Structural Disruption of Phospholipid Bilayers over a Range of Length

Feb 26, 2014 - Iwan Setiawan and G. J. Blanchard*. Department of Chemistry, Michigan State University, 578 South Shaw Lane, East Lansing, Michigan ...
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Structural Disruption of Phospholipid Bilayers over a Range of Length Scales by n‑Butanol Iwan Setiawan and G. J. Blanchard* Department of Chemistry, Michigan State University, 578 South Shaw Lane, East Lansing, Michigan 48824-1322, United States ABSTRACT: We report on the exposure of planar multicomponent lipid bilayers supported on mica to n-butanol. The bilayer contains 49 mol % 1,2-dioleoyl-sn-phosphatidylcholine (DOPC), 10 mol % cholesterol, 40 mol % sphingomyelin, and 1 mol % sulforhodamine-tagged 1,2-dioleoylsn-phosphatidylethanolamine (SR-DOPE). Phase separation of the cholesterol domains is seen within the bilayer structure, and exposure of this supported bilayer to controlled amounts of n-butanol in the aqueous overlayer produces morphological changes over a range of length scales. We report steady state fluorescence imaging, fluorescence lifetime imaging, and fluorescence anisotropy decay imaging for these bilayers. These data are consistent with literature reports on the interactions of lipid bilayers with n-butanol and provide molecular-scale insight relative to bilayer organization that has not been available to date. The exposure of these bilayers to n-butanol leads to more extensive disruption of the bilayer than is seen for their exposure to ethanol.

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INTRODUCTION Plasma membranes have been examined extensively for a number of reasons, ranging from their central role in cellular function to their utility in biomimetic sensing applications and the production of biofuel by fermentation.1 Plasma membranes are highly complex structures, containing in excess of 100 different molecular constituents distributed nonuniformly within the membrane. Indeed, the existence and role of lipid raft structures has been an area of intense scrutiny over the past decade, with the implication of their role in cell sorting and cellular signaling.2−4 For many applications where biomimetic behavior is desired, attempts have been made to use model bilayer systems containing a limited number of constituents, typically phospholipid(s), sphingolipid(s), and cholesterol. In such three-component systems, phase segregation is seen, with the characteristic domain size of the phase-segregated regions depending on the identity and amount of each constituent present. A great deal of effort has gone into the characterization of these phase-segregated model systems, with their existence and form being explained in the context of line tension.5−10 If such model systems are to be predictably useful for applications such as housing trans-membrane proteins in their active form(s), a molecular understanding of the interactions between domains and short-range organization in these systems will be needed. In plasma membrane structures, control over their function is achieved through variations in their chemical composition and thus their organization. For applications aimed at the use of biomimetic model bilayers, sometimes present on a planar support, control over their organization must be achieved in other ways. One such method is through the composition of an aqueous overlayer. It is well-known that the organization of supported bilayer structures depends on the composition of the © 2014 American Chemical Society

aqueous overlayer in contact with the bilayer. Control can be achieved through the pH, the ionic strength, or the presence of species such as short-chain alcohols.11−22 Much of the work aimed at controlling the organization of model bilayer structures has focused on changes in the micrometer-scale morphology of the bilayer as a function of a system variable, and less effort has been devoted to understanding the corresponding changes in organization at the molecular scale. We are interested in the molecular-scale organization of model bilayer systems and how this organization depends on the presence of selected species in the aqueous overlayer. It has been established from past studies that short-chain alcohols mediate bilayer organization through interactions between the alcohols and the hydrophilic headgroups of the lipids. Rowe et al. have used 13C NMR spectroscopy to show that for dipalmitoylphosphatidylcholine (DPPC) and L-αdimyristoylphosphatidylcholine (DMPC) bilayers, n-butanol is more soluble in the bilayer fluid phase than in the gel phase.20 For fluid phase bilayers, n-butanol exhibits two resonances, associated with its presence in the aqueous phase and in the lipid bilayer. For the gel phase only one (aqueous phase) nbutanol resonance is observed, implying limited interactions between n-butanol and the more ordered bilayer structure. Ly and Longo have shown that n-butanol decreases the structural integrity and increases the area per molecule of stearolyloleoylphosphatidylcholine (SOPC) lipid bilayers through their use of a micropipet aspiration technique designed to measure the area compressibility modulus, bending modulus, lysis tension, lysis strain, and area expansion.23 Westerman et al. have studied the Received: January 15, 2014 Revised: February 26, 2014 Published: February 26, 2014 3085

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Figure 1. Structures of the bilayer components used in this work: (a) DOPC, (b) cholesterol, (c) sphingomyelin, and (d) SR-DOPE.

acquired steady state fluorescence microscopy images to evaluate changes in micrometer-scale organization of these bilayers. The specific ternary system we have chosen contains cholesterol, DOPC, and egg sphingomyelin because this system is known to form microdomains where liquid ordered and liquid disordered lipid phases are known to coexist.26−29 In an earlier study we observed that this same ternary bilayer system, upon exposure to ethanol, exhibited changes in the characteristic lipid domain size and the dynamics of the same headgroup-bound chromophore revealed that, in the vicinity of 0.8 M ethanol, the chromophore revealed the presence of two distinct environments as seen in a bimodal distribution of the reorientation time data. These data underscored the different and complementary information available from time-resolved lifetime and anisotropy decay imaging and from steady state fluorescence imaging. Our findings for the exposure of this bilayer system to n-butanol differ somewhat from those for exposure to ethanol,21 owing to the lower solubility of nbutanol in water and the greater aliphatic character of the alcohol. The enhanced aliphatic character of n-butanol facilitates interactions with the hydrophobic acyl chain region of the bilayer, thereby disrupting hydrogen bonding interactions between the lipid headgroups.2 Exposure of the bilayer to n-butanol is seen to alter the organization of the bilayer at significantly lower concentrations than are observed for ethanol. Steady state fluorescence images of the supported bilayer show that the integrity of the bilayer is compromised at n-butanol concentrations of 0.4 M and above, but the size and shape of the remaining microdomains do not change greatly with increasing n-butanol concentration. It is possible that n-

interaction of n-alcohols with lipid bilayer membranes, and they showed that n-butanol creates greater disorder than other longer chain n-alcohols (n-octanol, n-dodecanol, and ntetradecanol) in DMPC bilayers.22 They reasoned that the amphiphilic nature of n-butanol combined with its water solubility were responsible for this finding. Other n-alcohols, which could interact significantly with the lipid acyl chain region, exhibited low solubility in water, with the achievable concentrations being too low to cause bilayer disruptions analogous to those seen for n-butanol. The interdigitation of lipid bilayer structures caused by the presence of alcohols has been observed and predicted using a variety of measurements and modeling methods.15−19,24 The interactions of bilayers with short-chain alcohols were believed to have a significant effect on membrane fluidity, permeability, and lateral mobility, thereby influencing the ability of the bilayers to mediate membrane protein conformation and functionality.2,11,25 In this study our primary interest lies in understanding the molecular-scale dynamics and longer range bilayer morphology of a supported heterogeneous bilayer as a function of its exposure to n-butanol. We probe the dynamics of a lipid headgroup-bound fluorophore incorporated in supported bilayers comprised of 1,2-dioleoyl-sn-phosphatidylcholine (DOPC), sphingomyelin, and cholesterol. We control the composition of the aqueous overlayer in contact with these bilayers. We use time correlated single-photon-counting (TCSPC) detection in concert with confocal scanning microscopy to image the sample by means of the spatial dependence of the fluorescence lifetime and the anisotropy decay time of the tethered chromophore. We have also 3086

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was used to obtain the time-domain data. The DCS-120 is equipped with a polarization-selective beam splitter and two avalanche photodiode detectors (ID-Quantique ID100) for the simultaneous acquisition of polarized emission transients. TCSPC detection electronics (Becker & Hickl SPC-152, PHD-400N reference diode) were used to collect time-resolved data. The instrument response function for this system is less than 100 ps full width at half-maximum (fwhm). The excitation source is a synchronously pumped cavity dumped dye laser (Coherent 702) excited by the second harmonic output of a passively mode locked Nd:YVO4 laser (Spectra Physics Vanguard). This laser produces 13 ps pulses with 80 MHz repetition rate with 2.5 W average output power at 355 nm and at 532 nm. The repetition rate of the dye laser is controlled by cavity dumping electronics (Gooch and Housego). The pulse from the dye laser is ca. 5 ps fwhm at a repetition rate of 4 MHz (250 ns pulse spacing), and the average power at the sample is less than 0.5 mW. 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 DCS-120 and the optics in the microscope. These wavelengths were selected based on the excitation and emission spectra of SR-DOPE. Steady state fluorescence images were acquired using a Nikon Intensilight C-HGFI illuminator and a QImaging cooled monochromatic CCD 1392 × 1040 pixel camera (model Q1C-F-M-12-C). Nonimaging TCSPC Measurements. For infinite time anisotropy (r(∞)) determinations we used a different TCSPC instrument, also described elsewhere.18 This system has a light source identical to that used for the imaging system (vide supra) and a detection system that records fluorescence transients polarized vertically and horizontally simultaneously. For vesicles in solution, the sample is prepared by freeze−thaw− vortex mixing and extrusion through a 2 μm diameter porecontaining membrane in aqueous buffer solution or in aqueous buffer containing n-butanol solution. Emission is collected using a reflective 40× microscope objective (Ealing), with polarization components separated using a polarizing cube beam splitter (Newport). The two polarized signal photons are passed through subtractive double monochromators (Spectral Products CM-112) to microchannel plate photomultiplier tube (PMT) detectors (Hamamatsu R3809U). The electronic signals are processed using commercial TCSPC electronics (Becker & Hickl SPC132) and recorded using software written in LabVIEW programming language. The instrument response function for this system is typically 40 ps fwhm. The excitation wavelength was 575 nm, and emission transients were collected at 590 nm.

butanol disrupts the DOPC liquid disordered regions of the bilayer preferentially, and because of its aliphatic chain length, n-butanol can penetrate more deeply than ethanol to the hydrophobic acyl chain region of the bilayer. These results are consistent with literature reports on similar systems.3,12 The information we report on the fluorescence lifetime and anisotropy decay dynamics for these supported bilayers indicates that the presence of n-butanol initially gives rise to less motional freedom for the chromophore and, at higher concentrations, an increase in the distribution of domains is seen, consistent with the physical disruption of the bilayer.

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EXPERIMENTAL METHODS Materials. 1,2-Dioleoyl-sn-phosphatidylcholine, cholesterol (ovine wool), sphingomyelin (egg chicken), 1,2-oleoyl-snglycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl ammonium salt) (SR-DOPE) were obtained from Avanti Polar Lipids (Alabaster, AL, USA), dissolved in chloroform, and used without further purification. The structures of these compounds are shown in Figure 1. Trizma Tris Buffer (Sigma Aldrich) was prepared to a concentration of 10 mM (pH 7.4−7.5, 100 mM NaCl) for bilayer deposition and 6.9 mM (pH 7.4−7.5, 69 mM NaCl) for rinsing. Lipid vesicle solutions were prepared using Milli Q water. A solution of 2 mM CaCl2 was used for sample deposition on mica support surfaces. High grade Muscovite Mica was used as the substrate (Ted Pella Inc.). n-Butanol was purchased from SAFC in >99.9% purity and was used as received. Lipid Bilayer Preparation and Deposition. Lipid bilayers were formed on mica by means of vesicle fusion, using a literature method with slight modifications.7,28,30 All lipids were dissolved in chloroform initially to produce a solution containing the bilayer constituents in predetermined amounts. The solvent was removed by drying under a N2(g) stream. The lipid mixture composition for this experiment is 10 mol % cholesterol, 40 mol % egg sphingomyelin, 49 mol % DOPC, and 1 mol % SR-DOPE. The dried lipid 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 a temperature of 50 °C (water bath), then vortexed briefly (1−2 min), and sonicated for 30 min at 50 °C (the solution changed from turbid to clear). After sonication, the clear lipid vesicle solution was cooled to room temperature before deposition. An aliquot of 20 μL of vesicle-containing solution was placed on the mica surface, followed by 60 μL of Tris buffer solution (10 mM Tris pH 7.4− 7.5, 100 mM NaCl) and 7 μL of 2 mM CaCl2. The vesicle deposition on mica was performed for ca. 10 min before rinsing with ∼3 mL of rinsing solution (6.9 mM Tris, pH 7.4−7.5, and 69 mM NaCl) or with rinsing solution containing n-butanol solution (0.1−0.6 M, depending on the experiment). The surface was hydrated with rinsing solution (aqueous buffer or aqueous buffer containing the appropriate amount of nbutanol) during spectroscopic measurements. Fluorescence Lifetime and Anisotropy Imaging Measurements. Fluorescence lifetime and anisotropy images were obtained using an instrument that has been described in detail elsewhere,21 and we provide a brief overview of its salient features. The sample is imaged using an inverted optical microscope (Nikon Eclipse Ti-U). Fluorescence lifetime and anisotropy images were collected with a 40× objective. A polarized dual channel confocal scanning instrument (Becker & Hickl DCS-120) attached to an output port of the microscope and controlled by a galvodrive unit (Becker & Hickl GDA-120)

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RESULTS AND DISCUSSION The primary 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 n-butanol in the aqueous overlayer. Ultimately, our goal is to achieve control over the morphology and fluidity of supported bilayer structures by means of the composition of the solvent overlayer because these bilayer properties have a direct influence on transmembrane protein functionality. To evaluate the extent to which this control can be achieved, we use steady state fluorescence imaging and time-resolved fluorescence lifetime and anisotropy decay imaging measurements. The supported structures we study are ca. 6 nm thick, and the depth of field of our instrument is in excess of 1 μm. Because of the nature of 3087

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the systems under investigation, rapid loss of the fluorescence signal due to dilution would result if the bilayer detached from the support. In fact, we observe such loss for n-butanol concentrations in excess of 0.4 M. Before considering the information content of our data, it is useful to provide some perspective on what is known about these systems. The exposure of lipid bilayers to n-butanol has been examined before.1,12−14,19,20 Key issues include whether or not there is a relationship between n-butanol-driven changes in short-range and long-range organization in the model bilayer, and how the presence of n-butanol in the aqueous medium above the bilayer influences the dynamics of the headgroupbound SR-DOPE chromophore. We examine these issues using steady state fluorescence imaging and time-domain fluorescence lifetime and anisotropy decay imaging measurements. We consider first the dependence of micrometer-scale bilayer organization on the presence of n-butanol. The data shown in Figure 2 are the steady state fluorescence images of the planar supported lipid bilayer exposed to n-butanol, revealing largescale disruption of the bilayer for n-butanol concentrations of 0.4 M and above. This behavior is not seen for exposure to ethanol.21 There are several interesting features contained in these data that warrant discussion. The first is that the n-

butanol-dependent change in the characteristic cholesterol domain size does not appear to occur at the same n-butanol concentration as the large-scale disruption of the bilayer structure. The fact that the disruption of the bilayer structure and the change of cholesterol domain size occur at different nbutanol concentrations indicates that the interaction of nbutanol with these two domains can be treated as distinct to some level of approximation. In other words, it appears that nbutanol interacts preferentially with phospholipids over cholesterol, a result that is not surprising based on the structures of the compounds. The second point of interest is the existence of bright spots in the bilayer with no n-butanol present, and the appearance of bright “lines” with increasing n-butanol concentration. We have seen the same bright spots before for investigations of ethanol interactions with bilayers,21 and while their cause remains under investigation, we do know that these spots typically appear at the boundary of phospholipid and cholesterol domains and that their size can be increased by laser illumination. For these reasons, we have postulated that the bright spots may be associated with chromophore J-aggregates,31−33 with the formation of the aggregates being mediated by annealing of these bilayer regions on irradiation. What is novel for exposing the bilayer to n-butanol is the formation of bright “lines”. One may be able to understand the formation of lines in the context of bilayer organization reflecting features such as scratches on the support surface, but such an explanation would not require n-butanol to be present. Given that the motional freedom of the chromophores decreases with increasing n-butanol concentration (vide inf ra), the formation of the lines cannot be mediated by enhanced fluidity within the bilayer upon exposure to n-butanol. For samples that are not exposed to n-butanol, bright spots are observed, and these spots appear to be the origin of the bright lines seen for samples exposed to n-butanol (Figure 2). As with the bright spots, the origin and properties of these bright lines requires further examination, but there does appear to be a connection between the lines and spots. The third issue of note in the steady state images is for nbutanol concentrations of 0.4 M and above. The structural integrity of the bilayer appears to be compromised for these nbutanol concentrations, and for 0.4 M n-butanol, in the boundary regions between void space and bilayers there appear to be regions of lower intensity emission from SR-DOPE. One explanation for this finding is that the lower intensity regions are lipid monolayers, but such a structure would necessarily expose the acyl chain region of the leaflet remaining at the mica surface to polar solution unless n-butanol plays an amphiphilic mediating role. It is also possible that the low-intensity regions are due to the bilayer in those regions being lower in density. This explanation implies that the bilayer is dilated significantly due to the presence of n-butanol. Such an explanation is qualitatively consistent with bilayer “thinning” seen with ethanol exposure34 and may be a precursor to bilayer interdigitation. For n-butanol concentrations of 0.5 M and above; these regions of intermediate intensity are not seen, suggesting that 0.4 M n-butanol represents a concentration where the loss of bilayer integrity is starting to become prominent. While the steady state fluorescence images shed light on changes in the micrometer-scale organization of the supported bilayers, such data do little in the way of providing a quantitative measure of any corresponding changes in bilayer organization at the molecular scale. In order to gain insight into

Figure 2. Steady state fluorescence images of phospholipid bilayers supported on mica, with images acquired for different concentrations of n-butanol 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× microscope objective. The scale bar is 10 μm for each image. 3088

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the role of n-butanol interacting with phospholipid bilayers, we have used fluorescence lifetime and anisotropy imaging measurements. The images presented in Figures 3 and 4 are 256 × 256 pixels, and each pixel contains fluorescence transients polarized parallel (I∥(t)) and perpendicular (I⊥(t)) to the excitation pulse. For the lifetime data presented in Figure 3, these transients are combined according to eq 1,

Figure 4. Fluorescence anisotropy decay time constant images of phospholipid bilayers supported on mica, with images acquired for different concentrations of n-butanol 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 indicating short lifetimes and red indicating longer lifetimes. Images were acquired using a 40× microscope objective.

Ifl(t ) = I (t ) + 2I⊥(t )

(1)

And the functional form of Ifl(t) is given by Figure 3. Fluorescence lifetime images of phospholipid bilayers supported on mica, with images acquired for different concentrations of n-butanol 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 indicating short lifetimes and red indicating longer lifetimes. Images were acquired using a 40× microscope objective.

Ifl(t ) = Ifl(0) exp( −t /τfl)

(2)

where Figure 3 is of the form τfl(x,y). The anisotropy decay image shown in Figure 4 is given by r (t , x , y ) = 3089

I (t , x , y) − I⊥(t , x , y) Ifl(t , x , y)

(3)

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It is the functional form of r(t) that contains information on the molecular motion of the (tethered) chromophore. Because the chromophore is tethered to the headgroup of a phosphoethanolamine, which incorporates into the supported bilayer, the anisotropy decay dynamics must be treated in the context of the hindered rotor model.35,36 In this model, the chromophore can execute diffusional motion within a conic volume of semiangle θ0 and also execute a “wobbling” motion (Dw) about the bond connecting it to the phosphoethanolamine moiety. The anisotropy decay function, r(t), is given by eq 4, r(t ) = r(∞) + (r(0) − r(∞)) exp( −t /τHR )

(4)

where r(∞), the infinite time anisotropy, is a consequence of the inability of the chromophore to access all possible orientations; r(0) is the zero-time anisotropy, related to the orientation of the emitting transition dipole moment relative to that of the absorption transition; and τHR is the decay time constant of the hindered rotor.

τHR ≈

7θ0 2 24Dw

⎞ ⎛ ⎛ ⎞1/2 ⎞1/2 ( ) ∞ r ⎟ θ0 = cos ⎜0.5⎜⎜8⎜ ⎟ + 1⎟⎟ − 0.5⎟ ⎜ ⎝ ⎝ r(0) ⎠ ⎟ ⎠ ⎝ ⎠

(5)

⎛

−1⎜

(6)

The quantities θ0 and Dw are relevant in evaluating the extent of order and motional freedom available to the chromophore, respectively. For the data presented in this work, multiple locations on each image were selected and the polarized fluorescence transients were summed for blocks of data (typically ca. 20 × 20 pixels) to provide a higher S/N ratio. The τfl, τHR, r(0), and r(∞) data were used to gain insight into the local dielectric response and motional freedom experienced by the chromophore. We find that the presentation of the fluorescence lifetime and anisotropy decay time data in tabular format with standard deviations is not the most informative way to show these data. Rather, because of the structural heterogeneity of the systems we study and the changes in this heterogeneity that occur as a consequence of introducing nbutanol, the data are not well approximated as a Gaussian distribution of values centered around a single value, and the nature of the distribution that characterizes τfl and τHR changes with n-butanol concentration. For this reason we show in Figures 5 and 6 the histogram distributions of the fluorescence lifetime and anisotropy decay time constant as a function of nbutanol concentration. The fluorescence lifetime images of the bilayers as a function of n-butanol concentration shown in Figure 3 reveal the expected distribution of the chromophore in phosphocholine regions and their absence in cholesterol regions, and that the fluorescence lifetime is relatively uniform over micrometer length scales. It is interesting to note that, in certain images, the bright spot and line features are associated with a slightly longer fluorescence lifetime. This finding is consistent with the bright features being associated with J-aggregates, but our inability to obtain spectrally resolved data hampers the positive identification of these features. What is not apparent from the images shown in Figure 3 is the change in the average lifetime with increasing n-butanol concentration. The lifetime histogram data in Figure 5 shows qualitatively an increase in lifetime from ca. 2200 ps to ca. 2600

Figure 5. Histogram plots of averaged fluorescence lifetime data over selected (ca. 20 × 20) regions of pixels. The x-axes (time) are in picoseconds, and the y-axes are the number of occurrences. From top to bottom the n-butanol concentration is varied from 0.0 to 0.6 M in the aqueous overlayer, as indicated.

ps, with an approximately Gaussian distribution of results, up to 0.4 M n-butanol. For 0.5 M n-butanol there is a pronounced bimodal distribution, with one maximum in the vicinity of 2200 ps and the other near 2600 ps, and for 0.6 M n-butanol, there appears also to be a bimodal distribution, although the 3090

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providing unambiguous information on the chromophore local environment. For this reason we have also acquired anisotropy decay imaging data. The anisotropy decay time constant data (Figure 4) do not exhibit a discernible difference between regions of normal emission intensity and bright features. This finding suggests that any aggregation that the chromophore may participate in is short-lived on the time scale of its (rotational) diffusive motion. The anisotropy decay histogram data are shown in Figure 6. In contrast to the lifetime data, there appears to be no clear trend in the anisotropy decay time constant(s) with increasing nbutanol concentration. We note that for all of the anisotropy decay data we recover a cone angle θ0 ∼ 54.7° to within the experimental uncertainty, indicating that there is substantial disorder in the bilayer structure in the vicinity of the phospholipid headgroups. The bilayer in contact with aqueous solution containing no n-butanol exhibits an approximately Gaussian distribution centered at ca. 850 ps. The addition of nbutanol, at concentrations as low as 0.1 M, broadens the distribution in τHR and increases the average to ca. 1500 ps. Because in all cases the value of θ0 is the same to within the experimental uncertainty, the values of τHR shown in Figure 6 all exhibit the same proportionality to Dw−1. In other words, the data presented in Figure 6 reflect a slowing of Dw by a factor of ∼2 upon addition of n-butanol. Such an increase in Dw with the addition of an alcohol has been seen before, for ethanol.21 In that work, however, there was a discernible trend in the ethanol anisotropy decay time constant with increasing ethanol concentration and that trend was consonant with fluorescence lifetime data. There has been a report showing that n-butanol can interact with lipid headgroups,5 and such an interaction would serve to alter the reorientation dynamics of a pendent chromophore. For n-butanol in the aqueous overlayer we see no such correlation between lifetime and anisotropy decay time constant. We attribute the decrease in Dw with the presence of n-butanol to be related to the change in the headgroup local environment that serves to increase the effective viscosity experienced by the chromophore. We believe that the presence of n-butanol serves to disrupt the organization of the bilayer significantly, and the anisotropy decay data indicate that this is the case, even for comparatively small amounts present. This finding is not inconsistent with the lifetime data (Figure 5) because the two processes sense different properties of the chromophore immediate environment, and while there may be some expected variation of the dielectric response of the bilayer interface due the increasing contribution from n-butanol, reorientation time constant data are not sensitive to environmental properties that do not contribute to the immediate chromophore environment. In other words, the molecular-scale disruption produced in the bilayer by the presence of n-butanol occurs before such changes manifest themselves measurably on micrometer length scales. To this point we have considered the dynamics of the SRDOPE chromophore to be the same regardless of whether the chromophore resides in the top or bottom leaflet. We believe this is justified because there is likely some amount of aqueous medium between the mica support and the bilayer bottom leaflet.37 It is not possible to quantitate the amount of aqueous medium in this region, but the functional form of the experimental data, i.e., single-exponential fluorescence lifetime and single-exponential anisotropy decays, indicates that, regardless of the leaflet in which the chromophore resides, it

Figure 6. Histogram plots of averaged fluorescence anisotropy decay time constant data over selected (ca. 20 × 20) regions of pixels. The xaxes (time) are in picoseconds, and the y-axes are the number of occurrences. From top to bottom the n-butanol concentration is varied from 0.0 to 0.6 M in the aqueous overlayer, as indicated.

separation of the two populations is less well resolved. On the basis of the steady state fluorescence images shown in Figure 2, we believe that the structural integrity of the bilayer is compromised significantly for 0.6 M n-butanol. These data point to there being a transition in the organization of the bilayer near 0.5 M n-butanol, moving from one phase to a condition where two phases appear to coexist in the phospholipid regions. While these data are interesting and suggestive of a structural transition, the theoretical framework for the interpretation of fluorescence lifetime data for such complex chromophores is not developed to the point of 3091

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to the ability of n-butanol to function as an amphiphile and partition efficiently into the lipid bilayer. With the addition of sufficient n-butanol the integrity of the bilayer is compromised, leading to large-scale disruption of the structure. Taken collectively, our data show that the influence of n-butanol on bilayer structure and organization can be used to mediate bilayer organization.

exhibits the same dynamics to within the experimental uncertainty. With these experimental data in mind, we consider them in the context of previous studies on the interaction(s) of lipid bilayers with short-chain alcohols.12,38 A common thread in these studies has been that n-butanol interacts to a greater extent with the phospholipid bilayers than shorter chain alcohols and that these interactions can either serve to mediate order within the bilayer structure38 or disrupt bilayer organization.12 It is also known that the ionic strength of the aqueous overlayer can play a role in controlling the partitioning of n-butanol into bilayers.13 In mixed DOPC/DPPC bilayers, the partitioning of n-butanol was shown to depend on the relative amount of each lipid present and that the partitioning into DOPC- and DPPC-rich regions of the bilayer was not the same.14 This finding is consistent with our data, which suggest that the partitioning of n-butanol with cholesterol domains was different than for DOPC-rich regions of the bilayer. Another unifying theme of these studies is that the interactions of nbutanol with lipid bilayers give rise to morphological changes on the micrometer scale. The work we present here shows that the molecular-scale organization of multicomponent bilayer structures is also affected by the presence of n-butanol and that the amount of n-butanol in the aqueous overlayer provides a means of controlling the organization of the bilayer. The anisotropy decay image data (Figure 4) show that the molecular-scale organizational heterogeneity of the bilayers increases upon exposure to n-butanol, but there is not a clear relationship between the chromophore Dw and the amount of n-butanol present. Fluorescence lifetime imaging data indicate partitioning of the chromophore into two coexisting structural domains at ca. 0.4 M n-butanol. The fact that there is not a discernible difference between the coexisting domains in the anisotropy decay data indicates that it is the dielectric response of the chromophore environment and not the local viscosity that are influenced most strongly by the presence of n-butanol.

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AUTHOR INFORMATION

Corresponding Author

*Tel.: +011 517 355 9715 x224. E-mail: blanchard@chemistry. msu.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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CONCLUSION The data presented here on the interactions of n-butanol with a planar supported lipid bilayer reveal a greater extent of structural disruption of the bilayer than is seen for interactions with ethanol.21 This is not a surprising result owing to the greater amphiphilic character of n-butanol. Fluorescence lifetime imaging data show an n-butanol-concentration-dependent increase in the fluorescence lifetime of SR-DOPE up to ca. 0.4 M n-butanol and the appearance of a bimodal fluorescence lifetime distribution for 0.5 M n-butanol. It is known that multicomponent lipid bilayer structures can exist with more than one phosphocholine domain simultaneously, and our lifetime imaging data are consistent with this condition. In contrast to data on ethanol interactions with model bilayers,21 we do not see a clear correlation between the fluorescence lifetime imaging data and fluorescence anisotropy decay imaging data. Rather, on exposure to n-butanol, there appears to be a broadening of the anisotropy decay distribution, indicative of short-range structural disruption of the bilayer. The interactions between n-butanol and the bilayer appear to influence the dielectric response of the bilayer, but this effect is not manifested clearly in the (rotational) diffusional properties of the chromophore. The steady state imaging data we present show that nbutanol produces a substantial micrometer-scale disruption of the lipid bilayer organization. We attribute this disruptive effect 3092

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