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* MEMPHYS-Center for Biomembrane Physics, Department of Chemistry, and LiPlasome Pharma A/S, Technical University of Denmark, DK-2800 Lyngby, Denmark; and Department of Physics, University of Southern Denmark, DK-5230 Odense M, Denmark
Correspondence: Address reprint requests to Chad Leidy, MEMPHYS-Center for Biomembrane Physics, Technical University of Denmark, Dept. of Chemistry, Bldg. 206, DK-2800 Lyngby, Denmark. Tel.: 45-4525-2431; Fax: 45-4588-3136; E-mail: cleidy{at}kemi.dtu.dk.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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). PLA2 is interfacially activated, showing higher activity toward lipids that are aggregated, for example into a bilayer lipid membrane structure, than toward lipids that are free in solution (Verger et al., 1973; Waite, 1991). PLA2 plays a prominent role in physiological processes involving lipid membrane modification and generation of bioactive lipids through hydrolysis. These processes include pro-inflammatory response (Nevalainen et al., 2000), neuronal lipid membrane repair and regeneration (Geddis and Rehder, 2003), and generation of the stratum corneum (Mao-Qiang et al., 1995). PLA2 has been suggested to be involved in these and other processes both as a generator of signaling lipids (Murakami and Kudo, 2002), and as a direct modifier of lipid membrane morphology (Brown et al., 2003). In either case, in order for PLA2 to induce the appropriate changes in the lipid membrane without compromising lipid membrane integrity, the enzyme activity needs to be regulated.
Several years of extensive work have shown that the physical properties of the lipid membrane play a major role in modulating PLA2 activity (Kensil and Dennis, 1979; Dennis et al., 1981; Burack and Biltonen, 1994; Hønger et al., 1996; Høyrup et al., 2002). Structural defects and fluctuations that occur near phase transition temperatures are known to increase PLA2 activity (Hønger et al., 1996). These perturbations in the lipid membrane structure are believed to induce packing defects that facilitate access of the enzyme to the cleavage site at the sn-2 position. Additionally, in giant unilamellar vesicles showing gel/fluid coexistence, PLA2 was shown to preferentially bind and hydrolyze the fluid-phase domains (Sanchez et al., 2002).
Curvature also has a strong influence on PLA2 activity. In the case of small unilamellar vesicles the enzyme shows sensitivity to different degrees of curvature (Bell and Biltonen, 1989). Furthermore, the enzyme has also been postulated to present increased sensitivity toward regions of high curvature achieved by product accumulation and phase separation (Burack and Biltonen, 1994; Burack et al., 1997b). Large unilamellar vesicles show under certain conditions a so-called lag-burst behavior where a long lag time of low activity is followed by a period of rapid hydrolysis (Apitz-Castro et al., 1982; Hønger et al., 1996; Nielsen et al., 1999). After this onset, the hydrolysis then occurs in a very rapid manner. The fast hydrolysis after the lag time has been attributed to product accumulation, thereby increasing the interfacial disorder that results in increased access for the enzyme to the cleavage site (Burack et al., 1993). This lag phase before hydrolysis, can be reduced with the exogenous addition of fatty acids and lysophospholipids (Burack and Biltonen, 1994). The fatty acids when ionized can contribute to increased affinity of PLA2 to the lipid membrane surface by electrostatic attraction of the positively charged enzyme. Negative charge is known to increase the affinity of PLA2 toward the membrane. However, the fatty acids produced are not necessarily fully ionized, and it is not clear how much they contribute to increasing the activity of the enzyme (Peters et al., 2000). Lysophospholipids, on the other hand, are known to induce positive lipid membrane curvature, which increases the affinity of PLA2 for the lipid membrane (Henshaw et al., 1998). Vesicles were observed to undergo sharp changes in morphology due to product accumulation, and these changes in morphology were correlated with an increase in PLA2 activity (Burack et al., 1997a). These results suggest that lipid membrane curvature can be an important regulator for PLA2 activity.
In this article we explore the sensitivity of PLA2 toward curved bilayers by studying the activity of PLA2 toward the ripple phase. The two most studied and well-characterized lipid phases are the gel phase and the fluid phase. Both of these lipid phases correspond to locally flat lipid membrane morphologies. The ripple phase, in contrast, has an intriguing surface morphology, characterized by corrugations in the lipid bilayer with a well-defined periodicity (Nagle and Tristram-Nagle, 2000). The origin of the ripple structure is still under investigation. Different models have been presented, attempting to explain the local spontaneous curvature observed in the ripple phase, including an electrostatic coupling between water molecules and the polar lipid headgroups (Doniach, 1979), a coupling between lipid membrane curvature and molecular tilt (Lubensky and Mackintosh, 1993), as well as the generation of curvature by linear arrays of fluid-state lipid molecules (Heimburg, 2000). Two different ripple structures can form depending on the history of the sample (Zasadzinski et al., 1988, Tenchov et al., 1989; Koynova et al., 1996; Katsaras et al., 2000). One is a stable ripple phase (
/2-ripples) with repeat distance around 130–150 Å, and the other is a metastable ripple phase (-ripples) with repeat distance of 260–300Å which is approximately double the spacing as the stable ripple phase. The proportion of each phase depends on the thermal history of the sample, and there is a slow conversion from the metastable to the stable ripple phase. We have previously shown that the ripple phase can be visualized by atomic force microscopy (AFM) in supported double bilayers composed of 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) and DMPC/1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC) mixtures, showing coexisting ripple structures (- and /2 ripples), ripple/fluid phase coexistence, and ripple/gel phase coexistence (Leidy et al., 2002; Kaasgaard et al., 2003). These results have shown that changes in the ripple structure can be carefully monitored by AFM in a time- and temperature-dependent manner.
The ripple phase presents an interesting model for studying lipid membrane curvature effects on PLA2 activity, and it provides the opportunity to visualize directly by AFM the enzymatic hydrolysis of a lipid membrane surface that presents a high degree of local curvature. In the first part of the study a comparison is done between the hydrolysis of the ripple-phase DMPC bilayers compared to the gel-phase DMPC bilayers, testing the expected higher sensitivity of a ripple-phase bilayer toward hydrolysis, due to the preference of PLA2 for curved lipid membranes. In the second part we monitor the hydrolysis of different ripple structures in 1:1 DMPC/DSPC supported double bilayers. The presence of DSPC reduces the rate of hydrolysis, making it possible to monitor in a time-dependent manner the changes in lipid membrane morphology due to both accumulation of products and variation in composition during hydrolysis. The final part of the study addresses evidence showing that hydrolysis of PLA2 occurs preferentially along the direction of the ripples.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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).
Atomic force microscopy
The mica supported lipid bilayers were imaged in contact mode using a PicoSPM atomic force microscope (Molecular Imaging, Phoenix, AZ). The cantilevers were oxide-sharpened silicon nitride cantilevers (ThermoMicroscopes, Sunnyvale, CA) with nominal spring constants of 0.01 N/m. To ensure that the force was kept minimal during scanning, the force was frequently decreased until the tip left the surface and subsequently slightly increased until just regaining contact. In general, ripples could only be resolved when the force was at an absolute minimum. Due to scanner hysteresis and small variations in temperature during scanning, precise statements about the scanning force are difficult to make. Even temperature fluctuations of the order of 10–2°C cause noticeable thermal bending of the gold-coated cantilevers. However, a conservative estimate of the force range would be 20–300 pN based on the nominal spring constant. PLA2 was added to give a total concentration of 
420 nM. The total lipid concentration of the solid supported lipid bilayer system in the AFM fluid cell was 2 µM. The protein to lipid ratio is then 1:5.
Fluorescence spectroscopy
PLA2 hydrolysis was studied in bulk by monitoring the release of a water-soluble fluorescent dye, calcein, encapsulated in DMPC large unilamellar vesicles (200 nm) in a self-quenching concentration (50 mM). Calcein was dissolved in water and pH adjusted with NaOH to pH = 7.5 before adding the calcein solution to the 10 mM HEPES buffer (10 mM HEPES, 1 mM KCl, 1 mM NaN3, 30 µM CaCl2, 10 µM NaEDTA). The lipid suspension was hydrated in the calcein solution as described above. The multilamellar vesicles were extruded 10 times through two stacked 200 nm pore size polycarbonate filters (Whatman, Clifton, NJ) forming large unilamellar vesicles (LUVs) with a narrow size distribution. Untrapped calcein was removed from the liposome suspension by gel filtration through a column packed with Sephadex G-50 using the HEPES buffer as eluent. Fluorescence measurements were performed using a SLM DMX 1100 spectrofluorometer. Excitation and emission channels were set at 490 nm and 515 nm respectively, and fluorescence was measured as a function of time. PLA2 was added after 100 s and the emission fluorescence is recorded after 1000 s. 100% release is obtained by adding 50 µl of 10% Triton X-100. Plots are corrected for passive release of the calcein. The total lipid concentration for each sample was 150 µM and the enzyme concentration used was 150 nM.
Differential scanning calorimetry
DSC was performed using a MicroCal MC-2 (Northhampton, MA) on samples of 50 mM DMPC LUVs at a scan rate of 10°C/ h. An appropriate baseline was substracted from the resulting thermograms.
High-performance liquid chromatography (HPLC)
HPLC quantification of the hydrolysis reaction (Høyrup et al., 2001a) was made using a Waters Millennium 2010 system (Milford, MA) equipped with a Waters 510 pump, a Waters 717 Plus autosampler, and a PL-EMD 950 evaporative light scattering mass detector from Polymer Laboratories (Cheshire, UK) using 5 µm Phenomenex (Torrance, CA) diol spherical column and a mixture of chloroform/methanol/water (73:23:3 v/v) as isocratic mobile phase. 100 µl lipid suspension samples were retrieved directly from the reaction chamber in the fluorometer several times during the PLA2 lipid hydrolysis time course. The extracted lipids were rapidly mixed in chloroform/methanol/acetic acid/water (2:4:1:1) to quench the PLA2 lipid hydrolysis effectively. After quenching the PLA2 activity it was confirmed that the composition of the sample did not change by storage. Salts were extracted from the sample by shaking thoroughly with 1 ml water. From the organic phase 20 µl was analyzed by HPLC. The degree of hydrolysis was measured by the reduction in the substrate fraction. Dose/response calibration curves were linear in the used concentration range.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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-ripples forming in a DMPC supported double bilayer at 17°C. The average ripple spacing is found by Fourier transform to be 312 ± 4 Å. Fig. 1 b shows an image of the same region after addition of PLA2. The scan is taken from bottom to top, and shows a sudden degradation of the lipid membrane as the scanning probe reaches the top one-third of the image. The initial stages of degradation are observed to occur within 5 min after the addition of PLA2. Fig. 1 c shows the same region 9 min after addition of PLA2. The image shows that the ripples have disappeared and the surface has become inhomogeneous, likely due to degradation of the lipid followed by incorporation of fatty acids and lysophospholipids. In Fig. 1 d, a hole is scratched down to the mica surface at a high force setting to corroborate that the lipid membrane has been degraded. The height profile of the hole in Fig. 1 e shows that the height difference between the surface features and the mica is 10 Å, which is much smaller than the expected height for a supported bilayer. For comparison, a control showing the height profile of a DPPC supported double bilayer is presented. Fig. 1 f shows a height profile of a hole that has been scratched on a DPPC double bilayer down to the mica surface. The 90 Å and 60 Å bars correspond to the height profiles of the second and first bilayers respectively. From this it is clear that the 10 Å height difference observed in Fig. 1 e does not correspond to a bilayer. The fact that the surface is not completely cleared during hydrolysis suggests that at least part of the lipids remain attached to the mica, forming rearranged structures. The remaining features on the surface most likely correspond to aggregates of hydrolysis byproducts (i.e., fatty acids, lysophospholipids), and some remaining DMPC. Fig. 1 d also indicates that the bottom bilayer is not present, indicating that the bottom bilayer in the double bilayer system has been degraded along with the top bilayer. In general, the remaining aggregate does not show a defined bilayer structure.
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; Giocondi et al., 2001; Leidy et al., 2002). The lipid membrane therefore maintains a flat morphology characteristic of a gel-phase lipid membrane. The undulations seen in the image are due to noise from the temperature control stage. After addition of PLA2, a much longer time is necessary for the lipid membrane to show signs of degradation. The first evidence of degradation is the formation of a significant number of small protrusions distributed on the surface, with a diameter of 20–40 nm (Fig. 2 b, 65 min), followed by the formation of holes (Fig. 2 c, 122 min) that grow continuously until the lipid membrane is fully degraded. Protrusions are first detected typically 1 h after addition of PLA2, which is significantly longer than the time it takes (5 min) for degradation of the ripple-phase lipid membrane. This long lag time before initial degradation by PLA2 is most likely related to the lag-phase studied intensively both in bulk measurements using liposomes (Apitz-Castro et al., 1982; Hønger et al., 1996; Nielsen et al., 1999) and in supported gel-phase bilayers (Nielsen et al., 1999). The duration of the lag phase in bulk measurements has been correlated with the presence of defects on the bilayer (among other factors such as composition and calcium concentration) (Fernandez et al., 1991; Hønger et al., 1996, 1998). These defects are proposed to provide the enzyme with easier access to the carbonyl backbone of the phospholipid for hydrolysis. Other AFM studies have shown that in the presence of preformed holes no lag phase is observed (Grandbois et al., 1998; Kaasgaard et al., 2001). In these cases, hydrolysis is shown to take place at the hole edges where the surface curvature is high, providing an exposed site for hydrolysis. The formation of protrusions in Fig. 2 b, which seem too large (20–40 nm) to correspond to individual proteins, which have a diameter of 4 nm (Han et al., 1997), are most likely a result of product accumulation, serving as sites for hydrolysis to progress, leading to formation and growth of holes.
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; Leidy et al., 2002). Probing of the DMPC-supported bilayer deposited through vesicle fusion (Fig. 2 a) shows no evidence of holes, which appears to indicate close to complete coverage of the mica. The lack of evident defects is therefore most likely the cause for the delay in the initiation of hydrolysis. On the other hand, when DMPC is in the ripple phase, hydrolysis begins almost immediately after addition of PLA2. The very fast kinetics of hydrolysis observed for DMPC supported double bilayers in the ripple phase, even in the absence of any apparent defects in the lipid membrane (Fig. 1), indicate that the undulated surface provides PLA2 with easier access to the substrate. These observations are also supported by bulk hydrolysis measurements. Fig. 3 a shows fluorescence measurements done on large unilamellar vesicles (200 nm) encapsulating a self-quenching dye. PLA2 activity is measured by monitoring an increase in fluorescence as the dye is released from the liposomes during hydrolysis. Percent release is plotted as a function of temperature. A differential scanning calorimetry trace for DMPC unilamellar vesicles (200 nm) is also plotted showing the gel-phase regime below the pretransition at 14°C, the ripple-phase regime between the pretransition at 14°C and the main phase transition at 24°C, and the fluid-phase regime above the main phase transition at 24°C. Although the highest activity is observed in the fluid-phase regime, a clear increase in activity is observed within the ripple-phase regime compared to the gel-phase regime. The inset graph compares fluorescence traces as a function of time for large unilamellar vesicles in the gel-phase temperature regime at 8°C, and in the ripple-phase temperature regime at 18°C. The sharp initial drop in fluorescence indicates the addition of the enzyme. Fluorescence is then observed to increase immediately after addition of PLA2 for the liposomes in the ripple-phase temperature regime, whereas the liposomes in the gel-phase temperature regime show very little increase in fluorescence in the initial stages.
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/2-ripple structure for DMPC, which has been resolved by x-ray diffraction (Sun et al., 1996). The crystal structure of the snake venom PLA2 enzyme (Han et al., 1997) is placed to scale next to the ripple structure. The illustration shows that the length scale of the ripples is relevant to the length scale of the enzyme. The illustration in Fig. 3 b indicates how the ripple phase can provide better access for PLA2 hydrolysis by exposing the lipid headgroups to the catalytic pocket.
Alternatively, increased PLA2 activity in the ripple phase could be explained by the presence of fluid-phase lipids in the ripple structure. Fluid-phase lipids show higher susceptibility toward hydrolysis than gel-phase lipids (Kaasgaard et al., 2001; Sanchez et al., 2002). A small fraction of fluid-phase lipids has been shown by x-ray diffraction to form at the pretransition temperature (Rappolt et al., 2000). Ripple formation is proposed to result from a rearrangement of this fluid-phase lipid fraction into highly ordered linear arrays, which induce the undulations in the lipid membrane (Heimburg, 2000). According to this model the ripple structure is proposed to be a band pattern of fluid- and gel-phase domains. It has been shown that PLA2 has higher activity in the presence of coexisting fluid- and gel-phase domains (Hønger et al., 1996). The enzyme is believed to act at the interface between these domains, so the activity of PLA2 in the ripple-phase regime could be accentuated by the presence of coexisting fluid- and gel-phase regions in the ripples. Also, a combination of both curvature effects and the presence of fluid-phase lipids at the sites of high curvature could contribute to the increased activity of PLA2 in the ripple-phase regime. The present study cannot distinguish between these two possibilities.
PLA2 hydrolysis induces changes in ripple morphology while preserving bilayer integrity in a two-component DMPC/DSPC supported double bilayer
PLA2 has very low activity toward gel phase DSPC at 24°C, resulting in a slower hydrolysis of DMPC/DSPC mixtures compared to pure DMPC bilayers (Høyrup et al., 2001a). In the following we explore how the more regulated hydrolysis of DMPC/DSPC bilayers leads to a more gradual rearrangement of the lipid membrane structure as seen through changes in ripple morphology for both the stable (/2) and metastable () ripples. This is in contrast with the fast and complete degradation of the one-component DMPC double bilayer shown in Fig. 1. This slower hydrolysis also allows the lipid membrane to preserve a bilayer structure during these morphological changes.
Fig. 4 a shows an AFM image of the /2 ripple phase forming in a 1:1 DMPC/DSPC supported double bilayer at 24°C. The ripple spacing for the DMPC/DSPC mixture is slightly larger (200 Å) compared to the ripple spacing reported for the one-component DMPC sample (150 Å) (Sun et al., 1996). After addition of PLA2 to the 1:1 DMPC/DSPC sample we observe a long time period (35 min) during which no changes in the surface are detected. As mentioned before, this period likely reflects lag time reported for bulk measurements of PLA2 activity (Apitz-Castro et al., 1982), and is expected for DMPC/DSPC mixtures due to the lower activity of the enzyme toward DSPC (Høyrup et al., 2001a). The long lag period with no visual evidence of hydrolysis (Fig. 4 a) is followed by a gradual increase in the ripple spacing (Fig. 4, b–e) until the ripple phase finally dissipates (Fig. 4 f). This result is in contrast to the fast and total degradation of the lipid membrane observed for DMPC in Fig. 1.
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200 Å) the ripple valleys are not reached during scanning, and a noisy ripple profile is observed. Therefore, it is not possible to obtain an accurate measure for the ripple height of the /2-ripple phase using this setup (Kaasgaard et al., 2003). However, the ripple spacing is accurately measured by AFM and a gradual increase in spacing during hydrolysis is clear from the results.
The increase in spacing is followed by dissipation of the ripple phase (Fig. 4 f). However, instead of hydrolysis resulting in complete degradation as observed for DMPC in Fig. 1 c, the lipid membrane continues to maintain a bilayer structure. In Fig. 5 images of the same DMPC/DSPC-supported double bilayers are taken after the ripple phase has dissipated during PLA2 hydrolysis. Fig. 5 a shows remnants of the ripples surrounding a large hole in the lipid membrane. This image corresponds to a zoom-out of Fig. 4 d. In Fig. 5 b, the membrane is completely flat, as the ripples have finally dissipated. The same hole as in Fig. 5 a appears at the bottom of the picture as reference. Fig. 5 b shows the formation of a series of holes in the top part of the image. These holes slowly grow with time, as would be expected for a gel-phase lipid membrane being hydrolyzed by PLA2. The enzyme has not been inactivated so hydrolysis continues. A line profile of Fig. 5 b is shown in Fig. 5 d. The height difference is 90 Å, corresponding to the distance between the second and the first bilayer as reported previously (Leidy et al., 2002). These results are clear indication that the membrane has undergone a structural transformation while preserving the bilayer integrity during hydrolysis.
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-ripple phase. Fig. 6 shows the presence of -ripples in a 1:1 DMPC/DSPC supported double bilayer. The -ripples have approximately double the spacing of the /2 ripples (Koynova et al., 1996), making them easier to resolve by AFM, and therefore making it easier to monitor the changes in the -ripple morphology during PLA2 hydrolysis. Fig. 6, a–d, show a gradual increase in ripple spacing during PLA2 hydrolysis, after a long lag phase (58 min) where no changes occur. This increase in ripple spacing is followed by a drastic collapse of the ripple structure resulting in a flat lipid membrane surface. Lipid membrane integrity is also maintained after ripple collapse (data not shown). Different ripple profiles during PLA2 hydrolysis are shown in Fig. 6, g–i, corresponding to the line profiles shown in Fig. 6, a, c, and e. Fig. 6 i shows the flattened lipid membrane surface after ripple collapse.
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- and /2-ripples. This behavior is plotted in Fig. 7 for both - and /2-ripples. In Fig. 7 a the increase in ripple spacing during PLA2 hydrolysis for the - and /2-ripples is plotted as a function of time. A lag period is observed for both - and /2-ripples before any changes in ripple morphology are detected. This is in contrast to Fig. 1, where complete hydrolysis occurred almost immediately for pure DMPC ripples. The presence of longer-chained lipids has been shown to increase the lag time for hydrolysis (Høyrup et al., 2001a), which would explain the difference. The lag period is followed by a linear increase in ripple spacing for both the /2- and -ripples, showing very similar rates of change 23 Å/min. The spacing is increased to 2.5 times the original spacing of the /2-ripples, before the ripples cannot be detected any more, and 2.7 times the original spacing of the -ripples before collapse of the ripple structure occurs (Fig. 6 e).
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-ripples, making it possible to monitor in a qualitative manner the ripple depth. Measurement of the ripple depth by AFM for the -ripples is plotted in Fig. 7 b as a function of time during PLA2 hydrolysis. The change in ripple depth is also linear, showing a drastic increase of four times the original depth.
Although the surface area cannot be measured accurately, the more than twofold increase in ripple spacing strongly suggests a large increase in lipid membrane surface area. The most likely explanation for the change in ripple morphology and increase in surface area is product accumulation, although a change in lateral membrane pressure during hydrolysis may also provide forces that undermine the ripple structure. Different additives, such as cholesterol, have been shown to increase ripple spacing (Mortensen et al., 1988). Accumulation of fatty acids and lysophospholipids, which are known to alter lipid packing (Weltzien, 1979; Senisterra et al., 1988), are likely to induce an increase in spacing as observed in Fig. 7. Since the calculated lipid concentration in the supported bilayer system is 2 µM, lysolipid and fatty acid partitioning coefficients would suggest that most of the lysolipids and fatty acids go into solution (Høyrup et al., 2001b). However, this is a time-dependent process, which may be slow. Fig. 7 b also shows that after 80 min, and after reaching maximum expansion of the ripple amplitude, the lipid membrane suddenly collapses and flattens. This suggests a drastic reorganization of the lipid membrane, which could be a result of the products of hydrolysis leaving the surface. We therefore suggest that the effects of product accumulation are likely responsible for the initial changes in ripple structure, and the eventual flattening of the membrane could be a result of these products leaving the surface. PLA2 also induces changes in the relative DMPC/DSPC composition due to preferential hydrolysis. The degree of PLA2 hydrolysis of DMPC and DSPC in 1:1 DMPC/DSPC unilamellar vesicles (200 nm) in the ripple-phase regime (24°C) was measured by HPLC chromatography as a function of time (data not shown). The results show that DMPC is preferentially hydrolyzed by PLA2 at this temperature. After 20 min, 80% of the DMPC in the sample was hydrolyzed, whereas DSPC showed no measurable degree of hydrolysis. Since hydrolysis products are expected to eventually leave the surface, the change in relative composition will be the dominating factor in the final state of the supported bilayer. The change in composition induced by PLA2 explains the conversion from a ripple-phase lipid membrane to a gel-phase lipid membrane. DSPC has a high main phase transition 55°C and a high pretransition 52°C. A gradual enrichment of DSPC during hydrolysis will drive the lipid membrane from a ripple-phase regime into a gel-phase regime. In addition, the transition toward a DSPC-enriched composition resulting in a gel-phase lipid membrane reduces the susceptibility of the lipid membrane toward further PLA2 hydrolysis (Høyrup et al., 2001a).
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/2 and -ripples in a two-component DMPC/DSPC supported double bilayer/2-ripples in a 1:1 DMPC/DSPC supported double bilayer as the lipid membrane is hydrolyzed by PLA2. The holes that are forming during hydrolysis stretch parallel to the ripple direction. Fig. 8 b shows the -ripple phase in a 1:1 DMPC/DSPC supported double bilayer, during continual PLA2 hydrolysis, showing that after collapse a hole has formed in parallel to the direction of the ripples. Fig. 8 c shows another example where holes form in parallel to the -ripple phase after the collapse of the ripple structure. The hydrolysis along the ripple direction appears to occur within the ripple valleys (Fig. 8, a and b), indicating that the high curvature regions are susceptible to hydrolysis. It is intriguing that the formation of holes occurs only at the valleys, and not at the cusps. As indicated in Fig. 3 b we suggest that both the ripple cusps and the ripple valleys could show higher susceptibility to hydrolysis. Since the holes are formed at the valleys, it appears that the valley regions are most sensitive to hydrolysis. What is clear from the results is that hole formation occurs along the direction of the ripples, indicating the presence of anisotropic regions (strips) within the ripple structure that show a higher susceptibility to hydrolysis. |
CONCLUSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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, which proposes that the ripples are a result of a nano-structural array of fluid-phase and gel-phase lipids. Fig. 8 shows evidence that holes resulting from hydrolysis form along the ripple direction. The formation of anisotropic holes implies that the ripple structure has regions or strips, running parallel to the ripple direction, that show higher sensitivity to hydrolysis. These holes appear for both - and /2- ripples, making it a general characteristic of the ripple structure. We suggest that these strips that are more sensitive to degradation could be characterized by having either high curvature or fluid-phase lipids. On the other hand, the strip regions that are not hydrolyzed could be resistant due to a high degree of hexagonal headgroup packing, and a decreased access to the ester linkage in the sn-2 position.
In the DMPC/DSPC mixtures hydrolysis is slower, so instead of complete hydrolysis of the lipid membrane we observe a gradual change in lipid membrane morphology that occurs while preserving lipid membrane integrity. In both - and /2-ripple structures an increase in ripple spacing is followed by the collapse of the ripples and the formation of a gel-phase lipid membrane. HPLC results show that DMPC is preferentially hydrolyzed within the ripple-phase temperature regime. DSPC has a main phase transition at 55°C and a pretransition at 52°C, so enrichment in DSPC explains the transformation of the lipid membrane into gel phase. These results provide direct evidence, within a model system, of morphological changes in the lipid membrane induced by PLA2 activity. It is shown that PLA2 activity can be regulated by composition, and that the more gradual hydrolysis observed in the DMPC/DSPC system results in changes in lipid membrane morphology through gradual changes in lipid membrane composition. It is also important to note that these morphological changes occur whereas the lipid membrane still maintains a bilayer structure, which points to the usefulness of PLA2 as a remodeling tool for the lipid membrane. A combination of preferential hydrolysis and the right enzyme kinetics leads to changes in the lipid membrane phase state and the lipid membrane morphology.
In general, these results are clear evidence that PLA2 can induce a compositional phase transition in a lipid membrane through preferential hydrolysis while still preserving lipid membrane integrity. At the cellular level, PLA2 has been implicated in inducing morphological changes in the lipid membrane during cell membrane remodeling (Brown et al., 2003; Geddis and Rehder, 2003). However, the focus of PLA2 activity has been on its role as a generator of signaling molecules, inducing these changes in an indirect manner (Murakami and Kudo, 2002). The current results show that preferential hydrolysis may be a mechanism by which PLA2 can induce changes in membrane morphology and phase state directly, independent of the generation of signaling molecules.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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MEMPHYS-Center for Biomembrane Physics is supported by the Danish National Research Foundation.
Submitted on October 17, 2003; accepted for publication March 31, 2004.
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). Superimposed is a differential scanning calorimetry trace of DMPC unilamellar vesicles (200 nm) showing the pretransition at 
) and metastable (•) ripple phase as measured by Fourier transform; (b) changes in ripple amplitude as a function of time after addition of PLA2 for the metastable ripple phase as measured by averaging height profiles of more than 20 ripples. The change in ripple amplitude for the stable ripple phase is not reported due to the low resolution caused by reduced access of the AFM tip into the ripple valleys.