A study of the interaction between cell membranes and small molecules derived from lignin, a protective phenolic biopolymer found in vascular plants, is crucial for identifying their potential as pharmacological and toxicological agents. In this work, the interactions of model cell membranes [supported 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) lipid bilayers] are compared for three βO4 dimers of coniferyl alcohol (G lignin monomer): guaiacylglycerol guaiacol ester with a hydroxypropenyl (HOC3H4-) tail (G-βO4′-G), a truncated GG dimer without HOC3H4- (G-βO4′-truncG), and a benzylated GG dimer (benzG-βO4′-G). The uptake of the lignin dimers (per mass of lipid) and the energy dissipation (a measure of bilayer disorder) are higher for benzG-βO4′-G and G-βO4′-truncG than those for G-βO4′-G in the gel-phase DPPC bilayer, as measured using quartz crystal microbalance with dissipation (QCM-D). A similar uptake of G-βO4′-truncG is observed for a fluid-phase bilayer of 1,2-dioleoyl-sn-glycero-3-phosphocholine, suggesting that the effect of the bilayer phase on dimer uptake is minimal. The effects of increasing lignin dimer concentration are examined through an analysis of density profiles, potential of mean force curves, lipid order parameters, and bilayer area compressibilities (disorder) in the lipid bilayers obtained from molecular dynamics simulations. Dimer distributions and potentials of mean force indicate that the penetration into bilayers is higher for benzG-βO4′-G and G-βO4′-truncG than that for G-βO4′-G, consistent with the QCM-D results. Increased lipid tail disorder due to dimer penetration leads to a thinning and softening of the bilayers. Minor differences in the structure of lignin derivatives (such as truncating the hydroxypropenyl tail) have significant impacts on their ability to penetrate lipid bilayers.

Lignin is a complex phenolic cross-linked polymer that, together with cellulose, acts as the structural material in plants and is important for rigidity and water transport within plant cell walls.1 Lignin is the second most abundant biopolymer after cellulose, comprising 15%−30% of biomass. However, unlike cellulose, lignin has limited current uses due to the heterogeneity of its structure. While comprising only three monomers, lignin has a complex structure due to a variation in both the sequence and the types of bonds between monomers. The goal of deconstructing lignin to commercially viable small molecules for use as chemicals or advanced materials has proved elusive because of this heterogeneity, leading to much of lignin being discarded as waste or burned for energy and only a small part being utilized for value-added products.2–4 Currently, the potential uses of products derived from lignin are being explored for a wide variety of applications. These include using lignin and its small-molecule derivatives as commodity chemical feedstocks, reinforcements for polymers, UV protectants, in tissue engineering materials, and as antioxidants and antimicrobials.2,5–7

Lignin is composed of three monolignols (the aromatic alcohols p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol) and every monomer includes a characteristic phenolic ring [p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S), respectively] and a hydroxypropenyl group.8 The monomers can be linked together in many different ways to form dimers, trimers, or more complex oligomers connected by various bonding motifs, including βO4, αO4, 4O5, 55, ββ, β5, and β1.9,10 This variety makes the structure of lignin very complicated and implies that when lignin is deconstructed by methods such as pyrolysis, hydrogenolysis, and hydrolysis, an extremely diverse mixture of lignin-derived molecules is acquired.11,12 Finding ways to analyze, separate, and identify small molecule lignin derivatives is arduous because of this complexity. In our previous published work, we proposed a generalized pathway for synthesizing different βO4 dimers that are potential model compounds in the development of applications, and analytical and separation methods for small lignin-derived molecules.13 

A study of the interaction between cell membranes and small molecules is crucial for recognizing the pharmacological mechanism of some compounds and to elucidate their toxicological impacts on biological systems.14,15 For instance, the mode of action of a substantial number of small pharmaceutical drugs such as tranquillizers, anesthetics, narcotics, and antidepressants is based on their transport through the hydrophobic interior of the cell membrane.16 Phenolic compounds such as lignans have structures similar to lignin oligomers and are of great interest for human therapeutic applications due to possible protective (e.g., antioxidant) or growth-inhibiting (e.g., anticancer) properties.17–21 Thus, the interaction of compounds derived from lignin with cell membranes is an indicator of lignin's therapeutic or toxic behavior10,22 and allows us to identify effective ways to use the oligomers of lignin (some of which overlap with lignans23) for therapeutic applications with the consideration of risk to humans and the ecosystem.24 

The outer leaflet of mammalian cell membranes, facing the extracellular space, is primarily made of phosphatidylcholine (PC), sphingomyelin (SM), phosphatidylethanolamine (PE), and cholesterol.25,26 Phosphatidylcholine (PC) and sphingomyelin (SM) are the most abundant phospholipids in the human red blood cell (erythrocyte) membrane, with PC constituting ∼35% of the membrane.27,28 Moreover, the pulmonary surfactant of all mammalian species contains substantial amounts (∼80%) of PC, with nearly 60% of it being in the dipalmitoylated form.29 Here, a 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) bilayer has been chosen as a simplified model system for determining the basic functions of biological mammalian membranes.30,31 Building on our previously published work in dilute systems, here, we study the interactions of three βO4 dimers of coniferyl alcohol (GG lignin dimer), namely, the novel G-βO4′-G and benzG-βO4′-G dimers and the commercially available G-βO4′-truncG dimer (Fig. 1) with DPPC lipid bilayers. All three monomers are built around two guaiacyl (G) units linked with a βO4 bond. G-βO4′-G has a hydroxypropenyl tail expected to be found in natural lignin, while G-βO4′-truncG is truncated, and benzG-βO4′-G has the hydroxypropenyl tail but is benzylated at the phenol end. In our previously published work, the effect of the incorporation of these dimers on the structure of a DPPC lipid bilayer was interpreted from changes in the gel-to-fluid phase transition at a limit of infinite dilution using differential scanning calorimetry (DSC) and molecular dynamics (MD) simulations.32benzG-βO4′-G and G-βO4′-truncG decreased the bilayer transition temperature significantly relative to pure DPPC, while G-βO4′-G had little effect on the transition temperature.32 Using MD simulations of a single dimer molecule in DPPC bilayers, the location of the dimer within the bilayer was examined and the gel−fluid phase transition temperatures were calculated, which showed the same trend as that of the DSC results. In the current contribution, the concentration dependence of the uptake of the G-βO4′-G, G-βO4′-truncG, and benzG-βO4′-G dimers on DPPC bilayers and the corresponding effect on lipid bilayer disorder are investigated using a quartz crystal microbalance with dissipation (QCM-D) and complementary MD simulations.

FIG. 1.

Chemical structures of the three βO4-linked lignin dimers used in the experiments and simulation, and the lipids DPPC and DOPC.

FIG. 1.

Chemical structures of the three βO4-linked lignin dimers used in the experiments and simulation, and the lipids DPPC and DOPC.

Close modal

QCM-D has been widely used as a robust method for investigating and quantifying the adsorption of small molecules to lipid bilayers supported on solid surfaces.33 Previously, Joshi et al. used QCM to study the interaction of aminoglycosides (kanamycin A and neomycin B) with model cell membranes supported on gold and observed similar rates of membrane penetration for both aminoglycosides followed by bilayer disruption when a critical concentration threshold was reached.21 Kannisto et al. studied the interaction of two small-molecule drugs, propranolol and tetracaine, with lipid bilayers supported on silica using QCM and successfully measured the adsorbed mass of the compounds as a function of their concentration in the aqueous phase, as well as the impact of the compounds on the bilayer viscoelasticity.34 Wargenau et al. evaluated the cell membrane penetration potential of low-molecular-weight lipid-soluble compounds (vanillin, gallic acid, and protocatechualdehyde) using supported phospholipid bilayers on QCM-D silica sensors. They observed that while the studied compounds did not cause any noticeable mass density changes in the bilayer, they caused significant changes in their gel−fluid phase transitions.35 Here, lipid bilayers are formed on gold QCM sensors by solvent-assisted deposition from isopropanol followed by an injection of lignin dimer aqueous solutions in a flow cell. Changes in dissipation (a measure of bilayer rigidity and disorder) and mass of the lipid bilayer are used to interpret the uptake of the solute into the lipid bilayer. The interactions of silica nanoparticles functionalized with a double-bond modified G-βO4′-truncG dimer and lignin-graft-PLGA nanoparticles with DPPC lipid bilayers have been studied using QCM-D and presented in our recent publications.36,37

In addition to experimental studies, we present the results of the MD simulations of the same lignin dimers interacting with DPPC lipid bilayers. Computational studies of small molecules interacting with lipid bilayers have provided a molecular understanding of their effects on cell membranes. Some examples include the work of Lin et al., who showed that 3 mol. % DMSO has significant structural and permeability effects on DMPC lipid bilayers,38 and that of Lyu et al., who observed variations in curcumin interactions with lipid bilayers that they related to differences in curcumin antimicrobial activity with different microorganisms.39 Recently, Vermaas et al. reported the permeabilities of various lignin monomer and dimer derivatives, including G-βO4′-truncG through lipid bilayers at low lignin concentrations.40 Other MD simulation studies of lignin dimers and oligomers focused on their conformational and mechanical properties and adsorption dynamics on cellulose/hemicellulose surfaces.41–43 In this study, we examine the concentration dependence of dimer incorporation on bilayer properties such as molecule positioning, bilayer thickness, and the compressibility of dimer/lipid systems in order to gain new insights into the mechanism of action of lignin oligomers with potential therapeutical or toxic properties at biological interfaces.

Isopropanol (99%) and de-ionized water (ASTM type II) were purchased from VWR International; phosphate-buffered saline (PBS) from Sigma Aldrich; 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) from Avanti Polar Lipids (Alabaster, USA); and guaiacylglycerol-β-guaiacyl ether, which is referred to as the G-βO4′-truncG dimer (≥97%), and 1,2-didecanoyl-sn-glycero-3-phosphocholine (DOPC) from TCI America. All chemicals were used as received without further purification.

The method of synthesis of 1-(4-hydroxy-3-methoxyphenyl)-2-[2-methoxy-4-(3-hydroxyprop-1-en-1-yl)phenoxy]propane-1,3-diol, which in this work is referred to as G-βO4′-G, followed the detailed description of Asare et al.13 This dimer shares the same βO4 linked units with the commercially available guaiacylglycerol-β-guaiacyl ether, which is referred to as the G-βO4′-truncG lignin dimer. However, G-βO4′-G possesses a 3-hydroxyprop-1-enyl tail. The synthesis of a benzyl-modified GG dimer 1-(4-benzyloxy-3-methoxyphenyl)-2-[2-methoxy-4-(3-hydroxyprop-1-en-1-yl)phenoxy]propane-1,3-diol (benzG-βO4′-G) was performed with a slight alteration in the G-βO4′-G synthesis method: a benzyl group was used to protect the phenol group in the A ring of vanillin rather than a triisopropylsilyl group, as detailed previously.32 The molecular structures of the G-βO4′-G and benzG-βO4′-G dimers were characterized by nuclear magnetic resonance (H NMR) and high-resolution mass spectrometry (HR-MS), as described in the supplementary material.69 

A QCM-D (Q-sense E4, Biolin Scientific, Sweden) was employed to monitor the interaction of the G-βO4′-G, G-βO4′-truncG, and benzG-βO4′-G dimers in PBS buffer (pH 7.4) doped with a small amount of isopropanol with DPPC- and DOPC-supported lipid bilayers on a gold substrate at 297 ± 0.2 K. The variations in oscillation frequency (Δf) and energy dissipation (ΔD) of the sensor were measured in real time during the formation of a supported lipid bilayer (SLB), the absorption of dimers from 0.1 mg/ml (unless otherwise indicated) aqueous solutions of dimers into the bilayer, and the rinsing of the bilayer, as discussed below.

Both DPPC and DOPC lipid bilayers were created on a QCM-D gold-coated sensor (Nanoscience Instruments, Phoenix, USA) using the solvent-assisted lipid bilayer (SALB) formation method reported by Tabaei et al. with slight changes.44 Initially, PBS solution was passed over the substrate until a steady baseline was reached, followed by isopropanol for ∼35 min, all at a flow rate of 30 ml/h. Then, a lipid solution in isopropanol (0.5 mg/ml for both DOPC and DPPC lipid bilayers) was flowed through the QCM-D cell for 60 min. Finally, the substrate was rinsed with PBS again, leading to a rearrangement of the lipids adsorbed on the substrate into an SLB.45–47 

After forming the lipid bilayers, solutions containing G-βO4′-G, G-βO4′-truncG, or benzG-βO4′-G dimers at a concentration of 0.1 mg/ml were passed through the QCM chamber for 60 min at a flow rate of 30 ml/h. To make the solutions of dimers, the dimers were first dissolved in 0.2 ml isopropanol and then added to 29.8 ml PBS to obtain a isopropanol/PBS solution (0.67% v/v). Finally, the substrate was rinsed with pure PBS for 60 min to remove unbound molecules from the surface. Under the conditions of this experiment (297 K), the supported DPPC bilayer was in its gel phase. The G-βO4′-truncG dimer solution (at 0.1 mg/ml) was also used to study the interaction with a phosphocholine lipid bilayer, DOPC, in its fluid phase.

QCM-D experimental data collected at the third overtone (n = 3; 15 MHz) were analyzed by using the Sauerbrey model [Eq. (1)], which assumes that the adjacent layer is thin, rigid, and homogenous, and is utilized to determine the mass of lipid bilayers formed on QCM sensors,45,48,49

ΔmA=CΔfn.
(1)

In this equation, the mass of lipid bilayer per area (Δm/A) was measured from the change in frequency (Δf). C is the material-specific Sauerbrey constant (17.7 ng cm−2 Hz−1), and n is the overtone number at which the frequency is being recorded.50 The penetration depth of the overtones decreases with an increasing overtone number.21,50 The fundamental frequency (n = 1, 5 MHz), which measures a penetration depth of ∼250 nm from the quartz crystal surface, was not considered because this resonance is very sensitive to variations in the bulk solution and prone to producing a noisy response. All the frequency values in this work are divided by the overtone number which is n = 3 unless otherwise specified. In pure water, the penetration depth for the third overtones is ∼145 nm.48,51 The thickness of a hydrated phospholipid bilayer is approximately 8 nm (much lower than the penetration depth at n = 3; d/λ3 = 0.055), so the Sauerbrey equation [Eq. (1)] is expected to be applicable.48,52

1. Simulation systems

For simulations, the DPPC lipid bilayers were composed of 128 lipids with 64 in each leaflet and were fully hydrated with water:lipid ratios of 50–55:1. The bilayers were placed in the center of the simulation box with periodic boundaries applied in all directions. The GROMOS 54A7 force field53 and the simple point charge54 water model were employed. The process of force field validation, as well as the procedure for adding dimer molecules to the system, was described in our previous work.32 The average temperature was controlled with the thermostat of Bussi et al. with a 0.1 ps time constant at 326 K and the average pressure was controlled with a Berendsen barostat with semi-isotropic coupling and a 2.5 ps time constant at 1 bar. The particle mesh Ewald algorithm was used for long-range electrostatics. The short-range cutoff distance was 1.4 nm for van der Waals and electrostatic interactions.

2. Potential of mean force

The potential of mean force (PMF) plays an important role in quantifying the partitioning of molecules into lipid bilayers. It describes the free energy landscape of compounds as a function of a collective coordinate (or coordinates). The collective coordinate that we used was the distance between the centers of mass of the lignin molecules and the center of mass of the bilayer in the bilayer normal direction. High-energy barriers along collective coordinates usually lead to insufficient sampling in some regions in unbiased simulations. In this work, we employed umbrella sampling to bias the simulations and alleviate this problem.55 PMF curves for the G-βO4′-G, G-βO4′-truncG, and benzG-βO4′-G dimers with 128 lipid DPPC bilayers were calculated. We performed the biased simulations for each umbrella sampling window for 80 ns with an equilibration time of 30 ns at 326 K and 1 bar. The initial configurations for 38 windows were prepared with distances ranging from 0.0 to 3.8 nm with a spacing of 0.1 nm. The force constants for the harmonic biasing potentials were 3000 kJ/(mol nm2). Most of the initial configurations were obtained from unbiased simulations. The initial configurations near the bilayer center that could not be extracted from the unbiased simulations were obtained by dragging one dimer to the desired distances at a rate of 3×104nm/ps. The weighted histogram analysis method56 was employed to combine the data from each window and correct the biased PMF to an unbiased one.

3. Area compressibility modulus

Area compressibility modulus, KA, was calculated using the average bilayer area, A, and the variance of the area σA2 determined from the simulation results according to Eq. (2),57 

KA=AkBTσA2.
(2)

Details on error estimation of KA are given in supplementary material.69 The compressibility is indirectly related to the viscoelastic behavior embodied by dissipation values obtained from QCM-D. We expect the dissipation to be higher in a more compressible bilayer.

Artificial membranes composed of SLBs have structures and dynamics that mimic the primary characteristics of cell membranes and are, thus, frequently used as simplified systems for studying membrane thermodynamics and transport processes.58,59 The SALB deposition method was used as the membrane preparation approach here because it has been shown to produce reproducible bilayers on unmodified gold QCM sensors.44 QCM-D is a highly sensitive method, especially for viscous layers containing large amounts of water (e.g., lipid bilayers), and has been used to investigate SLB formation and molecular interactions at lipid–bilayer interfaces.50 In QCM-D, the energy dissipation is recorded as the damping of the resonance when the driving voltage to the sensor is shut off. Variations in dissipation (ΔD) are related to changes in the rigidity or viscoelasticity of the layer adhered on the surface;24 the deformation of a soft or disordered adlayer on the substrate leads to large dissipation changes, whereas nondeformable or rigid adlayers are identified by small dissipation changes.

Here, QCM-D was utilized to examine and quantify the adsorption of lignin dimers to supported lipid bilayers. Representative frequency and dissipation plots for the formation of a DPPC-supported lipid bilayer using the SALB method at 297 ± 0.2 K are illustrated in Fig. 2, where DPPC is below its gel–fluid phase transition temperature (Tm = 315.3 ± 0.15 K). In Fig. 2, the time axis starts after reaching steady state, with PBS flowing over the sensor. PBS is followed by isopropanol, followed by a solution of DPPC in isopropanol (0.5 mg/ml), and finally PBS again. Rinsing with PBS in the final step causes the lipids adsorbed to the sensor from the isopropanol solution to reorganize into a bilayer,44 as shown by a sharp increase in frequency and a corresponding sharp decrease in dissipation. The Δf value for the bilayer in Fig. 2 before dividing by the overtone number (n = 3) is ∼ −62 Hz, which, using the Sauerbrey equation [Eq. (1)], corresponds to a mass value of ∼366 ng/cm2, falling within the mass range reported in the literature for a supported lipid bilayer.60,61 The ΔD for the gel-phase DPPC bilayer in Fig. 2(b) is 10.6 × 10−6, which is higher than the value reported for fluid-phase lipid bilayers (less than 0.5 × 10−6);44 however, it is consistent with the value reported by Lind et al. for gel-phase DPPC lipid bilayers in PBS at 298 K (∼6.5 × 10−6 at n = 7).62 The mass of the SLB formed on the sensor was measured from the difference in the frequency of the initial PBS baseline (t = 5 min) and the baseline in PBS after SLB formation (t = 150 min) using the Sauerbrey equation [Eq. (1)]. The DPPC lipid bilayer mass for all SLBs in this work was above 350 ng/cm2, which is in agreement with the reported mass for a rigid SLB that is completely covered and is well coupled to the sensor.47,63,64

FIG. 2.

Representative QCM-D results for solvent-assisted DPPC lipid bilayer formation on a gold sensor. (a) Frequency shifts divided by the overtone number (n= 3) and (b) dissipation shifts at the third overtone (n= 3). The parts of the graphs separated by the vertical dashed lines denote the introduction of flowing PBS, isopropanol (“i-PrOH”), 0.5 mg/ml DPPC lipid in isopropanol (“DPPC”), and finally PBS. The insets show the schematics of lipid deposition and bilayer formation during the process.

FIG. 2.

Representative QCM-D results for solvent-assisted DPPC lipid bilayer formation on a gold sensor. (a) Frequency shifts divided by the overtone number (n= 3) and (b) dissipation shifts at the third overtone (n= 3). The parts of the graphs separated by the vertical dashed lines denote the introduction of flowing PBS, isopropanol (“i-PrOH”), 0.5 mg/ml DPPC lipid in isopropanol (“DPPC”), and finally PBS. The insets show the schematics of lipid deposition and bilayer formation during the process.

Close modal

The amount of the lignin dimers interacting with the bilayer, and consequently the ratio of the mass of the bound dimer to the mass of the lipid bilayer, depends on the concentration of the dimer in solution. Because QCM sensors are sensitive to the density and viscosity of fluids, it is conventional to flow dilute solutions (with concentrations in the order of 1–1000 μM) over the sensor when studying the adsorption of molecules to lipid bilayers.21,34,65 Concentration-dependent studies of G-βO4′-truncG were conducted (Fig. 3) to understand the sensitivity of QCM to the contributions of uptake by the SLB as well as bulk solvent properties. QCM-D measurements were carried out using three different concentrations of G-βO4′-truncG solutions in isopropanol/PBS (0.67% v/v) in increasing order (0.01, 0.02, and 0.1 mg/ml). In Fig. 3, t = 0 corresponds to the initial-supported DPPC bilayer and the values of Δf/3 and ΔD are shifted to 0 at this initial condition. At t = 12 min, the flow of the 0.01 mg/ml dimer solution is initiated over the SLB for 1 h, resulting in a decrease in frequency (Fig. 3), which is indicative of dimer association and incorporation into the lipid bilayer. However, the frequency does not plateau over the 1 h period of injection, meaning that equilibrium with the SLB is not reached. Similarly, increasing the dimer solution concentration to 0.02 mg/ml for 1 h results in a further decrease in frequency without any evidence of membrane saturation. When the flow is switched to a much more concentrated solution of dimer (0.1 mg/ml), the frequency continues to decrease and then plateaus at ∼268 min. In the final step, the sensor was rinsed with PBS solution for ∼115 min. Thus, the concentration of 0.1 mg/ml was chosen to study the interaction of the dimers with the SLB within a time period of 1 h, with a goal of achieving measurable equilibriumlike interactions of the bilayer in semidilute solutions.

FIG. 3.

(a) Changes in frequency divided by the overtone (n = 3) and (b) changes in dissipation with time as G-βO4′-truncG dimer solutions of variable concentrations (0.01, 0.02, and 0.1 mg/ml) flow over a DPPC lipid bilayer. The vertical dashed lines denote time frames over which the solutions of lignin dimer with different concentrations were introduced, followed by PBS. Time of zero corresponds to the initial DPPC bilayer, the introduction of the 0.01 mg/ml dimer solution started at t = 12 min, followed by the introduction of 0.02 mg/ml solution after 60 min and 0.1 mg/ml after 60 additional minutes. The flow at 0.1 mg/ml was continued for 176 min before rinsing with PBS for 115 min. Data are presented for the third overtone (n = 3).

FIG. 3.

(a) Changes in frequency divided by the overtone (n = 3) and (b) changes in dissipation with time as G-βO4′-truncG dimer solutions of variable concentrations (0.01, 0.02, and 0.1 mg/ml) flow over a DPPC lipid bilayer. The vertical dashed lines denote time frames over which the solutions of lignin dimer with different concentrations were introduced, followed by PBS. Time of zero corresponds to the initial DPPC bilayer, the introduction of the 0.01 mg/ml dimer solution started at t = 12 min, followed by the introduction of 0.02 mg/ml solution after 60 min and 0.1 mg/ml after 60 additional minutes. The flow at 0.1 mg/ml was continued for 176 min before rinsing with PBS for 115 min. Data are presented for the third overtone (n = 3).

Close modal

To investigate the effect of the modifications of the lignin GG dimer chemical structures on the interactions with phospholipid membranes, 0.1 mg/ml solutions of benzG-βO4′-G, G-βO4′-truncG, and G-βO4′-G in isopropanol/PBS (0.67% v/v) were separately introduced to the supported DPPC bilayers on QCM sensors. The resulting QCM-D responses for Δf/3 and ΔD as functions of time are presented in Fig. 4, in which the time of zero has been set for the fully developed bilayer and a corresponding Δf/3 value of zero is set. As shown in Fig. 4(a), upon the introduction of the benzG-βO4′-G dimer to the bilayer at ∼7 min, Δf/3 drops rapidly due to a change in the solvent properties of the semidilute isopropanol-dope dimer solution (0.1 mg/ml). Concurrent with the decrease in frequency, a step change followed by a more gradual increase occurs in the dissipation profile [Fig. 4(b)]. Within 1 h of initiating the benzG-βO4′-G dimer flow over the bilayer, Δf/3 continues to slowly decrease due to the uptake of the dimer by the bilayer (causing an increase in film mass). A subsequent switch to pure PBS solution at ∼67 min results in a rapid increase in the frequency. This is mainly attributed to a difference in the bulk solvent properties and a possible redissolution of some of the dimers. Concurrently, ΔD undergoes a rapid drop immediately after switching to pure PBS and then gradually plateaus at a relatively constant value above the starting value, suggesting that the final SLB is more fluid than the initial bilayer and retains some amount of dimer.

FIG. 4.

Interactions of G-βO4′-G (red), G-βO4′-truncG (black), and benzG-βO4′-G (blue) dimers with supported DPPC lipid bilayers on gold-coated quartz crystal sensors obtained by QCM-D. (a) Changes in frequency divided by the overtone (n = 3) and (b) changes in dissipation. Time is measured starting from dimer injection onto an existing bilayer (“Lignin dimers”), followed by the PBS rinse (“PBS”). Δf/3 and ΔD are shifted to a value of “zero” at time “zero” corresponding to the initial DPPC bilayer. Data are presented for the third overtone (n = 3).

FIG. 4.

Interactions of G-βO4′-G (red), G-βO4′-truncG (black), and benzG-βO4′-G (blue) dimers with supported DPPC lipid bilayers on gold-coated quartz crystal sensors obtained by QCM-D. (a) Changes in frequency divided by the overtone (n = 3) and (b) changes in dissipation. Time is measured starting from dimer injection onto an existing bilayer (“Lignin dimers”), followed by the PBS rinse (“PBS”). Δf/3 and ΔD are shifted to a value of “zero” at time “zero” corresponding to the initial DPPC bilayer. Data are presented for the third overtone (n = 3).

Close modal

The net frequency change in the bilayer due to the binding and incorporation of the lignin dimers was interpreted from the change in frequency representing the initial DPPC-supported bilayer (before ∼7 min) and the bilayer after rinsing the sensor with pure PBS (at ∼130 min). benzG-βO4′-G and G-βO4′-truncG displayed similar trends. The net Δf/3 changes for the benzG-βO4′-G and G-βO4′-truncG were ∼ −10.13 and ∼ −3.17 Hz, respectively. Similarly, the net ΔD changes of the bilayer for the benzG-βO4′-G and G-βO4′-truncG dimers were ∼3.4 × 10−6 and ∼11.5 × 10−6, respectively [Fig. 4(b)]. The dissipation shifts measured by QCM-D are an indirect measurement of bilayer fluidity and its viscous losses.61 The increase of dissipation with the introduction of benzG-βO4′-G and G-βO4′-truncG is consistent with dimer incorporation into the bilayer and suggests that these hydrophobic dimers ultimately perturb the bilayer's rigidity and correspond to an increase in dissipation loss to the surrounding medium.66 The results of a separate control experiment presented in Figs. S1(a) and S1(b)69 show that the use of a small amount of isopropanol to solubilize the dimers (0.67 vol. % in the PBS solution) has a minimal effect on the net frequency (−0.66 Hz) and net dissipation (less than 0.5 × 10−6) relative to the dimer uptake in the SLBs.

At a concentration of 0.1 mg/ml, the interaction of G-βO4′-G with SLBs is different from that of benzG-βO4′-G and G-βO4′-truncG, as interpreted from their respective frequency and dissipation responses (Fig. 4). As seen in Fig. 4(a), G-βO4′-G incorporates more slowly into the membrane. Interestingly, Δf/3 increases when switching to the PBS buffer but does not plateau over the 1-h period of buffer rinse. Instead, it trends toward the initial frequency. The reduced changes in net frequency for the G-βO4′-G dimer (∼ −1.94 Hz) relative to benzG-βO4′-G and G-βO4′-truncG and the ability to resolubilize it during rinsing suggest that G-βO4′-G did not associate appreciably with the bilayer interior. The small net changes in ΔD (less than 1 × 10−6) indicate that the bilayer fluidity of the SLB was largely unaffected by the G-βO4′-G dimer addition and subsequent rinsing with PBS. Thus, the time-dependent desorption behavior suggests that the G-βO4′-G dimer is not well incorporated into the SLB relative to the G-βO4′-G dimer, G-βO4′-truncG.

Table I summarizes the mass of dimers associated with the bilayers after the final PBS rinse relative to the mass of the supported bilayer initially deposited onto the sensor. The results are normalized with respect to bilayer mass because the bilayer mass varied slightly with each bilayer formed using the SALB method (an average of 400 ± 96 ng/cm2), which is consistent with a range reported in the literature for the mass of the supported phosphatidylcholine lipid bilayers60,61 and with the observation that DPPC-supported lipid bilayers deposited in the gel phase have more variability than those deposited in the fluid phase.62 The ratio of adsorbed dimer mass to DPPC bilayer mass follows the following trend: G-βO4′-G < G-βO4′-truncG < benzG-βO4′-G. This is consistent with the order of increasing equilibrium partitioning of these dilute lignin compounds into phospholipid bilayers67 as well as their estimated hydrophobicity. Our previous differential scanning calorimetry and MD studies of dimer incorporation into dilute systems also showed enhanced partitioning of the benzylated and truncated GG dimers into DPPC relative to a naturally occurring G-βO4′-G dimer.32 G-βO4′-G has the most hydroxyl groups (four), one of which is a part of the hydroxypropenyl (HOC3H4) tail. The free hydroxyl groups increase the hydrophilicity of the dimer and consequently decrease its penetration depth into the interior of the lipid bilayer, leading to slow diffusion away from the bilayer following the PBS rinse. In a study by Tammela et al., flavonoids with more than three free hydroxyl groups (e.g., luteolin, quercetin, and morin) were reported to initially bind strongly to the heads of the DPPC lipid bilayer via hydrogen bonding, but not to diffuse and transport deeply into the hydrophobic interior of the bilayers compared with more hydrophobic flavonoids.68 Here, G-βO4′-truncG lacks the hydroxypropenyl tail, which increases its hydrophobicity and ultimately leads to more penetration. Similarly, benzG-βO4′-G has an added benzyl group at the phenolic end of the dimer, which causes an increase in hydrophobicity, allowing it to penetrate deeper into the bilayer. Our previously published PMF calculations for dilute dimeric systems in fluid-phase DPPC bilayers illustrated that a major part of the G-βO4′-G dimer resides at or close to the exterior bilayer surface with only a small probability of finding this dimer inside the bilayer.32 In contrast, a major part of G-βO4′-truncG is embedded in the bilayer with only a small probability of finding it at the exterior bilayer surfaces. Our PMF studies also showed that benzG-βO4′-G is expected to reside almost exclusively in the bilayer interior.32 

TABLE I.

Quantitative QCM-D results of the lignin GG dimers' interactions with the synthetic DPPC lipid bilayers with 1 h of exposure at a concentration of 0.1 mg/ml of dimer solutions.

DimerG-βO4′-GaG-βO4′-truncGbbenzG-βO4′-Gb
Average ratio of the mass of the bound dimer to the mass of the pure DPPC bilayer 0.10 ± 0.009 0.16 ± 0.03 0.25 ± 0.08 
DimerG-βO4′-GaG-βO4′-truncGbbenzG-βO4′-Gb
Average ratio of the mass of the bound dimer to the mass of the pure DPPC bilayer 0.10 ± 0.009 0.16 ± 0.03 0.25 ± 0.08 
a

Three replicate measurements constitute the basis for these data.

b

Five replicate measurements constitute the basis for these data.

The MD simulation studies reported here were performed at T = 326 K to allow for equilibration within the time scale of the simulations. However, the QCM-D experiments were performed at 297 K, which is below Tm of DPPC (Tm = 315.3 ± 0.15 K), because preliminary experiments indicated that SALB formation of a DPPC bilayer was unsuccessful at 318 K, a temperature above its Tm (data not given). To show that our QCM results were also valid for fluid bilayers, the interaction of G-βO4′-truncG as the model compound with the phosphatidylcholine lipid bilayer, DOPC (Tm = 256.65 K) was investigated at 297 K. Representative QCM-D results for solvent-assisted DOPC lipid bilayer formation on a gold sensor and the resulting frequency and dissipation profiles after the introduction of G-βO4′-truncG to the bilayer are included in Figs. S2 and S3 (Ref. 69), respectively. The frequency and dissipation values for a DOPC lipid bilayer formed on gold were found to be −15 ± 1.5 Hz and (0.94 ± 0.2) × 10−6, respectively [Figs. S2(a) and S2(b)]69. The net change in frequency due to the incorporation of G-βO4′-truncG into the DOPC lipid bilayer in the fluid phase and the average ratio of the mass of the adsorbed G-βO4′-truncG dimer to the mass of the DOPC bilayer were ∼ −2.33 Hz [Fig. S3(a)]69 and 0.20 ± 0.09 (Table S169), respectively. These values are consistent with our measurements of G-βO4′-truncG dimer interactions with DPPC bilayers in the gel phase. This suggests that the phase of the lipid bilayer (gel or fluid) does not drastically affect the amount of dimers associated with the membrane. However, the net ΔD changes for the G-βO4′-truncG dimer and the DOPC bilayer were ∼1 × 10−6 [Fig. S3(b)]69, which is substantially lower than what was measured for the G-βO4′-truncG dimer and DPPC. The uptake of the G-βO4′-truncG dimer has a minimal effect on the fluidity and viscoelastic properties of a bilayer that is already in the fluid phase.

QCM-D provides a bulk description of the mass uptake of lignin dimers adsorbed in the lipid bilayers as well as their effect on the fluidity of the bilayer (manifested by changes in dissipation). MD simulations provide complementary information on how the lignin molecules are distributed in the bilayer and how they orient themselves relative to the bilayer normal. As detailed below, we computed partial density profiles for the three lignin dimers and lipid chemical groups at several dimer concentrations. We also calculated the PMF curves that show the distribution of dimers inside the bilayer but also give an indication of the strength of the association of the dimers with the bilayer relative to the bulk solution. We analyzed the ordering of the lipid tails, bilayer thickness and area per lipid, and bilayer area compressibility in the presence of lignin dimers, all of which are related to the dimer concentration dependence of the bilayer fluidity.

1. Partial density profiles

Partial density profiles are a straightforward way of depicting how different atom groups in the lignin dimers locate within the lipid bilayer and allow the determination of the average orientations of the dimers in the bilayer. Partial density profiles for lignin dimer and lipid atom groups with a mole fraction of dimer Xdimer = 0.289 are shown in Fig. 5. The profiles for other concentrations are presented in supplementary material (Fig. S4)69. The entire G-βO4′-G dimer tended to sit on the bilayer surface since the peaks in the density distributions for the center of mass (COM), hydroxypropenyl hydroxyl (OH-T), and phenol ring (OH-B) were all well outside the peak for the lipid ester carbons. For G-βO4′-truncG and benzG-βO4′-G, the unhydroxylated aromatic terminal groups can easily access the interior of the bilayer due to their hydrophobicity. Hydroxylated terminal groups, on the other hand, point more toward the water phase and the most preferable positions are closer to the bilayer surface, near the location of the lipid ester carbon atoms. benzG-βO4′-G is significantly larger than the other two dimers, which explains the larger width for its dimer density distribution. Due to the increased size of the hydrophobic region on the end of benzG-βO4′-G with the added benzene ring, benzG-βO4′-G groups also showed much more penetration into the center of the lipid bilayer than G-βO4′-truncG groups. We note that in the G-βO4′-truncG and benzG-βO4′-G systems, the lipid ester carbons sampled distances much smaller than in a pure lipid bilayer, indicating that lipid flip-flop was probably more likely, and the bilayers were less stable.

FIG. 5.

Partial density profiles of lignin and lipid groups as a function of distance (dL-B) from the lipid bilayer center. (a) G-βO4′-G, (b) G-βO4′-truncG, and (c) benzG-βO4′-G. The mole fraction of dimers is Xdimer = 0.289 in all cases. Ester-C refers to the ester carbons in the lipids located near the boundary between the hydrophobic and the hydrophilic bilayer regions. Dimer refers to the whole lignin dimer molecule. Benzene-T and Benzene-B are terminal benzene rings in the lignin dimers. OH-T stands for the hydroxyl group in the hydroxypropenyl group. OH-B is the phenol hydroxyl group. Benzene and hydroxyl groups are circled in the inset structures with the same color as the corresponding density profile curves. (d) Average distance corresponding to the whole dimer minus the average distance corresponding to the lipid ester carbons as a function of dimer concentration.

FIG. 5.

Partial density profiles of lignin and lipid groups as a function of distance (dL-B) from the lipid bilayer center. (a) G-βO4′-G, (b) G-βO4′-truncG, and (c) benzG-βO4′-G. The mole fraction of dimers is Xdimer = 0.289 in all cases. Ester-C refers to the ester carbons in the lipids located near the boundary between the hydrophobic and the hydrophilic bilayer regions. Dimer refers to the whole lignin dimer molecule. Benzene-T and Benzene-B are terminal benzene rings in the lignin dimers. OH-T stands for the hydroxyl group in the hydroxypropenyl group. OH-B is the phenol hydroxyl group. Benzene and hydroxyl groups are circled in the inset structures with the same color as the corresponding density profile curves. (d) Average distance corresponding to the whole dimer minus the average distance corresponding to the lipid ester carbons as a function of dimer concentration.

Close modal

Figure 5(d) shows the differences between the average distances between the dimer and the ester carbon [each determined from density profiles as in Figs. 5(a)5(c)], dL-C, as a function of dimer concentration. This difference dL-C does not change significantly for benzG-βO4′-G with increasing dimer concentration up to Xdimer = 0.289, which indicates that the interior of the bilayer is not yet saturated with dimers. For G-βO4′-truncG, dL-C begins to increase above a critical concentration, indicating a saturation of the bilayer between Xdimer = 0.289 and 0.385. The value of dL-C for G-βO4′-G increases monotonically with concentration for all concentrations considered, indicating that the dimers are stacking on the bilayer surface with increasing concentration instead of partitioning into the bilayer interior; therefore, the bilayer interior is saturated with G-βO4′-G at a concentration lower than Xdimer = 0.086.

2. Potential of mean force

Figure 6 shows the PMF curves for all three dimers at Xdimer = 0.289. A comparison of these PMFs with the low-concentration PMFs for single dimers in 128 lipid bilayers from our previous work32 shows that the PMF minima became shallower for higher concentrations of G-βO4′-truncG and benzG-βO4′-G, but the minimum was deeper for G-βO4′-G at a higher concentration. At Xdimer = 0.289, there is a significant amount of G-βO4′-G on the bilayer surface, so the deeper PMF minimum relative to a lower concentration was probably due to an affinity of G-βO4′-G with itself and not due to an increased affinity of G-βO4′-G with DPPC. The barrier for flipping from one leaflet to the other is obtained from the difference between the PMF minima and the values at the center of the bilayer. These barriers dropped significantly (more than 50%) for both XG-βO4′-truncG = 0.289 and XbenzG-βO4′-G = 0.289 compared with the single dimer case. Based on the values of these barriers, it seemed flipping might occur spontaneously during unbiased simulations. However, flipping was not observed for Xdimer = 0.289 or for simulations with XG-βO4′-truncG = 0.385 and XG-βO4′-truncG = 0.458. However, using just the dimer center of mass to calculate the energy of crossing the bilayer likely leads to an underestimation of the barrier for the flip-flop, especially for benzG-βO4′-G. This is because the hydrophilic hydroxyl groups are the most difficult to get across the bilayer, but the center of mass can cross the bilayer center while the hydroxyl groups are still in the original leaflet. The barrier based on the benzG-βO4′-G center of mass was only 3kBT. We did an extra PMF calculation (Fig. S9)69 of XbenzG-βO4-G= 0.289 based on the distance from the phenol hydroxyl group to the bilayer center. The PMF minimum was located much closer to the water phase and the barrier for flipping was about two times larger than the estimate based on the COM. To obtain an accurate flipping barrier for these relatively large molecules would probably require two or more coordinates to adequately sample the orientations of the dimers as they cross the bilayer. However, we can say that the mean time for flipping leaflets was significantly longer than our unbiased simulation times.

FIG. 6.

PMF profiles for the lignin dimer center of mass to bilayer center of mass distance along the bilayer normal direction (dL-B). The mole fractions of dimers relative to lipids are Xdimer = 0.289 in all cases. The circular markers show the lipid ester carbon mean positions for reference.

FIG. 6.

PMF profiles for the lignin dimer center of mass to bilayer center of mass distance along the bilayer normal direction (dL-B). The mole fractions of dimers relative to lipids are Xdimer = 0.289 in all cases. The circular markers show the lipid ester carbon mean positions for reference.

Close modal

The QCM-D frequency and dissipation shifts as functions of time during the buffer rinse step (Fig. 4) indicate that G-βO4′-G is removed from the supported bilayer more slowly than G-βO4′-truncG and benzG-βO4′-G. The shifts did not even reach steady values after an hour for G-βO4′-G, while they plateaued at relatively steady values for G-βO4′-truncG and benzG-βO4′-G much faster. This is consistent with the simulated PMF curves. Although G-βO4′-G did not associate much with the bilayer interior, it actually had a stronger affinity for the bilayer headgroups than the other dimers due to hydrogen bonding and possibly self-association at the bilayer surface.68 This is shown by the more negative minimum in the PMF curve for G-βO4′-G of about −23 kBT compared with about −15 kBT for the other two dimers. The mean time for a dimer to move from the bilayer into bulk solution increases approximately exponentially as the PMF minimum value decreases, which explains why it was difficult to remove G-βO4′-G from the bilayer surface.

The final mass ratios of lignin dimers relative to pure DPPC bilayers obtained from QCM-D are given in Table I, and these can be compared with the density profile results (Fig. 5) and PMF curves (Fig. 6) from simulations. The mass ratios from QCM-D for the G-βO4′-truncG and benzG-βO4′-G systems are 1.6 and 2.5 times higher than for G-βO4′-G, respectively. Direct comparisons of mass ratios between MD and QCM-D are inexact because the QCM-D results may be kinetically limited, and we do not know the mass of water in the QCM-D-supported lipid bilayers. However, comparisons of the mass ratios for different dimers are valid since the bilayer mass cancels out. The dimer:lipid ratios at saturation from MD [Fig. 5(d), discussed above] are Xdimer < 0.086 (mass ratio < 0.048) for G-βO4′-G and 0.289 < Xdimer < 0.385 (0.177 < mass ratio < 0.273) for G-βO4′-truncG. The increase by a factor of 3.7 of the mass ratio of G-βO4′-truncG to the mass ratio of G-βO4′-G is consistent with the dimer uptake ratio of 1.6 from QCM-D, considering that the experiments may not have reached full saturation of the lipid bilayers. Also, consistent with the ratios of dimer uptake >1 observed by QCM-D, the density profiles and PMF curves indicate that G-βO4′-truncG and benzG-βO4′-G were found deeper in the bilayer than G-βO4′-G.

3. Deuterium order parameter for lipid tail atoms

The deuterium order parameter is a measure of order in the lipid tails defined to be comparable with the 2H NMR measurements of lipid bilayers.32 As lignin dimer is added, we expected this parameter to decrease due to the increased amount of lignin in the bilayer creating disorder. The time-averaged order parameters were calculated for the three dimers at Xdimer = 0.0, 0.086, 0.158, and 0.289 and the results for one carbon atom in the sn1 tail are given in Fig. 7 [more details given in Figs. S6–S8 (Ref. 69)]. Pure lipids with no dimers or a low concentration of dimers had more ordered lipid acyl chains. There was a trend of decreasing order parameter with an increasing concentration of G-βO4′-truncG and benzG-βO4′-G. The order parameter for G-βO4′-G decreases initially with increasing concentration, but the order parameter for Xdimer = 0.289 increased relative to Xdimer = 0.158 back to a value nearly as high as pure DPPC. This behavior is likely related to the fact that G-βO4′-G already saturated the interior of the bilayer by Xdimer = 0.158. Increasing the concentration further led to a significant portion of the bilayer being covered by dimers that presumably restricted the motion of the lipids in the bilayer plane and promoted order. See snapshots in Fig. S5 (Ref. 69) showing an increased coverage of the bilayer by G-βO4′-G at the higher concentration.

FIG. 7.

Deuterium order parameters, |SCD|, for lipid carbon atom 7 (see Fig. 1) as a function of lignin dimer concentration, Xdimer.

FIG. 7.

Deuterium order parameters, |SCD|, for lipid carbon atom 7 (see Fig. 1) as a function of lignin dimer concentration, Xdimer.

Close modal

4. DPPC bilayer thickness

Table II shows the bilayer thickness, defined as the difference in the average positions of the phosphorus atoms in the upper and lower leaflets, as a function of lignin concentration (see Table S269 for data at higher G-βO4′-truncG concentrations). Lipid bilayer thickness decreased and the bilayer area increased with increasing concentrations of benzG-βO4′-G or G-βO4′-truncG. As more dimers inserted themselves into the spaces between the lipid head groups, they expanded the bilayer horizontally. Accordingly, the bilayer contracted in the vertical direction to maintain approximately the same volume. The G-βO4′-G dimer did not follow the same trend. The bilayer thickness increased, going from Xdimer = 0.086 to 0.158, but decreased as even more G-βO4′-G was added. This result is consistent with the deuterium order parameter results where the order parameter decreased initially followed by an increase with increasing Xdimer.

TABLE II.

Bilayer thickness with different concentrations of the dimers. The pure DPPC bilayer thickness is 3.733 ± 0.126 nm.

Xdimer (mole fraction)DPPC bilayer P-P distance (nm)
G-βO4′-GG-βO4′-truncGbenzG-βO4′-G
0.086 3.695 ± 0.109 3.727 ± 0.021 3.693 ± 0.036 
0.158 3.589 ± 0.145 3.606 ± 0.061 3.604 ± 0.017 
0.289 3.699 ± 0.007 3.463 ± 0.068 3.402 ± 0.126 
Xdimer (mole fraction)DPPC bilayer P-P distance (nm)
G-βO4′-GG-βO4′-truncGbenzG-βO4′-G
0.086 3.695 ± 0.109 3.727 ± 0.021 3.693 ± 0.036 
0.158 3.589 ± 0.145 3.606 ± 0.061 3.604 ± 0.017 
0.289 3.699 ± 0.007 3.463 ± 0.068 3.402 ± 0.126 

5. Bilayer area compressibility

Bilayer area compressibilities are shown in Table III and were obtained from Eqs. (2) and (3) using mean bilayer areas and variance in the bilayer area. Note that decreasing KA indicates a less rigid, more fluid bilayer. The value of KA is the largest for pure DPPC. KA decreases with increasing concentrations of G-βO4′-truncG and benzG-βO4′-G, indicating that both dimers penetrate into and fluidize the bilayer to reduce its compressibility. Consistent with the density profile and deuterium order parameter results, the compressibility of the G-βO4′-G system was much lower than that of pure DPPC for Xdimer = 0.158 and then increased back to slightly lower than the value of pure DPPC for Xdimer = 0.289.

TABLE III.

Bilayer area compressibility KA (N/m) as a function of lignin dimer concentration.

SystemXdimerKA (N/m)
Pure DPPC 1.98 ± 0.46 
DPPC + G-βO4′-G 0.086 0.38 ± 0.21 
0.158 0.35 ± 0.15 
0.289 1.21 ± 0.31 
DPPC + G-βO4′-trunc0.086 0.63 ± 0.14 
0.158 0.49 ± 0.20 
0.289 0.46 ± 0.15 
DPPC + benzG-βO4′-G 0.086 0.53 ± 0.10 
0.158 0.68 ± 0.10 
0.289 0.39 ± 0.10 
SystemXdimerKA (N/m)
Pure DPPC 1.98 ± 0.46 
DPPC + G-βO4′-G 0.086 0.38 ± 0.21 
0.158 0.35 ± 0.15 
0.289 1.21 ± 0.31 
DPPC + G-βO4′-trunc0.086 0.63 ± 0.14 
0.158 0.49 ± 0.20 
0.289 0.46 ± 0.15 
DPPC + benzG-βO4′-G 0.086 0.53 ± 0.10 
0.158 0.68 ± 0.10 
0.289 0.39 ± 0.10 

Herein, the interactions of supported DPPC lipid bilayers with concentrated amounts of the three βO4′ linked dimers of coniferyl alcohol and its derivatives (GG lignin dimers) are investigated using results from QCM-D and from molecular dynamics simulations of unsupported DPPC bilayers.

QCM-D provides a bulk description of the effect of the increased amounts of dimers on the bilayer and revealed that a truncated GG dimer with the hydroxypropenyl tail removed, G-βO4′-truncG, and GG dimers with an added benzyl group, benzG-βO4′-G, penetrated more deeply into the bilayer interior. This conclusion is based on the greater increases in the mass of dimer incorporated and the dissipation (viscoelasticity) of the thin film on the gold support for these two variants. On the contrary, the guaiacylglycerol guaiacol ester with a hydroxypropenyl (HOC3H4-) tail (G-βO4′-G) demonstrated relatively weak affinity toward the interior of the lipid bilayer as inferred from smaller shifts in frequency (mass) and dissipation (viscoelasticity) measured before and after dimer exposure.

Molecular dynamics simulations of lignin GG dimers with DPPC lipid bilayers qualitatively explain the QCM-D results, indicating that G-βO4′-G does not accumulate to a significant extent in the bilayer interior, while the more hydrophobic G-βO4′-truncG and benzG-βO4′-G are taken up to a significant extent. Density profiles for different concentrations of dimers and the PMF results for one concentration indicate that G-βO4′-truncG and benzG-βO4′-G were found deeper in the bilayer than G-βO4′-G. Furthermore, the results for the ordering of the lipid tails, bilayer thickness, and area per lipid, and bilayer compressibility showed that by promoting disorder in the lipid tails, G-βO4′-truncG and benzG-βO4′-G increase the bilayer fluidity more than G-βO4′-G, which is consistent with the QCM-D dissipation results and those of our previous study on the gel–fluid transition temperature at low concentrations.32 

The results from this study enhance our understanding of pharmacological and toxicological effects of lignin derivatives as well as properties required for them to be developed as potential antigrowth and therapeutic agents by providing details of their activity for a model cell membrane. These data suggest that the hydroxypropenyl tail may present a barrier to the penetration of lignin oligomers into the cell and, thus, may affect their pharmacology.

The authors gratefully acknowledge the support from the U.S. National Science Foundation under EPSCoR RII Track-2 Program (Project No. OIA-1632854). The computer resources were provided by Louisiana Optical Network Infrastructure (LONI) and High Performance Computing (HPC) at LSU. This work was performed in part at the Center for Nanoscale Science and Engineering (CeNSE), which belongs to the National Science Foundation NNCI Kentucky Multiscale Manufacturing and Nano Integration Node, supported by Grant No. ECCS-1542174.

The data that support the findings of this study are available within the article and its supplementary material.

1.
B. M.
Upton
and
A. M.
Kasko
,
Chem. Rev.
116
,
2275
(
2016
).
2.
D.
Kai
,
M. J.
Tan
,
P. L.
Chee
,
Y. K.
Chua
,
Y. L.
Yap
, and
X. J.
Loh
,
Green Chem.
18
,
1175
(
2016
).
3.
M.
Azadfar
,
A. H.
Gao
,
M. V.
Bule
, and
S.
Chen
,
Int. J. Biol. Macromol.
75
,
58
(
2015
).
4.
C. A.
Lekelefac
,
N.
Busse
,
M.
Herrenbauer
, and
P.
Czermak
,
Int. J. Photoenergy
2015
,
1
18
.
5.
J. A.
Belgodere
 et al,
ACS Appl. Bio Mater.
2
,
3562
(
2019
).
6.
J.
Chen
,
A. E.
Kazzaz
,
N.
AlipoorMazandarani
,
Z. H.
Feizi
, and
P.
Fatehi
,
Molecules
23
,
868
(
2018
).
7.
Z.
Sun
,
B.
Fridrich
,
A.
de Santi
,
S.
Elangovan
, and
K.
Barta
,
Chem. Rev.
118
,
614
(
2018
).
8.
T. D. H.
Bugg
,
M.
Ahmad
,
E. M.
Hardiman
, and
R.
Rahmanpour
,
Nat. Prod. Rep.
28
,
1883
(
2011
).
9.
Y. S.
Choi
,
R.
Singh
,
J.
Zhang
,
G.
Balasubramanian
,
M. R.
Sturgeon
,
R.
Katahira
,
G.
Chupka
,
G. T.
Beckham
, and
B. H.
Shanks
,
Green Chem.
18
,
1762
(
2016
).
10.
11.
P.
Figueiredo
,
K.
Lintinen
,
J. T.
Hirvonen
,
M. A.
Kostiainen
, and
H. A.
Santos
,
Prog. Mater. Sci.
93
,
233
(
2018
).
12.
A. E.
Kazzaz
,
Z. H.
Feizi
, and
P.
Fatehi
,
Green Chem.
21
,
5714
(
2019
).
13.
S. O.
Asare
,
P.
Kamali
,
F.
Huang
, and
B. C.
Lynn
,
Energy Fuel
32
,
5990
(
2018
).
14.
T.
Inoue
,
K.
Miyakawa
, and
R.
Shimozawa
,
Chem. Phys. Lipids
42
,
261
(
1986
).
15.
S.
Fujisawa
,
Y.
Kadoma
, and
E.
Masuhara
,
J. Biomed. Mater. Res.
21
,
89
(
1987
).
16.
M. K.
Jain
and
N. M.
Wu
,
J. Membr. Biol.
34
,
157
(
1977
).
17.
R. R. J.
Arroo
,
A. W.
Alfermann
,
M.
Medarde
,
M.
Petersen
,
N.
Pras
, and
J. G.
Woolley
,
Phytochem. Rev.
1
,
27
(
2002
).
18.
R.
Kiyama
,
Trends Food Sci. Technol.
54
,
186
(
2016
).
19.
J. M.
Landete
,
Food Res. Int.
46
,
410
(
2012
).
20.
J.
Zhang
,
J.
Chen
,
Z.
Liang
, and
C.
Zhao
,
Chem. Biodivers.
11
,
1
(
2014
).
21.
T.
Joshi
,
Z. X.
Voo
,
B.
Graham
,
L.
Spiccia
, and
L. L.
Martin
,
Biochim. Biophys. Acta Biomembr.
1848
,
385
(
2015
).
22.
S. M.
Cragg
 et al,
Curr. Opin. Chem. Biol.
29
,
108
(
2015
).
23.
H.
Lange
,
S.
Decina
, and
C.
Crestini
,
Eur. Polym. J.
49
,
1151
(
2013
).
24.
C. M.
Bailey
,
E.
Kamaloo
,
K. L.
Waterman
,
K. F.
Wang
,
R.
Nagarajan
, and
T. A.
Camesano
,
Biophys. Chem.
203-204
,
51
(
2015
).
25.
F.
Neville
,
M.
Cahuzac
,
O.
Konovalov
,
Y.
Ishitsuka
,
K. Y. C.
Lee
,
I.
Kuzmenko
,
G. M.
Kale
, and
D.
Gidalevitz
,
Biophys. J.
90
,
1275
(
2006
).
26.
A.
Vahedi
,
P.
Bigdelou
, and
A. M.
Farnoud
,
Sci. Rep.
10
,
15111
(
2020
).
27.
M. H.
Nouri-Sorkhabi
,
L. C.
Wright
,
D. R.
Sullivan
, and
P. W.
Kuchel
,
Lipids
31
,
765
(
1996
).
28.
S. L.
Keller
,
W. H.
Pitcher
 III
,
W. H.
Huestis
, and
H. M.
McConnell
,
Phys. Rev. Lett.
81
,
5019
(
1998
).
29.
E. J. A.
Veldhuizen
and
H. P.
Haagsman
,
Biochim. Biophys. Acta Biomembr.
1467
,
255
(
2000
).
30.
F.
Zhao
,
J. P.
Holmberg
,
Z.
Abbas
,
R.
Frost
,
T.
Sirkka
,
B.
Kasemo
,
M.
Hassellöv
, and
S.
Svedhem
,
RSC Adv.
6
,
91102
(
2016
).
31.
P.
Bigdelou
,
A.
Vahedi
,
E.
Kiosidou
, and
A. M.
Farnoud
,
Biointerphases
15
,
041001
(
2020
).
32.
X.
Tong
,
M.
Moradipour
,
B.
Novak
,
P.
Kamali
,
S. O.
Asare
,
B. L.
Knutson
,
S. E.
Rankin
,
B. C.
Lynn
, and
D.
Moldovan
,
J. Phys. Chem. B
123
,
8247
(
2019
).
33.
A. M.
Farnoud
and
S.
Nazemidashtarjandi
,
Environ. Sci.: Nano
6
,
13
(
2019
).
34.
K.
Kannisto
,
L.
Murtomäki
, and
T.
Viitala
,
Colloids Surf. B
86
,
298
(
2011
).
35.
A.
Wargenau
,
S.
Schulz
,
A.
Hakimzadeh
, and
N.
Tufenkji
,
J. Anal. Chem.
90
,
11174
(
2018
).
36.
C. E.
Byrne
,
C. E.
Astete
,
M.
Vaithiyanathan
,
A. T.
Melvin
,
M.
Moradipour
,
S. E.
Rankin
,
B. L.
Knutson
,
C. M.
Sabliov
, and
E. C.
Martin
,
Nanomedicine
15
,
981
(
2020
).
37.
M.
Moradipour
,
E. K.
Chase
,
M. A.
Khan
,
S. O.
Asare
,
B. C.
Lynn
,
S. E.
Rankin
, and
B. L.
Knutson
,
Colloids Surf. B
191
,
111028
(
2020
).
38.
J.
Lin
,
B.
Novak
, and
D.
Moldovan
,
J. Phys. Chem. B
116
,
1299
(
2012
).
39.
Y.
Lyu
,
N.
Xiang
,
J.
Mondal
,
X.
Zhu
, and
G.
Narsimhan
,
J. Phys. Chem. B
122
,
2341
(
2018
).
40.
J. V.
Vermaas
,
R. A.
Dixon
,
F.
Chen
,
S. D.
Mansfield
,
W.
Boerjan
,
J.
Ralph
,
M. F.
Crowley
, and
G. T.
Beckham
,
Proc. Natl. Acad. Sci. U.S.A.
116
,
23117
(
2019
).
41.
B.
Lindner
,
L.
Petridis
,
R.
Schulz
, and
J. C.
Smith
,
Biomacromolecules
14
,
3390
(
2013
).
42.
Y.
Zhang
,
H.
He
,
Y.
Liu
,
Y.
Wang
,
F.
Huo
,
M.
Fan
,
H.
Adidharma
,
X.
Li
, and
S.
Zhang
,
Green Chem.
21
,
9
(
2019
).
43.
S.
Besombes
,
D.
Robert
,
J.-P.
Utille
,
F. R.
Taravel
, and
K.
Mazeau
,
J. Agric. Food Chem.
51
,
34
(
2003
).
44.
S. R.
Tabaei
,
J.-H.
Choi
,
G.
Haw Zan
,
V. P.
Zhdanov
, and
N.-J.
Cho
,
Langmuir
30
,
10363
(
2014
).
45.
A. R.
Patel
and
C. W.
Frank
,
Langmuir
22
,
7587
(
2006
).
46.
N.-J.
Cho
,
C. W.
Frank
,
B.
Kasemo
, and
F.
Höök
,
Nat. Protocols
5
,
1096
(
2010
).
47.
48.
M.
Andersson
,
A.
Sellborn
,
C.
Fant
,
C.
Gretzer
, and
H.
Elwing
,
J. Biomater. Sci. Polym. Ed.
13
,
907
(
2002
).
49.
M. J.
Santos-Martinez
,
I.
Inkielewicz-Stepniak
,
C.
Medina
,
K.
Rahme
,
D.
D’Arcy
,
D.
Fox
,
J.
Holmes
,
H.
Zhang
, and
M. W.
Radomski
,
Int. J. Nanomed.
7
,
243
(
2012
).
50.
T. K.
Lind
and
M.
Cardenas
,
Biointerphases
11
,
020801
(
2016
).
51.
N.
Shpigel
,
M. D.
Levi
,
S.
Sigalov
,
L.
Daikhin
, and
D.
Aurbach
,
Acc. Chem. Res.
51
,
69
(
2018
).
52.
K.
Sadman
,
C. G.
Wiener
,
R. A.
Weiss
,
C. C.
White
,
K. R.
Shull
, and
B. D.
Vogt
,
Anal. Chem.
90
,
4079
(
2018
).
53.
N.
Schmid
,
A. P.
Eichenberger
,
A.
Choutko
,
S.
Riniker
,
M.
Winger
,
A. E.
Mark
, and
W. F.
van Gunsteren
,
Eur. Biophys. J.
40
,
843
(
2011
).
54.
55.
J.
Kästner
,
Wiley Interdiscip. Rev.: Comput. Mol. Sci.
1
,
932
(
2011
).
56.
M.
Souaille
and
B.
Roux
,
Comput. Phys. Commun.
135
,
40
(
2001
).
57.
S. E.
Feller
and
R. W.
Pastor
,
J. Chem. Phys.
111
,
1281
(
1999
).
58.
R. P.
Richter
,
R.
Bérat
, and
A. R.
Brisson
,
Langmuir
22
,
3497
(
2006
).
59.
A. A.
Adib
,
S.
Nazemidashtarjandi
,
A.
Kelly
,
A.
Kruse
,
K.
Cimatu
,
A. E.
David
, and
A. M.
Farnoud
,
Environ. Sci. Nano
5
,
289
(
2018
).
60.
C. A.
Keller
and
B.
Kasemo
,
Biophys. J.
75
,
1397
(
1998
).
61.
R.
Richter
,
A.
Mukhopadhyay
, and
A.
Brisson
,
Biophys. J.
85
,
3035
(
2003
).
62.
T. K.
Lind
, M. Cárdenas, and
H. P.
Wacklin
,
Langmuir
30
,
7259
(
2014
).
63.
T. J.
Zwang
,
W. R.
Fletcher
,
T. J.
Lane
, and
M. S.
Johal
,
Langmuir
26
,
4598
(
2010
).
64.
J. T.
Marquês
,
A. S.
Viana
, and
R. F. M.
de Almeida
,
Langmuir
30
,
12627
(
2014
).
65.
H.-H.
Shen
,
P. G.
Hartley
,
M.
James
,
A.
Nelson
,
H.
Defendi
, and
K. M.
McLean
,
Soft Matter
7
,
8041
(
2011
).
66.
E.
Tellechea
,
D.
Johannsmann
,
N. F.
Steinmetz
,
R. P.
Richter
, and
I.
Reviakine
,
Langmuir
25
,
5177
(
2009
).
67.
E.
Boija
and
G.
Johansson
,
Biochim. Biophys. Acta Biomembr.
1758
,
620
(
2006
).
68.
P.
Tammela
,
L.
Laitinen
,
A.
Galkin
,
T.
Wennberg
,
R.
Heczko
,
H.
Vuorela
,
J. P.
Slotte
, and
P.
Vuorela
,
Arch. Biochem. Biophys.
425
,
193
(
2004
).
69.
See supplementary material at https://www.scitation.org/doi/suppl/10.1116/6.0001029 for 1H NMR and mass spectrometry (HR-MS) of the G-βO4′-G and benzG-βO4′-G dimers structures, QCM-D results for the interaction of isopropanol with supported DPPC lipid bilayers, solvent-assisted formation of DOPC bilayers and G-bO4′-truncG dimer interaction with DOPC lipid bilayer. Supplementary material for molecular dynamics includes error estimation for partial density profiles for lignin and lipid groups as a function of dimer distance from the lipid bilayer center, bilayer thickness with high concentrations of G-bO4′-truncG dimer, top view of DPPC lipid bilayer with G-bO4′-G dimer, deuterium order parameters for carbons in the lipid tails for G-bO4′-G, benzG-bO4′-G, and G-bO4′-truncG systems, and PMF profile as a function of the distance (dL-B) from the bilayer center to the terminal –OH group of a benzG-βO4′-G.

Supplementary Material