Article pubs.acs.org/JPCB
Ethanol-Induced Perturbations to Planar Lipid Bilayer Structures Iwan Setiawan and G. J. Blanchard* Department of Chemistry, Michigan State University, 578 S. Shaw Lane, East Lansing, Michigan 48824, United States ABSTRACT: We report on the formation of planar lipid bilayer structures on mica where the bilayer contains the phosphocholine 1,2-dioleoyl-snphosphatidylcholine (DOPC), cholesterol, sphingomyelin and sulforhodamine-tagged-1,2-dioleoyl-sn-phosphatidylethanolamine (SR-DOPE). Phase separation is seen for the cholesterol domains within the bilayer structure, and exposure of this supported bilayer to controlled concentrations of ethanol reveals organizational changes on both the micrometer- and molecular-length scales. We report steady state fluorescence imaging, fluorescence lifetime imaging, and fluorescence anisotropy decay imaging for these bilayers. These data are complementary to existing information on the interactions of lipid bilayers with ethanol and point to subtle but important changes in the molecular-scale organization of these structures.
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INTRODUCTION Mammalian plasma membranes are uniquely complex structures. In a typical mammalian lipid bilayer structure there can be several hundred distinct molecular species, dominated by phospholipids, sterols, sphingomyelin, and transmembrane proteins.1 These multicomponent systems are known to manifest heterogeneity, with structures termed “lipid rafts” thought to play a key role in the function of plasma membranes.2−7 The plasma membrane plays an essential role in life function at the cellular level, mediating the transport of specific species between intracellular and extracellular environments. A great deal of research effort has gone into understanding the reasons for the existence and characteristic sizes of lipid raft structures. While phase separation is observed readily for model systems containing a limited number of constituents due to the size of the phase segregated domains,8−21 in actual plasma membranes the size of these structures has been found recently to be small, on the order of tens of nanometers.22 We are interested in the use of model plasma membrane systems for the purpose of creating supported biomimetic structures. Uses for such structures include supporting transmembrane proteins in their active form so that they may be used for chemical sensing or for the examination of viral attachment and infection processes in vitro. If this family of structures is to be used for these or other applications, it will be important to address the issue of structural control over model systems comprised of constituents fewer in number than the full complement of a natural plasma membrane. In addition to direct control over the composition of the model bilayer structure, it is also possible to exert structural control through both the support on which the bilayer is mounted and through the composition of the (aqueous) overlayer. Ultimately, for a bilayer structure to function in a biomimetic manner, it must retain its fluid properties,1 and this fact constrains the options available for control through the support. Arguably greater © 2013 American Chemical Society
structural control can be achieved through the composition of the liquid overlayer, and it is this issue we begin to address in this paper. The interaction of lipid bilayer structures with ethanol has been examined in some detail.7,23−38 It is known that the forces responsible for the organization of lipid bilayer structures involve hydrogen bonding between adjacent phospholipid head groups and between headgroups and the aqueous environment in closest proximity. These polar interactions function cooperatively with van der Waals interactions between the acyl chain regions of the lipid bilayer, producing two distinct layers of phospholipids with opposing orientations. For typical lipid bilayer structures, there exists a “gallery” region between lipid leaflets, and this structure allows for quasi-independent fluid behavior for each leaflet. Indeed, if this fluidity did not exist, diffusion-mediated processes related to the operation of trans-membrane protein structures could not occur. If there are chemical means of controlling the fluid and/or morphological properties of lipid bilayers through the composition of the aqueous overlayer, such control would open the door to new options in mediating bilayer function. Ethanol is known to interfere with the hydrogen bonding behavior between lipid headgroups and the aqueous environment in immediate contact with them. In addition, by virtue of its aliphatic functionality, ethanol is capable of interacting to some extent with the acyl chain region of phosphocholines. For these reasons, sufficiently high concentrations of ethanol have been shown to produce bilayer interdigitation, where the lipid acyl chains from both leaflets interpenetrate, eliminating the interleaflet gallery and dilating the headgroup region.23,25−32 Short of this extensive restructuring of the bilayer, however, it has been shown that the organization of these structures is Received: October 17, 2013 Revised: December 27, 2013 Published: December 27, 2013 537
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Figure 1. Structures of the bilayer components used in this work. (a) DOPC, (b) SR-DOPE, (c) sphingomyelin, and (d) cholesterol.
influenced by the presence of ethanol.27 Previous work focusing on the rotational diffusion dynamics of the nonpolar chromophore perylene imbedded in the acyl chain region of a C14 phosphocholine (DMPC) bilayer have shown that the motional freedom of the chromophore within this region is indeed altered by the presence of ethanol in the aqueous environment.27 In that work 100 nm diameter DMPC vesicles were the bilayers studied. Those results revealed, among other things, that the acyl chain region of the bilayer structure is sensitive to the composition of the aqueous extra-vesicular medium. For ethanol concentrations between 0.6 and 0.7 M, a change in the rotational dynamic of perylene was found, and this finding was seen for both the gel and fluid phases of the bilayer. In this work we are interested in the examination of planar bilayer structures because of their potential utility in the formation of biomimetic structures for chemical sensing. We are also interested in the effect of ethanol on somewhat more complex bilayer systems. We have investigated planar supported bilayers composed of the phosphocholine DOPC, cholesterol, and sphingomyelin, with a small amount of the
headgroup-tagged lipid SR-DOPE added as an optical probe. Exposing this supported bilayer structure to ethanol in the aqueous overlayer gives rise to structural and organizational changes that can be sensed by the time-domain response of the sulforhodamine tethered probe. The data show that in the vicinity of 0.8 M ethanol there is a distinct change in the organization of the membrane that leads to bimodal distributions of fluorescence lifetime and anisotropy decays, and that for ethanol concentrations above ca. 1.5 M the structural integrity of the bilayer structure is degraded significantly. These data underscore the ability of ethanol to mediate the organization of multicomponent bilayer systems.
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EXPERIMENTAL METHODS Materials. 1,2-Dioleoyl-sn-phosphatidylcholine (DOPC), cholesterol (ovine wool), sphyngomyelin (chicken egg), and sulforhodamine tagged 1,2-dioleyol-sn-phospatidylethanolamine (SR-DOPE), dissolved in chloroform, were obtained from Avanti Polar Lipids and used without further purification. The structures of these compounds are presented in Figure 1. Trizma Tris buffer (Sigma Aldrich) was prepared at a 538
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Figure 2. Schematic of fluorescence lifetime and anisotropy imaging instrument. The inverted microscope is a model Nikon Ti−U, equipped with a broadband illuminator and a confocal scanning apparatus (Becker & Hickl). Detection ports are equipped with a two channel, polarization-selective time correlated single photon counting (TCSPC) detection system (Becker & Hickl) and a CCD camera (Q-imaging).
of rinsing solution (6.9 mM Tris and 69 mM NaCl) or with rinsing solution containing ethanol (0.3 M-1.5 M), depending on the ethanol concentration to be studied. The interface was hydrated with the same rinsing solution (aqueous buffer or aqueous buffer containing ethanol) during the measurements. Fluorescence Lifetime and Anisotropy Imaging Measurements. The instrument used to acquire fluorescence lifetime and anisotropy images is based on an inverted microscope optical configuration (Nikon Eclipse Ti−U) shown in Figure 2. The microscope is equipped with a mercury lamp illuminator for the acquisition of steady state fluorescence images and with 10×, 20×, 40×, 60×, and 100× objectives (Nikon). Detection of time-resolved data is achieved with a polarized dual channel confocal scanning instrument (Becker & Hickl DCS-120) attached to an output port of the microscope and controlled by a galvo-drive unit (Becker & Hickl GDA120). The DCS-120 is equipped with a polarizing beam splitter and two avalanche photodiode detectors (ID-Quantique ID100) for the acquisition of fluorescence lifetime and anisotropy decay images. Polarized fluorescence transients used in the generation of images presented in this work are acquired using timecorrelated single photon counting detection electronics (Becker & Hickl SPC-152, PHD-400N reference diode). This system is characterized by an instrument response function of less than 100 ps fwhm. The time-correlated single photon counting (TCSPC) and confocal scanning instruments are controlled by
concentration of 10 mM (pH 7.5,100 mM NaCl) for bilayer deposition and 6.9 mM (pH 7.3−7.5, 69 mM NaCl) for rinsing of the deposited bilayers. Milli Q water was used to prepared lipid vesicle solutions. CaCl2 (2 mM aqueous solution) was used for vesicle fusion deposition of the bilayer onto the support. High grade muscovite mica (Ted Pella, Inc.) was used as the support. Ethanol used in the aqueous overlayer was purchased from Sigma Aldrich (190 proof ACS spectrometric grade) and used as received. Lipid Bilayer Preparation and Deposition. Lipid bilayers were formed by vesicle fusion, a technique used widely to form bilayer structures on mica.39−41 The initial step is the formation of a mixture of the bilayer constituents in chloroform, followed by drying the mixture under a stream of N2 (g) to remove chloroform. The composition of the mixture used in this work is 10 mol % cholesterol, 40 mol % egg-sphingomyelin, 49 mol % DOPC, and 1 mol % SR-DOPE. The dried mixture was taken up in Milli Q water to a final lipid concentration of 1 mg/ mL. The mixture was hydrated for 30 min at 50 °C, then vortexed briefly (1−2 min), and sonicated for 30 min at 50 °C (during which time the solution went from turbid to clear). The resulting vesicle-containing solution was allowed to cool to room temperature before deposition. Twenty μL of vesiclecontaining solution was placed on mica, followed by 60 μL of Trizma buffer (10 mM Tris, 100 mM NaCl) and 7 μL of 2 mM CaCl2. The vesicle containing solution remained in contact with the mica support for ca. 10 min before rinsing with ∼3 mL 539
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SPC132) and recorded using software written in LabVIEW programming language. The typical instrument response function for this system is 40 ps fwhm. The excitation wavelength was 573 nm and emission transients were collected at 587 nm. Steady State Excitation and Emission Measurement. Steady state excitation and emission spectra of vesicles of the same composition used for the formation of supported planar bilayers, formed by sonication in aqueous buffer solution and containing CaCl2, were measured using a Spex Fluorolog 3 emission spectrometer. These spectra were used to determine the excitation and emission wavelengths for the time-resolved measurements. The bandpass for both the excitation and emission monochromators was 1 nm for all measurements.
commercial software (Becker & Hickl) run on a windows-based PC. The light source for this instrument is a synchronously pumped cavity dumped dye laser (Coherent 702) excited by the output of a passively mode locked Nd:YVO4 laser (Spectra Physics Vanguard). The Vanguard pump laser produces 13 ps pulses at 80 MHz repetition rate, with 2.5 W average power at both 355 and 532 nm. The dye laser is cavity dumped (Gooch and Housego 64389.5-SYN-9.5−1 cavity dumper driver) to control the repetition rate. The output of the dye laser is characterized by ca. 5 ps pulses at a repetition rate that is variable between 80 kHz and 80 MHz. Typically the repetition rate is 4 MHz (250 ns interpulse spacing), and the power at the sample is less than 0.5 mW average. The dye laser output can be tuned from 430 to 850 nm depending on the dye and optics used and the excitation wavelength. For this work the dye laser output was set to 575 nm and the emission collection window of the microscope optics was 630 ± 30 nm, determined by the bandpass filter used in the confocal scanning apparatus and the bandpass optics in the microscope. These wavelengths were selected based on the excitation and emission spectra of the SRDOPE chromophore used in this work, presented in Figure 3.
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RESULTS AND DISCUSSION The central focus of this work is on understanding how the organization and dynamics of a planar lipid bilayer supported on mica are influenced by the presence of ethanol in the aqueous overlayer. The larger motivation for this work is to achieve control over bilayer properties through the composition of the solvent overlayer. To evaluate the organization and dynamic behavior of these supported bilayers we use steady state fluorescence imaging as well as time-resolved fluorescence lifetime and anisotropy decay imaging measurements. It is important to keep in mind that the bilayer structures we are examining are on the order of 6 nm thick. While the microscope objectives we use for collection and imaging have a depth of field well in excess of the bilayer thickness, rapid loss of signal would result if the bilayer detached from the support due to dilution. We observe no such loss of signal in any of the experiments we have performed. As is well established in the literature, for the bilayer composition used in this work we observe phase segregation of the cholesterol and phosphocholine domains.3,5,7 In the images shown in Figure 4, the dark circular regions are comprised of cholesterol and the bright regions, into which the SR-DOPE partitions selectively, are primarily phosphocholine. While the distribution of sphingomyelin is not evident in these images, it is known that this constituent is required for the formation of the phase segregated structures. In two-component bilayers containing only cholesterol and phosphocholine or only phosphocholine and sphingomyelin, similar microscale phase segregation is not observed. Of particular importance to this work is that the size and distribution of cholesterol domains depends on the amount of ethanol present in the aqueous overlayer. It is clear based on the steady state fluorescence intensity images alone, shown in Figure 4, that ethanol in the overlayer exerts a structural influence on the bilayer. We also note the presence of bright spots in our steady state images. These bright spots are seen for all samples (with either aqueous or ethanolic solution overlayers), and comprise ca. 1% of the total area of the images. Due to the small fractional area of the images occupied by the bright spots, it is not possible to explain their presence simply on the basis of probability. There are two features of the bright spots that are noteworthy. The first is that many of these spots are seen to form at the phase boundary between the lipid-rich and sterol-rich regions within the bilayers (Figure 4), and the second is that the size of the bright spots can be increased by exposure to high intensity light (data not shown). We note in some instances the presence of such spots in the lifetime images, as seen in Figure 5. Comparison of the time domain data for these bright spots with
Figure 3. Excitation and emission spectra of SR-DOPE in aqueous vesicle solution, normalized to unit intensity.
Steady state fluorescence images were acquired using a Nikon Intensilight C-HGFI illuminator and a Q-Imaging cooled monochromatic CCD 1392 × 1040 pixel2 camera (model Q1CF-M-12-C), both mounted on the inverted microscope. Nonimaging TCSPC Measurements. For the determination of the infinite-time anisotropy, a characteristic feature of the reorientation data for tethered chromophores (vide infra), we have used a TCSPC instrument that has been described elsewhere.27 This system, which utilizes a light source identical to that described above for the imaging microscope, has a detection system that records fluorescence transients polarized vertically and horizontally at the same time. The sample is 2 μm diameter vesicles prepared by freeze−thaw-vortex mixing and subsequent extrusion through a 2 μm filter in aqueous buffer solution or in aqueous buffer containing ethanol.42 Emission is collected using a reflective 40× microscope objective (Ealing), with polarization components separated using a polarizing cube beam splitter (Newport). The two polarized transients are passed through subtractive double monochromators (Spectral Products CM-112) to microchannel plate PMT detectors (Hamamatsu R3809U). The electronic signals are processed using commercial TCSPC electronics (Becker & Hickl 540
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Figure 4. Steady state fluorescence images of phospholipid bilayers supported on mica, with images acquired for different concentrations of ethanol in the aqueous overlayer, as indicated in each image. Red regions are phospholipid-rich, and dark circular regions are cholesterol domains. Images were acquired using a 40× objective and the scale bar is 10 μm for each image.
regions adjacent to them shows no difference outside of experimental uncertainty for the anisotropy decay time, but the lifetime images show bright spots characterized by a longer lifetime. One possible explanation for these data is the thermally induced formation of chromophore aggregates. It is known that the formation of H-aggregates typically gives rise to lower fluorescence quantum yields but J-aggregates can exhibit a higher quantum yield than the corresponding monomer.43−48 Because of the inability to acquire spectrally resolved dynamical information with our inverted microscope system, we are unable to evaluate this possibility at the present time. The steady state fluorescence images alone, however, cannot elucidate the details of the interactions between the bilayer and ethanol. To gain greater insight into the effect of ethanol on bilayer structure and dynamics, we have acquired fluorescence lifetime and anisotropy images of our bilayers as a function of ethanol concentration. We present the fluorescence lifetime images in Figure 5 and the anisotropy decay images in Figure 6. These data provide insight into the polarity and environmental freedom that characterizes these interfaces as the ethanol concentration is varied. Before considering these results in
Figure 5. Fluorescence lifetime images of phospholipid bilayers supported on mica, with images acquired for different concentrations of ethanol in the aqueous overlayer, as indicated in each image. Colored regions are phospholipid-rich and contain SR-DOPE, and the dark circular regions are cholesterol domains. The colors shown are set by the imaging software, with blue being short lifetimes and red being longer lifetimes. Images were acquired using a 40× objective.
detail, however, some discussion is required regarding the information we report and its interpretation. The time-domain images presented in Figures 5 and 6 are 256 × 256 pixel2 images, where each pixel represents either the lifetime (Figure 5) or anisotropy decay time constant (Figure 6) extracted from emission transients polarized parallel (I∥(t)) and perpendicular (I⊥(t)) to the (polarized) excitation pulse. The fluorescence lifetime data are calculated from these emission transients according to eq 1.
Ifl(t ) = I (t ) + 2I⊥(t ) 541
(1)
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data of sufficiently high S/N ratio to allow for quantitative determinations of either Ifl(t) or r(t). In order to gain more quantitative insight into the information contained in these data, we have taken selected uniform regions of pixels from the acquired images (several different images were acquired for each bilayer/ethanol concentration and selected regions from all images yielded the same averaged results to within the experimental uncertainty). Transients of the same polarization for individual pixels within the selected regions were averaged and the fluorescence lifetime decay and anisotropy decay were calculated for these averaged, higher S/N ratio data. The results for these processed data are presented in the histogram plots accompanying the lifetime (Figure 7) and anisotropy (Figure 8) data. It is important to note that it is more useful to present these results in histogram format than simply as averages and associated uncertainties in tabular form because the systems under investigation are heterogeneous and the assumption of a single Gaussian distribution of results is not justified, a priori. Fluorescence lifetime data are useful because they provide some information on spatial variations in polarity at the bilayer interface. While the fluorescence lifetime is expected to depend on the dielectric response of the chromophore local environment, the actual spectral response of the probe is complicated by inhomogeneous broadening, low symmetry and the presence of heteroatoms. As a result, the fluorescence lifetime images shown in Figure 5 and the associated ethanol-dependent changes in lifetime presented in Figure 7 are useful as gauges of variations in the relative polarity of bilayer interface, but quantitative information cannot be extracted reliably from them. Quantitative information on the chromophore local environment is available through anisotropy decay measurements, where a well-established body of theory allows for significant detail to be extracted from experimental data. We treat the motion of the SR-DOPE chromophore in the context of the hindered rotor model because of its attachment to phosphoethanolamine.49,50 In this formalism the chromophore is constrained to rotate within a cone of confinement defined by the semiangle θ0. The chromophore is also able to rotate about its tethering bond, with a characteristic (rotational) diffusion constant Dw. The anisotropy decay recovered from the experimental data contains a zero-time anisotropy, R(0), a decay time constant, τHR, and an infinite-time anisotropy, R(∞). We measure the quantities R(0), R(∞), and τHR, and from these data we extract the parameters θ0 and Dw, according to eqs 3 and 4.
Figure 6. Fluorescence anisotropy decay time constant images of phospholipid bilayers supported on mica, with images acquired for different concentrations of ethanol in the aqueous overlayer, as indicated in each image. Colored regions are phospholipid-rich and contain SR-DOPE and the dark circular regions are cholesterol domains. The colors shown are set by the imaging software, with blue being short lifetimes and red being longer lifetimes. Images were acquired using a 40× objective.
τHR =
(4)
We recognize that eq 3 is an approximation of the full equation.49,50 Given the uncertainty in the experimental data we report here, we do not compromise the information presented with the use of this approximate form. The quantities θ0 and Dw are characteristic of the local environment for the SR-DOPE chromophore. The cone semiangle, reflective of the cone of restriction in which the chromophore resides is found to be 55° to within the experimental uncertainty for all ethanol concentrations. This cone angle is the magic angle for linear spectroscopic measurements, and it indicates that, averaged
I (t ) − I⊥(t ) I (t ) + 2I⊥(t )
(3)
⎛ ⎛ ⎞1/2 ⎞1/2 ( ) R ∞ cos θ0 = 0.5⎜⎜8⎜ ⎟ + 1⎟⎟ − 0.5 ⎝ ⎝ R(0) ⎠ ⎠
and the anisotropy decay data are calculated according to eq 2
r (t ) =
7θ0 2 24Dw
(2)
The images, calculated using commercial (Becker & Hickl) software, provide substantial insight into variations in the morphology and dynamics of the bilayers as a function of exposure to ethanol. While the overall images provide very useful qualitative information, individual pixels do not contain 542
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Figure 7. Histogram plots of averaged fluorescence lifetime data over selected regions of pixels (typically ca. 20 × 20). The x axes are time in picoseconds, and the y axes are the number of measurements. From top to bottom the ethanol concentration is varied from 0.0 to 1.5 M in the aqueous overlayer.
Figure 8. Histogram plots of averaged fluorescence anisotropy decay time constant data over selected regions of pixels (typically ca. 20 × 20). The x axes are time in picoseconds and the y axes are the number of measurements. From top to bottom the ethanol concentration is varied from 0.0 to 1.5 M in the aqueous overlayer.
over the pixel size, there is a nominally random distribution of chromophore angles. In other words, for all ethanol concentrations there exists substantial disorder at the lipid headgroup-aqueous overlayer interface. This is not a surprising result, and it confirms what has been established previously from both experimental and computational perspectives.23,25−30,32,51
The cone angle values we recover experimentally are essentially identical to within the experimental uncertainty to that calculated for this same chromophore using a molecular dynamics model.52 The value we recover can be attributed to disorder in the system, and the calculated findings of Kyrychenko indicate that this tilt angle is a consequence of favorable interactions between the chromophore and the 543
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headgroups of adjacent phospohocholines.52 We believe that the experimental and calculated results are fully consistent with one another and that the interactions between the chromophore and adjacent headgroups serves to mediate the anisotropy decay time constants we measure. The data shown in Figure 8 are the values of the individual determinations of τHR from the averaged data for each ethanol concentration. Because the θ0 values are the same for all ethanol concentrations, the τHR decays are all proportional in the same manner to Dw and are thus directly comparable to one another. We note that the SR-DOPE chromophore is presumably distributed equally between the top and bottom leaflets of the bilayer, and the bottom leaflet is in contact with a presumably hydrated layer on the mica support. The environment presented to the inner leaflet is expected to change to the extent that ethanol can access this region, either by means of defects in the bilayer film or by diffusion through the bilayer structure. Our experimental data, which exhibit ethanoldependent reorientation dynamics, are sensitive to chromophores present in both leaflets. Interleaflet exchange (translocation) is known to occur in lipid bilayers and the presence of two different environments on each side of the supported bilayer may provide a sufficient driving force to cause bilayer asymmetry, where the chromophore resides predominantly in one leaflet or the other. Based on our experimental data and the higher solubility of rhodamines in ethanol than in water, we believe that any bilayer asymmetry that may arise in our systems would result in a higher concentration of probe in the outer leaflet. We recognize that the size of the headgroupbound rhodamine chromophore may serve to hinder translocation,53 but the presence of defects and phase boundaries in our planar supported bilayers could act to mediate any translocation that does proceed in this system. Understanding and characterizing interleaflet exchange in this system is beyond the scope of the present work and will likely require the use of a second-order nonlinear optical technique.54−59 As the ethanol concentration in the overlayer is increased from 0 to 0.3 M, the rotational time constant becomes longer, i.e., Dw becomes smaller, indicating that for low ethanol concentrations some local organization is seen in the bilayer headgroup region. With further increase in ethanol concentration to 0.5 M, we observe a slight increase in τHR but with an associated broadening of the distribution of τHR values. At 0.8 M ethanol concentration, we observe what appears as a bimodal distribution of τHR times, indicative of a change in the morphology of the bilayer structure and the coexistence of two distinct environments sensed by the SR-DOPE probe. This finding is consistent with the steady state fluorescence images (Figure 4) and the anisotropy decay images (Figure 6), where the characteristic cholesterol domain size is seen to change. For 1.0 M ethanol a unimodal distribution of τHR is seen and for 1.5 M ethanol it is likely that integrity of the bilayer structure is compromised.27 The anisotropy decay time data are qualitatively similar to the lifetime data, with an increasing trend in fluorescence lifetime up to 0.8 M ethanol and a broadening distribution width in lifetime with higher ethanol concentration. When the steady state fluorescence images are viewed with the perspective of the fluorescence lifetime and anisotropy decay images, it becomes clear that the presence of ethanol in the aqueous overlayer is causing a change in the organization of the bilayer and in the vicinity of 0.8 M ethanol concentration the bilayer experiences a macroscopic change in its morphology.
We have observed an analogous change in the organization of DMPC unilamellar vesicles where perylene was used as a fluorescent probe of the nonpolar lipid acyl chain region. The fact that the ethanol concentration at which the structural transition was seen for DMPC was in the 0.6 to 0.7 M range is not surprising based on the different compositions of the bilayers used in that work and those on which we report here. What is significant, however, is that a change in the organization of the bilayer is seen for sub 1 M ethanol for both bilayer systems and that this change is observed both in the headgroup region (vide infra) and in the acyl chain region.27 One structural change in the bilayer system that will affect both the headgroup and acyl chain regions is the interdigitation of the two leaflets. Such a change would certainly give rise to changes in the local environment of both regions of the bilayer and interdigitation is a known phenomenon for lipid bilayers exposed to ethanol.25,26,28−32 If interdigitation is occurring near 0.8 M ethanol, one would expect to see a dilation of the headgroup functionalities with a consequent reduction in interheadgroup interactions. This is seen qualitatively in the anisotropy decay data (Figure 8), as the τHR data for 1.3 M ethanol is faster than that for 1.0 M ethanol. It is interesting that the interdigitation of bilayer leaflets should give rise to a reduction in the cholesterol domain size in these supported bilayers. It is expected that ethanol will interact with both the phospholipid-rich and sterol-rich regions of the bilayer although the interactions of cholesterol and ethanol are not well understood. It is clear, however, that the addition of ethanol to the aqueous overlayer does indeed affect the lipid bilayer morphology. The interaction of ethanol with phospholipids has been studied extensively, with a substantial effort being made in the area of molecular dynamics because of their ability to elucidate molecular interactions and provide modeling support for experimental data.7,33−36 Complex, multicomponent systems, some including a headgroup-bound chromophore,52 have been examined computationally, and the results of these efforts have been useful in understanding experimental data. A common result of many of these studies is that regions of the bilayer appear to diminish somewhat in thickness upon interaction with ethanol, and this so-called thinning occurs more readily in bilayer regions where the lipids exist in a disordered phase.7,33 It has also been noted that the presence of cholesterol in these model lipid bilayers tends to make the bilayer more rigid.33 With exposure to increasing concentrations of ethanol, nonbilayer structures have been predicted to form,34 consistent with the destruction of bilayer structures seen experimentally under this condition. Taken collectively, these studies point to the ability of ethanol to mediate and modify the organization of lipid bilayer structures, and our experimental data support this assertion. Our data show that the measured anisotropy decay time becomes longer with increasing ethanol concentration, consistent with an alteration of the lipid headgroup organization that facilitates interactions with the SR-DOPE chromophore. In the vicinity of 0.8 M ethanol, we observe the onset of a bimodal distribution in the anisotropy decay time, suggesting the existence of two distinct environments for the chromophore. The fluorescence lifetime data show an increase in lifetime with ethanol concentration and a bimodal distribution for 1 M ethanol. At 1.5 M both the lifetime and anisotropy decay data suggest that the organization of the bilayer has been compromised. Experimental data and 544
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molecular dynamics simulations are consistent and provide much insight into the effects of ethanol on bilayer structures.
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CONCLUSIONS We have examined the interactions of supported four component bilayers (DOPC, sphingomyelin, cholesterol, and SR-DOPE) with ethanol present in the aqueous overlayer. Our data demonstrate ethanol concentration-dependent changes in the morphology of the supported bilayer. Steady state fluorescence images demonstrate that, in the vicinity of 0.8 M ethanol, the characteristic domain size of the sterol-rich domains becomes significantly smaller. These findings are corroborated by changes in the fluorescence lifetime and anisotropy images of the bilayers with increasing ethanol concentration. The fluorescence anisotropy decay images and decay time constants reveal the presence of two distinct domains for 0.8 M ethanol, indicating a structural transition occurring in this ethanol concentration range. Our data indicate that the integrity of the supported bilayer is degraded significantly at 1.5 M ethanol concentration. These data are consistent with those reported previously for unilamellar vesicles of DMPC in which perylene was incorporated. In that work, analogous bilayer behavior was seen via perylene reorientation dynamics to occur in the range of 0.6−0.7 M ethanol. The similarity in ethanol concentration for which a structural transition is seen in such different bilayer systems suggests that the dominant driving force for the structural transition is the interaction of ethanol with the polar headgroup region of the bilayers. It is known that ethanol disrupts hydrogen bonding in the lipid headgroup region and that interdigitation of the bilayer leaflets can occur.25−32 Our data provide insight into the changes in polarity and molecular scale organization that are associated with ethanol-induced structural changes in lipid bilayer structures. We thus have a means of mediating the organization of lipid bilayer structures through intermolecular interactions in the lipid headgroup region. Future work in this area lies in understanding the role of the aliphatic functionality of the alcohol in mediating lipid bilayer organization.
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AUTHOR INFORMATION
Corresponding Author
*Tel: (517) 355 9715 x224. E-mail:
[email protected]. edu. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS We are grateful to the Donors of the Petroleum Research Fund for their support of this work through Grant 52692-ND6 and to the National Science Foundation for their support of the instrument construction through Grant 1048548.
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