1. Introduction
Lipid peroxidation is a complex process associated with the oxidative deterioration of lipids and the production of various breakdown products [
1,
2]. Lipid hydroperoxides and conjugated dienes or trienes are considered primary oxidation products, which, due to their instability, break down and form secondary oxidation products, among which are aldehydes, ketones, hydrocarbons, alcohols, and others [
3]. Lipid peroxidation occurrence in the human body is a consequence of oxidative stress, which has been correlated with various diseases and health conditions, including neurodegenerative diseases, heart and cardiovascular system conditions, inflammatory immune injuries, and others [
4]. The presence of polyunsaturated fatty acids (PUFAs) in a phospholipid bilayer makes it highly susceptible to oxidative damage, resulting in changes in the membrane properties. For example, a decrease in the fluidity leads to the loss of its functionality as a barrier. Furthermore, it has been suggested that some peroxidation products, in particular malondialdehyde (MDA), 4-hydroxy-2,3-nonenal (HNE), and other 4-hydroxy-2,3-alkenals (HAKs) of different chain lengths can affect several cell functions, including signal transduction, gene expression, cell proliferation, and the response of the target cell [
5]. For example, Cajone and Bernelli-Zazzera [
6] showed that HNE caused an increase in the expression of the hsp 70 gene in human hepatoma cells. Moreover, it was shown that HNE can cause the activation of heat shock factor in vitro [
7]. To maintain homeostasis, it is crucial to achieve balance between a steady formation of pro-oxidants and a similar rate of their consumption by antioxidants. If the continuous regeneration of antioxidants is not sufficient, oxidative damage occurs, resulting in pathophysiological events [
8,
9].
Flavonoids are natural antioxidants with the ability to act as reducing agents, hydrogen donors, and singlet oxygen quenchers [
10]. Additionally, they possess a metal-chelating potential [
11,
12]. They are polyphenolic compounds, consisting of fifteen carbon atoms arranged in a C6-C3-C6 configuration. Two aromatic rings are connected through a 3-carbon bridge, usually in the form of a heterocyclic ring [
13]. The structural characteristics that are presumably most important for their antioxidant activity are a hydroxylated B-ring (
Figure 1) and the presence of the double bonds C2=C3 and C=O in the C-ring.
Depending on their substituents, flavonoids can further be classified into several subcategories: flavonols, flavones, catechins, flavanones, anthocyanidins, and isoflavonols [
14]. Various antioxidants, including polyphenols, are commonly found in foods and beverages of plant origin, including fruits, vegetables, cocoa, tea, and wine. Therefore, they play an essential role in promoting preventive health care through diet [
13,
15]. The antioxidant activity of a polyphenolic compound is determined by the ability of the phenolic hydrogen to scavenge free radicals [
16]. Although almost all subcategories of flavonoids exhibit antioxidative activity, it has been reported that flavones and catechins demonstrate the most powerful protection against reactive oxygen species [
15]. In relation to their free radical scavenging and metal ions-chelating activities, a multitude of health-promoting effects have been observed, including anti-inflammatory, anti-mutagenic, and anti-carcinogenic. Furthermore, they are studied as key cellular enzyme functions modulators [
15].
Flavonol protection against oxidative stress can be achieved through different mechanisms, with the direct scavenging of free radicals being one of them. Highly reactive hydroxy groups can react with the radical and stabilize the reactive oxygen species, which is followed by the inactivation of the radical [
15]. Apart from direct scavenging, flavonoids can inhibit lipid peroxidation by the chelation of metal ions which usually catalyze the reaction. The impact of transition metal ions on lipid peroxidation has been extensively studied [
17,
18,
19,
20,
21]. Fenton and Fenton-like reactions are often used to explain the production of hydroxyl radicals, since it was demonstrated that even trace levels of cellular transition metal ions can catalyze a Fenton reaction in vivo at the physiological level of hydrogen peroxide [
18,
22].
The interaction of polyphenolic compounds and lipid membranes could be one of the mechanisms in the protection against peroxidation. Depending on the lipid and flavonol chemical structure and composition, the interaction between flavonols and biological or model membranes can result in the binding of flavonol at the lipid–water interface or the distribution of flavonol in the hydrophobic part of the bilayer [
23]. For example, Van Dijk et al. [
24] investigated the relationship between the relative hydrophobicity of flavonoids and their binding with the liposomes using fluorescence quenching. The affinity of flavonoids from the subcategory of flavonols for liposomes was determined to be much higher than the one of flavanones, which was explained by different structural characteristics. It was determined that the planar configuration of flavonols, in contrast to the tilted configuration of flavanones, favors intercalation into vesicle membranes. If the flavonols partition in the non-polar region of the bilayer, they can influence the membrane fluidity. If the membrane is rigid, the probability of lipid radical interactions with other fatty acids is increased, due to the limited motion of fatty acid chains [
25]. The more hydrophilic flavonols can form hydrogen bonds with the polar head groups at the lipid–water interface of the membrane and provide a level of protection for the bilayer. Furthermore, different subgroups of flavonols can have different effects on the phase transition temperature of various lipids, making membranes more or less ordered [
23].
Within this study, we used three different flavonols: quercetin (QUE), myricetin (MCE), and myricitrin (MCI) (
Figure 1), which are all found in various plants. For example, QUE can be found in onions, apples, and berries. MCE is also found in various berries and vegetables, as well as in teas and wines produced from various plants [
26]. MCI has been extracted from numerous plants, such as
Manilkara zapota and
Eugenia uniflora [
27]. QUE is one of the most abundant and most studied flavonols, owing to its antioxidant and anti-inflammatory properties [
28,
29]. Furthermore, it has been shown that QUE exhibits a hepatoprotective, antifibrotic [
30], anti-coagulative [
31], and antimicrobial [
32] activity. MCE has, among others, been suggested as a good candidate for the development of new drugs for the treatment of Alzheimer’s disease due to its strong free radical-scavenging activity, which can block Aβ-induced neuronal death [
28]. Its rhamnose glycoside, MCI, shows numerous beneficial effects as well, including anti-allergic [
33], antioxidant, anti-inflammatory, antifibrotic [
34], and antinociceptive activity [
35]. The three flavonols differ in substituents and, consequently, hydrophobicity.
To investigate the protective effect of three structurally different flavonols on the molecular structure, integrity, and elasticity of lipid membranes under oxidative attack, we used a combination of techniques. The membrane integrity was measured by its crucial structural parameters, such as elasticity, surface roughness, thickness, and fluidity. We hypothesized that the insertion of antioxidative flavonols would suppress the disintegration of the lipid membranes during the lipid peroxidation. Three different molar fractions (
x = 0.01, 0.05, 0.1) of flavonols were chosen. Additionally, we wanted to highlight the dependence of the preservation of the membrane on the flavonol hydrophobicity, i.e., on their localization inside the bilayer. The most hydrophobic flavonol was expected to be hosted deep within the membrane and vice versa. As the model membrane, 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) was chosen, which possesses two chains of monounsaturated fatty acids. Due to the absence of adjacent double bonds with methylene bridges between them, which occur in PUFAs, the initiation phase of the lipid peroxidation is slower, while the overall mechanism is essentially the same in mono- and polyunsaturated fatty acids [
36]. The main difference is in the occurrence of different peroxidation products, which are formed as a consequence of conjugation. Mass Spectrometry (MS) was used to identify and compare the lipid peroxidation products before and after the incorporation of flavonols in different molar fractions, while Fourier Transform Infrared Spectroscopy, Attenuated Total Reflectance technique, (FTIR-ATR) was used to determine the extent of the lipid peroxidation reaction and quantify the inhibition in the samples with flavonols. Atomic Force Microscopy (AFM) and Force Spectroscopy (FS) were used to analyze the nanomechanical and topological properties of the lipid bilayers, such as roughness (determined from the AFM), thickness, and elasticity defined via Young’s modulus (determined from force–distance curves (FS)). Dynamic Light Scattering and electrophoretic measurements (DLS/ELS), as well as Electron Paramagnetic Resonance (EPR) and Small Angle X-Ray Scattering (SAXS) measurements provided further information on the flavonol antioxidative role in the preservation of membrane integrity. Our results demonstrate the unique capability of this multi-technique approach and indicate its potential to deeply enhance the understanding of cellular oxidative injury.
2. Materials and Methods
2.1. Chemicals
Iron(II) chloride tetrahydrate (98%) was purchased from Alfa Aesar, (Ottawa, ON, Canada). Myricetin (>97%) and myricitrin (>98%) were purchased from TCI Chemicals Pvt. Ltd. (Chennai, India). Quercetin (≥95%) and phosphate-buffered saline (PBS) buffer (PBS tablets, pH 7.4, Ic = 150 mM) were purchased from Sigma-Aldrich (St. Louis, MO, USA). 1-2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) was supplied by Avanti Lipids (Industrial Park Drive Alabaster, AL, USA). Coumarin hydrazine (CHH) was synthesized at the UL-Faculty of Pharmacy. Methanol (99.5%) and hydrogen peroxide (30% p.a.) were purchased from Kemika, Zagreb. Chloroform (99.93%) p.a., was purchased from Lach-ner Ltd. (Neratovice, Czech). The EPR probe 5-DOXYL-stearic acid and 2,2-diphenyl-1-picrylhydrazyl (DPPH) were purchased from Sigma Aldrich (St. Louis, MO, USA). All the chemicals were used without further purification.
2.2. Preparation and Oxidation of Liposomes with and without Inserted Flavonols
The DOPC liposomes were prepared by dissolving DOPC in chloroform. The QUE, MCE, and MCI were inserted in three different molar fractions (0.01, 0.05, 0.1) by mixing methanol solution of each flavonol with chloroform solution of DOPC. Solvents were evaporated using a rotary evaporator and the remaining films were dried in a vacuum. The dried films were dispersed in PBS by manual shaking at room temperature. During rehydration, the lipid film was gradually scraped off the wall of the glass bottle by alternately immersing the flask in the ice and hot water. The liposome suspension was left to stabilize overnight. Multilamellar liposomes were used to avoid the loss of lipids and flavonols during the process of extrusion, except for the DLS measurements. The concentration of lipids was adjusted for different methods and will be stated in the corresponding sections. The lipid peroxidation was initiated by the addition of FeCl2 × 4H2O and H2O2. The final concentrations of FeCl2 × 4H2O and H2O2 in the suspension were 1 mM, and the reaction was advanced for 1 h before measurements.
Hydrogen peroxide emerged as a major redox metabolite operative in redox sensing, signaling, and regulation. The concentration of extracellular H
2O
2 in redox signaling under physiological conditions was between 0.1 μM and 10 μM in oxidative eustress conditions. Higher extracellular concentrations (5–500 μM) led to adaptive stress responses. Supraphysiological extracellular concentrations in oxidative distress (5 μM <
c < 1 mM) led to the irreparable damage of biomolecules [
37]. Since we wanted to initiate damage of the lipid molecules, we used the 1 mM of H
2O
2 as it was in the extracellular solution under oxidative distress conditions.
In our experiments, the lipid peroxidation process was initiated by the addition of hydrogen peroxide and iron(II) ions (Fenton reaction) to the liposome suspension, where hydroxyl radicals were formed mainly by one-electron redox reactions between the H
2O
2 and the pre-existing hydroperoxides with transition metal ions. The reactions of ferrous (Fe
2+) ions with hydrogen peroxide and oxygen can generate ferryl and perferryl species, which are strong oxidants and have also been suggested to be capable of initiating radical reactions. In relation to the present approach, it must be mentioned that there are other physiological conditions that induce lipid peroxidation, for example systems including ascorbate and Fe
2+ ions. Even vitamin C, which is a known antioxidant, can act as a prooxidant. Under favourable conditions, it contributes to the oxidative damage of lipids by reducing Fe
3+ to Fe
2+ ions (or Cu
2+ to Cu
+) [
38]. The ascorbate/Fe
2+ system was used in a study of peroxidation and was found to cause a significant degradation of ethanolamine phosphoglycerides [
39]. Moreover, it has been suggested that the prooxidant effect of vitamin C could not have relevance in vivo [
40]. Therefore, by performing experiments using a H
2O
2/Fe
2+ system to initiate lipid peroxidation, two opposite (antioxidant and prooxidant) effects were avoided.
2.3. EPR Spectroscopy
The EPR spectra were collected by a home-modified Varian E-109 spectrometer (Ruđer Bošković Institute, Zagreb, Croatia) using a Bruker ER 041 XG microwave bridge working at a microwave frequency of 9.3 GHz (i.e., X-band) at room temperature (25 °C). The temperature in the EPR cavity was controlled by a Bruker ER 4111 temperature controller using a nitrogen gas flow accurate to 1 °C.
2.3.1. Antioxidant Activity
For the determination of the antioxidant activity, the spectrometry settings were: magnetic field modulation frequency 100 kHz, central field 331.0 mT, sweep range 10 mT, sweep time 20 s, microwave power 10 mW, and modulation amplitude 0.1 mT. A 2,2-diphenyl-1-picrylhydrazyl (DPPH) stable free radical was used to monitor the scavenging capability of flavonols using EPR spectroscopy. The stability of the freshly prepared ethanol solution was checked, and no significant loss of signal in the EPR signal was recorded within 24 h. A 4050 μL volume of a DPPH stock ethanol solution (0.5 mM) was added to 450 μL of a flavonol solution and mixed. The final solution was promptly inserted into the EPR capillary, which was then placed in a standard quartz tube. The EPR spectra were collected as a function of time initiated by contact with the sample and radical solution. The scavenging effect of the flavonol samples on DPPH radicals was obtained from the EPR signal intensities calculated by the double integration of the EPR spectra and expressed in arbitrary units. The signal intensity of the pure 0.5 mM DPPH solution in PBS, recorded just before starting the sample evaluation, was set as the reference signal intensity (I0) for the reaction time t = 0 min. The EPR signal intensity of DPPH radicals was decreased upon the flavonol addition and monitored for 20 min in recording intervals of 0.5 min and 1.0 min, depending on the sample activity. The remaining signal intensity, i.e., the remaining DPPH radicals after the reaction time, t, normalized and expressed as a percentage, IN, was calculated as: IN = (I/I0) × 100, where I is the signal intensity of the DPPH radicals in the flavonol solution measured at time, t. Each sample was analyzed in triplicate. The results are presented as mean values.
2.3.2. Fluidity Change during Lipid Peroxidation
The experimental parameters for monitoring the fluidity change upon the initiated lipid peroxidation were: central magnetic field 331.0 mT, sweep width 10 mT, modulation amplitude 0.1 mT, and microwave power 4.9 mW. A standard Bruker ER 4111 VT temperature controller with a nitrogen gas flow was used to control the temperature within 1 °C. A manganese standard reference, Mn2+ in MnO, was used to calibrate the magnetic field of the EPR spectrometer. The EPR spectra were simulated with a custom-built program in MATLAB (The MathWorks Inc., Natick, MA, USA) using the EasySpin program package (Stoll and Schweiger, 2006) to extract the spectral parameters—either one component with a slow dynamic or two components with different dynamics. A three-line, narrow EPR spectra is typical for nitroxide free radicals undergoing rapid isotropic motion, which can be characterized with aoN. The value of aoN was taken to be one half of the difference in the resonance fields of the high- and low-field lines. For the slow component, the distance between the outer peaks (2Azz) was monitored.
All the samples for EPR spectroscopy were prepared with spin probe 5-doxyl stearic acid (5-DSA) dissolved in ethanol (1% v/v of 200 mM).
2.4. High Resolution Mass Spectrometry
2.4.1. Derivatization with 7-(Diethylamino)coumarin-3-carbohydrazide (CHH)
Due to the low concentrations of lipid peroxidation products (LPPs) as well as the low ionization efficiencies—i.e., their low proton affinities—their chemical derivatization is necessary. It provides high proton affinities and enhances their ionization in positive ion mode and thus provides the detection of numerous LPPs. We used CHH for derivatization, assuming a rapid and specific reaction between CHH and aldehydes and ketones. Additionally, the hydrophobicity and relatively high mass of CHH were expected to enable the simultaneous extraction of both short aliphatic and nonpolar high molecular weight carbonyls with organic solvents. The CHH derivatization enhanced the ionization of both aliphatic and lipid-bound carbonyl-containing LPPs, giving access to both small, aliphatic, and water-soluble and large, nonpolar, lipid-esterified carbonylated species. The oxidized liposomes (1.5 mM) were individually derivatized with CHH (50 μL, 10 mM) at 37 °C for 1 h (
Figure 2). After derivatization, the samples were extracted in the same volume of chloroform, diluted with a mixture of methanol and chloroform (2:1, v/v), and analysed immediately.
2.4.2. Measurement Parameters
The samples were diluted (to 10 pmol/μL) in a mixture of methanol and chloroform (2:1, v/v) and analyzed by direct infusion using a Q Exactive™ Plus Hybrid Quadrupole-Orbitrap™ Mass Spectrometer (Thermo Scientific™). Type of mass detector: Orbitrap measuring range m/z: 50–6000 m/z; mass resolution: 140,000 FWHM (full width at half maximum); mass accuracy: <1 ppm with internal calibration, <3 ppm with external calibration. The MS spectra were acquired in Fourier Transform Mass Spectrometry (FT-MS) scan mode with a target mass resolution of 100,000 at m/z 400. The acquisition period was 15 min. The recorded spectra were analyzed with a Thermo Xcalibur Qual Browser (Xcalibur 4.2 SP1, Thermo Fisher Scientific Inc., Waltham, MA, USA). All the spectra were manually searched throughout the whole timeframe for all the suspected aliphatic carbonyl compounds with a mass accuracy of 5 ppm.
2.5. FTIR-ATR Spectroscopy
The concentration of DOPC for the FTIR measurements was adjusted to 20 mg mL
−1. The spectroscopic measurements were performed using a PerkinElmer “Spectrum 400 Series” spectrometer (Jožef Stefan Institute, Ljubljana, Slovenia) equipped with a Horizon ATR accessory (Harrick Scientific) with a trapezoidal germanium crystal. Each spectrum was collected at a nominal resolution of 4 cm
−1 resolution and as mean value of 32 spectra. A special holder for the ATR crystal was used. It was placed in contact with an aluminum block and the temperature was controlled by a circulating water bath. All the spectra were collected at 40 °C. A quantity of 200 μL of each sample was placed onto the ATR crystal and spread over the whole area. The sample was dried by purging with dry nitrogen until there was no significant change in the broad band at 3200–3600 cm
−1, which corresponded to ν(O–H) of the solvent [
41].
Data Analysis—The Extent of Lipid Peroxidation
A data analysis was performed by modifying the procedure used by Oleszko et al. [
42]. Since lipid peroxidation should lead to a change in the integral absorbance of the ν(C=O) band at 1737 cm
−1, it was analyzed with respect to the ν
as(CH
3) at 2959 cm
−1 band, where we expect no changes in the absorbance after the reaction [
42]. The values of the integral absorbances of both bands were calculated using an Origin Pro 9. According to Bradley and Kretch [
43], the IR spectra of solids usually consist of peaks which can be described using a Gaussian function, while gases are dominantly fitted with a Lorentzian function. Since the lipid membranes were in the fluid phase, the peak shape was expected to be a combination of Gaussian and Lorentzian curves. When fitted with a linear combination of Gaussian and Lorentzian curves, the bands in the interval 2800–3100 cm
−1 showed a large Lorentzian character, while the band in the interval 1600–1800 cm
−1 showed a large Gaussian character. This can be explained by the fact that this peak is composed of several slightly shifted Lorentzian peaks belonging to different species formed during the lipid peroxidation, which cannot be resolved. Therefore, peaks in the interval 2800–3100 cm
−1 were fitted to a pure Lorentzian curve, while the peak in the interval 1600–1800 cm
−1 was fitted to a pure Gaussian curve. In the first iteration, all the peak parameters were included in the fit. The obtained band widths were then used to compute the average band widths for the systems containing the selected flavonol. Finally, the integral absorbances were recomputed using fixed peak widths in order to reduce the parameter dependencies.
The ratio of the integrated absorbances
Ai of the
i-th sample (
i = 0 without flavonols,
i = 1, 2, 3 with flavonols) was calculated according to:
for all the samples before and after lipid peroxidation. The change in that ratio after lipid peroxidation (LP),
was determined relatively to that of the control sample, which is the liposome suspension before the occurrence of the reaction:
If the number of C=O bonds increases, the ratio
increases. Finally, the inhibition for each flavonol and each molar ratio was calculated using the formula:
where
ρ0 corresponds to the relative ratio of the integrated absorbances for the case of DOPC without inserted flavonols (
i = 0). If added flavonols hinder the lipid peroxidation,
Ri exhibits negative values with a minimum value of −1, corresponding to the total inhibition of the reaction. In that case, the value
Ri is a measure of the inhibition of the lipid peroxidation reaction (
Appendix A,
SI).
2.6. Dynamic Light Scattering (DLS) and Electrophoretic (ELS) Measurement
A photon correlation spectrophotometer equipped with a 532 nm green laser (Zetasizer Nano ZS, Malvern Instruments, Worcestershire, UK) was used for the determination of the size distribution and zeta potential of the unilamellar DOPC liposomes (Avanti’s Mini-Extruder with 100 nm membrane) in PBS at (25.0 ± 0.1) °C. The final concentration of the suspension was 0.2 mg mL−1. The intensity of the scattered light was measured at a 173° angle. The hydrodynamic diameter (dH) was determined from the peak maximum of the volume size function. The zeta potential (ζ) was calculated from the electrophoretic mobility using a Smoluchowski approximation (f(κa) = 1.5). The hydrodynamic radius values were reported as an average value of 10 measurements, while the zeta potential values were reported as an average of 3 independent measurements. The data processing was carried out using Zetasizer Software 7.13 (Malvern Instruments LTD, Malvern, Worcestershire, UK).
2.7. Small Angle X-ray Scattering (SAXS)
For the SAXS measurements, dried films were dispersed in water instead of PBS to avoid a decrease in the electronic contrast and final signal that could appear using the PBS buffer solution. To initiate lipid peroxidation, the dried films were resuspended in the water solutions of FeCl2 × 4H2O (10 mM, 250 μL) and H2O2 (10 mM, 250 μL). The lipid peroxidation reaction advanced for 1 h before recording. The final concentration of the samples for the SAXS measurements was 50 mg mL−1.
The SAXS measurements were carried out in transmission mode at defined temperatures by a laboratory SAXS instrument (SAXS-Point 2.0, Anton Paar, Graz, Austria). The SAXS camera was equipped with a micro-X-ray source operating at 50 W (point-focus) using Cu-Kα-radiation (λ = 0.1542 nm) and a 2D X-ray detector (EIGER2 R 500K, Dectris, Switzerland). The SAXS patterns were recorded at 571 mm sample-to-detector distance. All the isotropic 2-dimensional SAXS patterns were azimuthally averaged to 1-dimensional SAXS-curves. The SAXS curves of pure water were taken for background subtraction. The angular q-range was 0.01 nm−1 to 6 nm−1, with q being the magnitude of the scattering vector, which corresponds to a total 2θ region of 0.14° to 7° applying the conversion q [nm−1] to 2θ(°) using Equation (5). The sample cell in the X-ray beam was a quartz capillary (1 mm diameter, wall thickness of 10 µm) with two vacuum tight screwcaps on both ends inserted into a thermostatted sample-stage set to a defined temperature (30 °C). The vacuum in the camera during the measurement was kept at ≈1 mbar. The exposure time was 300 s times 3.
The analysis of the scattering data of liposome structures after the lipid peroxidation was performed using the programs GIFT [
44] and DECON [
45], developed by Otto Glatter. GIFT (Generalized Indirect Fourier Transformation) is based on the simultaneous determination of the form and the structure factor.
The scattering intensity is expressed by the following equation:
where
P(
q) and
S(
q) are the form and structure factor, respectively, and n is the number density of the particles.
P(
q) describes the internal structure of the particles, while
S(
q) describes the interaction between the particles. The value
q is the magnitude of the scattering vector and is related to the scattering angle by the following equation:
where
λ is the wavelength of the X-ray and 2
θ is the angle of the scattered X-rays. For the lamellar structure, the form factor
P(
q) can be expressed as:
where
A is the area of the lamellar phase. The relation between
Pl(
q) and the normal bilayer pair distribution function
pl(
r) is the Fourier transformation shown in (7), while the relation between
Pl(
q) and the self-correlation function of electron density function Δ
ρ(
r) is the Fourier transformation in (8):
A Fourier analysis of the multilamellar SAXS patterns (with the sharp Bragg peaks) was performed using the in-house Javascript program available online [
46]. The input parameters were: the number of the visible first Bragg peaks (2); the intensities of the two peaks; the lamellar
d-spacing of the peaks (6.1 nm); and the sign of the amplitudes (square roots of the intensities) of the two peaks, which can be either + or −. Only the combination of − for the two peak amplitudes gave a reasonable result for the electron density.
2.8. Atomic Force Microscopy (AFM) and Force Spectroscopy (FS)
2.8.1. Preparation of the Supported Lipid Bilayers (SLBs)
The procedure for the preparation of SLBs is the drop deposition method, which has already been reported [
47]. Briefly, a drop of multilamellar vesicles (MLVs) suspension (100 µL) was added to the fluid cell with a freshly cleaved mica plate and thermostated at 25 °C. After 10 min incubation, due to electrostatic interactions between the DOPC liposomes and mica, the liposomes adsorbed on the mica surface and formed SLB [
48]. The unadsorbed liposomes were removed by washing the surface with the filtered (0.22 µm Whatman) PBS solution.
2.8.2. AFM Imaging in Fluid and FS Measurements before and after Lipid Peroxidation
AFM images were obtained by scanning the SLBs on the mica surface in the fluid using an AFM FastScan Dimension (Bruker Billerica, USA), operated using the PeakForce QNM mode sing Scanfastsyst—Fluid + Bruker probes, with the spring constant (Nom.
k = 0.7 Nm
−1; Nom. resonant freq.
ν = 150 kHz). The imaging was performed at 25.0 °C, allowing thermal equilibration before each sample imaging. The thermal tune method was used for the cantilever calibration as previously described [
47,
49]. AFM images were collected using random spot surface sampling (at least two areas per sample) for each analyzed sample. The quantitative mechanical data was obtained by employing DMT modulus within the Bruker software. All the images were processed by first-order two-dimensional flattening only and analyzed using the NanoScope Analysis software (Version 1.9).
2.9. Statistical Data Analysis
Where applicable, the obtained data have been presented as the mean value and the standard deviation. A general linear model (GLM) for ANOVA has been used for the statistical comparison. The influence of each flavonol and its molar fraction, as well as the interaction between the molar fraction and flavonol typ were tested. Tukey’s post hoc HSD (honestly significant difference) test was performed to test the differences between the groups. Differences were considered statistically significant if p ≤ 0.05. The statistical analysis was performed using the software STATISTICA (data analysis software system), version 12.0 (StatSoft, Tulsa, OK, USA).
4. Conclusions
We have used a multi-technique approach to study the structural changes of the model lipid membrane caused by lipid peroxidation and the antioxidative role of three structurally and chemically different flavonols, QUE, MCE, and MCI.
Lipid peroxidation leads to structural alterations that violate the membrane integrity, as indicated by the changes in the crucial properties of the membranes, such as elasticity, surface roughness, thickness, and fluidity. All these properties are essential for the function of biological membranes.
The antioxidative activity of the three investigated flavonols strongly depends on the environment in which they are located. Specifically, when the antioxidative assay was performed in the solution, MCE showed the highest activity in the scavenging of free radicals, while the most hydrophobic flavonol, QUE, showed the lowest. In contrast to this, the comparison between the flavonols when they were incorporated within the liposome membrane displayed different results. The most hydrophilic flavonol, MCI, showed a higher protection than QUE and MCE, which are more hydrophobic and located deep within the bilayer. This was concluded based on the fact that, while the number of identified peroxidation products (taken as the measure of flavonol inhibition) decreased with an increase in the molar fraction for all the flavonols, the effect was the most pronounced in the case of MCI. Since MCI is located closer to the surface of the membrane, oriented towards the aqueous medium, it is more exposed to the incoming radicals and is able to scavenge them before they reach the reactive site.
The loss of multilamellar structure and the loss of lipid material upon lipid peroxidation indicated multiple surface processes and the rearrangement of the membrane. As a result of these processes, changes in the surface roughness and elasticity were observed. Since these changes are less pronounced in the presence of flavonols, it can be concluded that they preserved the supramolecular and mechanical properties of the membrane. Finally, the significant degree of inhibition of peroxidation process of all investigated flavonols has also been confirmed by the preservation of the bilayer fluidity.
Our study suggests that all the techniques employed can be used as a highly valuable tool in other biomedical applications aimed at screening and monitoring the lipid peroxidation effects at the cellular level. Furthermore, it was demonstrated that, when studying the protective activity of various antioxidants, it is necessary to consider their environment along with their chemical properties.