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Originally published In Press as doi:10.1074/jbc.M606031200 on July 25, 2006

J. Biol. Chem., Vol. 281, Issue 39, 28565-28574, September 29, 2006
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A Synergism between Temporins toward Gram-negative Bacteria Overcomes Resistance Imposed by the Lipopolysaccharide Protective Layer*

Yosef Rosenfeld{ddagger}, Donatella Barra§, Maurizio Simmaco§, Yechiel Shai{ddagger}, and Maria Luisa Mangoni{ddagger}1

From the {ddagger}Department of Biological Chemistry, Weizmann Institute of Science, Rehovot 76100, Israel and the §Istituto Pasteur-Fondazione Cenci Bolognetti, Dipartimento di Scienze Biochimiche and Unità di Diagnostica Molecolare Avanzata, II Facoltà di Medicina e Chirurgia, Università di Roma La Sapienza, Azienda Ospedaliera S. Andrea, 00189 Roma, Italy

Received for publication, June 23, 2006 , and in revised form, July 20, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Temporins are short and homologous antimicrobial peptides (AMPs) isolated from the frog skin of Rana genus. To date, very little is known about the biological significance of the presence of closely related AMPs in single living organisms. Here we addressed this question using temporins A, B, and L isolated from Rana temporaria. We found that temporins A and B are only weakly active toward Gram-negative bacteria. However, a marked synergism occurs when each is mixed with temporin L. To shed light on the underlying mechanisms involved in these activities, we used various experimental strategies to investigate: (i) the effect of the peptides' interaction on both the viability and membrane permeability of intact bacteria and spheroplasts; (ii) their interaction with lipopolysaccharides (LPS) and the effect of LPS on the oligomeric state of temporins, alone or combining one with another; (iii) their structure in solution and when bound to LPS, by using circular dichroism and ATR-FTIR spectroscopies. Our data reveal that temporin L synergizes with A and B by preventing their oligomerization in LPS. This should promote their translocation across the outer membrane into the cytoplasmic membrane. To the best of our knowledge, this is the first study that explains how a combination of native AMPs from the same species can overcome bacterial resistance imposed by the LPS leaflet.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In addition to their highly specific cell-mediated adaptive immune response, vertebrates possess an additional protection mechanism consisting of different classes of gene-encoded host-defense peptides (1–5). These peptides represent key components of the innate immune system found throughout the living world. Most of them, also termed antimicrobial peptides (AMPs),2 are generally composed of about 12 to 50 amino acids (6, 7). Although many AMPs differ in their chain lengths, hydrophobicity, and distribution of basic and acidic residues, they have common structural features, such as a net positive charge and the capacity to adopt an amphiphatic {alpha}-helix or a beta-sheet structure upon their association with the pathogen membrane (8–10). These properties allow many of them to initially bind the negatively charged microbial membrane and alter its integrity, leading to cell death (9, 11, 12). However, a few AMPs have been shown to have additional/alternative intracellular targets besides the cytoplasmic membrane (13–18).

Interestingly, several kinds of plants and animals can synthesize a considerable number of AMPs with relatively minor sequences and structural differences among them. For example, all frog species produce their own unique set(s) of defense peptides constituting families of 2–40 closely related members (19). These families encompass bombinins and bombinins H, from the European toads Bombina variegata and Bombina orientalis (20–23), magainins from the African clawed frog Xenopus laevis (24), dermaseptins from the South American arboreal frog Phyllomedusa sauvagii (25, 26), and those from the Rana genus (e.g. brevinins, ranalexins, esculentins, and temporins) (19, 27–30). Other examples of families of AMPs with high structural similarities include cecropins from insects (31, 32), the mammalian defensins and cathelicidins, isolated from circulating leukocytes and epithelia (33, 34), and the human salivary histatins (35). In Amphibia, temporins represent the largest family of AMPs (more than 40 members), and up to 10 isoforms have been identified in a single specimen (30). They are among the smallest amphipathic {alpha}-helical AMPs found in nature to date (10 to 14 amino acids), containing only a few positively charged amino acids (the net charge at a neutral pH ranging from 0 to +3) and are active particularly toward Gram-positive bacteria. Interestingly, a member of this family, temporin L, is an exception, being highly potent against both Gram-positive and Gram-negative strains. So far, biological and mode of action studies have been carried out mainly with temporins A (FLPLIGRVLSGIL-NH2), B (LLPIVGNLLKSLL-NH2), and L (FVQWFSKFLGRIL-NH2) from Rana temporaria. These peptides modify the permeability of the microbial membrane, allowing the passage of small and large size molecules (e.g. the enzyme beta-galactosidase), in a rapid and dose-dependent manner (36–38).

The physiological significance of the existence of multiple forms of AMPs within the same organism has been addressed in only a few cases, showing a synergism between them, against several bacteria. These include the combination between magainin 2 and PGLa from X. laevis (39), between different isomers of dermaseptins (40), cathelicidins and defensins (41), and between hepcidin and moronecidin, isolated from bass gill tissue (42). However, except for the pair magainin-2/PGLa, the molecular mechanism accounting for the synergistic effects is not yet known.

Here, we report on the synergism of temporin L when combined with temporins A or B to overcome resistance imposed by the anionic lipopolysaccharide (LPS or endotoxin), the major component of the outer membrane of Gram-negative bacteria (43), along with a plausible new mode of action underlying this effect. Among these bacteria is Aeromonas hydrophila, which is associated with the global decline of amphibians (44, 45), and which is resistant to the majority of natural AMPs (36, 45). Our data suggest that temporin L prevents the aggregation of temporins A and B within LPS, thus allowing their translocation across the outer membrane into the cytoplasmic membrane.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Rink amide 4-methyl benzhydrylamine resin and 9-fluorenylmethoxycarbonyl (Fmoc) protected amino acids were obtained from Calbiochem-Novabiochem AG (Switzerland). Other reagents used for peptide synthesis included trifluoroacetic acid (Sigma), piperidine (Merck), N,N-diisopropylethylamine (Sigma), N-hydroxybenzotriazole hydrate (Aldrich), 2-(1H-benzotriazole-1-yl)-1,1,3,3 tetramethyluronium hexafluorophosphate, and dimethylformamide (peptide synthesis grade, Biolabs). Proteinase-K and LPS (from Escherichia coli O111:B4) were purchased from Sigma. Buffers were prepared in double glass-distilled water.

Peptide Synthesis, Fluorescent Labeling, and Purification— Peptides were synthesized by a Fmoc solid phase method on Rink amide 4-methyl benzhydrylamine resin, using an ABI 433A automatic peptide synthesizer. Cleavage of the peptides from the 4-methyl benzhydrylamine resin resulted in C terminus-amidated peptides. To label the peptides, the Fmoc protecting group was removed from the N terminus of the resin-bound peptides by incubation with piperidine for 12 min, whereas all the other reactive amine groups of the attached peptides were kept protected. The resin-bound peptides were washed twice with dimethylformamide, and then treated with rhodamine-N-hydroxysuccinimide (2 eq), in anhydrous dimethylformamide containing 2% N,N-diisopropylethylamine, leading to the formation of a resin-bound N-rhodamine peptide. After 24 h, the resin was washed thoroughly with dimethylformamide and then with methylene chloride. The three rhodamine-labeled temporins A (rhodamine-temporin A), B (rhodamine-temporin B), and L (rhodamine-temporin L) were then cleaved from the resin. All the peptides were purified by reverse phase high performance liquid chromatography on a C18 reverse phase Bio-Rad semi-preparative column (250 x 10 mm, 300-Å pore size, 5-cm particle size). The column was eluted in 40 min, using a linear gradient of 20–60% acetonitrile in water, containing 0.05% trifluoroacetic acid (v/v), at a flow rate of 1.8 ml/min. The purified peptides were further subjected to amino acid analysis and electrospray mass spectrometry to confirm their composition and molecular weights.

Antibacterial Activity of the Peptides—Susceptibility testing was performed by the microbroth dilution method according to the procedures outlined by the National Committee for Clinical Laboratory Standards (2001) using sterile 96-well plates. Aliquots (50 µl) of bacteria in mid-log phase at a concentration of 2 x 106 colony forming units/ml in culture medium (Mueller-Hinton, MH) were added to 50 µl of MH broth containing the peptide in serial 2-fold dilutions in 20% ethanol. Inhibition of growth was determined by measuring the absorbance at 600 nm with a 450 Bio-Rad Microplate Reader after an incubation of 18–20 h at 30 °C. Antibacterial activities were expressed as the minimal inhibitory concentration (MIC), the concentration of peptide at which 100% inhibition of growth was observed after 18–20 h of incubation.

In addition, a synergism between temporins was evaluated by adding combinations of two temporins, in a serial 2-fold dilution, to wells containing 1 x 105 colony forming units in a final volume of 100 µl. The ranges of peptide dilutions used were 0.15–40 µM for temporin L and 0.2–400 µM for temporins A and B. The fractional inhibitory concentration (FIC) index for combinations of two peptides was calculated according to the equation: FIC index = FICA + FICB = A/MICA + B/MICB, where A and B are the MICs of drug A and drug B in the combination, MICA and MICB are the MICs of drug A and drug B alone, and FICA and FICB are the FICs of drug A and drug B. The FIC indices were interpreted as follows: ≤0.5, synergy; 0.51–4.0, no interaction; >4.0, antagonism.

The following bacterial strains were used: the standard Gram-negatives E. coli D21, E. coli ATCC 25922, Pseudomonas aeruginosa ATCC 15692, and A. hydrophila Rt-6, isolated from the natural flora of R. temporaria (46); the standard Gram-positives Staphylococcus aureus Cowan I, S. aureus ATCC 25923; Bacillus megaterium Bm11, and Staphylococcus epidermidis ATCC 12228.

The Effect of LPS on the Killing Activity of Temporins—The effect of E. coli O111:B4 LPS on the killing activity of temporins was also measured. Briefly, each peptide (4 µM temporin L, 10 and 15 µM temporins A and B, respectively) was incubated with LPS, and dissolved in water (peptide:lipid ratio = 1:1 and 1:2) for 30 min at 37 °C. This mixture was then added to a suspension of exponentially growing S. aureus Cowan I cells in MH broth and incubated at 30 °C for up to 45 min. After 20 min, aliquots were withdrawn, diluted in MH, and plated onto MH agar plates for the counting of colony forming units.

Preparation of Bacterial Spheroplasts—Spheroplasts of E. coli D21 were prepared according to the procedure described by Sullivan et al. (47). Briefly, bacteria from a log-phase culture were collected by centrifugation at 3,000 x g and washed in 0.01 M phosphate buffer, pH 7.0 (PB). The cells were then pleated by centrifugation and resuspended in a half-volume of 0.5 M sucrose solution in PB to induce plasmolysis. Lysozyme (Sigma) was added to the cell suspension at a final concentration of 50 µg/ml. After incubation at 37 °C for 90 min with moderate shaking, the sample was diluted 1:1 with PB and EDTA was added to a final concentration of 10 mM. The cell suspension was incubated again at 37 °C for an additional 15 min. Thereafter, transition of the rod-shaped bacteria into spheres was determined by light microscopy. When ~80% of the cells were spheroplasted, the reaction was stopped by pelleting the cells at 500 x g for 15 min. The spheroplasts were then washed in 0.25 M sucrose in PB (SPB2), centrifuged at 500 x g for 15 min, and resuspended in SPB2.

The Effect of Combining Temporins on the Viability of Intact E. coli and Spheroplasts—Aliquots of 100 µl of SPB2 containing 2 x 107 intact bacteria or spheroplasts were treated with a combination of a sublethal concentration of temporin A (25 and 5 µM for intact cells and spheroplasts, respectively) and temporin L (4 and 2 µM for intact cells and spheroplasts, respectively) as well as with the peptides alone at 30 °C for 30 min. Afterward, bacterial viability was assayed by the reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) to insoluble formazan. Briefly, 100 µl of 1 mg/ml MTT was added to the bacterial suspension, transferred into a 96-well culture microplate, which was incubated for 1 h at 37°C. The reduced formazan was then solubilized by the addition of an equal volume of 10% (w/v) SDS, and measured in a 450 Bio-Rad Microplate Reader equipped with a 595-nm filter, after an overnight incubation at 37 °C. All assays were performed in triplicate, and the experiments were repeated twice. Cells without adding peptides were used as a control.

The Effect of Combining Temporins on the Plasma Membrane Permeation—To assess the ability of a combination of subinhibitory concentrations of temporins A or B and L to alter the membrane permeability of whole E. coli D21 cells and spheroplasts, 2 x 107 cells were mixed with 1 µM SYTOXTM Green in SPB2 for 5 min in the dark. After adding peptides, the increase of fluorescence, due to the binding of the dye to intracellular DNA, was measured at 30 °C in a microplate counter (Wallac 1420 Victor3TM, PerkinElmer Life Sciences) using 485- and 535-nm filters for excitation and emission wavelengths, respectively. Controls used were given by cells without adding peptides.

The Effect of LPS on the Oligomeric State of the Peptides as Determined by Rhodamine Fluorescence Dequenching Measurements—Rhodamine-labeled peptides (final concentration, 3 µM for temporin A and 1.5 µM for both temporins B and L) were added to 100 µl of PBS and changes in the intensity of the fluorescence emission were followed upon the addition of different concentrations of LPS, by using the same microplate counter with excitation and emission wavelengths set at 485 and 590 nm, respectively. Proteinase-K (80 µg/ml in PBS) was then added, and the resulting fluorescence was monitored. An increase in fluorescence indicates that the peptide exists as an oligomer (48). All fluorescence measurements were performed at 30 °C.

The Effect of the Interactions between the Peptides on Their Structure in Buffer or in LPS Solution—To determine whether the interaction between the peptides affects their structure, we followed the CD spectra of temporins A, B, and L using a Jasco J-500A spectropolarimeter after calibrating the instrument with (+)-10-camphorsulfonic acid. The spectra were scanned at 25 °C in a capped, quartz optical cell with a 0.5-mm path length. Spectra were obtained at wavelengths of 190 to 250 nm. Eight scans were taken for each peptide at a scan rate of 20 nm/min. Mean residue ellipticities were expressed as [{theta}] (deg cm2/dmol). The peptides (100 µM) were scanned alone or combined with two peptides in the presence or absence of LPS (25 µM, using Mr of 4,000) dissolved in PBS. The experimental results of peptide combinations were compared with the calculated average of the two signals of each peptide alone. Fractional helicities were calculated from the dichroic minimum at 222 nm, as previously described (49, 50).

The Structure of the Peptides within LPS and Their Effect on LPS Phosphate Groups—To obtain information on the influence of LPS on the secondary structure of the peptides, we used attenuated total reflectance Fourier-transform infrared (ATR-FTIR) spectroscopy. Spectra were obtained with a Bruker equinox 55 FTIR spectrometer equipped with a deuterated triglyceride sulfate detector and coupled to an ATR device as previously described (51). Briefly, peptides (final concentration 25 µM) were dissolved in 400 µl of LPS solution in double distilled water (25 µM), deposited on a ZnSe horizontal ATR prism, and dried under vacuum at 37 °C. Spectra were recorded and the respective pure phospholipid spectra were subtracted to yield the difference spectra. The background for each spectrum was a clean ZnSe prism. Hydration of the sample was achieved by introducing excess deuterium oxide (2H2O) into a chamber placed on top of the ZnSe prism in the ATR casting, and incubating for 15 min before acquisition of the spectra. The H/D exchange was considered complete after a total shift of the amide II band. Any contribution of 2H2O vapor to the absorbance spectra near the amide I peak region was eliminated by subtracting the spectra of pure lipids equilibrated with 2H2O under the same conditions.

ATR-FTIR Data Analysis—Prior to curve fitting, a straight baseline passing through the ordinates at 1700 and 1600 cm–1 was subtracted. To resolve overlapping bands, we processed the spectra using PEAKFITTM software (Jandel Scientific, San Rafael, CA). Second-derivative spectra were calculated to identify the positions of the component bands in the spectra. These wave numbers were used as initial parameters for curve fitting with Gaussian component peaks. Position, band widths, and amplitudes of the peaks were varied until (i) the resulting bands shifted by no more than 2 cm–1 from the initial parameters, (ii) all the peaks had reasonable half-widths (<20–25 cm–1), and (iii) good agreement between the calculated sum of all the components and the experimental spectra was achieved (r2 > 0.99). The relative contents of the different secondary structure elements were estimated by dividing the areas of individual peaks, assigned to a specific secondary structure, by the whole area of the resulting amide I band. The results of two independent experiments were averaged. The interactions of the peptides with LPS head groups were studied by monitoring the asymmetric stretching vibration of the negatively charged phosphate groups, {nu}as(PO2), in the range of ~1220 cm–1 (52).


Figure 1
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  		FIGURE 1.
Synergistic effect of temporins A and L on membrane permeation of E. coli D21 cells by following the intracellular influx of the vital dye SYTOXTM Green after adding peptides. Bacteria (2 x 107 cells in 100 µl) were incubated with 1 µM SYTOX Green in PBS. Once basal fluorescence reached a constant value, peptides were added (arrow, t = 0), and changes in fluorescence were monitored ({lambda}exc = 485 nm, {lambda}ems = 535 nm) and plotted as arbitrary units. Control bacteria, bold line. Peptide concentrations used were 4 µM temporin A ({blacksquare}), 0.7 µM temporin L ({circ}), and 4 µM temporin A + 0.7 µM temporin L ({blacktriangleup}). Data points are the mean of triplicate samples with S.D. values not exceeding 2% from a single experiment, representative of three different experiments. Results similar to those obtained with temporin A were found with temporin B and therefore are not shown.

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Synergism between Temporins toward Bacteria—We measured the ability of all combinations of temporins A, B, and L to inhibit the growth of several Gram-negative and Gram-positive bacterial strains, using the checkerboard titration method. The results are illustrated in Table 1. The data revealed FIC indices of 0.48 and 0.50 for temporins A+L and B+L pairs, respectively, against E. coli D21. This corresponds to a synergistic effect (an FIC index of ≤0.5 indicates synergy (53, 54)). Similar results were obtained against the Gram-negative E. coli ATCC 25922 and P. aeruginosa ATCC15692, but the greatest synergism was found against A. hydrophila Rt-6, with FIC values of 0.34 and 0.41 for the combination of temporins A + L and B + L, respectively. In contrast, an indifferent interaction was found when temporin A was used together with temporin B on all the tested Gram-negative bacterial strains (Table 1). Notably, when the peptides were assayed on Gram-positive bacteria, FIC values ranging from 0.55 to 2 were observed, pointing out the absence of any synergistic activity.


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  		TABLE 1
MICs of temporins A, B, L, and FIC indices of their combination against several bacterial strains

MICs are presented as average values from three independent measurements.

 
The Effect of Combining Temporins on the Plasma Membrane Permeation—We monitored the influx of SYTOX Green into E. coli cells, after the addition of temporins at their sublethal dose. This cationic dye (Mr = 600), which is excluded by intact membranes (55), but not from those having lesions of a size large enough to allow its entrance, increases its fluorescence when bound to intracellular nucleic acids. A marked and rapid enhancement of fluorescence intensity was detected after the addition of temporins A + L (Fig. 1) or B + L (data not shown) to the bacterial cells. This suggests that temporin L can synergize with the other two temporins by assisting them to increase the permeability of the membrane, which is otherwise slightly altered when microbes are exposed to the individual peptides (Fig. 1).

To understand whether this increase in activity occurred at the level of the inner membrane, we studied the effect of subinhibitory peptide concentrations on the viability of both E. coli intact cells and spheroplasts that lack a cell wall (Fig. 2A). Because only a weak improvement in bactericidal activity was detected when combinations of temporins were used against spheroplasts with respect to intact cells, the described synergic effect had to be associated with the cell wall of Gram-negative bacteria and presumably to their outer LPS membrane. This notion was also supported by the fact that no significant effects regarding the permeability of the spheroplasts' membrane were discerned after treating them with a combination of sublethal concentrations of both temporins A and L (Fig. 2B).

The Effect of LPS on the Organization of Temporins—To ascertain differences in the structural organization of the three temporins when in contact with LPS, we labeled the peptides at their N terminus with rhodamine, a fluorescent probe that is only slightly sensitive to the polarity of its environment. Note that this labeling did not interfere with the biological function of temporins (data not shown). When rhodamine-labeled monomers are self-associated and the rhodamine probes are in proximity, the result is self-quenching of the emission fluorescence. Moreover, a significant increase in the fluorescence intensity of the peptide after treatment with a proteolytic enzyme determines whether the peptide is self-associated or not (56). The following experiments suggest that LPS triggers the aggregation of rhodamine-temporins A and B, which are slightly assembled in solution, and in contrast, it induces the disaggregation of rhodamine-temporin L, which is highly self-associated in solution.


Figure 2
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  		FIGURE 2.
A, effect of temporins A and L on the viability of intact cells and spheroplasts of E. coli D21. Cells (2 x 107 in 100 µl) were incubated with a combination of sublethal concentrations of temporin A (25 and 5 µM for intact cells and spheroplasts, respectively) and temporin L (4 and 2 µM for intact cells and spheroplasts, respectively) as well as with peptides alone at 30 °C for 30 min. Afterward, bacterial viability was tested by the inhibition of MTT reduction. The values are the mean of triplicate samples ± S.D. values (error bars) from a single experiment, representative of two different experiments. B, effect of temporins A and L on the membrane permeation of spheroplasts of E. coli D21. Cells (2 x 107 in 100 µl) were incubated with 1 µM SYTOX Green in SPB2. Once basal fluorescence reached a constant value, peptides were added (arrow, t = 0), and changes in fluorescence were monitored ({lambda}exc = 485 nm, {lambda}ems = 535 nm) and plotted as arbitrary units. Control spheroplasts, bold line. Peptide concentrations used were 1 µM temporin A ({blacksquare}), 0.4 µM temporin L ({circ}), and 1 µM temporin A + 0.4 µM temporin L ({blacktriangleup}). Data points represent the mean of triplicate samples with S.D. values not exceeding 3% from a single experiment, representative of three different experiments. Results similar to those obtained with temporin A were found for temporin B and therefore are not shown.

 
The fluorescence of the rhodamine peptides was measured before and after the addition of different concentrations of LPS to each rhodamine-temporin. Fig. 3, A–C, shows the results obtained with rhodamine-temporin A, rhodamine-temporin B, and rhodamine-temporin L, respectively. The data reveal a clear difference between temporin L and the other two isomers as follows. (i) Adding LPS to labeled temporins A or B rapidly decreased the fluorescence of the peptide, in a dose-dependent manner, suggesting that oligomerization of the peptides takes place immediately after contact with LPS. The strongest effect is achieved when the peptide:LPS ratio is 1:2 and 1:4. The addition of proteinase-K (indicated by the right arrow in Fig. 3, A and B) to LPS-treated rhodamine-temporins A and B, induced a rapid increase of fluorescence up to the initial level measured prior to the addition of LPS, thus confirming the induction of peptide assembly by LPS. (ii) The addition of LPS to rhodamine-temporin L increased its fluorescent signal in a dose-dependent manner. This increase in signal envisages that the labeled peptide disaggregates partially in the presence of LPS. Accordingly, proteolytic digestion of LPS-treated rhodamine-temporin L (indicated by the right arrow in Fig. 3C) resulted in an additional increase of fluorescence.

The Effect of LPS on the Antimicrobial Activity of Temporins—The difference regarding the effect of LPS on the organization of the peptides was also demonstrated by the loss of the bactericidal activity of temporin L, when preincubated with LPS, on S. aureus Cowan I, whereas temporins A and B hardly lost any activity (Fig. 4). We used S. aureus, because being a Gram-positive bacterium, it lacks the LPS outer membrane and therefore it is sensitive to all temporins. A plausible explanation for the loss of activity by temporin L and not A and B is that temporins A and B are assembled on the outer surface of LPS head groups, compared with temporin L, which, being less aggregated in LPS, inserts better into the hydrophobic chains of the LPS micelles (supported by the FTIR data and will be discussed later). This should reduce the concentration of temporin L available for the antimicrobial activity. Note that this explanation does not contradict with the high potency of temporin L against Gram-negative bacteria, because in that case the peptide would insert into the LPS hydrophobic core and partition between it and the target adjacent phospholipid inner membrane.

Temporin L Induces Disaggregation of Temporins A and B—To get insight into the mechanism of the synergic activity between the temporin isomers, the effect of temporin L on the oligomerization of temporins A and B in the presence of LPS was analyzed. We recorded the fluorescence of each rhodamine peptide, alone and in combination with each of the unlabeled temporins (Fig. 5), before and after the addition of LPS (arrow at time 0). Proteinase-K was then added after 30 min (right arrow). Fig. 5, A–C, shows the results found with rhodamine-temporins A, B, and L, respectively. The data reveal significantly less fluorescence quenching upon addition of LPS, when rhodamine-temporins A and B were mixed with unlabeled temporin L, compared with the results obtained with rhodamine peptides alone or in combination with unlabeled A or B (Fig. 5, A and B). This indicates that temporin L inhibits LPS-induced oligomerization of both temporins A and B. In comparison, when unlabeled temporins A and B were mixed with rhodamine-temporin L, the fluorescence of the peptide only slightly increased or decreased, respectively (Fig. 5C).

Combining Temporins Has No Effect on Their Structure in Buffer and in LPS Solution—To determine whether the mechanism of synergism involves changes in the structure of temporins, we used CD spectroscopy to determine the secondary structure of temporins, alone or in combination with another, in buffer and LPS. The interaction between peptides can be assessed by comparing the spectrum of the mixed peptides (experimental signal) with the average spectrum of each peptide alone (theoretical summation) (57). The two signals are identical if the peptides do not interact with each other. When the peptides were tested alone, temporin L exhibited a 32 and 47% {alpha}-helical structure (in PBS and LPS solution, respectively), whereas temporins A and B were predominantly unstructured in both media (Fig. 6A). The actual experimental signal for each mixture of peptides (temporin A + L or temporin A + B) is shown in Fig. 6B. The experimental and the theoretical signals were similar in both PBS and LPS, indicating that the synergistic effect is not due to the ability of temporin L to induce structural changes on temporins A and B.


Figure 3
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  		FIGURE 3.
The effect of LPS on rhodamine-labeled temporins. Once basal fluorescence of the labeled peptide reached a constant value, E. coli O111:B4 LPS was added at the corresponding concentration (first arrow, t = 0), and changes in fluorescence were monitored ({lambda}exc = 485 nm, {lambda}ems = 590 nm) and plotted as arbitrary units. Control ({blacksquare}) is given by the peptide alone, without LPS treatment. LPS concentrations used were 0.7 ({square}), 1.5 ({triangleup}), 3({circ}), 6({blacktriangleup}), and 12 µM (•). The second arrow indicates the addition of proteinase-K (80 µg/ml) to all samples. A, rhodamine-temporin A (3 µM); B, rhodamine-temporin B (1.5 µM); C, rhodamine-temporin L (1.5 µM). The values represent the mean of triplicate samples with S.D. values not exceeding 2% from a single experiment, representative of three different experiments.

 
The Structure of the Peptides within LPS and Their Effect on LPS Phosphate Groups Determined by ATR-FTIR—ATR-FTIR was used to determine the secondary structure of the peptides when bound to LPS. The secondary structure components of a peptide in a membrane-bound state can be determined from the wave numbers of the amide I vibration (followed by deconvolution) either before or after deuteration. Here we determined the secondary structure of the peptide after complete deuteration. The amide I region spectra as well as the fitted band components (obtained by deconvolution) of the peptides that are bound to LPS are shown in Fig. 7A. Assignment of the different secondary structures to the various amide I regions was calculated according to the values previously used (58–61). The data indicate a strong band at 1660 cm–1 (75%) for all the peptides, which is typical for dynamic helical structures. In addition, they all have a small fraction of aggregated beta-sheet, characterized by a band at 1625 cm–1 (25%).


Figure 4
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  		FIGURE 4.
The effect of LPS on the killing activity of temporins against S. aureus Cowan I. Temporins A (10 µM), B (15 µM), and L (4 µM) were preincubated with E. coli O111:B4 LPS, solubilized in water (peptide:LPS ratio equal to 1:1 and 1:2), for 30 min at 37 °C. These mixtures were then added to exponential phase bacteria (~106 cells/ml) in MH and incubated at 30 °C. After an incubation time of 20 min, aliquots were withdrawn, diluted in MH broth, and plated on an agar plate for colony forming units counting. The number of colony forming units is reported as the percentage relative to the control (bacteria without peptide, 9.9 x 105 ± 0.1 cells/ml). The values are the average of three independent experiments ± S.D. values (error bars).

 
The mode of interaction of the peptides with the phosphate groups of LPS correlates with their ability to traverse it (43, 62). The peptide-phosphate interaction was determined by following the asymmetric stretching vibration of the negatively charged phosphate {nu} (PO–2as). In the range of this vibration, namely ~1220 cm–1, a change in the band shape can be detected as a result of an interaction between the negatively charged phosphates and the cationic peptides (52). The data (Fig. 7B) revealed a stronger effect of temporin L compared with temporin A or B. This suggests peptide binding to both the outer and inner phosphates, because of its deeper insertion into the LPS. This is in agreement with the ability of temporin L to interact with lipid A, the highly conserved inner portion of LPS (63), and its high antimicrobial activity toward Gram-negative bacteria. Indeed, the deeper insertion of temporin L into the LPS layer should make it easier for this peptide to partition from the bacterial outer membrane into the adjacent cytoplasmic membrane and disrupt it.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
To date, very little is known about the biological significance of the presence of closely related AMPs in single living organisms. Mor and colleagues (40) investigated the role of structurally similar dermaseptins in frog skin and showed a synergistic effect between several of them, suggesting that a combination of various AMPs provides the animals with maximum coverage over a wide range of possible pathogenic microbes. However, the molecular basis of this synergism is not known. Temporins represent another important group of structurally related peptides. Initially isolated from R. temporaria skin secretion (30), they were subsequently discovered in other Rana species of American and Euroasian origin (19, 64), and to date they are among the largest family of natural AMPs with the shortest amphiphatic {alpha}-helix. Their killing mechanism involves the alteration of the permeability of the microbial membrane, although membrane permeation is not per se a lethal event (38).


Figure 5
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  		FIGURE 5.
The effect of unlabeled temporin A ({blacksquare}), temporin B ({blacktriangleup}), and temporin L (•) on the fluorescence of rhodamine-labeled temporins in the presence of E. coli O111:B4 LPS. Three µM rhodamine-temporin A (A), 1.5 µM rhodamine-temporin B (B), and 1.5 µM rhodamine-temporin L (C) were incubated with the same concentration of unlabeled temporins. Once the fluorescence reached a constant value, LPS was added (first arrow, t = 0) and changes in fluorescence were monitored ({lambda}exc = 485 nm, {lambda}ems = 590 nm) and plotted as arbitrary units. The peptide:LPS ratio was 1:4. The second arrow indicates the addition of proteinase-K (80 µg/ml) to all samples. Control (bold line) is given by the fluorescent peptide without the addition of unlabeled temporins. In panels A and B the results obtained with the unlabeled temporin B and A, respectively, are not reported, being similar to those shown when the unlabeled temporin A and B were used instead, in the respective cases. The values represent the mean of triplicate samples with S.D. values not exceeding 2% from a single experiment, representative of three different experiments.

 


Figure 6
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  		FIGURE 6.
The effect of peptide-peptide interaction on their structure in suspension and in LPS. A, CD spectra of temporin A ({diamondsuit}), B ({blacksquare}), or L ({blacktriangleup}) (100 µM) in PBS or LPS suspension. B, experimental CD spectra of temporin A + temporin B ({blacksquare}) or temporin A + temporin L ({blacktriangleup}) in PBS or LPS suspension compared with those of the empirically added structures (empty squares and triangles, respectively).

 
In this paper we report two important findings. First, we described the biophysical properties of temporins A and B that are most likely responsible for their inactivity against Gram-negative bacteria; second, we demonstrated, for the first time, a synergistic effect between temporins on Gram-negative bacteria, together with a plausible new mechanism for this effect.


Figure 7
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  		FIGURE 7.
The structure of temporins in LPS and their effect on LPS phosphate groups. A, FTIR spectra deconvolution of the fully deuterated amide I band (1600 to 1700 cm–1) of peptides in LPS. Second derivatives were calculated to identify the positions of the component bands in the spectra. The component peaks are the result of curve fitting using a Gaussian line shape. The sums of the fitted components are superimposed on the experimental amide I region spectra. Thick lines represent the experimental FTIR spectra; thin lines represent the fitted components. B, the influence of temporins on the head groups of LPS. IR absorbance spectra are shown in the range of the asymmetric stretching vibration of the negatively charged phosphates {nu}as(PO2) for the different temporins. Spectra are representatives of three independent experiments. The differences between the ratios of the deconvoluted areas within each experiment did not exceed 2%.

 
The cell wall of Gram-negative bacterial strains is composed of the heteropolysaccharide peptidoglycan, which is surrounded by an outer membrane consisting predominantly of the anionic LPS (65). Therefore, before reaching the cytoplasmic membrane, the peptides need to interact and traverse both the LPS and the peptidoglycan layers (66, 67). The outer cell envelope was previously considered to be a barrier against temporins, because E. coli strains with the progressively decreased LPS-polysaccharide chain were found to be concomitantly more sensitive to these peptides (30, 68). Here, we provide an explanation for the inactivity of temporins A and B toward Gram-negative bacteria. (i) The two peptides oligomerize when in contact with the outer membrane, as envisioned by the fast quenching of fluorescence intensity of rhodamine-temporins A and B, after the addition of purified E. coli LPS (Fig. 3, A and B). This is in contrast with the highly active antibacterial temporin L, which disassembles when in contact with LPS (Fig. 3C). This is in line with recent studies showing that oligomerization of AMPs gives rise to a dramatic reduction in their antimicrobial activity on Gram-negative bacteria, because they cannot diffuse efficiently through the cell wall because of their large size (43, 69). (ii) In contrast to temporin L, which adopts an {alpha}-helix structure when added to LPS micelles in solution, isomers A and B do not (Fig. 6A). Because the hydrophobic environment of membranes induces an {alpha}-helical structure to peptides that have this propensity, our CD spectroscopy data suggest that temporins A and B bind mainly to those portions of LPS facing the solution (phosphates or sugar residues) and not to those in proximity with the inner lipid moiety. This is also supported by the FTIR data showing that the overall effect on all the phosphate groups is less pronounced for temporins A and B, compared with that obtained with temporin L (Fig. 7B). Note, however, that temporins A and B have structures similar to that of temporin L when incorporated into LPS and measured by FTIR spectroscopy. This confirms that once temporins A and B are inserted within the LPS molecules, they can adopt the active structure (a dynamic {alpha}-helix).

The second interesting outcome of the present work is the synergistic effect of temporin L when combined with either temporin A or B, and the new molecular mechanism underlying this effect. This mode of action is different from that previously described for the pair magainin/PGLa, which was found to be associated with increasing perturbation of the cytoplasmic membrane (70, 71). In the case of temporins, the synergistic effect takes place at the level of the outer membrane of Gram-negative bacteria. This is demonstrated by the findings that, in contrast with intact bacteria, temporins do not display synergistic antibacterial activity on E. coli spheroplasts, which lack the cell wall (Fig. 2A). Furthermore, they also do not synergize to increase the permeability of the cytoplasmic membrane of spheroplasts to a low molecular mass fluorescent marker (Fig. 2B). Our data suggest that this synergism is related to the ability of temporin L to assist temporins A and B to traverse the LPS layer. After crossing it, they would have a capability, similar to that of temporin L, to reach and permeate the inner bacterial membrane.

As discussed above, isomers A and B oligomerize when bound to LPS. Previous studies have shown that LPS serves as a protective layer against peptides that bind to it as oligomers (43, 69). However, in the presence of temporin L, the aggregation of both temporins A and B is prevented, as manifested by only a slight quenching of fluorescence in the corresponding rhodamine-labeled peptides upon the addition of endotoxins, compared with the quenching when the peptides are alone (Fig. 5). Therefore, temporin L should promote the translocation of temporins A and B through the LPS leaflet, allowing them to easily get into the cytoplasmic membrane, which serves as their major target. To the best of our knowledge, this is the first example showing how peptides synergize to overcome bacterial resistance caused by the physical barrier of LPS.

Note that the strongest synergism was observed against A. hydrophila, considered to be part of the natural flora of a multiplicity of animals including humans (46, 72–74). A. hydrophila is an opportunistic pathogen found in healthy frogs, but capable of inducing diseases, such as the natural outbreaks of "red-leg" in Amphibia (45), including the species of R. temporaria (44). Beside causing high mortality in amphibian populations, this bacterium is also responsible for variety of infections in humans (75–77), especially in immunocompromised individuals, and it has been found to be insensitive to several classical antibiotics (72, 74, 78). Furthermore, Rollins-Smith and colleagues (45) reported that A. hydrophila is resistant to different peptides of frog skin origin such as magainin I, magainin II, PGLa, CPF, ranalexin, and dermaseptin, when tested in growth inhibition assays. Importantly, the synergism on different bacteria, between similar innate defense peptides that coexist within the same organism, should provide the animals with a quick and readily available weapon to combat infections by a wide range of invading microorganisms, including those species belonging to the natural flora, without harming the host.

In summary, we have demonstrated that a combination of natural small-sized and structurally related peptides can increase their spectrum of antimicrobial activity by inducing changes in the biophysical properties of the peptides (e.g. preventing their oligomerization) when bound to the LPS layer. Such findings should stimulate additional studies to better understand this new molecular mechanism of a synergism between homologue AMPs from the same species, and whether such a synergism exists in other families as well. Besides their scientific importance, studies along this line should contribute for the future development of a new peptide-based anti-infective therapeutic strategy, urgently needed because of the increasing resistance of microbes to the commonly used drugs.


   FOOTNOTES
 
* This work was supported by the Italian Ministero dell'Università e della Ricerca Grant 2005062410 and by grants from the Università di Roma La Sapienza and Istituto di Biologia e Patologia Molecolari of the National Research Council. 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

1 To whom correspondence should be addressed: Unità di Diagnostica Molecolare Avanzata, II Facoltà di Medicina e Chirurgia, Azienda Ospedaliera S. Andrea, Via di Grottarossa, 1035, 00189 Roma, Italy. Tel.: 39-06337-75457; Fax: 39-06337-75405; E-mail: marialuisa.mangoni{at}uniroma1.it.

2 The abbreviations used are: AMP, antimicrobial peptide; ATR-FTIR, attenuated total reflectance Fourier-transform infrared; Fmoc, fluorenylmethoxycarbonyl; FIC, fractional inhibitory concentration; LPS, lipopolysaccharide; MIC, minimal inhibitory concentration; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; PB, phosphate buffer; PBS, phosphate-buffered saline; MH, Mueller-Hinton. Back


   ACKNOWLEDGMENTS
 
We thank Dr. John Mayer from Eli Lilly Company for the generous gift of synthetic temporins.



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Antimicrob. Agents Chemother., January 1, 2008; 52(1): 85 - 91.
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