Studies on the interactions of neutral Galleria mellonella cecropin D with living bacterial cells
Cecropins constitute an important family of insect antimicrobial peptides involved in humoral innate immune response. In comparison with the highly basic cecropins A and B, cecropins D are less cationic and more hydrophobic. Interestingly, cecropins D were described only in lepidopteran insects, e.g., the greater wax moth Galleria mellonella. In the present study, interactions of neutral cecropin D (pI 6.47) purified from hemolymph of G. mellonella with living Escherichia coli cells were investigated. Fluorescence lifetime imaging microscopy using fluorescein isothiocyanate-labeled cecropin D revealed very fast binding of the peptide to E. coli cells. Fourier transform infrared spectroscopy analyses showed that G. mellonella cecropin D interacted especially with E. coli LPS and probably other lipid components of the bacterial cell envelope and exhibited an ordering effect with regard to lipid chains. This effect is consistent with the peptide binding mechanism based upon its incorporation into the lipid phase of the cell membrane. The interaction resulted in permeabilization of the bacterial cell membrane. Upon cecropin D binding, the cells lost characteristic surface topography, which was accompanied by altered nanomechanical properties, as revealed by atomic force microscopy. The interaction of the peptide with the bacterial cells also led to intracellular damage, i.e., loss of the cell envelope multilayer structure, formation of membrane vesicles, and enlargement of periplasmic space, which eventually caused death of the bacteria. In summary, it can be concluded that amphipathic character of α-helices, exposure of small positively charged patches on their polar surfaces and hydrophobic interactions are important physicochemical characteristics related to effective binding to E. coli cells and antibacterial activity of neutral G. mellonella cecropin D.
KeywordsGalleria mellonella Cecropin D Atomic force microscopy Transmission electron microscopy Fourier transform infrared spectroscopy Fluorescence lifetime imaging microscopy
Antimicrobial peptides (AMPs) are small, amphipathic, and mainly cationic molecules that upon interaction with microbial membranes lead to disruption of the membrane structure and function, which eventually results in microbial cell death. Several models have been proposed to explain the interactions of AMPs with microbial membranes, including the barrel-stave, carpet-like, aggregate, and toroidal pore model. Nevertheless, irrespective of the model, it is generally accepted that the initial step of the binding is mediated by electrostatic interactions between the cationic peptide and negatively charged membrane components followed by hydrophobic interactions between amphipathic domains in the peptide molecule and lipid components of the membrane (Nguyen et al. 2011; Scocchi et al. 2011; Cytryńska and Zdybicka-Barabas 2015).
Cecropins constitute a family of invertebrate AMPs, found especially in insects. They are amphipathic and mostly highly basic linear peptides composed of 31–39 amino acids, which can form two flexible α-helices bound with a hinge region in a hydrophobic environment. The distribution of hydrophilic and hydrophobic groups is conserved among all cecropins. They possess a hydrophilic N-terminal part containing many charged residues, a very hydrophobic stretch between residues 22–30, and a more hydrophilic C-terminal end. Usually, the N-terminal part of a cecropin molecule is strongly basic, whereas the C-terminal part is hydrophobic. The peptides were first characterized in Hyalophora cecropia (H. cecropia) (Lepidoptera) and classified into several subfamilies: A, B, C, D, E (Hultmark et al. 1982). Cecropins A, B, and C were then found in species belonging to other insect orders, i.e., Diptera and Coleoptera (Otvos 2000; Bulet and Stöcklin 2005; Yi et al. 2014). Interestingly, cecropins D were described only in lepidopteran insects, suggesting that this subfamily appeared relatively late in the evolution of insects (Gudmundsson et al. 1991).
A great arsenal of diverse AMPs was reported in the greater wax moth Galleria mellonella (G. mellonella), a model insect species widely used in studies on innate immunity mechanisms, host–pathogen interactions, and pathogenicity and virulence factors of different microorganisms (Cytryńska et al. 2007; Brown et al. 2008, 2009; Vogel et al. 2011). Among G. mellonella AMPs, two cecropins have been described to date, i.e., a highly basic cecropin A (pI 10.38) (Kim et al. 2004) and a neutral cecropin D (pI 6.47) (Mak et al. 2001; Cytryńska et al. 2007). Cecropin D, purified from G. mellonella immune hemolymph, was active against selected Gram-negative and Gram-positive bacteria as well as a filamentous fungus Aspergillus niger (Cytryńska et al. 2007). More recently, anti-Salmonella enterica and anti-Proteus mirabilis activity of synthetic G. mellonella cecropin D, named Gm1, has been demonstrated (Correa et al. 2014a). The interactions of Gm1 and its positively charged analog ∆Gm1 with model lipid membranes have also been analyzed (Correa et al. 2014b). In addition, Oñate-Garzón et al. (2017a; b) investigated the antibacterial activity and binding to model lipid membranes of two cationic peptides, ∆M1 and ∆M2, with a net charge +5 and +9, respectively, derived from G. mellonella cecropin D.
To provide more data on the interactions of the native G. mellonella cecropin D with living bacterial cells, the effects of the peptide action on Escherichia coli (E. coli) cells were investigated in the present study. The alterations in bacterial cell surface components that occur upon the peptide binding were investigated in detail by Fourier transform infrared (FTIR) spectroscopy. Furthermore, fluorescently labeled G. mellonella cecropin D was used for analysis of the peptide binding to bacterial cells by fluorescence lifetime imaging microscopy (FLIM). The cell surface topography and properties were analyzed by atomic force microscopy (AFM), whereas the intracellular damage was imaged by transmission electron microscopy (TEM).
Materials and methods
Bacterial strains and culture conditions
Gram-negative bacteria Escherichia coli D31 and E. coli JM83 and Gram-positive bacterium Micrococcus luteus (M. luteus) ATCC 10,240 were grown in 2.5% Lysogeny Broth (LB; Sigma-Aldrich) at 37 and 28 °C, respectively. In the experiments, bacteria in the logarithmic phase of growth were used.
Insect immune challenge, hemolymph collection, and preparation of hemolymph methanolic extracts
The larvae of G. mellonella (Lepidoptera: Pyralidae) were reared on a natural diet—honeybee nest debris at 30 °C in the dark. For the immune challenge, the last instar larvae (250–300 mg in weight) were pierced with a needle dipped into a pellet containing live E. coli D31 and M. luteus ATCC 10,240, and the hemolymph was collected 24 h after the immunization. Acidic/methanolic extracts rich in antimicrobial peptides and proteins below 30 kDa were prepared from the hemocyte-free hemolymph as described previously (Cytryńska et al. 2007; Mak et al. 2010).
Purification and fluorescent labeling of G. mellonella cecropin D
The cecropin D peptide was purified from the immune hemolymph extracts using a modified procedure described in our previous papers (Cytryńska et al. 2007; Palusińska-Szysz et al. 2012; Zdybicka-Barabas et al. 2012) and a P680 high-performance liquid chromatography (HPLC) system (Dionex). In brief, the lyophilized and lipid-free hemolymph extract was reconstituted in 0.1% (v/v) trifluoroacetic acid (TFA), filtered through a 0.45 μm filter, and subjected to HPLC using a Discovery Bio Wide Pore C18 250 mm × 4.6 mm column (Sigma-Aldrich) and two buffer sets: A—0.1% TFA (v/v), B—0.07% TFA, 80% acetonitrile (v/v). A linear 30–70% gradient of buffer B over 35 min, spectrophotometric detection at 220/280 nm, and a 1 ml/min flow rate were applied. The fraction of cecropin was collected and then purified to homogeneity using the same column as above and a linear 49–52% B gradient over 20 min. The purity of the collected final peptide was confirmed independently by analytical HPLC separation on the C18 column as above and by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) (Schägger and von Jagow 1987), while the identity was confirmed by determination of the N-terminal sequence on a Procise 491 automatic protein sequencer (Applied Biosystems). The concentration of the peptide was determined by amino acid analysis (Bocheńska et al. 2013).
Labeling of cecropin D with fluorescein isothiocyanate (FITC, isomer 1; Sigma-Aldrich) was performed according to a procedure described in our previous paper (Zdybicka-Barabas et al. 2014). Briefly, 4 nmol of the peptide was dissolved in 50 µl of sodium borate, pH 9.0, and mixed with 100-times molar excess of FITC dissolved in dimethylsulfoxide (DMSO). The solution was incubated for 20 h at room temperature in darkness and then the unreacted FITC was quenched by addition of 10 μl of 1 M glycine water solution. After additional 0.5-h incubation in darkness, the solution was subjected to gel filtration on a Superdex Peptide 10/300 GL column (GE Healthcare). The separation was carried out at a 0.7 ml/min flow rate in 10 mM ammonium acetate buffer, pH 5.8, containing 30% (v/v) acetonitrile. The peak containing the monosubstituted FITC-cecropin D peptide was collected and freeze-dried. Examination of the labeled peptide by SDS–PAGE revealed a single yellow band at approx. 5 kDa.
Bacterial membrane permeabilization assay
The E. coli JM83 strain bearing a pCH110 plasmid was used in the assay (Pharmacia-Amersham). The plasmid encodes constitutively synthesized cytoplasmic β-galactosidase and ampicillin resistance (Mak et al. 2007). The assay was performed using a suspension of mid-logarithmic phase cells prepared in 20 mM phosphate buffer, pH 6.8, essentially as described previously (Zdybicka-Barabas et al. 2012). Briefly, 2 μl of the suspension (5 × 105 colony forming units, CFU) was added to 23 μl of pre-incubated (37 °C, 15 min) cecropin D in the phosphate buffer (final concentrations of peptide: 0.625–20.0 μM). The samples were then incubated at 37 °C for 5–60 min. Subsequently, 20 mM HEPES/150 mM NaCl buffer, pH 7.5, and 50 mM p-nitrophenyl-β-d-galactopyranoside were added, the samples were incubated for 1.5 h at 37 °C, and absorbance was measured at 405 nm. Simultaneously, two types of the control samples were analyzed: (1) live bacteria incubated without the peptide and (2) bacteria completely killed by treatment with 5 µM cecropin B (Sigma-Aldrich). At the given time point, the perforation level of the live bacteria was subtracted from the results of the tested samples. Then, the perforation percent was calculated assuming the dead bacteria perforation level as 100%. Three independent experiments with three repetitions for each type of sample were performed.
Atomic force microscopy (AFM)
Preparation of the bacterial samples
100 μl of the suspension containing log-phase E. coli JM83 cells (OD600 = 0.2) cultured in the LB medium was incubated without (control) and in the presence of cecropin D (final concentrations 2.5 and 1 μM) for 0.5, 1, and 1.5 h at 37 °C. The bacterial suspensions were centrifuged at 8000×g for 10 min at 4 °C. The bacterial pellets were gently washed once with 20 mM phosphate buffer, pH 6.8, and next twice with non-pyrogenic water (100 μl). After final centrifugation, the bacteria were suspended in 5 μl of non-pyrogenic water and applied on the surface of freshly cleaved mica disks. Before imaging, the samples were allowed to dry at 28 °C (Zdybicka-Barabas et al. 2011, 2012).
AFM imaging and analysis
All AFM measurements in the tapping or Peak Force QNM operation modes were carried out using a NanoScope V AFM (Veeco) equipped with NanoScope 8.10 software and a piezoscanner with a maximum scan range of 150 × 150 μm (Analytical Laboratory, Faculty of Chemistry, UMCS, Lublin, Poland). A rectangular Si cantilever/tip (Veeco) with a spring constant of 20–80 N/m and resonance frequency of 300 kHz was used. The resolution of the scans obtained was 256 × 256 pixels. The height and peak force error images were obtained simultaneously. Four fields on each mica disk were imaged. The data were analyzed with WSxM 5.0 software (Nanotec). The roughness values were measured over the entire bacterial cell surface on 400 × 400 nm areas. The average surface root-mean-square (RMS) roughness of the cells was calculated from forty fields examined during two independent experiments.
For estimation of the G. mellonella cecropin D influence on the elasticity and adhesion properties of bacterial cells, Derjaguin–Muller–Toporov (DMT) modulus and adhesion forces were determined, respectively. The force measurements were performed in the Peak Force QNM operation mode using a silicon tip at the nitride lever, SCANASYST-AIR, with a spring constant of 0.4 N/m (Veeco). The data were analyzed with Nanoscope Analysis ver. 1.40 software (Veeco).
Transmission electron microscopy (TEM)
The bacteria (18 Eppendorf tubes with 100 µl suspension in LB medium; OD600 = 0.4) were incubated for 1 h at 37 °C without (control) or with cecropin D (final concentration 2.5 µM). Immediately after the incubation, the cells were directly fixed in the culture medium to avoid osmotic problems in the cells. The cells were flooded with the first fixative—2.5% (v/v) glutaraldehyde dissolved in 0.1 M phosphate-buffered saline (PBS, pH 7.4). After 30 min, the cells were gently centrifuged at 3000×g for 30 min at 4 °C. After rinsing several times with 0.1 M PBS, the cell pellets were post-fixed in a 1% (v/v) osmium tetroxide solution in 0.1 M phosphate buffer (pH 7.4) for 2 h at 4 °C. The bacterial cells were then dehydrated in a series of alcohol and acetone, and embedded in LR White resin. Ultrathin sections (65 nm) were cut with a diamond knife on a microtome RMC MT-XL (Tuscon, USA). The samples were collected on copper grids and contrasted with the use of uranyl acetate and Reynold’s liquid. The samples were observed under a LEO-Zeiss 912 AB electron microscope (Oberkohen, Germany).
Confocal laser scanning microscopy imaging (CLSM)
For CLSM imaging, the procedure was carried out essentially as described previously (Zdybicka-Barabas et al. 2014). Briefly, 60 μl of the suspension containing log-phase E. coli JM83 cells (OD600 = 0.2) was incubated with 10 µM FITC-labeled cecropin D at 37 °C for 15 min. Then, the bacterial suspension was centrifuged (2000×g, 10 min, 4 °C) and washed twice with 20 mM phosphate buffer, pH 7.4, containing 0.9% NaCl (100 μl). Next, the bacterial cells were suspended in 10 μl of 20 mM phosphate buffer, pH 7.4, and imaged using a laser scanning confocal microscope LSM 5 PASCAL (Carl Zeiss Microscopy) (excitation and emission wavelengths were 470 and 520 nm, respectively; excitation time was 600 ms).
Fluorescence lifetime imaging microscopy (FLIM)
For FLIM analysis, the log-phase E. coli JM83 cells (100 μl of suspension; OD600 = 0.2) in 20 mM phosphate buffer, pH 7.4, were incubated without (control) or with 2.5 μM FITC-labeled cecropin D at 37 °C for up to 60 min. The bacterial suspensions were centrifuged at 4000×g for 10 min at 4 °C. The cells were gently washed once with 20 mM phosphate buffer, pH 7.4 (500 μl), twice with non-pyrogenic water (300 μl), and finally they were suspended in 100 μl of non-pyrogenic water. Then, 10 μl of the resulting suspension was applied on a polylysine-coated cover slip and after 10 s carefully rinsed three times with water.
Fourier transform infrared (FTIR) spectroscopy
For FTIR analysis of cecropin D interactions with bacterial cells, 3 μl from 10 µl of the E. coli JM83 suspension (OD600 = 0.2) prepared in PBS, pH 7.4, supplemented with 5% D2O (v/v), immediately after cecropin D addition (final concentration 2.5 μM) was placed onto the attenuated total reflection (ATR) crystal element. For analyses of cecropin D-LPS interactions, cecropin D (10 µM) was incubated with E. coli K-235 lipopolysaccharide (LPS) (final concentration 1.5 mg/ml; Sigma-Aldrich, cat. no L-2143) in non-pyrogenic water containing 5% D2O (v/v) for 1 h at 37 °C (final volume 4 µl). The samples were deposited on the ATR crystal element by 5-min evaporation and the absorption spectra were recorded. The absorption spectra of the control cells, the cells exposed to cecropin D, cecropin D alone, as well as the LPS and PBS used were collected. The time period of 5 min was selected and standardized for all the experiments on the basis of preliminary optimization experiments where IR absorption spectra were recorded repeatedly till the moment at which no spectral changes were observed in the O–H stretching region. Such a procedure allowed to remove only the bulk water before recording of the spectra, whereas the water molecules inside cells and the water fraction tightly bound to the cells were preserved to maintain conditions as close as possible to the physiological ones.
Infrared absorption spectra were recorded using a Fourier transform infrared absorption spectrometer equipped with an attenuated total reflection set-up (ATR–FTIR). The internal reflection element (diamond prism) was used as an attenuated total reflection element. Continuous purging with argon during the measurements was applied. The spectra were recorded with a Nicolet iS50R spectrometer (Thermo Scientific). Typically, 10 scans were collected, Fourier transformed, and averaged for each measurement. Absorption spectra at a resolution of one data point every 2 cm−1 were obtained in the region between 4000 and 400 cm−1 using a clean crystal as the background. Taking into consideration that spectroscopic measurements in the phase transition region of the lipid may affect interpretation, all the spectroscopic experiments were performed at 21 °C, relatively far from the phase transition temperature of LPS (~ 35 °C) (Brandenburg and Seydel 1998; Hassan and Ilev 2014; Heinbockel et al. 2015). Spectral analysis was performed with Grams Al software (ThermoGalactic).
The protein concentration in the hemolymph and methanolic extracts was estimated with the Bradford method using bovine serum albumin (BSA) as a standard (Bradford 1976).
To compare two means, statistical analysis was performed using Student’s t test. The data were presented as the means ± standard deviation (± S.D.). The differences were considered statistically significant at *p < 0.05, **p ≤ 0.01, ***p ≤ 0.001.
Permeabilization of E. coli cell membrane
Effects of G. mellonella cecropin D on the cell surface and ultrastructure of E. coli
AFM imaging and analysis
Biophysical properties of E. coli cell surface after treatment with G. mellonella cecropin D
RMS roughness (nm)
4.98 (± 1.49)
6.92 (± 1.99)*
5.76 (± 2.07)
8.11 (± 2.17)**
DMT modulus (GPa)
3.31 (± 3.07)
11.28 (± 2.4)***
3.67 (± 2.49)
13.25 (± 3.94)***
Adhesion force (nN)
3.74 (± 1.13)
1.9 (± 1.2)***
1.13 (± 0.44)
1.43 (± 0.54)*
Effect of cecropin D on the ultrastructure of E. coli cells
Analysis of interactions of G. mellonella cecropin D with E. coli cells
FLIM analysis of cecropin D binding to bacterial cells
FTIR study of cecropin D interaction with LPS and live E. coli cells
Figure 7 presents the infrared absorption spectra of cecropin D, LPS, LPS modified by interaction with the peptide, and a difference spectrum. To calculate the difference spectrum [(LPS + cecropin) minus cecropin], the pure cecropin D spectrum was multiplied by the factor selected to minimize the intensity of the spectral band centered at 1204 cm−1 present exclusively in the cecropin D spectrum. Further, the LPS spectrum was subtracted from the resulting spectrum [(LPS + cecropin) minus cecropin]. To do so, the LPS spectrum was taken with the factor calculated to minimize the band centered at 1070 cm−1, particularly intensive in the spectrum of pure LPS. Such a procedure yields a spectrum that reflects selectively and exclusively spectral changes basically free of the contributions from the individual components. Some changes are observed in the difference spectrum as a result of the cecropin D and LPS interaction. A relatively broad band centered at 3382 cm−1 is observed and can be an indication of the water fraction bound via hydrogen bonds to the LPS. The spectral shift towards lower frequencies of the bands centered at 2850 and 2922 cm−1 assigned to the symmetric and antisymmetric C–H stretching in the CH2 groups of alkyl chains of lipids is observed as well. This indicates an ordering effect of cecropin D with respect to lipid chains directly involved in the interaction with the peptide. In the spectral band centered at 1649 cm−1, corresponding to both the α-helix structure of cecropin D and the stretching vibration of the carbonyl group of LPS, a considerable increase in the oscillator strength is noted. Such an effect can be interpreted in terms of the interaction between LPS and cecropin D via water bridges. It has been noted that deformation vibrations of water molecules may also contribute to the spectrum in this region.
E. coli cells exposed to cecropin D were also examined with application of the FTIR technique (Fig. 8). The broad band at 3291 cm−1 represents O–H stretching in water molecules in a bulk phase present in living cells. The results of our measurements show that cecropin D caused a decrease in the water fraction bound to the cells. In the spectral band centered at 1061 cm−1, representing C–O–P–O–C and C–O–C stretching modes of lipids, the very strong effect observed can be interpreted in terms of breakage of hydrogen bonds (a strong negative band at 1061 cm−1 accompanied by a positive band at 1203 cm−1). Such combined changes in a difference spectrum represent a hypsochromic spectral shift but this effect has to be interpreted with caution due to the fact that spectral contribution of the phosphate group of the PBS buffer may interfere with the spectra of endogenous phospholipids, despite precise calculation of the difference spectra recorded from the samples containing the same buffer. The principal band between 2800 and 3000 cm−1 represents the C–H stretching modes with the maxima of peaks at 2852 and 2922 cm−1, corresponding to the symmetric and antisymmetric stretching in the CH2 groups of alkyl chains. The spectral shift towards lower frequencies is attributed to the ordering effect with regard to the lipid fraction of the E. coli cells. It is possible that the certain ordering effect with respect to lipid acyl chains observed is directly related to incorporation of the peptide or its part into the lipid phase of the membrane.
Cecropins constitute a family of defense peptides characterized by two α-helices bound with a flexible hinge. In contrast to the highly basic cecropins A and B, cecropins D are less basic and more hydrophobic (Hultmark et al. 1982; Sidén and Boman 1983). So far, much attention has been paid to the antimicrobial action of cecropins A and B, most probably because of their highly cationic character, effective killing of pathogens, and much broader occurrence among insect species (Bulet and Stöcklin 2005; Yi et al. 2014).
In the present study, we analyzed the anti-E. coli activity of native G. mellonella cecropin D and interactions of this peptide with live E. coli cells as well as LPS. As revealed by the cell membrane permeabilization assay, cecropin D used at the 2.5 µM concentration effectively permeabilized E. coli JM83 cell membranes. It is worth mentioning that the concentration of the peptide used during the experiments corresponded to the cecropin D concentration determined in the hemolymph of naive G. mellonella larvae (Mak et al. 2010). Thus, our results indicate that, to some extent, the peptide can effectively protect the larvae against Gram-negative bacteria invading the hemocoel. In another study, the concentration below 2.5 µM of synthetic G. mellonella cecropin D completely inhibited the growth of the E. coli Re mutant (Correa et al. 2014a), whereas the growth of E. coli ATCC 25,922 was inhibited only at the 40 µM concentration of this peptide (Oñate-Garzón et al. 2017b). On the other hand, it was reported that 8.6 µM native G. mellonella cecropin D killed E. coli D31 cells (Cytryńska et al. 2007). The differences in the susceptibility of the E. coli strains to G. mellonella cecropin D may result from the different assays and growth conditions, e.g., different initial numbers of bacterial CFU (1 × 105, 5 × 105, 2 × 107 CFU/ml) used for determination of antimicrobial activity. However, there is strong literature evidence that the most important factor contributing to such differences is the properties of the bacterial cell envelope. In accordance with this, E. coli SB1004, D21e7, and D22 mutants with increased hydrophobicity of the surface layer were more susceptible to H. cecropia cecropin D than wild-type strains (Sidén and Boman 1983). These authors suggested that hydrophobicity alterations in the bacterial surface facilitated the better penetration of cecropin D through the outer membrane, which is limited in comparison with cecropins A and B due to the dissimilar properties of cecropin D.
There are studies indicating that the LPS layer plays a crucial role in the attraction and control of AMPs binding to the surface of Gram-negative bacteria (Ding et al. 2003; Papo and Shai 2005; Agrawal and Weisshaar 2018). Ebbensgaard et al. (2015) demonstrated protective effect of LPS against antimicrobial activity of many AMPs (including cecropin B and cecropin P1) manifested by a higher susceptibility of the E. coli LPS defective mutants in comparison with their parental strains: O6 serotype E. coli ATCC 25,922 and rough E. coli BW25113K-12. Moreover, comparison of the two parental E. coli strains revealed less susceptibility of smooth E. coli ATCC 25,922 to selected AMPs than E. coli K-12, further supporting important role of LPS structure in interactions with antimicrobial peptides (Ebbensgaard et al. 2015). Furthermore, Agrawal and Weisshaar (2018) using single-cell fluorescence microscopy showed that deletion of sugar layers or/and phosphate groups from the core oligosaccharide in E. coli K-12 LPS accelerated permeabilization of the outer membrane and cytoplasmic membrane by cecropin A. They postulated that the core oligosaccharide part of LPS functions as a kinetic barrier slowing cecropin A passage to the lipid A leaflet (Agrawal and Weisshaar 2018).
As revealed by the FTIR analyses, G. mellonella cecropin D interacted with E. coli LPS and exhibited an ordering effect with regard to lipid chains. No changes in the carbohydrate region of the spectrum were observed, suggesting that the sugar chain had no influence on the peptide binding. A similar effect was detected when the interactions of the peptide with live E. coli cells were analyzed, indicating that G. mellonella cecropin D interacted especially with the lipid part of LPS and probably with other lipid components of the bacterial cell envelope. It is known that the energy (spectral position) of the νsCH2 band reflects the conformational order and mobility of the lipid acyl chains. Owing to this dependency, sharp changes in a position of this particular band on the energy (wavenumber) scale, accompanying phase transitions, can be applied to monitor the gel to liquid phase transitions induced by temperature (Lewis and McElhaney 2002, 2013). On the other hand, any other factors affecting the dynamic properties of a lipid phase, such as interaction with sterols, proteins, flavonoids, etc. (Pawlikowska-Pawlęga et al. 2013), influence remarkably energy of the νsCH2 vibrations and therefore vibrational spectroscopy analysis appears a sensitive tool in monitoring fine effects of different modifiers on the dynamic and structural properties of the hydrophobic core of the lipid bilayers. A consequence of the ordering effect observed can be disturbance in membrane fluidity and, finally, loss of its integrity. Given that ordering of lipid acyl chains leads to stiffening of the membrane structure (Frolov et al. 2011; Fowler et al. 2016), these changes also contributed to the demonstrated increase in DMT modulus of E. coli cell surface upon cecropin D treatment. Using synthetic G. mellonella cecropin D named Gm1 and the rough type LPS of P. mirabilis and S. enterica, Correa et al. (2014b) demonstrated that the acyl chains of LPS became disordered upon Gm1 binding, leading to fluidization of the structure. The ordering effect observed in our study versus the disordering action described by Correa et al. (2014b) can be explained by the differences in the structure of the LPS used. These authors reported that the lipid A moiety was the most important part of the LPS for interaction with the peptide, whereas the length of the LPS sugar chain had no effect on the cecropin binding. Their results also suggested that cecropin D molecules were located in the lipid head group regions and then underwent intercalation into the P. mirabilis LPS aggregates (Correa et al. 2014b). The lipid A moiety of LPS was also important for synthetic H. cecropia cecropin A binding that occurred due to hydrophobic interactions between C-terminal helices of the peptide and the lipid A (De Lucca et al. 1995). Dissociation of LPS aggregates upon H. cecropia cecropin A binding was also reported (Lee et al. 2015). Interestingly, using circular dichroism Avitabile et al. (2014) demonstrated that folding of α-helical peptides, cecropin A and magainin 2, was induced by interaction with LPS present in the outer membrane of E. coli cells.
Correa et al. (2014a) studied the morphological changes in the S. enterica and P. mirabilis cell surface caused by synthetic G. mellonella cecropin D used at a 10-µM concentration. These alterations were described as indentations, leakage of cytoplasm, clustering of the cells, and fusion of the membranes. As demonstrated by AFM analysis in the present work, binding of G. mellonella cecropin D (applied at a four-times lower concentration than that used in the above study) to E. coli cells resulted in loss of characteristic cell surface topography and showed significant changes in nanomechanical properties. The alterations in the E. coli cell envelope were undoubtedly confirmed by TEM imaging, which revealed loss of the multilayer structure of the cell envelope, formation of membrane vesicles, and enlargement of the periplasmic space. Nevertheless, the E. coli cells exposed to G. mellonella cecropin D seemed to retain their shape to some extent, as no disintegrated cells were observed. In contrast, the cell membrane of E. coli treated with Heliothis virescens cecropin B exhibited formation of pores that eventually led to disruption of membrane continuity and cell disintegration (Lockey and Ourth 1996). A comparison of TEM images obtained in this study and those presented by Lockey and Ourth (1996) suggests that the mode of G. mellonella cecropin D action against E. coli cells may be based on a mechanism other than pore formation. On the other hand, Vaara and Vaara (1994) demonstrated that the mode of the antibacterial action of synthetic H. cecropia cecropin B resembled that of cationic amphiphilic detergents, i.e., benzalkonium chloride and cetylpyridinium chloride, known to disorganize the outer membrane resulting in lysis of Gram-negative bacteria cells. In this context, considering the TEM images of E. coli cells that exhibited symptoms of partial membrane dissolution, the action of G. mellonella cecropin D might be explained by detergent-like activity; however, this hypothesis needs further studies. Interestingly, it was postulated that cecropins A and B used at sub-inhibitory concentrations might induce a stress response in viable E. coli cells by activation of the hyperosmotic stress gene osmY (Oh et al. 1998).
In summary, the antibacterial activity of native G. mellonella cecropin D against E. coli is determined by initial interaction with LPS in the outer membrane and other lipid components of the cell envelope. This interaction affects the membrane fluidity and integrity by an ordering effect in regard to lipid chains. The effective binding demonstrated by FLIM leads to damage to the cell envelope reflected by disruption of the multilayered structure, changes in the nanomechanical properties, and loss of characteristic surface topography. Disturbance in the membrane integrity results in permeabilization of the cell membrane and leakage of the intracellular content, as confirmed by the cytoplasmic β-galactosidase activity assay. Although they retained their shape to some extent, E. coli cells experiencing these changes eventually die. Considering the neutral character of G. mellonella cecropin D, our results provide further evidence that the overall cationic character is not a necessary factor for antibacterial activity of AMPs and for effective binding to bacterial cells.
The work was financially supported by grant No 2012/05/B/NZ1/00,033 from the National Science Centre (Poland). The research was carried out with the equipment purchased thanks to the financial support of the European Regional Development Fund in the framework of the Polish Innovation Economy Operational Program (contract no. POIG.02.01.00-06-024/09 Centre of Functional Nanomaterials) as well as European Union structural funds (grants POIG.02.01.00-12-064/08 and POIG.02.01.00-12-167/08). The Faculty of Biochemistry, Biophysics and Biotechnology of Jagiellonian University in Krakow is a partner of the Leading National Research Centre (KNOW) supported by the Ministry of Science and Higher Education (Warszawa, Poland).
Compliance with ethical standards
Conflict of interest
The authors declare that they have no conflict of interest.
All applicable international, national, and/or institutional guidelines for the care and use of animals were followed.
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