INTERACTION OF ANTIBIOTICS WITH LIPID VESICLES ON THIN FILM POROUS SILICON USING REFLECTANCE INTERFEROMETRIC FOURIER TRANSFORM SPECTROSCOPY

SUMMARY Reflectance interferometric Fourier transform spectroscopy (RIFTS) method was employed to monitor lipid vesicles rupture and formation of planar lipid bilayers induced by various antibiotics on an oxidized porous silicon (pSi) surface. We intended to demonstrate that RIFTS on pSi is an analytical platform suitable for investigation of the mechanism of antibiotic action on cell membranes. 1 1. INTRODUCTION The ability to observe interactions of drugs with cell membranes is an important area in pharmaceutical research. However, these processes are often difficult to understand due to the dynamic nature of cell membranes. Therefore, artificial systems composed of lipids have been used to study membrane properties and their interaction with drugs. 2 They are usually designed in the form of vesicles (large or giant unilamellar vesicles, LUV and GUV, respectively), or in the form of supported lipid bilayers (SLB) depending on the analytical techniques that are used to probe their structure, composition, and phase properties. Classical techniques to investigate the way drugs interact with membranes include fluorescence techniques usually combined with NMR spectroscopy, dynamic light scattering (DLS), X-ray or neutron reflection and diffraction methods. UV−vis spectroscopy, Fourier transform infrared (FTIR), and calorimetry methods are also commonly used to characterize changes in the lipids organization when exposed to membrane-active drugs. Other techniques are more suitable to probe SLB films, such as ellipsometry, X-ray photoelectron spectroscopy (XPS), surface plasmon resonance (SPR), quartz crystal microbalance with dissipation monitoring (QCM-D) and atomic force microscopy (AFM).The main advantage of label-free sensor methods such as SPR or QCM-D is the possibility of investigating the kinetics of interactions and the possibility for multiplexing. However, all of the above methods require very expensive instrumentation and are therefore not necessarily suitable for laboratories, which do not have access to these tools. Here, lipid vesicle adsorption, rupture, and formation of planar lipid bilayers induced by various antibiotics (surfactin, azithromycin, gramicidin, melittin and ciprofloxacin) and the detergent dodecyl-b-D-thiomaltoside (DOTM) were studied using reflective interferometric Fourier transform spectroscopy on an oxidized porous silicon surface as a transducer. 2. EXPERIMENTAL RESULTS AND DISCUSSIONS The pSi films were prepared as thin films of 3 μm thickness with pore dimensions of a few nanometers in diameter by electrochemical etching of p++ type crystalline silicon at 30 mA/cm 2 for 325 s. The pSi film were then thermally oxidized at 600 °C to form a negatively charged stable hydrophilic surface that has been noted to be favorable for phospholipid vesicle deposition and bilayer formation. 3,4 Suspensions of LUVs with diameters of approximately 100 nm were generated using a mixture of DPPC doped with 1% DOPE-Rhod, and deposited on the oxidized pSi surface. Due to the phase transition temperature (Tm = 41 °C) of DPPC lipids, bilayer formation was not favorable at room temperature via lipid vesicle rupture and fusion. AFM imaging was conducted on vesicles adsorbed to an oxidized pSi surface and exposed to the antibiotics and to DOTM for 16 h. Representative tapping mode AFM images taken at room temperature with adsorbed vesicles are shown in Figure 1A. The AFM data confirmed that DOTM caused the complete solubilization of the vesicles (Figure 1C). In contrast to DOTM, introduction of surfactin did not result in complete solubilization of the lipids, but resulted in the formation of a bilayer with holes at the pSi surface (Figure 1B). A section analysis confirmed that the bilayer thickness was 5.5 nm, consistent with a DPPC bilayer. Surfactin here was observed to strongly destabilize the vesicles. Figure 1. AFM images of (A) DPPC vesicles on pSi surface, and after addition of (B) surfactin (0.05 mM) and (C) DOTM (2.15 mM) after 16 h. The image was performed in tapping mode in Tris:NaCl buffer. The RIFTS method was then employed to monitor the destabilization of the DPPC + DOPE-Rhod 1% vesicles at the pSi surface, induced by the antibiotics surfactin, azithromycin, gramicidin, melittin, and ciprofloxacin, and by DOTM. The RIFTS method makes it possible to monitor real time changes in the conformation of lipid vesicles adsorbed to the pSi surface by monitoring index contrast changes at the surface of pSi. We have monitored the amplitude and the position (2nL) of the FFT of the reflectivity of the DPPC +DOPE-Rhod 1% lipid vesicles/pSi system (figure 2). When the lipid vesicles are solubilized and disappear from the pSi surface, the pSi/vesicle and the vesicle/PBS interfaces disappear in favor of the pSi/PBS interface. The refractive index contrast at the pSi surface increases consequently causing the increase of the amplitude of the FFT of the reflectivity the signal. Figure 2. DPPC + 1% DOPE-Rhod vesicles and exposed to DOTM (2.15 mM). (A) Value of 2 nL as a function of time. (B) Percent change in the amplitude of the FFT peak as a function of time. (C) Possible situations during vesicle dissolution. 3. CONCLUSIONS The RIFTS method was used to investigate in real time the effect induced by various antibiotics (surfactin, azithromycin, gramicidin, melittin, and ciprofloxacin) and DOTM on lipid vesicles adsorbed on an oxidized porous silicon (pSi) surface. The RIFTS method applied in this work appeared to be robust and is cost-effective in comparison with techniques such as AFM and DLS. Moreover, it correlated very well with the results obtained from DLS and from AFM, suggesting that it can constitute an interesting preliminary technique to monitor time and concentration dependent changes in membrane model systems, induced by antibiotics without the need for advanced technology. REFERENCES T. Guinan,  C. Geodfrey, N. Lautredou, S. Pace, P. E. Millhiet, N. H. Voelcker and F. Cunin, Langmuir, 2013, 29, 10279-10286. Seddon, A. M.; Casey, D.; Law, R. V.; Gee, A.; Templer, R. H.; Ces, O. Chem. Soc. Rev. 2009, 38 (9), 2509−2519. Cunin, F.; Milhiet, P.-E.; Anglin, E.; Sailor, M. 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