J. Phys. Chem. B 2003, 107, 1251-1257
1251
Nature of the Main Transition of Dipalmitoylphosphocholine Bilayers Inferred from Fluorescence Spectroscopy Antti J. Metso,† Arimatti Jutila,‡ Juha-Pekka Mattila,† Juha M. Holopainen,† and Paavo K. J. Kinnunen*,§ Helsinki Biophysics & Biomembrane Group, Institute of Biomedicine, UniVersity of Helsinki, Helsinki, Finland, and MemphyssCenter for Biomembrane Physics ReceiVed: December 13, 2001; In Final Form: September 25, 2002
The structural dynamics of the main phase transition of large unilamellar dipalmitoylphosphocholine (DPPC) vesicles were studied using differential scanning calorimetry (DSC) as well as steady-state and time-resolved fluorescence spectroscopy. Two membrane-incorporated fluorescent lipid analogues, 1-palmitoyl-2-[10-(pyren1-yl)]decanoyl-sn-glycero-3-phosphocholine (PPDPC) and 1-palmitoyl-2-(3-(diphenylhexatrienyl) propanoyl)sn-glycero-3-phosphocholine (DPHPC), were employed. In gel-state DPPC, the excimer formation times (τR) for PPDPC (mole fraction XPPDPC < 0.02) decrease with increasing temperature up to (T - Tm) ≈ -10 and together with high values for the excimer/monomer emission ratio (Ie/Im) suggest an enrichment of this probe in the clusters. A pronounced prolongation of τR observed with further increases in temperature and starting at (T - Tm) ≈ -10 is accompanied by only a modest decrement in Ie. This behavior is interpreted as with the dispersion of the clusters and the enrichment of PPDPC into the interfacial boundary separating emerging “fluid” domains from the bulk gel phase. Rapidly decreasing excimer intensity, fluorescence quantum yield, and acyl chain order within the temperature range of -4 < (T - Tm) < 0 together with a slight increment in τR suggest the gradual disappearance of the phase boundary upon approaching Tm. Accordingly, our data indicate the possibility that there is an intermediate phase of a strongly fluctuating lattice of fluidlike (“excited”) and gel-like (“ground”) lipids, its formation starting at (T - Tm) ) -4 and being complete at Tm. With further increases in temperature (T - Tm > 0), this intermediate phase would transform into the liquid disordered phase as a second-order process with an increment in trans f gauche isomerization corresponding to approximately one-third of the total transition enthalpy.
Introduction One of the pertinent current questions regarding biomembrane function concerns the 2D ordering of their components.1 A large fraction of all biological membranes are believed to be in a fluid state under physiological conditions. Accordingly, it is of particular importance to establish mechanisms generating the lateral heterogeneity and membrane ordering of this phase. Furthermore, as most cellular functions of eukaryotes apparently take place on membrane surfaces,2 the elucidation of the coupling between the organization and function of biomembranes is clearly of fundamental importance.3 Characteristic of the liquid crystalline state of matter, biomembrane lipids exhibit a range of phases and phase transitions.4 The significance of these properties of lipids is beginning to be recognized, and phospholipid phase transitions are considered to be important in regulating the activities of membrane-associated proteins,1,3,5-7 for instance. Dynamic lateral heterogeneity due to coexisting fluctuating gel and liquid crystalline domains accompanies the main transition of phospholipids.8-11 Upon T f Tm, the intensity of these fluctuations has been suggested to become enhanced and * To whom correspondence should be addressed. E-mail:
[email protected]. Fax: 358-9-19125444. † University of Helsinki. ‡ Present address: Mechanical Engineering and Material Science Department, Duke University, Durham, North Carolina 27708. § Memphys sCenter for Biomembrane Physics.
cause the bending elasticity and both the lateral (area) and transversal compressibilities as well as the heat capacity of the bilayer to have a maximum at Tm.8,10,12-15 The permeability maximum of bilayers and the augmented activity of phospholipases A2 near Tm have been attributed to the length of the phase boundary8,11,16-20 having a maximum at Tm.10 We addressed the molecular-level events of the main phase transition of 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) using fluorescent phospholipid analogues 1-palmitoyl-2[10(pyren-1-yl)]decanoyl-sn-glycero-3-phosphocholine (PPDPC) and 2-(3-(diphenylhexatrienyl) propanoyl)-1-hexadecanoyl-snglycero-3-phosphocholine (DPHPC). Pyrene-labeled lipids, such as PPDPC, incorporated into model membranes form excimers (excited dimers) in a concentration-dependent manner.21,22 In lipid membranes, the efficiency of excimer formation is controlled by both the rate of lateral diffusion and local enrichment.21 In brief, excited pyrene may either relax back to the ground state by emitting at ∼400 nm or collide with a ground-state pyrene so as to yield an excited dimer (excimer). The latter relaxes back to two ground-state pyrenes while emitting at ∼480 nm. In the absence of possible quantum mechanical effects,21 the ratio of excimer and monomer emission intensities (Ie/Im) depends on the collision rate between pyrene moieties. Accordingly, pyrene-labeled lipids have been utilized to measure the rate of lateral diffusion of lipids23 as well as the formation of domains24 and domain boundaries25,26 in membranes. The variation of Ie/Im as a function of the content of the
10.1021/jp0145098 CCC: $25.00 © 2003 American Chemical Society Published on Web 01/10/2003
1252 J. Phys. Chem. B, Vol. 107, No. 5, 2003 probe has been interpreted as the formation of superlattices,27-31 the driving force being steric repulsion between the bulky probe moieties covalently linked to the phospholipid structure. Complementing the Ie/Im data for PPDPC in DPPC, we measured the fluorescence anisotropy for the phospholipid analogue 2-(3(diphenylhexatrienyl)propanoyl)-1-hexadecanoyl-sn-glycero-3phosphocholine (DPHPC). The contained probe 1,6-diphenyl1,3,5-hexatriene (DPH) has been used as such to monitor the acyl chain order of lipid membranes.32 This phospholipid analogue is reported to have an approximately 3-fold preference for the fluid relative to the gel phase, with the orientation of the long axis of the fluorophore moiety of DPHPC being restricted to a position perpendicular to the bilayer plane.33 The fluorescence measurements were complemented by differential scanning calorimetry (DSC), which was used to record transition temperatures and enthalpies. On the basis of the data presented, a detailed mechanism of the phospholipid main phase transition is suggested. Materials and Methods Materials. Hepes and EDTA were purchased from Sigma, DPPC, from Coatsome (Amagasaki, Hyogo, Japan), DPHPC, from Molecular Probes (Eugene, OR), and PPDPC, from K&V Bioware (Espoo, Finland). No impurities in the above lipids were detected by thin-layer chromatography upon examination of the silicic acid-coated plates after iodine staining using chloroform/methanol/water/ammonia (65:20:2:2 by vol) as the eluent. Lipids were dissolved and stored in chloroform without further purification. The concentration of DPPC was determined gravimetrically using a high-precision electrobalance (Cahn Instruments Inc., Cerritos, CA). The concentrations of the labeled lipids were determined spectrometrically from ethanol solutions using a Perkin-Elmer Lambda Bio 40 UV/vis spectrometer (Norwalk, CT). Molar extinction coefficients of 80 000 cm-1 at 356 nm and 42 000 cm-1 at 342 nm were employed for DPHPC and PPDPC, respectively. The buffer used in all experiments was 20 mM hepes, 0.1 mM EDTA, pH 7.0 prepared using freshly deionized Milli RO/Milli Q water (Millipore, Bedford, MA). The pH of the buffer was adjusted to 7.0 with NaOH. Preparation of Liposomes. Lipids were dissolved and mixed in chloroform to obtain the indicated compositions. In steadystate measurements, the fluorescent lipid analogues PPDPC and DPHPC were present at mole fractions of 0.01 and 0.005, respectively. To improve the signal-to-noise ratio and shorten the time required for the measurements, XPPDPC ) 0.02 was used in the time-resolved experiments. After mixing the lipids, the solvent was removed under a gentle stream of nitrogen. The lipid residue was subsequently maintained under reduced pressure for at least 4 h and then hydrated above the main transition temperature of DPPC to yield a lipid concentration of 0.4 mM. The resulting multilamellar vesicles were used as such in the DSC measurements. To obtain unilamellar vesicles, the hydrated lipid mixtures were extruded with a LiposoFast small-volume homogenizer (Avestin, Ottawa, Canada) above the transition temperature. Samples were subjected to 19 passes through one polycarbonate filter (100-nm pore size, Nucleopore, Pleasanton, CA). Both LUVs and MLVs were annealed by taking them through the transition five times by repeated heating and cooling between zero and 60 °C. Minimal exposure of the lipids to light was ensured throughout the procedure. Subsequently, the liposome solution was divided into proper aliquots and diluted with the above buffer for DSC and fluorescence spectroscopy. The final total lipid concentrations used in the
Metso et al. steady-state and time-resolved fluorescence experiments were 25 and 100 µM, respectively. Differential Scanning Calorimetry. The heating scans were recorded using a VP-DSC microcalorimeter (Microcal Inc., Northampton, MA). The heating rate was 30 °/h, and the final lipid concentration in the DSC cell was 0.4 mM. Analyses of individual samples were repeated once to verify reproducibility. The obtained endotherms were analyzed using the routines in the software provided by the manufacturer. Steady-State Fluorescence Spectroscopy. Steady-state fluorescence measurements were carried out with an SLM 4800S (Urbana, IL) spectrofluorometer. The instrument is equipped with two photomultiplier tubes arranged in a T format, thus allowing the simultaneous recording of both the pyrene monomer and excimer emission or alternatively both the vertically and horizontally polarized components of the DPHPC emission and avoiding artifacts due to lamp intensity fluctuations. Excitation wavelengths of 344 and 354 nm were utilized for PPDPC and DPHPC, respectively. A pyrene-monomer emission wavelength of 398 nm was selected by a monochromator whereas a long-pass filter was utilized in the other channel to isolate the excimer emission. Consequently, the intensity readings are relative, and the obtained Ie/Im values do not represent absolute values. A similar setup was used in the anisotropy measurements, where the vertical component I| was detected at 428 nm using a monochromator and the horizontal component I+ was detected using a long-pass filter. Fluorescence anisotropy r is defined as
r ) (I| - I⊥)/(I| + 2I⊥) The readings were corrected using an appropriate G factor.34 Both the excitation and emission bandwidths were 16 and 4 nm for DPHPC and PPDPC, respectively. To improve the signal-to-noise ratio in each measurement, a time average of 15 s was used. The cuvette holder of the spectrofluorometer is equipped with a magnetic stirrer and is thermostated with a circulating waterbath. The average scanning rate was about 3 °/h, and the temperature was monitored continuously with a digital thermometer (Omega HH42, Stamford, CT) placed in a cuvette that was adjacent to the sample cuvette in the holder. Time-Resolved Fluorescence Spectroscopy. Time-resolved fluorescence measurements were performed with a commercially available system (PTI, Ontario, Canada). A train of 500-ps excitation pulses at 337 nm at a repetition rate of 10 Hz was produced by a nitrogen laser. Time-resolved fluorescence intensities of pyrene monomers and excimers were detected at 398 and 480 nm, respectively, by a photomultiplier tube (Hamamatsu, Japan). The minimum lifetime accessible to the instrument is 200 ps. Each intensity decay curve represents an average of five subsequent measurements, and the reproducibility of the essential features was checked with another sample. The decay curves were fitted to a sum of exponentials and analyzed by the nonlinear least-squares method so that a good fit typically produced a reduced χ2 value of 0.9-1.2. The average rate of the temperature increase was ∼2 °/h. The excimer fluorescence lifetimes and rise times were measured every 5 ° when (T - Tm) < -10, every 2 ° when (T - Tm) < -6 or (T - Tm) > 0.7, and every 1 ° when -6 < (T - Tm) < 0.7. The monomer fluorescence decay was monitored at -4 < (T - Tm) < 1 every 0.2 °, every 2 ° at (T - T) > 2, and every 3 ° at (T - T) < -4. The instrument is equipped with a magnetic stirrer and a circulating waterbath to control the temperature. The temperature was measured continuously by a probe
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TABLE 1: Values for Tm (°C) Measured by DSC for Both MLVs and LUVs with Different Lipid Compositionsa vesicle composition
MLV Tm
∆H
LUV Tm
∆H
DPPC DPPC/PPDPC (0.99/0.01) DPPC/PPDPC (0.98/0.02) DPPC/DPHPC (0.995/0.005)
41.3 41.0
27.7 25.8
41.0 40.7
23.1 22.0
40.9
25.4
40.3
19.7
40.5
26.1
40.2
19.5
a The total lipid concentration in the DSC cell was 0.4 mM in 20 mM hepes, 0.1 mM EDTA, pH 7.0. The numbers within parentheses refer to the respective lipid compositions in mole fractions.
immersed in a cuvette adjacent to the sample in the cuvette holder of the spectrofluorometer. Results and Discussion Differential Scanning Calorimetry. The fluorescent probes used in this study represent perturbing substitutional impurities in the DPPC matrix and are thus expected to decrease the Tm of DPPC. Accordingly, we first characterized their effects on the thermal phase behavior of the liposomes by DSC. Because LUVs were used in fluorescence spectroscopy, both MLVs and LUVs were investigated. Compared to MLVs, the corresponding LUVs exhibit a broader endotherm. This broadening is likely to reflect reduced cooperativity and coherence of the membrane due to the lack of interbilayer coupling35 as well as both the increased curvature of the vesicles9 and the smaller size of the LUVs limiting the maximum size of the cooperative unit. The extrusion of MLVs to yield LUVs also decreases the Tm values for both neat DPPC liposomes and those containing the fluorescent probes (Table 1). As observed previously, pretransition is absent for LUVs (Figure 1). The presence of PPDPC (XPPDPC ) 0.01) in DPPC MLVs lowers Tm from 41.3 to 41 °C and also broadens the main transition peak. The enthalpy peak remains asymmetric, with 18.9 kJ/mol at T < Tm and 8.8 kJ/ mol above Tm. For neat DPPC LUVs, the corresponding values are 14.1 kJ/mol at T < Tm and 9.0 kJ/mol above Tm. Broadening is also observed for MLVs containing DPHPC (X ) 0.005), with Tm decreasing to 40.5 °C. The main phase transition of lamellar lipid bilayers is considered to be a pseudocritical, weakly first-order process.36 The decrement in Tm could be partly due to the partitioning of the probes into the interfacial boundary between the coexisting fluid and gel-state domains in the transition region,8-10 as reported earlier for PPDPC in DMPC.25 This would stabilize the interfacial boundary and thus favor the formation of fluid domains at lower temperatures. The decrement in the overall transition enthalpy caused by the probes is in keeping with their chain-disordering effect at T < Tm, thus resulting in a smaller remaining increment for the extent of trans f gauche isomerization upon chain melting in the transition. Steady-State Fluorescence Spectroscopy. For DPPCPPDPC (XPPDPC ) 0.01) LUVs, the values for Ie/Im increase with temperature up to a local maximum at (T - Tm) ≈ -3, followed by a decline and a local minimum about 1 ° above Tm (Figure 2A). When (T - Tm) < -3.4 or (T - Tm) > 1, Ie increases slightly, and Im decreases with temperature (Figure 2B), in keeping with enhanced lateral diffusion. Upon approaching Tm and starting at (T - Tm) ≈ -3, the value for Ie declines steeply whereas at (T - Tm) ≈ 1 a minor increase is evident. Yet, despite the decline of Ie, there is no increase in Im, but a progressive decrease is evident. The decrement in Ie/Im close to Tm (Figure 2B) is thus solely caused by the decline in Ie.
Figure 1. Differential scanning calorimetry (DSC) excess heat capacity scan for neat DPPC LUVs (A) and with XPPDPC ) 0.02 (B). The total lipid concentration was 0.4 mM in 20 mM hepes, 0.1 mM EDTA, pH 7.0. The calibration bars in A and B correspond to 2 and 1 kJ/degree mol-1, respectively. The dotted lines in this and in subsequent figures are intended to guide the eye.
Accordingly, in addition to the augmented thermal de-excitation, the quantum yield for pyrene is diminishing due to other processes. This is also indicated by the slightly enhanced transient reduction in Im in the temperature interval of approximately -5.5 e (T - Tm) e -2.5 (Figure 2B). The fluorescence anisotropy r of DPHPC assessing acyl chain order decreases with temperature, with the midpoint for the decline at about Tm (Figure 3). Time-Resolved Fluorescence Spectroscopy. To obtain a better understanding of the molecular-level processes underlying the above steady-state fluorescence data, pyrene-monomer lifetimes (τM) as well as excimer formation (τR) and decay times (τD) were measured for PPDPC in the vicinity of Tm. The PPDPC-monomer intensity decay is a two-exponential process with the shorter-lifetime component varying between 7 and 16 ns and the longer-lifetime component, between 73 and 119 ns. The fractional intensity of the shorter-lifetime component is maximally only 7%, and it is thus only a minor contributor to the weighted-average lifetime (τjM). Similar to the steady-state measurements for PPDPC in DPPC LUVs, the time-resolved monomer fluorescence at 398 nm shows no significant anomalies in the vicinity of Tm (Figure 4), and an almost steady overall decrement by 15 ns in the pyrene-monomer lifetime is observed when (T - Tm) increases from -4 to 0. Time-resolved excimer fluorescence at 480 nm is characterized by two rise times of ∼10 and 56 ns and a single decay time τD of ∼73 ns. The fractional intensity of the shorter rise time is maximally only about 1% and can thus be neglected when considering the changes in the overall rise time τR (Figure 5A). As expected, the integrated excimer intensity intIe of the time-resolved
1254 J. Phys. Chem. B, Vol. 107, No. 5, 2003
Metso et al.
Figure 4. Weighted-average decay time τjM for monomer emission obtained from time-resolved data for DPPC LUVs with XPPDPC ) 0.02 vs (T - Tm). The total lipid concentration was 100 mM in 20 mM hepes, 0.1 mM EDTA, pH 7.0.
Figure 2. (A) Ratio of pyrene excimer and monomer steady-state emission intensities Ie/Im for DPPC LUVs at XPPDPC ) 0.01 as a function of (T - Tm) and (B) monomer (9) and excimer (b) intensities vs (T - Tm). The total lipid concentration was 25 mM in 20 mM hepes, 0.1 mM EDTA, pH 7.0.
Figure 3. Anisotropy of DPHPC (XDPHPC ) 0.005) in DPPC LUVs as a function of (T - Tm). The total lipid concentration was 25 mM in 20 mM hepes, 0.1 mM EDTA, pH 7.0.
fluorescence emission at 480 nm behaves similarly to the steadystate Ie (Figure 5B). To facilitate the analysis of the data contained in Figures 2-5, we divided the processes into five temperature ranges, viz., I (T - Tm < -10), II (-10 e T - Tm e -4), III (-4 < T - Tm e 0), IV (0 < T - Tm e 3), and V (T - Tm > 3 ), where Tm corresponds to the peak of the endotherm determined by DSC for LUVs with XPPDPC ) 0.02. In the so phase (gel phase) of DPPC, the pyrene-labeled analogue PPDPC has been suggested to form clusters so as to minimize the perturbation that it imparts to the packing of the acyl chains of DPPC.27,30 Our present data support this view. Accordingly, the temperature-induced increase in Ie/Im is steeper at T < Tm than above the transition. Likewise, in the first temperature range (region I), the excimer rise time
τR decreases with temperature (Figure 5A), in keeping with the enhanced collisional rate due to augmented lateral diffusion. In the second temperature interval (region II), the value for the excimer rise time τR is dramatically prolonged from 46 to 60 ns, with only a minor decrement in the integrated intensity of the excimer emission. The kinetics of excimer formation thus must become fundamentally different when entering this temperature range. The value for τD continues to decrease progressively from ∼90 to ∼65 ns (Figure 5C), this accelerated decay of excimer emission being consistent with increased thermal mobility. The above changes are in keeping with the dispersion of PPDPC clusters. We have previously provided evidence for the preferential localization of PPDPC into the phase boundaries between the coexisting gel and fluid phases of DMPC as an edge actant.25 A similar conclusion was reached in our study on the lateral distribution of the negatively charged pyrenelabeled phospholipid analogue, PPDPG in DPPC.26 Our present data are in agreement with the above interpretation. More specifically, assuming the probe to become accommodated into the emerging phase boundaries, the mode of lateral diffusion determining the rate of excimer formation would change from 2D to more 1D. Accordingly, τR reflecting lateral diffusion in the interfacial boundary should increase. Concomitantly, the value for τD continues to decrease in a progressive manner with no drastic changes in monomer decay. In keeping with the enrichment of PPDPC into the interfacial boundary, this would comply with a temperature-induced acceleration in excimer decay. In region II, the area of fluidlike domains would increase with temperature at the expense of gel-like domains. The next temperature range of interest is region III, between -4 e (T - Tm) e 0. In this temperature interval, a modest increase in τR and a minor decrease in τD occur (Figure 5), with a simultaneous, small decrease in monomer emission intensity and a drastic decrement in Ie/Im (Figure 2). This range coincides with a pronounced reduction in membrane acyl chain order signaled by DPHPC anisotropy (Figure 3) as well as a significant fraction (∼65%, 13 kJ/mol) of the transition enthalpy for DPPC LUVs (at XPPCPC ) 0.02) (Figure 1). A possible mechanism for the decline in Ie/Im in temperature range III would be the release of the probe into a 2D lattice from the disappearing domain boundaries with the formation of a time-averaged regular distribution of the probe molecules as a superlattice.30,25 An interesting finding evident from the data (Figures 2 and 5) is that the quantum yield for pyrene emission decreases steeply in region III. More specifically, as the pronounced decrement
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Figure 6. Schematic illustration of the DPPC membrane undergoing the main transition. The depicted values for the mole fractions of DPPC in gel (blue) and fluid (green) states as well as in the boundary (yellow) in the temperature range -10 < (T - Tm) < -4 should be considered as tentative only. The gradual changes in the colors correspond to phase changes, the intermediate phase being depicted by orange. See the text for details.
Figure 5. (A) Excimer emission rise time τR vs (T - Tm), (B) integrated PPDPC excimer emission intensity intIe vs (T - Tm), and (C) PPDPC excimer emission decay time τD vs (T - Tm) calculated from timeresolved data for DPPC LUVs with XPPDPC ) 0.02. The data in B were normalized using the highest intIe value. The excitation wavelength was 337 nm, and emission was recorded at 480 nm. The graphs in A and C were smoothed using Savitzky-Golay filtering with a window of two data points. The total lipid concentration was 100 mM in 20 mM hepes, 0.1 mM EDTA, pH 7.0.
in Ie is not accompanied by a reciprocal change (increase) in Im, we may conclude that the decrease in Ie is mainly caused by a decrease in the fluorescence quantum yield. Despite the release of the pyrene lipid from the vanishing boundaries and the considerable degree of acyl chain disorder, the value for excimer formation time τR increases only modestly with temperature, this increment being significantly smaller than in region II. It is possible that this is caused by a critical slowing of lateral diffusion due to intense fluctuations with a long coherence range, with the probes as substitutional impurities being trapped in the lattice at nodes corresponding to the resonance frequencies of lattice phonons. The present data is thus in keeping with the model by Chong et al.30 and Jutila et al.,25 suggesting the formation of hexagonal superlattices in the course of the main transition as the reason for the sharp dip in Ie/Im. Accordingly, perturbing bulky pyrene moieties of PPDPC become maximally separated from each other to minimize the system’s free energy. In other words, the
formation of superlattices would thus depend on the fine balance between energy minimization due to maximal separation of the bulky pyrene rings and entropy-driven randomization. Chong et al. suggested that at Tm of DMPC hexagonal superlattices and clusters of PPDPC would coexist so that excimer formation would not vanish completely because of the superlattices.30 Yet, for the DPPC matrix with a thicker membrane hydrocarbon region, we do consider it to be more likely that superlattices are time-averaged structures at equilibrium with randomly dispersed PPDPC. To this end, several studies have suggested the involvement of superlattices in the phospholipid main transition.25,28-30. The existence of superlattices in the course of the transition raises a question concerning the photophysics of excimer formation in this structure. Moreover, we need to explain the reduced quantum yield evident from the steady-state data. It is possible that the regular organization of the pyrenes causes longrange interactions between several pyrenes, resulting in their quantum mechanical coupling.31 If several pyrenes moieties were resonating, then the decay would occur at longer wavelengths and would be evident as decreased quantum yield. In other words, it is possible that in the highly phase-coherent lattice the mechanism of excimer formation also involves long-range interactions and the formation of pseudo-oligomers, together with a normal collision-controlled mechanism. Importantly, at T > Tm, no more discontinuities are signaled by the pyrene-labeled probe. The remaining enthalpy of approximately 9 kJ mol-1 and the further increase in DPHPC anisotropy necessitate consideration of the processes in the temperature range Tm < (T - Tm) < 3 (region IV). With increasing temperature, the properties of the coexisting fluid and gel phases approach each other (regions II and III), and fluctuations between the states become more intense. The overall process is schematically illustrated in Figure 6. Our data thus suggest that at the end of the processes occurring in temperature interval III, at Tm only a single homogeneous, intermediate phase would be present (Figure 6). The lack of a phase boundary would readily be alleviated if we assumed that in temperature range IV, 0 < (T - Tm) < 3, the intermediate phase would convert to the liquid disordered phase as a second-order process, with an increase in acyl chain trans f gauche isomerization (i.e., with a further decrease in acyl chain order). This complies with both DPHPC anisotropy measurements and the DSC data, with a peak in enthalpy at Tm (i.e., (T - Tm) ) 0). Upon exceeding Tm, approximately one-third of the transition enthalpy (∼9 kJ/mol) is required to complete the melting of the acyl
1256 J. Phys. Chem. B, Vol. 107, No. 5, 2003 chains in the transition toward the upper temperature limit of region IV (Figure 1). Importantly, although the main endotherm is broadened and the total enthalpy is reduced by approximately 17%, in the presence of PPDPC, a similar asymmetry with respect to Tm is also evident for neat DPPC LUVs. The above mechanism would also comply with results derived from other studies on the phospholipid main phase transition. The minimum at Tm in the lag time preceding the onset of the activity of phospholipase A2 toward phosphatidylcholines has been attributed to a maximum in the length of the boundary.20 Our model implies that the boundary emerging between the coexisting gel and fluid phases is equivalent to the intermediate phase, which has a maximum at Tm, with the entire bilayer becoming a “boundary”. Upon exceeding Tm, the latter would transform into the liquid disordered phase as a second-order process. Although the above model contradicts the conventional view of the phospholipid main phase transition as a first-order process, it readily complies with X-ray data on fully hydrated DPPC. More specifically, whereas the “large-scale” coexistence of gel and fluid phases characteristic of a first-order transition is seen at low water content, X-ray studies on fully hydrated DPPC at low scan rates have revealed the lack of this two-phase region,37,38 the transition progressing as a continuous process. Tenchov et al. further interpreted their data on cooling scans to comply with the presence of an intermediate phase with “smallscale” coexistence. In this context, it is also mandatory to consider observations on phospholipid Langmuir films by fluorescence microscopy and AFM studies on LangmuirBlodgett films and supported bilayers undergoing phase transition, revealing the large-scale coexistence of gel and fluid phases.39-42 In these studies, the macroscopic gel/fluid domains are segregated on a scale extending to tens of micrometers. Importantly, the average diameter of LUV studied here is approximately 100 nm, requiring domains to be significantly smaller. It is possible that the truly macroscopic dimensions of monolayers, Langmuir-Blodgett films, and supported bilayers together with the mica-membrane interactions provide constraints to the transition process so as to alter its characteristics in a fundamental manner. Such a difference is suggested by the contradiction of the data derived from the X-ray diffraction and those from the planar-model membranes. Summary In this study, we have continued to explore the detailed molecular-level processes in phospholipid bilayers undergoing the main transition by using both steady-state and time-resolved fluorescence spectroscopy and by monitoring the signals from two lipid analogues, PPDPC and DPHPC, incorporated in a DPPC membrane. These probes are structurally very close to the matrix lipid, and we may assume that some of the inherent features of the transition are reported by their fluorescence behavior.9 Yet, it must be emphasized that we are observing the transition of an impure DPPC matrix, and in essence, the mechanism that is forwarded applies in a strict sense to the bilayer melting in the presence of the contained probes. In the solid ordered (gel) phase PPDPC seems to be enriched in clusters, which become dispersed at (T - Tm) ≈ -10. Our data further indicate a concomitant enrichment of PPDPC into the emerging interfacial boundary upon the formation of fluid domains within the bulk gel phase, the length of the boundary increasing with the progression of the transition into the fluid state. Upon approaching Tm, the phase boundary characteristic of a first-order transition disappears, and an intermediate phase is formed (Figure 6). The disappearance of the domain boundary
Metso et al. is suggested to result from the properties of the coexisting fluid and gel phases approaching each other as parallel second-order processes developing with temperature, causing diminishing line tension and a hydrophobic mismatch. Accordingly, the energy barrier between the phospholipid molecules in fluid and gel states approaches kT, and the intermediate phase would thus be characterized by intense fluctuations. Our data further suggest that the intermediate phase subsequently transforms into the liquid disordered phase as a second-order transition (Figure 6), with a further increase in trans f gauche isomerization. Upon the completion of this process, the bilayer is in the liquid disordered phase. Abbreviations DMPC, 1,2-dimyristoyl-sn-glycero-3-phosphocholine; DPHPC, 1-palmitoyl-2-(3-(diphenylhexatrienyl) propanoyl)-sn-glycero-3-phosphocholine; DPPC, 1,2-dipalmitoyl-sn-glycero-3phosphocholine; ∆H, enthalpy change; Ie/Im, ratio of pyrene excimer and monomer fluorescence intensity; intIe, integrated excimer intensity of the time-resolved fluorescence emission; k, Boltzmann constant; LUV, large unilamellar vesicle; MLV, multilamellar vesicle; PPDPC, 1-palmitoyl-2[10-(pyren-1-yl)]decanoyl-sn-glycero-3-phosphocholine; SL, superlattice; T, temperature; Tm, main phase-transition temperature; Xlipid, mole fraction of the indicated lipid; τR, rise time (excimer formation time); τD, excimer decay time; τjM weighted-average monomer lifetime. Acknowledgment. We thank Drs. Tim So¨derlund, JuhaMatti Alakoskela, and Roy Siddall for their critical reading of the manuscript and for technical assistance. This study was supported by grants from the Research and Science Foundation of Farmos (A.J.M., J.M.H.), the Paulo Research Foundation, and the Emil Aaltonen Foundation (J.M.H.). Memphys is supported by The Danish National Research Foundation. HBBG is supported by the Tekes and Finnish Academy. References and Notes (1) Mouritsen, O. G.; Kinnunen, P. K. J. In Biological Membranes; Merz, K. M., Jr., Roux, B., Eds.; Birkha¨user Publishing: Boston, MA, 1996; pp 465-504. (2) Kinnunen, P. K. J. Cell Physiol. Biochem. 2000, 10, 243. (3) Kinnunen, P. K. J. Chem. Phys. Lipids 1991, 57, 357. (4) Kinnunen, P. K. J., Laggner, P., Eds. Special Issue of Chem. Phys. Lipids 1991, 57, 109. (5) Kinnunen, P. K. J., Mouritsen, O. G., Eds. Special Issue of Chem. Phys. Lipids 1994, 73, 1. (6) Tocanne, J.-F.; Ce´zanne, L.; Lopez, A.; Piknova, B.; Schram, V.; Tournier, J.-F.; Welby, M. Chem. Phys. Lipids 1994, 73, 139. (7) Epand, R., Ed. Special Issue of Chem. Phys. Lipids 1996, 81, 101. (8) Doniach, S. J. Chem. Phys. 1978, 68, 4912. (9) Marsh, D.; Watts, A.; Knowles, P. F. Biochim. Biophys. Acta 1977, 465, 500. (10) Freire, E.; Biltonen, R. Biochim. Biophys. Acta 1978, 514, 54. (11) Mouritsen, O. G.; Jørgensen, K.; Hønger, T. In Permeability and Stability of Lipid Bilayers; Disalvo, E. A., Simon, S. A., Eds.; CRC Press: Boca Raton, FL, 1995; pp 137-160. (12) Nagle, J. F.; Scott, H. L. Biochim. Biophys. Acta 1978, 513, 236. (13) Evans, E.; Kwok, R. Biochemistry 1982, 21, 4874. (14) Alakoskela, J.-M., Kinnunen, P. K. J. J. Phys. Chem. B 2001, 105, 11294-11301. (15) Bloom, M.; Evans, E.; Mouritsen, O. G. Q. ReV. Biophys. 1991, 24, 293. (16) Papahadjopoulos, D.; Jacobson, K.; Nir, S.; Isac, T. Biochim. Biophys. Acta 1974, 311, 310. (17) Op den Kamp, J. A. F.; Kauertz, M. T.; van Deenen, L. L. M. Biochim. Biophys. Acta 1975, 406, 169. (18) Maynard, V. M.; Magin, R. L.; Dunn, F. Chem. Phys. Lipids 1985, 37, 1. (19) Menashe, M.; Romero, G.; Biltonen, R. L.; Lichtenberg, D. J. Biol. Chem. 1986, 261, 5328.
Phospholipid Main Transition (20) Hønger, T.; Jørgensen, K.; Biltonen, R. L.; Mouritsen, O. G. Biochemistry 1996, 35, 9003. (21) Kinnunen, P. K. J.; Ko˜iv, A.; Mustonen, P. In Fluorescence Spectroscopy; Wolfbeis, O. S., Ed.; Springer-Verlag: Berlin, 1993; pp 159171. (22) Duportail, G.; Lianos, P. In Vesicles; Rosoff, M., Ed.; Marcel Dekker: New York, 1996; pp 296-372. (23) Galla, H.-J.; Theilen, U.; Hartmann, W. Chem. Phys. Lipids 1979, 23, 239. (24) Lehtonen, J. Y. A.; Holopainen, J. M.; Kinnunen, P. K. J. Biophys. J. 1996, 70, 1753. (25) Jutila, A.; Kinnunen, P. K. J. J. Phys. Chem. B 1997, 101, 7635. (26) So¨derlund, T.; Jutila, A.; Kinnunen P. K. J. Biophys. J. 1999, 76, 896. (27) Somerharju, P. J.; Virtanen, J. A.; Eklund, K. K.; Vainio, P.; Kinnunen, P. K. J. Biochemistry 1985, 24, 2773. (28) Tang, D.; Chong, P. L.-G. Biophys. J. 1992, 63, 903. (29) Sugar, I. P.; Tang, D.; Chong, P. L.-G. J. Phys. Chem. B 1994, 98, 7201. (30) Chong, P. L.-G.; Tang, D.; Sugar, I. P. Biophys. J. 1994, 66, 2029.
J. Phys. Chem. B, Vol. 107, No. 5, 2003 1257 (31) Kinnunen, P. K. J.; Tulkki, A.; Lemmetyinen, H.; Paakkola, J.; Virtanen, A. Chem. Phys. Lett. 1987, 136, 539. (32) Lentz, B. R. Chem. Phys. Lipids 1993, 64, 99. (33) Ameloot, M.; Hendrick, H.; Herreman, W.; Pottel, H.; van Cauwelaert, F.; van de Meer, W. Biophys. J. 1984, 46, 525. (34) Parente, R. A.; Lentz, B. R. Biochemistry 1985, 24, 6178. (35) Lotta, T. I.; Laakkonen, L. J.; Virtanen, J. A.; Kinnunen, P. K. J. Chem. Phys. Lipids 1988, 46, 1-12. (36) Mouritsen, O. G. Chem. Phys. Lipids 1991, 57, 179. (37) Tenchov, B. G.; Yao, H.; Hatta, I. Biophys. J. 1989, 56, 757. (38) Caffrey, M.; Fanger, G.; Magin, R. L.; Zhang, J. Biophys. J. 1990, 58, 677. (39) Tokumasu, F.; Jin, A. J.; Dvorak, J. A. J. Electron Microsc. 2002, 51, 1. (40) Nielsen, L. K.; Bjørnholm, T.; Mouritsen, O. G. Nature (London) 2000, 404, 352. (41) McConnell, H. M.; Keller, D. J. Proc. Natl. Acad. Sci. U.S.A. 1987, 84, 4706. (42) von Tscharner, V.; McConnell, H. M. Biophys. J. 1981, 36, 409.
