.5 μM in 10 mM Hepes buffer, pH 7.4, were obtained at 25 °C in the absence or presence of 200 μM liposome suspension, 200 μM LPS, assuming a 90000 g/mol molecular mass, or 200 μM LTA, as calculated from a 2200 molecular mass reference value. Fluorescence measurements were performed on a Cary Eclipse spectrofluorimeter. Spectra in the presence of liposomes were corrected for light scattering by subtracting the corresponding LUV background. For each condition three spectra were averaged. The fluorescence spectra were also calculated as a function of the frequency scale (wave number) and adjusted using a log-normal function as detailed in .
Depth-dependent fluorescence quenching experiments using labelled brominated lipids
Quenching of protein tryptophan residues by brominated lipids was introduced to analyse the relative location of both Trp10 and Trp35 residues upon membrane interaction. LUVs of DOPC/DOPG (3:2 molar ratio) were prepared as described in , containing either (6,7)-, (9,10)-, or (11,12)-Br2PC. Brominated lipids were mixed at increasing percentages, from 20 up to 80%, with unlabelled DOPC/DOPG vesicles. The emission fluorescence spectra were recorded at 25 °C using the following conditions: 10 mM Tris/HCl buffer, pH 7.4, 0.5 μM protein concentration and 200 μM liposome concentration, excitation wavelength of 280 nm, with excitation and emission slits set to 2 nm and 10 nm respectively. For each condition three spectra were averaged. The quenching efficiency of brominated lipids on protein tryptophan fluorescence was determined by calculating the area under the fluorescence spectra in range of 320 to 380 nm. Relative fluorescence intensities (F0/F) were compared, where F0 is the fluorescence intensity in the absence of quencher. To calculate the depth of insertion of the protein in the lipid bilayer the distribution analysis (DA) was employed, as detailed in .
ANTS/DPX liposome leakage assay
The ANTS/DPX liposome leakage fluorescence assay was performed as described in [16,28]. Briefly, a unique population of LUVs of DOPC/DOPG (3:2 molar ratio) lipids was obtained containing 12.5 mM ANTS, 45 mM DPX, 20 mM NaCl and 10 mM Tris/HCl, pH 7.5. The ANTS/DPX liposome suspension was diluted to 30 μM and was incubated at room temperature (25 °C) in the presence of ECP or the synthetic peptides. The leakage activity was assayed at different polypeptide concentrations, up to 8 μM, by following the release of the liposome content. Fluorescence was measured using a 386 nm excitation wavelength and 535 nm emission wavelength. Slits were set at 5 nm and 10 nm for excitation and emission respectively. The percentage of leakage (%L) produced by the proteins after 1 h of incubation with the liposomes was calculated with the following equation: %L=100(Fp−F0)/(F100−F0), where Fp is the final fluorescence intensity after addition of the protein (1 h), F0 and F100 are the fluorescence intensities before addition of the protein and after addition of 0.5% Triton X100. For each protein concentration three calculated leakage values were averaged. Leakage could not be quantified for those samples where the assay conditions triggered LUV precipitation, such as LUV samples incubated with high ECP protein concentrations.
LUV liposome aggregation
Aggregation of LUV lipid vesicles was monitored by measuring the scattering intensity with the excitation and emission wavelengths set at 470 nm using a Cary Eclypse spectrofluorimeter and cuvettes with an optical path of 1 cm. Prior to the addition of ECP and peptides, the vesicles were allowed to equilibrate for 15 min at room temperature. The buffer used was 10 mM Hepes, pH 7.4. Final assay conditions were 200 μM lipid and from 0.04 μM to 4 μM polypeptide range, in 10 mM Hepes buffer, pH 7.4. After 30 min of incubation at room temperature, the signal was read at 90° from the excitation beam with slits at 5 nm and 10 nm.
The far-UV CD spectra were collected with a JASCO J-715 spectropolarimeter. Spectra were recorded from 250 nm up to 190 nm, at 1 nm intervals, 1 nm bandwidth, with a scan speed of 10 nm/min, at 25 °C. 0.2 cm pathlength cuvettes were used. Mean-residue ellipticity [θ] (deg·cm2·dmol−1) was calculated using the formula: where MRW is the mean residue molecular mass of the protein, c is the protein concentration and l is the cell pathlength. Data of four consecutive scans were averaged and a linear smoothing from each five consecutive measurements was applied. ECP and peptides spectra in the absence and presence of SDS and LPS were recorded. ECP or peptide at 4–8 μM, diluted in 5 mM sodium phosphate buffer, pH 7.5, 1 mM SDS and 1 mM LPS (assuming a 90000 g/mol molecular mass), were used. Samples were centrifuged for 5 min at 10000-g before use. For near-UV CD, spectra were recorded from 340 nm up to 255 nm, at 1 nm intervals, 1 nm bandwidth, with a scan speed of 10 nm/min, at 25 °C in 1 cm pathlength cuvettes. Data from four consecutive scans were averaged. ECP and peptides spectra in the absence and presence of LPS were recorded. ECP or peptide at 80–100 μM, diluted in 5 mM sodium phosphate buffer at pH 7.5, and 1 mM LPS (assuming a 90000 g/mol molecular mass), were used. Percentage of each secondary structure was estimated using the JASCO software, based on the calculation method described by Yang et al. .
Bioinformatic analysis tools for peptide characterization
Physicochemical properties of peptides (pI and net charge) were predicted using the Swiss-Prot expasy server (http://www.expasy.ch/sprot/). Transmembrane-prediction domain analysis was performed using the Membrane Protein Explorer server (MPEx; http://blanco.biomol.uci.edu/mpex). Sequences with aggregation propensity were predicted using the Aggrescan server .
Bacterial cytoplasmic membrane depolarization assay
Membrane depolarization was monitored as described in , using the DiSC3(5) lipophilic dye, which changes its fluorescence intensity in response to changes in transmembrane potential. E. coli and S. aureus cells were grown to mid-exponential phase and resuspended in 5 mM Hepes/KOH, 20 mM glucose, and 100 mM KCl at pH 7.2 to an attenuance (D600) of ∼0.05. DiSC3(5) was added. Changes in the fluorescence due to the alteration of the cytoplasmic membrane potential were continuously monitored at 20 °C by fluorescence emission at 670 nm using an excitation wavelength of 620 nm. When the dye uptake was maximal, as indicated by a stable reduction in the fluorescence because of quenching of the accumulated dye at the membrane, polypeptide samples were added at a final concentration of 4 μM. All conditions were assayed in duplicate. The time necessary to reach a stabilized maximum fluorescence reading was recorded for each condition.
Fluorescent probe displacement assay for lipolysaccharide binding
LPS binding was assessed using the fluorescent probe BC as described in . BC binds strongly to native LPS, specifically recognizing the lipid A portion. LPS-binding assays were carried out in 5 mM Hepes buffer, pH 7.5. The displacement assay was performed by the addition of 1–2 μl aliquots of a stock solution of ECP and peptides to 1 ml of a continuously stirred mixture of LPS at 10 μg/ml and BC (10 μM). The BC excitation wavelength was 580 nm and emission wavelength was 620 nm. Excitation and emission slits were set at 2.5 nm and 20 nm respectively. Final values correspond to an average of four replicates. Quantitative effective displacement values (ED50) were calculated. The ED50 was computed at the midpoint of the fluorescent signal against the protein concentration of the displacement curve by a curve-fitting of the data to the equation: where OF is the occupancy factor, F0 the fluorescence intensity of BC alone, Fmax is the intensity in the presence of LPS at saturation concentration and F is the intensities of the LPS/BC mixtures at each displacer concentration. Polymyxin B and RNase A were used as positive and negative controls respectively.
SEM (Scanning electron microscopy)
Cultures of E. coli and S. aureus (1 ml) were grown at 37 °C to mid-exponential phase (D600 ∼0.4) and incubated with 10 μM ECP and the derived peptides in PBS at room temperature. Sample aliquots (500 μl) were taken after up to 4 h of incubation and prepared for SEM analysis as described in . The micrographs were viewed at a 15 kV accelerating voltage on a Hitachi S-570 scanning electron microscope, and a secondary electron image of the cells for topography contrast was collected at several magnifications.
TEM (Transmission electron microscopy)
Mid-logarithmic phase E. coli and S. aureus cells (D600 ∼0.4) were incubated with 10 μM ECP and peptides for 4 h. After treatment, bacterial pellets were pre-fixed with 2.5% (w/v) glutaraldehyde and 2% (w/v) paraformaldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, for 2 h at 4 °C, and post-fixed in 1% (w/v) osmium tetroxide in 0.1 M sodium cacodylate buffer, pH 7.4, for 2 h at 4 °C. The samples were dehydrated in acetone (50, 70, 90, 95 and 100%). The cells were immersed in EPON (liquid epoxy) resin, and ultrathin sections were examined on a JEOL JEM 2011 microscope.
Sequence analysis of ECP suggests that the N-terminus might contain an antimicrobial domain. Applying a predictive strategy based on previous high-throughput peptide library screening results , a bactericidal propensity value was assigned to each amino acid (M. Torrent, V. Nogués and E. Boix, unpublished work). On ECP a main potential active region at the N-terminus (residues 30–45) was identified. To test the activity of this region of ECP, the full N-terminal domain, ECP-(1–45), as well as two smaller representative peptides, ECP-(1–19) and ECP-(24–45), were chosen for the present study (Figure 1). ECP-(1–45) includes the first ECP α-helix elements (α1, α2 and α3) and the first β-region (β1). The ECP-(24–45) peptide spans the main predicted sequence and includes the Trp35–Arg36 residues previously identified as the determinant for cytotoxicity and membrane lysis activity [16,28,32]. The ECP-(1–19) peptide, which corresponds to the first α1 domain, represents the ECP homologue of the S-peptide in RNase A .
The bactericidal properties of the three peptides against two representative Gram-negative and -positive strains (E. coli and S. aureus respectively) have been compared with those of the full ECP sequence. Analysis of the IC50 and the MIC values (Table 1) shows the ECP-(1–45) segment to be practically equivalent to the entire native sequence in terms of antimicrobial activity against both strains. Within the ECP-(1–45) segment, activity is shown to be mostly confined to residues ECP-(24–45), whereas the ECP-(1–19) peptide requires approx. 10–20-fold higher concentrations to display significant bactericidal activity. As shown in Table 1, ECP and the three tested peptides displayed similar active ranges against both E. coli and S. aureus.
The bactericidal effects of the peptides on either strain have been further analysed to better characterize their distinct properties. ECP can depolarize the cytoplasmic membrane of both E. coli and S. aureus cells . The evaluation of the protein capacity to depolarize the bacteria cytoplasmic cell is an indirect method to assess its ability to interact at the bacteria surface and alter the inner membrane. Previous results on ECP depolarization capacity indicated that its activity is higher in the Gram-negative-tested strain, probably as a consequence of the easier access of the protein to the cytoplasmic membrane . In fact, ECP can alter by itself the Gram-negative strain outer membrane structure . We compared the protein depolarization capacity with the corresponding N-terminal peptides (Figure 2). ECP-(1–45) is the most active segment, whereas ECP-(24–45) retains an intermediate activity and ECP-(1–19) shows the worst performance, as previously observed for the corresponding bactericidal values (Table 1). However, significant differences between the two strains of bacteria are found. Whereas in E. coli ECP-(1–45) and ECP-(24–45) both have intermediate activity and ECP-(1–19) is quite inactive, in S. aureus ECP-(1–45) is nearly equipotent with ECP, and ECP-(1–19) is totally inactive.
For a better understanding of the cytotoxic activities of the peptides on bacteria we analysed their effects by electron microscopy. Potential damage on both the bacteria wall surface and the cytoplasmic membrane can be visualized by TEM. In E. coli, ECP promotes outer membrane detachment, alteration of the overall cell shape and partial loss of cell content . Comparison of the peptides with ECP confirms that both ECP-(1–45) and ECP-(24–45) are very active against E. coli (Figure 3). TEM micrographs include many hypodense damaged cells with vacuolization, local outer membrane detachment and loss of characteristic baton-shaped morphology. In contrast, practically no damaged cells can be seen upon treatment with ECP-(1–19). For S. aureus (see Supplementary Figure S2 available at http://www.BiochemJ.org/bj/421/bj4210425add.htm), TEM micrographs also show some hypodense cells for ECP-(1–45) and ECP-(24–45), although the overall damage is comparatively much less for this strain. ECP-(1–45) relative damage is similar to ECP, while ECP-(24–45) is slightly less active. As above, no significant effect is visible for ECP-(1–19).
Complementary analysis by SEM further illustrated the effect of the peptides on the bacterial surface. For ECP, previous data indicated a high ability to aggregate E.coli cells and to damage their wall envelope . This cell agglutination ability is retained by ECP-(1–45) (Figure 3), whereas no aggregates are observed upon incubation with the shorter peptides ECP-(24–45) and ECP-(1–19). On the other hand, significant surface wall damage is evident for cells treated with both ECP-(1–45) and ECP-(24–45), whereas little effect is seen upon incubation with ECP-(1–19). SEM micrographs of ECP-treated S. aureus cells did not reveal any aggregate, as reported previously . Surface wall damage for S. aureus cells is only mild compared with the treated E. coli cells but some local blebs appear. Cells incubated with ECP-(1–45) show nearly the same pattern, with ECP-(24–45) wall damage is quite mild and with ECP-(1–19) it is undetectable (see Figure S2).
Outer membrane detachment, visualized by electron microscopy, is considered to be related to an uncommonly high ability of ECP to bind LPS, the main component of Gram-negative bacterial surfaces, and its antigenic portion, the lipid A . We compared the LPS binding capacity of ECP and its N-terminal fragments. A decreasing order of LPS binding abilities, from ECP-(1–45) to ECP-(1–19), is registered (Table 2, and see Supplementary Figure S3 available at http://www.BiochemJ.org/bj/421/bj4210425add.htm). The displacement ability of ECP-(1–45) is considerably lower than that of ECP, but still high if compared with the positive control polymyxin B (Figure S3). On the other hand, ECP-(1–19) does not have any significant affinity to LPS. The LPS binding process can also be followed by the analysis of the intrinsic fluorescence spectra (Table 2). The recorded blue-shifts suggest that tryptophan residues are involved in the interaction and that Trp35 might be the residue that alters its environment most upon binding. Whereas the native protein does not modify its overall structure upon LPS binding, as assessed by CD spectra, we observe significant changes for the peptides, with an increase in their secondary structure content (see Figure 6). The near-UV spectra indicate that changes in the local environments of the aromatic residues take place in the presence of LPS. A careful inspection reveals that ECP-(1–19) does not significantly modify its overall profile, but that both ECP-(1–45) and ECP-(24–45) register an altered profile. In particular, ECP-(1–45) drastically shifts the bands in the characteristic ranges for tyrosine and tryptophan, suggesting the involvement of the exposed Tyr33 and Trp35 residues in LPS binding. Interestingly, both ECP-(1–45) and ECP-(24–45) peptide far-UV spectra in the presence of LPS indicate a much more structured conformation upon binding (see Figure 6).
Additionally, in order to identify potential interactions between the protein and the N-terminal peptides at the Gram-positive bacteria surface, we have also analysed the changes in the intrinsic fluorescence spectra in the presence of LTA. Electrostatic interactions between the polypeptide cationic residues and the negative-charged groups of LTA are expected. Results confirmed the binding of LTA and the contribution of tryptophan residues (Table 2). Pronounced shifts are mainly recorded for ECP-(1–45) and ECP-(24–45) peptides, with comparable values, whereas the blue-shift corresponding to ECP-(1–19) is only minor, suggesting the involvement of Trp35 in the interaction.
Interaction with lipid bilayers
The correlation between ECP cytotoxic activity and its membrane-destabilizing capacity has been previously reported [28,32]. Detailed analysis of ECP action using LUV and giant unilamellar vesicles as membrane models revealed aggregation and leakage activities in the nanomolar range [16,18]. In the present study, we characterized the interaction of ECP N-terminal peptides with LUV made up of DOPC/DOPG phospholipids.
The intrinsic fluorescence signals of the peptides were first analysed. ECP includes two tryptophan residues, Trp10 and Trp35, both present in the ECP peptides analysed, which have been classified into different spectral classes . Comparison of ECP with single tryptophan mutants indicates that most of the blue-shift in the fluorescence spectrum is due to the association of solvent-exposed Trp35 with the lipid bilayer . The intrinsic fluorescence spectra of the peptides (see Supplementary Figure S4 available at http://www.BiochemJ.org/bj/421/bj4210425add.htm) indicate that, as expected, both tryptophan residues are far more highly solvent-exposed than in their respective locations within full-length ECP. Of the two, Trp10 is the residue experiencing a more pronounced change, from a buried location in ECP  to a more exposed position in ECP-(1–19).
When the peptide spectra are recorded in the presence of the DOPC/DOPG LUV, a significant blue-shift is observed in all cases, confirming the interaction with the lipid bilayer. Shifts for ECP-(24–45) and ECP-(1–45) are more pronounced than for ECP-(1–19), suggesting that Trp35 does undergo a more pronounced change on its microenvironment when the peptides associate with lipid bilayers. In fact, the similarity between spectra of ECP-(24–45) and ECP-(1–45) suggests that the Trp10 contribution to the final signal is minor.
The interaction with lipid bilayers has been further investigated by fluorescence quenching experiments using brominated phospholipids. For that purpose, LUV made up of DOPC/DOPG with PC dibromo derivates at several positions remote from the head group (6,7, 9,10 and 11,12 respectively) have been prepared. This methodology allows an estimation of the degree of peptide insertion into the lipid bilayers. The quenching efficiency of the differently brominated lipids has been recorded by comparing the respective fluorescence quenching slopes. For all peptides, the presence of either Trp10 or Trp35 allowed the monitoring of changes upon membrane interaction. The Stern–Volmer quenching slopes show a steep decrease from the 6,7- to the 11,12-dibrominated LUV, indicating a diminishing interaction between the tryptophan residues and the bromine-labelled phospholipid (see Supplementary Figure S5 available at http://www.BiochemJ.org/bj/421/bj4210425add.htm). From these results an average insertion of ∼10Å (1Å=0.1 nm), from the bilayer centre, can be estimated for the tryptophan residues of both ECP-(24–45) and ECP-(1–45), similar to that in ECP , while a slightly deeper insertion (∼8Å) is found for the ECP-(1–19) peptide (Figure 4).
Another way to evaluate lipid bilayer destabilization is by determining the ability to aggregate LUV, triggering leakage and release of vesicles' aqueous content. Thus ECP promotes the release of encapsuled ANTS/DPX at a 1:300 protein/lipid ratio and above . Consistent with these trends, the leakage ability of ECP-(1–45) is similar to ECP, that of ECP-(24–45) is slightly lower, whereas ECP-(1–19) requires much higher peptide/lipid ratios to induce significant leakage (Figure 5A).
Liposome aggregation can also be monitored by the increase in scattering intensity at 470 nm. Using this method, ECP has detectable LUV aggregation above a 1:1500 protein/lipid molar ratio , which is mostly retained by ECP-(1–45) but, surprisingly, lost in the ECP-(24–45), whereas partial aggregating behaviour is observed for ECP-(1–19) (Figure 5B).
Finally, we have used CD to investigate the conformational changes brought about by peptide–membrane interaction (Figure 6). The CD spectrum of ECP in aqueous solution has been reported , with the inferred secondary structure values showing some deviation from those derived from X-ray crystallography . These differences were mainly attributed to the contribution of aromatic amino acid side chains to the spectral profile. Comparison of the near-UV spectra of ECP and the peptides (Figure 6), reveals significant changes in local environment for aromatic residues in ECP-(1–45) and ECP-(24–45).
We have also analysed the far-UV spectra for ECP and the derived peptides in the absence and presence of SDS (Figure 6). The results for ECP indicate that no overall conformation changes have taken place (Figure 6A). In contrast, for the three peptides some significant differences were observed (Figures 6B–6D). SDS induced the transition to more structured conformations for the three peptides, where a reduction in the β-sheet content and a significant increase in the α-helical content is registered. Thus we can conclude that when exposed to a lipid environment the peptides undergo conformational reorganization, with a considerable increase in their structured content. Comparison of the α-helix content for each peptide indicates an increase of 25, 35 and 70% for ECP-(1–19), ECP-(24–45) and ECP-(1–45) respectively. This is consistent with their respective abilities to disrupt lipid bilayers (Figure 5A). On the other hand, prediction of potential transmembrane segments using the MPEx software identifies a segment from residues 9 to 27 with a high tendency to form an α-helix in a lipid environment. This might explain why the peptide with the highest activity on lipid bilayers is ECP-(1–45), which includes this segment in its entirety.
ECP is an eosinophil-secreted protein involved in inflammation processes with diverse antipathogen properties [14,37]. In previous studies we have characterized its bactericidal activity and identified some structural determinants for its cytotoxic capacity [16–18,28]. ECP diverged from the other eosinophil RNase, EDN, under an unusual evolutive pressure and acquired a higher cationicity and cytotoxicity . We have now identified an N-terminal region that retains both the bactericidal and membrane-disrupting activities of ECP. The selected 1–45 segment fulfils the common structural and chemical criteria for AMPs [1,39,40]. It includes a high number of arginine residues, which confer the three synthetic peptides with a net positive charge [+2, +5 and +8 for ECP-(1–19), ECP-(24–45) and ECP-(1–45) respectively] as well as a high pI (predicted as 12, 11.5 and 11.9). On the other hand, the studied N-terminus includes two cysteine residues, which were substituted in the synthetic peptides by serine to avoid potential formation of intra- or inter-molecular disulfide bridges. Although the peptides cannot reproduce this full scenario (i.e. the context of the disulfide-bonded protein structure) the use of synthetic peptides with cysteine substitutions or deletions is a useful approach, as reported for other antimicrobial proteins . In fact, some antimicrobial proteins do retain activity in their denatured state . Furthermore, there is also a report of an antimicrobial RNase A homologue, the salmon RNase, which preserves activity when fully denatured . Moreover, the characterization of protein-derived peptides was also successfully applied to avian leucocyte RNase A homologues (see Figure 1S), where one of the peptides outside the protein scaffold retained bactericidal activity . A cationic cluster at the N-terminus was also identified for the skin-derived RNase 7, another human RNase A homologue (see Supplementary Figure S1) that contributes to the innate immunity at the skin–chemical barrier [43,44].
Comparison of the bactericidal and membrane-destabilizing activities of ECP and the three synthetic peptides indicates that the ECP N-terminus retains some sequence determinants involved at the different levels of ECP's cytotoxic capacity. The longest, ECP-(1–45), displays nearly all of the bactericidal activity of the whole protein against E. coli and S. aureus strains, and partially retains the LPS binding and depolarization abilities. Additionally, ECP-(1–45) has almost the same membrane interaction capacity as ECP, with nearly equivalent leakage and liposome aggregation activities. The inner ECP-(24–45) peptide can partially, but not fully, reproduce the bactericidal and membrane destabilization capacities of the intact protein, and the short ECP-(1–19) peptide has almost no toxicity for bacterial cells.
Altogether, the results obtained reveal, once again, that membrane disruption cannot by itself explain the toxicity of ECP towards bacterial cells. Furthermore, the observed differences in the activity of ECP against Gram-negative and Gram-positive strains also suggest the involvement of strain-specific determinants in the killing events. Electron microscopy illustrates diverse abilities of the peptides at the bacterial wall surface. Similar bactericidal activities do not readily translate to bacteria wall damage, suggesting that cell viability is lost before any cell surface destruction is visualized. Although ECP and its derived peptides are bactericidal against both E. coli and S. aureus, bacterial wall damage is much more pronounced in the Gram-negative strain. ECP shows a specific capacity to destroy the E. coli outer membrane and to agglutinate the cells , and this ability is only retained by the full ECP-(1–45) fragment.
The propensity of ECP and its derived peptides to bind lipid bilayers may also be facilitated by their hydrophobicity and aggregation tendency. We used the Aggrescan software  to screen the ECP sequence for the presence of hydrophobic patches that can favour aggregation. One such sequence is found at the ECP N-terminus (residues 8–16), and is not present in EDN. An N-terminal aggregation domain has also been identified in the skin-derived RNase 7, although not in other screened human RNases (results not shown). An exposed hydrophobic patch may promote both protein–protein and also protein–lipid bilayer interactions. The fact that ECP-(1–19) partially retains a lipid aggregation ability, whereas the internal segment ECP-(24–45) does not, also supports the presence of a potential ‘aggregation-promoting’ domain at the ECP N-terminus.
The short peptide ECP-(24–45) seems to include a key region for bactericidal action, as well as for specific binding to the bacteria wall. Both ECP-(1–45) and ECP-(24–45) retain part of ECP's affinity for LPS, with Tyr33 and Trp35 residues that might participate in the interaction. Recent results on the ECP–heparin binding affinity indicate that charged and aromatic residues 34–38 are involved in this selective ability , identifying a key domain responsible for protein anchoring to the cell surface. Recent work by Döring and co-workers  on the tyrosine nitration process activated during eosinophil maturation, identified in ECP a unique tyrosine nitration at residue 33 in ECP, and suggested that tyrosine nitration may modulate ECP aggregation and/or its interaction with other eosinophil proteins.
In view of all of the above results, we suggest that residues 1–45 of ECP embrace the main functional domain of the protein. In the present study we have shown that the ECP-(1–45) peptide reproduces most of ECP's antimicrobial properties. The inner ECP-(24–45) sequence retains the key bactericidal region, but loses some of ECP's membrane-destabilizing capacities. One can plausibly propose that the first 19 residues are also necessary either for preserving native-like conformation or for promoting aggregation, or both. Although devoid of significant bactericidal action, this 1–19 region retains membrane-destabilizing properties, consistent with data suggesting that it includes an aggregation patch.
In any event, ECP-derived peptides are one model of novel sources of AMPs as alternative antibiotics against resistant strains. The ECP E. coli cells agglutinating activity is indeed a promising property that is being investigated in the search for alternative antibiotics as a mechanism to induce bacteria clearance by the host phagocytic cells . Given the pharmaceutical interest in the development of new LPS-binding substances for treatment of immune disorders, the partial LPS-binding ability in residues 24–45 also offers a starting template for the design of new peptide-derived immunomodulator drugs.
Marc Torrent carried out all the functional and structural characterization of the peptides, prepared all the corresponding artwork and actively participated in the methodology development, experiment setup and discussion and interpretation of results. David Andreu contributed to the peptides' design and final writing of the manuscript. Beatriz de la Torre synthesized the peptides. Victòria Nogués contributed to the paper's discussion and correction. Ester Boix conceived the experimental setup and drafted the manuscript. All authors read and approved to the final manuscript.
This work was supported by the Ministerio de Educación y Cultura [grant numbers BMC2003–08485-C02–01, BFU2006–15543-C02–01]; and by the Fundació La Marató de TV3 [grant number TV3–031110]. M.T. was the recipient of a predoctoral fellowship from the Generalitat de Catalunya. Work at Universitat Pompeu Fabra was supported by the Spanish Ministries of Health (Fondo de Investigaciones Sanitarias) [grant number PI040885]; Spanish Ministries of Education and Science [grant numbers BIO2005–07592-CO2–02, PET2006–00139–00]; and by Generalitat de Catalunya [grant number SGR00494].
SEM was performed at the Servei de Microscopia of the Universitat Autònoma de Barcelona (UAB), Barcelona, Spain. We thank Francisca Cardoso, Alejandro Sánchez and Francesc Bohils for their assistance in electron micrography preparation. Fluorescence assays were done at the Laboratori d’Anàlisi i Fotodocumentació, Fac. Biociències, UAB, Barcelona, Spain. CD spectra were recorded at the Servei d’Anàlisi Química, Fac. Ciències, UAB, Barcelona, Spain.
Abbreviations: AMP, antimicrobial peptide; ANTS, 8-aminonaphthalene-1,3,6-trisulfonic acid disodium salt; BODIPY®, boron dipyrromethane (4,4-difluoro-4-bora-3a,4a-diaza-s-indacene; BC, BODIPY® TR cadaverine, BPI, bactericidal permeability-increasing protein; Br2PC, 1-palmitoyl-2-stearoyl-dibromo-sn-glycero-3-phosphocholine; CFU(s), colony-forming unit(s); DiSC3(5), 3,3-dipropylthiacar-bocyanine; DOPC, dioleoyl phosphatidylcholine; DOPG, dioleoyl phosphatidylglycerol; DPX, p-xylenebispyridinium bromide; ECP, eosinophil cationic protein; ED50, quantitative effective displacement values; EDN, eosinophil-derived neurotoxin; LB, Luria–Bertani; LPS, lipopolysaccharide(s); LTA, lipoteichoic acid(s); LUV, large unilamellar vesicles; MALDI–TOF-MS, matrix-assisted laser-desorption ionization–time-of-flight MS; MIC, minimal inhibitory concentration; SEM, scanning electron microscopy; TEM, transmission electron microscopy
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