Ether- versus Ester-Linked Phospholipid Bilayers Containing either Linear or Branched Apolar Chains

Sample preparation

All lipids were purchased from Avanti Polar Lipids (Alabaster, AL) and used without further purification. We obtained 2-dimethylamino-6-lauroylnaphthalene (Laurdan) and 1-(4-trimethylammoniumphenyl)-6-phenyl-1,3,5-hexatriene p-toluenesulfonate (TMA-DPH) from Molecular Probes (Alabaster, AL). We prepared phospholipid multilamellar vesicles (MLVs) by dissolving each phospholipid in chloroform/methanol (2:1), mixing them in the appropriate proportions (binary mixtures at 1:1 ratios were typically used), and drying the samples for 2 h under vacuum. The dry lipids were resuspended in a PIPES-EDTA or choline chloride buffer (depending on the type of measurement to be carried out (calorimetry or spectrophotometry); see details below) at 65°C, well above all lipid transition temperatures, for ∼20 min (gentle mixing was applied to favor vesicle formation). The samples were then allowed to cool down to room temperature (or ∼10°C for differential scanning calorimetry (DSC)).

DSC

We observed the phase transitions of phospholipid bilayer vesicles under 24 psi pressure using a VP-DSC high-sensitivity differential scanning calorimeter (MicroCal, Northhampton, MA). Measurements were carried out in 20 mM PIPES, 150 mM NaCl, 1 mM EDTA, pH 7.4 buffer. Vesicle solutions were prepared at 1.0 or 2.0 mmol L⁻¹. The heating rate was 45°C/h. Thermograms were recorded after sample equilibration at the starting temperature. We performed buffer subtraction, baseline correction, and integration of the calorimetric endotherms using the Microcal Origin software.

Fluorescence spectroscopy

Fluorescence spectroscopy was performed in a PTI-MP1 spectrofluorometer (Photon Technology International) equipped with a thermoregulated cell holder for measurements at 20°C and 45°C, and with polarizers for the anisotropy of TMA-DPH fluorescence.

The Laurdan generalized polarization excitation (GP_EX) parameter was calculated according to

where I₄₄₀ and I₄₉₀ are the emission intensities obtained at 440 and 490 nm, respectively, with excitation at 360 nm. The final probe/lipid molar ratio was 1:500.

For anisotropy of TMA-DPH fluorescence measurements, the instrument software computes anisotropies from the averages of ten measurements for each experimental point, correcting for the G factor. TMA-DPH fluorescence was excited at 360 nm and emission was recorded at 430 nm. The final probe/lipid molar ratio was 1:250. To avoid light scattering and inner filter effects, fluorescence anisotropy was measured on increasingly diluted samples. Only when anisotropy values remained constant with further dilution were they recorded.

Supported planar bilayer preparation

Supported planar bilayers (SPBs) were prepared on high-quality, V-2 grade, scratch-free mica substrates (Asheville-Schoonmaker Mica, Newport News, VA) that were previously attached to round 24 mm glass coverslips with the use of a two-component optical epoxy resin (EPO-TEK 301-2FL; Epoxy Technology, Billerica, MA). SPBs were prepared according to the vesicle adsorption method (18). For this purpose, multilamellar vesicles (MLVs) were initially prepared by mixing the appropriate amounts of synthetic pure lipids in chloroform/methanol (2:1, v/v) solutions. Samples were then dried by evaporating the solvent under a stream of nitrogen and placing them into high vacuum for 2 h. The samples were then hydrated in assay buffer and highly vortexed at a temperature above that of the sample lipids’ highest phase transition. After the lipids became completely detached from the test-tube bottom, the formed MLVs were introduced into a FB-15049 (Fisher Scientific, Waltham, MA) bath sonicator and kept at 60°C for 1 h. In this way, a proportion of small unilamellar vesicles (SUVs) were generated. Then, 120 μl of assay buffer containing 3 mM CaCl₂ was added onto a previously prepared 1.2 cm² freshly cleaved mica substrate mounted onto a BioCell coverslip-based liquid cell for atomic force microscopy (AFM) measurements (JPK Instruments, Berlin, Germany). Then, 60 μl of the sonicated vesicles was added on top of the mica. Final lipid concentration was 150 μM. Vesicles were left to adsorb and extend for 30 min maintaining the sample temperature at 70°C. The samples were left to equilibrate at room temperature for 30 min, and then the nonadsorbed vesicles were discarded by washing the samples 10 times with assay buffer in the absence of CaCl_2. A small amount of buffer was always left on top of the substrate to keep the SPBs hydrated at all times. The BioCell was set to 23°C and the planar bilayers were left to equilibrate for 1 h prior to AFM measurements.

AFM imaging

Planar bilayer topography was performed under contact-mode AFM scanning (constant vertical deflection) in a NanoWizard II AFM (JPK Instruments, Berlin, Germany). For proper measurements, the AFM was coupled to a Leica microscope and mounted onto a Halcyonics Micro 40 antivibration table (Halcyonics, Menlo Park, CA) and inside an acoustic enclosure (JPK Instruments). The Biocell liquid sample holder (JPK Instruments) was used to control the assay temperature at 23°C. V-shaped MLCT Si₃N₄ cantilevers (Veeco, Plainview, NY) with nominal spring constants of 0.1 N/m were used for bilayer imaging, always keeping the minimum possible force. Then, 512 × 512 pixel resolution images were collected at a scanning rate of 1 Hz and line fitted using the JPK Data Processing software.

Force spectroscopy

Before imaging, V-shaped MLCT Si₃N₄ cantilevers (Veeco, Plainview, NY) with nominal spring constants of 0.1 N/m were individually calibrated in a lipid-free mica substrate in assay buffer using the thermal-noise method. After proper AFM imaging to check for correct lipid extension of the SPBs, force spectroscopy was performed at a speed of 1 μm/s in not less than 500 × 500 nm bilayer areas in the form of 10 × 10 or 15 × 15 grids. Force curves were batch analyzed with JPK Data Processing software and bilayer breakthrough events were determined for each of the indentation curves as reproducible force steps or jumps within the extended traces. Histograms were generated from at least three independent sample preparations with at least three independently calibrated cantilevers (n = 647–2767).

Preparation of GUVs

GUVs were prepared at 65°C according to the electroformation method (19) with small modifications. Pure lipids and mixtures were suspended in chloroform/methanol (2:1, v/v) and GUVs were prepared using 10 mM Tris, pH 7, as buffer. Stock solutions of lipids (0.2 mg/ml total lipid containing 0.2% DiI or Rhodamine-DOPE) were transferred to Pt wires and connected to an electric wave generator under AC field conditions (frequency, 500 Hz; amplitude, 5.3 Vp-p) for 1.5 h at 65°C. The GUVs were then observed on an inverted confocal fluorescence microscope at 20°C.

Permeability measurements

All permeability measurements were performed at 20°C by spectrophotometry. MLVs (prepared at 2.5 mM in 50 mM choline chloride) were diluted (1:50 v/v) in a nonelectrolyte solution (100 mM) of glycol (62.07 gmol⁻¹), glycerol (92.09 gmol⁻¹), erythritol (122.12 gmol⁻¹), or glycine betaine (117.15 gmol⁻¹). The subsequent swelling of the vesicles and permeation of nonelectrolytes through the membrane caused a decrease in turbidity (A₄₅₀) that was measured as a function of time (20). The measurements are expressed as a normalized relative volume decrease, reflecting the entry of water and solutes into the vesicles and the consequent increase in translucency of the samples.

Thermotropic phase transitions

Fig. 1 shows representative DSC thermograms of the four phospholipids under study and their binary mixtures. The thermogram for pure DPPC, in agreement with previous reports, shows a small endotherm around 35°C (the pretransition temperature T_p) and a large endotherm at ∼41.5°C (T_m). The first transition reflects a reorganization in the order of lipids in the membrane, in which the gel phase (L_β′) changes to a pseudo-melted rippled phase (P_β′), whereas beyond the T_m the liquid crystalline (L_α) phase is reached (21). The pure DHPC thermogram shows a very similar profile, with only slight differences. In the latter case, the pretransition was attributed to the change from an interdigitated gel phase (L_βI) to a P_β′ one (22). In our experimental setting, this transition takes place at ∼26°C, whereas T_m occurs at 43°C (Fig. 1).

When DPhoPC or DPhPC were added to DPPC at equimolar ratios, the pretransition peak disappeared and the main transition peak became wider and asymmetric, and shifted toward lower temperatures (26–27°C). When the former lipids were mixed at the same equimolar ratios but with DHPC instead of DPPC, a similar effect was observed, with asymmetric main transitions at ∼29°C. In contrast, when DPhoPC and DPhPC systems were employed, either pure or in an equimolecular mixture (Fig. 1), no detectable gel-to-liquid crystalline phase transition was registered over a broad temperature range (10–100°C).

Domain formation in mixed bilayers

We used the above-described lipid mixtures to prepare GUVs and study their possible segregation into well-defined domains by confocal fluorescence microscopy. Representative examples are shown in Fig. 2, corresponding to N-Rh-DOPE-labeled images taken at 20°C (∼1 h after preparation, to allow sample cooling from formation temperature, T = 65°C). Independently of GUV size, which was somewhat variable (ranging from d = 5–10 μm up to 60 μm), fluid/solid phase coexistence was observed only in those mixtures in which asymmetric DSC thermograms were seen. Nevertheless, some interesting differences were detected in the process of phase separation. In binary mixtures containing DPPC (Fig. 2, A and B), lateral separation of dark and bright domains was clearly observed. However, when DHPC (ether linked) was used, finely reticulated domains appeared in the first minutes of observation (Fig. 2, C and D). These DHPC-specific reticulated domains were transient and disappeared soon (∼20 min) after formation, allowing the formation of clearly separated dark and bright areas (Fig. 2, E and F). Since the large headgroup of rhodamine-DOPE hinders insertion into gel phases, it is assumed that these fluorescent molecules segregate into fluid domains (23). Hence, dark zones are most probably DPPC or DHPC rich. No phase separation was observed in mixtures that did not contain DPPC or DHPC (i.e., those that did not exhibit an endothermic signal in Fig. 1).

Fluorescent probes at the bilayer interfacial region

Further information on the properties of ether- versus ester-containing bilayers may be obtained with the use of fluorescent probes that are known to partition preferentially at the bilayer lipid/water interface. In particular, TMA-DPH and Laurdan have been used in bilayers formed by each of the four lipids under study. The values of the TMA-DPH anisotropies and Laurdan GPs obtained at 20°C and 45°C are shown in Fig. 3. The fluid-phase states at 20°C, according to the data in Figs. 1 and 2, are as follows: pure linear lipids, gel; pure branched lipids, fluid; linear/branched mixtures, gel/fluid coexistence; branched/branched mixture, fluid. At 45°C, all samples are in the fluid state. The measurements shown in Fig. 3 were made after a 5 min thermal equilibration, but preliminary experiments had shown no changes over a period of 25 min. TMA-DPH anisotropy reports on the molecular order at the interface (Fig. 3, A and C). As expected, the order decreased when going from the low to the high temperature. It can also be seen that even if the differences are small, there is a higher degree of order for the ether than for the ester lipids, particularly at 45°C. For the binary mixtures (Fig. 3), these differences are blurred, and in general the anisotropy (molecular order) of the mixtures is lower than that of the pure components. Laurdan fluorescence relaxes through water dipoles. A low hydration in the region will mean poor relaxation and high GP values. When Laurdan GP is measured in our systems (Fig. 3, B and D), the results are consistent with the ether lipids being more hydrated than the ester lipids at their interfaces, and the branched ones being more hydrated than the linear ones. As expected from the lack of a detectable calorimetric transition for the branched lipids, their corresponding GP values are rather similar at 20°C and 45°C. In general, the binary mixtures (Fig. 3 D) reflect the combined properties of the components. The branched-chain mixture DPhoPC/DPhPC exhibits similarly low GP values (high hydration) at both temperatures.

AFM on SPBs

The SPBs that formed with each of the four lipids under study were examined by AFM in the force-spectroscopy mode. In this mode, the nanomechanical bilayers’ resistance to piercing by the AFM tip is measured. Typically, the AFM-tip indentation process on planar bilayers provides a force-distance record with a 4–6 nm jump in the extension trace. This jump reflects the exact force at which tip penetration through the bilayers occurs and is known as the breakthrough force (F_b). Representative AFM-tip indentation curves are shown in Fig. 4. DPPC samples (Fig. 4, curve a) gave a single signal, with a jump at ∼16 nN, similar to previously published values (24). However, for the other three lipids, two kinds of force curves were found in each case. Specifically, in a given SPB preparation, different indentation experiments provided either one or the other results (see, e.g., curves b and b′ for DHPC in Fig. 4). The alternative results were highly reproducible within each class. The statistics for the experiments on DPPC and DHPC are shown in Fig. 5. The two groups of results found with DHPC appear clearly separated in the histograms (Fig. 5 B), with F_b values of ∼14 and 22 nN.

Bilayer breakthrough force histograms for branched-chain lipids are shown in Fig. 6. Two populations of F_b values are found for both DPhPC and DPhoPC; however, the forces are much lower for the ether (DPhPC; Fig. 6 B) than for the ester (DPhoPC; Fig. 6 A) lipid bilayers. The origin of this dual behavior is discussed below. The DPhoPC histogram is very similar to the one published by García-Manyes et al. (25) (DFPC in their nomenclature), but our F_b values are lower (by ∼3 nN) than theirs, which we attribute to the different ionic composition of the buffer used. The same authors have shown that certain divalent cations, in particular Mg²⁺, reinforce the nanomechanical resistance of lipid bilayers in SPB preparations.

In a different series of experiments, contact-mode AFM imaging was performed on DHPC SPBs. In contrast to the height-homogeneous extensions that are commonly found for DPPC (24), the DHPC bilayers had a heterogeneous appearance (Fig. 7 A), with two coexisting phases 3.8 and 5.7 nm in height, respectively (see histograms in Fig. 7 E). However, these two phases were not stable at 23°C and the lower one shrank with time, disappearing after ∼30 min (Fig. 7, A, D, and F). This would be in agreement with the transient appearance of a reticulated phase in GUV made of mixtures containing DHPC (Fig. 2, C–F). In the light of studies by Guler et al. (16) and Maruyama et al. (22), it could be proposed that the lower phase corresponds to an interdigitated gel that is gradually transformed into a taller, tilted gel. The height of the tall DHPC phase is very similar to that of gel DPPC at 23°C (24). It appears that DHPC bilayers are not in equilibrium at 23°C, and this could explain the dual F_b values. Note that the TMA-DPH or Laurdan GP values of DHPC (or of DPPC) did not change with time in the first 30 min, as stated above. This is in agreement with changes taking place at the hydrophobic region of the bilayers (interdigitation) rather than those involving the lipid/water interface (e.g., gel-to-fluid phase transitions).

With respect to the dual F_b values found with DPhoPC and DPhPC, no time changes in AFM topography were seen. A possible explanation for this is that the lipid chain branching influences the piercing process so that there are two possible pathways for the tip. Alternatively, the SPB formed by branched-chain lipids could contain domains that are small and/or transient enough to be invisible in the AFM topographic images yet detectable by the force curves.

Assays for permeability to nonelectrolytes

To obtain a more complete picture of the properties of the bilayer membranes composed of these lipid mixtures, we tested the permeability of these membranes to nonelectrolytic compounds of various sizes: ethylene-glycol (e-glycol; 2-C), glycerol (3-C), and erythritol (4-C). In these experiments, the entry of water due to soluble diffusion inside the vesicles results in an increase of liposomal volume, which is measured through changes in turbidity. Because the mass of membrane lipids in each vesicle is constant but the volume is increased, the suspension turbidity decreases. Liposomes behave as ideal osmometers under these conditions (20). The time-dependent decrease in absorbance (A₄₅₀) was monitored and translated into relative volume changes, as shown in Fig. 8 for the most permeable species (e-glycol) in selected experiments at 20°C. At this temperature, DPPC forms a gel phase (L_β′), and consequently no e-glycol flux was observed with the pure lipid. However, when liposomes were heated above the T_m for this lipid (at 45°C), e-glycol permeation was detectable (Fig. 8, A and G). These results are consistent with the water permeability (P_f) reported for DPPC in gel (0.012 × 10⁻³ cm/s) and fluid (0.027 cm/s) phases, respectively (16).

The branched-chain lipids DPhPC and DPhoPC gave rise to bilayers that were clearly permeable to e-glycol (Fig. 8, D and E), and the same was observed for equimolecular DPhPC/DPhoPC mixtures (Fig. 8 F; Table 1). Guler et al. (16) reported that water permeability for fluid-phase DHPC was slightly lower than that for DPPC, consistent with Nagle et al.’s (26) triple-slab theory. We measured e-glycol permeability at 48°C for the four lipids under study. The initial rates of vesicle volume change (i.e., solute + water entrance), τ, were 0.43 (DPPC), 0.34 (DHPC), 0.15 (DPhoPC), and 0.19 (DPhPC) (Table 1). Thus, our results confirm Nagle et al.’s predictions and observations for linear-chain lipids, but do not appear to do so for the branched-chain lipids.

DPhoPC and DPhPC in equimolar mixtures with DPPC show 1.6 and 2.4 times faster permeation rates than each pure lipid, respectively. Taking into account that these mixtures show formation of solid domains in GUVs (Fig. 2), the previous results indicate that permeation could be enhanced at the boundaries with disordered phases, rich in branched lipids. This is confirmed by the observations of mixtures containing DPPC (or DHPC) and a branched lipid at 45°C. In these cases, the permeability rates are similar to those of the linear lipid, DPPC or DHPC, at 45°C, showing that when the lipid mixture is in the fluid phase, lipid composition is less important for permeability. Glycerol and erythritol showed a qualitative behavior similar to that of e-glycol, although their permeation capacity was significantly lower due to their larger molecular masses (see, e.g., Fig. 8 L). In all of these experiments, glycine-betaine (GB), an N-trimethylated amino acid, was used as a control and showed no sign of permeation in any of the cases tried.