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Department of Biochemistry and Pharmacy, Åbo Akademi University, FIN 20521 Turku, Finland
Correspondence: Address reprint requests to J. Peter Slotte, Dept. of Biochemistry and Pharmacy, Åbo Akademi University, PO Box 66, FIN 20521 Turku Finland. Tel.: +358-2-2154689; Fax: +358-2-2154010; E-mail: jpslotte{at}abo.fi0.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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; Ohvo-Rekila et al., 2002). It has been suggested that the favorable interactions between SM and cholesterol gives rise to cholesterol and SM rich domains or rafts. The rafts are believed to be important in protein and lipid transport and sorting, as well as in several signaling cascades (Simons and Ikonen, 1997). The nature of the interactions between SM and cholesterol remains unclear. However, it seems that the interactions are stabilized by hydrogen bonding (Sankaram and Thompson, 1990; Bittman et al., 1994; Veiga et al., 2001).
Most of the naturally occurring SMs have the phosphocholine headgroup linked to the hydroxyl group on carbon 1 of a long-chain base (most often an 18-carbon amine diol), and have a long and highly saturated acyl chain linked to the amide group on carbon 2 of the long-chain base (for a review, see Barenholz, 1984). These SMs have the D-erythro-(2S,3R) configuration of the long-chain base (Sarmientos et al., 1985). In cultured cells (i.e., human skin fibroblast and baby hamster kidney cells) 
90–95% of the SMs contain sphingosine (1,3-dihydroxy-2-amino-4-octadecene) as the long-chain base, whereas the remainder have sphinganine (1,3-dihydroxy-2-amino-4-octadecane) as the base (Ramstedt et al., 1999). The latter SMs are also called dihydrosphingomyelins (DHSM). Although it is currently not known why cells need both SMs and DHSMs, it is remarkable that DHSM accounts for 50% of all phospholipids in human lens membranes (Byrdwell and Borchman, 1997).
The physico-chemical properties of DHSM are not nearly as well documented as those of SM, but it is known that the lack of the trans double bond between carbons 4 and 5 in DHSM leads to a higher melting temperature (Tm) compared to acyl-chain matched SM (Barenholz et al., 1976). As a consequence, DHSMs are even more likely than SM to undergo lateral phase separation in bilayer membranes, and may thus contribute to the formation of laterally condensed domains in biomembranes (Brown, 1998; Kuikka et al., 2001). The sphingolipid and cholesterol-rich lipid rafts of cell membranes are known to be resistant to Triton X-100 induced solubilization at low temperatures (Brown and Rose, 1992; Schroeder et al., 1994; London and Brown, 2000). In model membrane systems, we have shown that Triton X-100 partitioning into unilamellar vesicles containing 16:0-SM was described by a lower partition coefficient, and a lower Gibbs free energy as compared to vesicles prepared from DPPC (Nyholm and Slotte, 2001). Partitioning of detergents into vesicles prepared from egg DHSM was also accompanied by lower partition coefficients and lower Gibbs free energies as compared to studies on vesicles made from egg SM (Ollila and Slotte, 2002).
In the present work, we have chosen to characterize both the interfacial properties and the dynamics of the bulk membrane interior of bilayer vesicles containing either D-erythro-N-palmitoyl-sphingomyelin, D-erythro-N-palmitoyl-dihydrosphingomyelin, or 1,2-dipalmitoyl-sn-glycero-3-phosphocholine. The interfacial properties of the bilayer membranes were studied by measuring the spectral shifts of Laurdan and Prodan (Weber and Farris, 1979; Massey and Pownall, 1998; Krasnowska et al., 1998), and by determining the fluorescence lifetime and deuterium isotope exchange of dansyl-phosphatidylethanolamine (Abbott and Nelsestuen, 1987; Epand and Leon, 1992; Asuncion-Punzalan et al., 1998). Information about order in the bulk hydrophobic core of the vesicle membranes was derived from steady-state and time-resolved fluorescence measurements of free and phospholipid-bound diphenylhexatriene (Parente and Lentz, 1985; Williams and Stubbs, 1988; Kalb et al., 1989; Lentz, 1989).
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EXPERIMENTAL PROCEDURES |
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), and purified as described for 16:0-SM. DPPC was purchased from Avanti Polar Lipids. DPH, DPH-PC, DPH propionic acid, Laurdan, Prodan, and dansyl-PE were obtained from Molecular Probes (Leiden, the Netherlands). Deuterium oxide (D2O) was obtained from Aldrich Chemical.
DPH-SM was prepared from D-erythro-lysosphingomyelin (Avanti Polar Lipids) and DPH propionic acid as previously described (Cohen et al., 1984; Ramstedt and Slotte, 1999). The DPH-SM formed was initially purified using bonded phase columns according to Kaluzny and co-workers (Kaluzny et al., 1985), followed by reversed phase chromatography on a LiChrospher 100 RP-18 column (5 µm particle size, 250 x 4 mm column dimensions) using methanol at a flow of 1 ml/min. DPH-DHSM was produced from D-erythro-lysodihydrosphingomyelin and DPH propionic acid as described for DPH-SM. The hydrogenation of D-erythro-lysosphingomyelin was performed as described for 16:0-SM above. Stock solutions of lipids were prepared in hexane:2-propanol (3:2, v/v), stored in the dark at -20°C, and warmed to ambient temperature before use.
Lipid vesicles were prepared in Dulbecco's phosphate buffered saline. In short, the dry lipids and fluorophore were rehydrated at 60°C in phosphate buffered saline (saturated with argon) and sonicated in a Bransonic 2510 (Branson Ultrasonics, CT, USA) bath sonicator for 20 min at 60°C. To protect the fluorophores, the sonication was performed in darkness and in an argon atmosphere. The concentrations of all fluorescent probe solutions were determined from the molar extinction coefficients as provided by Molecular Probes. For DPH-DHSM and DPH-SM solutions, we used the molar extinction coefficients of DPH-PC.
Steady state fluorescence measurements
All steady-state fluorescence experiments presented in this paper were performed on a PTI QuantaMaster 1 (Photon Technology International, Lawrenceville, NJ, USA) spectrofluorimeter operating in the T-format, with both the excitation and emission slits set to 5 nm (unless otherwise specified). The temperature was controlled by a Peltier element, with a temperature probe immersed in the sample solution. All experiments were made in quartz cuvettes and the sample solutions were kept at constant stirring (260 rpm) throughout the measurements. Background fluorescence was measured using the corresponding lipid solutions without fluorophores, and subtracted from the sample spectra. All emission spectra have been corrected to compensate for the wavelength dependence of the detector.
Steady-state fluorescence anisotropy
Steady-state fluorescence anisotropy of DPH-labeled phospholipids was determined in vesicles containing either DPPC, 16:0-SM, or 16:0-DHSM (50 µM) and 1 mol % of DPH-PC, DPH-SM, or DPH-DHSM, respectively. The vesicles were prepared by bath sonication as described above. DPH-labeled phospholipids were included with the bulk lipid before sonication. Excitation was carried out at 360 nm and the emission was recorded at 430 as the temperature was scanned between 20 and 60°C. The steady-state anisotropy, r, is defined as:
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are the fluorescence intensities with the analyzer parallel and perpendicular, respectively, to the vertical polarizer, and G represents the ratio of the sensitivity of the system for vertically and horizontally polarized light (Lakowicz, 1999).
Laurdan emission spectra
Laurdan emission spectra were recorded at 5°C below and above the gel-liquid phase transition of DPPC (36 and 46°C, respectively), 16:0-SM (36 and 46°C) and 16:0-DHSM (43 and 53°C). The total lipid concentration in the cuvette was 100 µM, out of which 1 mol % was Laurdan. Laurdan was included with the lipids before hydration and bath sonication (60°C) of the hydrated lipids. The emission spectra were collected between 390 and 550 nm, whereas the sample was excited at 360 nm. All spectra were recorded as the average of three scans.
Prodan emission spectra
Emission spectra of Prodan were measured in 50 µM phospholipid solutions containing 1 mol % Prodan. Excitation was performed at 360 nm and the emission intensity was recorded between 390 and 590 nm. Vesicles were produced with bath sonication at 60°C. Prodan was added from an ethanolic stock solution (1.7 mM) to the preformed vesicles and the solution was allowed to equilibrate (10 min at 25°C) before the emission spectra were measured at 5°C on both sides of Tm for the bulk phospholipids in the vesicles.
Time-resolved fluorescence measurements
All time-resolved fluorescence measurements were performed on a PTI TimeMaster (Photon Technology International) instrument. The light source in these experiments was an N2 laser. Data analysis was performed with the software (TimeMaster 1.2) supplied by the instrument manufacturers. Goodness of fit was determined from the reduced Chi2, Durbin-Watson, z value, and the weighted residuals.
Lifetimes of free and phospholipid-bound DPH in bilayer membranes
For these experiments, phospholipid vesicles contained either DPPC, 16:0-SM, or 16:0-DHSM (50 µM) and 0.33 mol % of either DPH-PC, DPH-SM, or DPH-DHSM, respectively. When free DPH was included, it was added to 1 mol %. The samples were excited at 337 nm, which is the wavelength generated by the N2 laser, and the emission was measured at 430 nm. The slits were set to 8 nm (except with samples containing free DPH when 12 nm was used). Time-resolved anisotropies were calculated using the following equation
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i. The order parameter, S, and the cone angle, 
C, were calculated from the fundamental anisotropy, r0, and the limiting anisotropy r
(Lakowicz, 1999).
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Dansyl-phosphatidylethanolamine fluorescence lifetimes in phospholipid membranes
Fluorescence lifetimes of dansyl-PE were measured in vesicles containing one of the three phospholipids together with 1 mol % dansyl-PE (total lipid concentration 50 µM). The samples were excited at 337 nm and emission was measured at 516 nm with a slit width of 8 nm. Dansyl-PE emission lifetimes were measured in buffer prepared either in water (H2O) or D2O.
Time-resolved fluorescence of Prodan in phospholipid membranes
Emission lifetimes of Prodan were measured in sonicated vesicles containing 0.33 mol % Prodan (150 µM total lipid concentration). The vesicles were prepared as previously described. The excitation wavelength was set to 360 nm. This wavelength was achieved with a dye laser, containing BPBD-365 (Exciton, Dayton, Ohio, USA) as the laser dye. The emission monochromator was removed and the emission was recorded through a 400CFLP (Melles Griot, Zevenaar, The Netherlands) cut-off filter. To minimize the amount of scattered light reaching the detector the emitted light was also filtered through a polarizer set at a 90° angle relative to the excitation light.
The partition coefficient (Cp) of Prodan between bilayer and water was calculated using a method described by Krasnowska and co-workers (Krasnowska et al., 1998, 2001), where Cp is calculated according to
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Here it is assumed that the lifetime is proportional to the quantum yield, which allows the use of the lifetime of Prodan in water (
W) and membrane (M) in the calculation of RM. RF is the ratio between Prodan fluorescence arising from the membrane and from the aqueous phase. RF is calculated according to
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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). In this study we have done a thorough examination of the interfacial and dynamic properties of acyl-chain defined SMs together with an acyl-chain matched phosphatidylcholine.
The steady-state anisotropy (rss) of DPH-PC, DPH-SM, and DPH-DHSM was recorded as a function of temperature in DPPC, 16:0-SM, and 16:0-DHSM vesicles, respectively. In all three vesicle types the gel to liquid-crystalline phase transition was clearly reported by a rapidly decreasing anisotropy of the DPH-labeled phospholipids close to the transition temperature of the bulk vesicle phospholipids. In the gel phase (between 20 and 32°C), the rss was highest for DPH-PC in DPPC, followed by DPH-DHSM in 16:0-DHSM and DPH-SM in 16:0-SM. Although all three bulk lipids have been shown to go through a pretransition when warmed from their gel phase to their liquid phase (McMullen et al., 1993; Kuikka et al., 2001), only DPH-DHSM in 16:0-DHSM vesicles appeared to report this transition, as evidenced by the increased rss before the main transition (Fig. 1).
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5°C below and above the Tm of the bulk lipid. The results from the experiments with free DPH are summarized in Table 1. In the analysis, the best fits were gained with a model containing two exponential decays, a shorter component with decay time of 2.9 ns (2), and a longer component with a more varied decay time (1). As the temperature was raised above the phase transition temperature, 1 got shorter due to the decreased packing density in the liquid-crystalline phase, whereas the 2 component tended to increase slightly after the transition. Below the phase transition, where the phospholipid membranes were in the gel phase, 1 was longer and appeared to be more sensitive to the composition of the membrane than in the liquid-crystalline phase. 1 for DPH in DPPC membranes was markedly shorter than 1 for DPH in 16:0-SM or 16:0-DHSM membranes. To determine whether a higher degree of H2O access to DPH in DPPC membranes shortened the lifetime, we performed corresponding experiments in D2O (results not shown). Because changing the solvent to D2O had no significant effect on the fluorescence lifetime of free DPH in any of the membrane types (above or below Tm), other factors than water probably induced the shortening of the 1 in DPPC membranes as compared to in SM membranes.
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1 and 2) decreased as the bulk lipid went from a gel phase to a liquid-crystalline phase. The 2 was shorter for all three lipid environments when the DPH was phospholipid-linked as compared with situation of the free probe. The differences in lifetimes among the three different phospholipid-bound DPH probes were much smaller when compared to free DPH. To study whether water affected the lifetime of phospholipid-bound DPH in the different membrane types, we performed corresponding experiments in D2O solution. The phospholipid-bound DPH lifetimes were similar in all membrane types irrespective of whether the membranes were hydrated with D2O or H2O (data not shown).
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), the order parameter (S), the residual anisotropy (r), and the cone angle (C). Measurements were performed 5°C below and above the Tm of the bulk lipid. Raising the temperature above the phase transition temperature of the bulk phospholipids led to decreased order and restriction of molecular motions, as seen from the increased cone angle and decreased order parameter. Of the phospholipids studied, 16:0-DHSM was the most ordered, both in the gel and the liquid-crystalline phase. DPPC bilayers appeared to be slightly more ordered than the 16:0-SM bilayers, both above and below the phase transition.
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; Asuncion-Punzalan et al., 1998), and its lifetime is primarily sensitive to the local environment, to which the major contribution comes from water (Epand and Leon, 1992). In addition, fluorophores with an exchangeable hydrogen, such as dansyl, are known to have a greater quantum yield in D2O relative to H2O due to a reduced rate of proton transfer (Stryer, 1966). An increase in the lifetime of dansyl-PE in D2O compared to in H2O therefore indicates exposure to water. Hence, the D2O/H2O lifetime ratio reflects the properties in the headgroup region of membranes (Ho et al., 1995; Ho and Stubbs, 1997; Saito et al., 2000). All intensity decays recorded with dansyl-PE were best represented by a single exponential fit (data not shown). The D2O/H2O lifetime ratios (Table 4) were all larger than 1, indicating that dansyl-PE was exposed to water in all membrane types. The D2O/H2O lifetime ratio increased as the bulk lipid went from gel phase to liquid-crystalline phase, which is expected inasmuch as the degree of hydration is known to increase as the bilayer melts (Kinnunen, 1991). The change in D2O/H2O lifetime ratio when going from gel to liquid-crystalline phase was similar for DPPC and 16:0-SM, but was smallest for 16:0-DHSM, indicating that the amount of water in the vicinity of the dansyl group was not changed markedly in 16:0-DHSM membrane upon phase transition.
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, 2001). Thus, the relative intensities in the two bands describe the degree to which Prodan partitioned into the membrane. The highest intensity from Prodan in the membrane (blue emission) was seen with 16:0-SM vesicles, whereas the blue emission was markedly reduced for 16:0-DHSM membranes. For DPPC membranes, the blue emission signal was intermediate to the two SMs, and was slightly less blue shifted, indicative of a more polar surrounding in the DPPC membrane compared to the two SM membranes. In the liquid-crystalline phase (5°C above the transition temperature; Fig. 3 B), the emission spectra of Prodan in 16:0-SM and 16:0-DHSM vesicles were superimposable, whereas the emission band from DPPC membranes was more red shifted.
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, 2001). The two measured lifetime components were used to calculate the partition coefficient (Cp) of Prodan partitioning between phospholipid membranes and bulk water. As expected, Prodan partitioned more efficiently into liquid-crystalline membranes than in gel phase membranes (Table 5). When the three phospholipids were compared, it was observed that Prodan partitioned best into 16:0-SM vesicles. 16:0-DHSM membranes were most resistant to Prodan partitioning, whereas DPPC vesicles had intermediate Cp:s.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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). These observations indicate that there are stronger cohesive forces between DHSMs than between SMs in the model membranes, probably largely due to better lateral packing of DHSMs. The finding that DHSM molecules are brought closer to each other compared to SM molecules have been shown to affect the hydrogen bond network in the headgroup region of membranes composed of DHSMs (Ferguson-Yankey et al., 2000; Talbott et al., 2000). The conclusions from the aforementioned studies were that SMs form more stable intramolecular hydrogen bonds than DHSMs, whereas DHSMs tend to form more stable intermolecular hydrogen bonds. The aim of this study was to obtain more information about the membrane properties of DHSMs to deepen the understanding of their role in cellular membranes, and especially in the membranes of human eye lens cells.
Steady-state and time-resolved fluorescence emission from free and phospholipid-bound DPH
DPPC and 16:0-SM have almost the same phase transition temperature, whereas 16:0-DHSM has a phase transition temperature that is 7°C higher (Small, 1986; Kuikka et al., 2001). The properties of phospholipid membranes are to a high degree governed by how the experimental temperature is related to the phase transition temperature. Therefore, we chose to study the properties of the phospholipid bilayers at 5°C on both sides of the phase transition. Hence, the phospholipid membranes were studied both in the gel phase and in the liquid-crystalline phase. The hydrophobic core region of the bilayers was studied by means of two fluorescent probes; free DPH and phospholipid-bound DPH, of which free DPH has been shown to locate slightly closer to surface of the membrane (Kaiser and London, 1998). The lifetime of DPH is extremely sensitive to the dielectric constant of the medium (Fiorini et al., 1987). The extent to which water is allowed to penetrate the membrane is known to affect the lifetime of DPH (Shinitzky and Barenholz, 1978; Barrow and Lentz, 1985; Lentz, 1989; Pap et al., 1994; Bernsdorff et al., 1997). The lifetime of free DPH in DPPC vesicles was shorter in comparison to SM and DHSM membranes (below the phase transition). Because the lifetimes of DPH were similar irrespective of whether the vesicles were in a D2O- or H2O-based solution, it seems that water does not affect the lifetime of DPH. Rather some other membrane property seems to be involved. It was also observed that free DPH had a shorter lifetime in 16:0-DHSM vesicles than in 16:0-SM membranes.
Linking DPH to phospholipids leads to fluorescent probes with different properties than free DPH. For instance, free DPH have been shown to be homogeneously distributed in membranes, whereas the phosphatidylcholine-bound derivative has been shown to partition preferentially into loosely packed membrane regions (Lentz et al., 1976; Andrich and Vanderkooi, 1976; Parente and Lentz, 1985). Similarly the DHSM- and SM-bound DPH has been observed to partition preferentially in fluid membrane regions (unpublished observation; Nyholm and Slotte, 2002). Because the fluorophore replaced an acyl chain in DPPC, 16:0-SM, and 16:0-DHSM, the DPH moiety should have a more defined position in the membrane than the free DPH. Therefore it seems reasonable that these probes should be good reporters of the acyl chain dynamics in these respective bilayer compositions. When phospholipid-bound DPH was used as a reporter molecule in phospholipid bilayers in the gel phase, the lifetimes of DPH in DPPC, 16:0-SM, and 16:0-DHSM were different in relation to those obtained with free DPH. Again we assessed the water access to the probe the different membrane types by comparing the lifetimes measured in H2O- and D2O-based solutions. Inasmuch as phospholipid-bound DPH proved to have similar lifetimes in H2O and D2O in all membranes types, the differences in lifetime could be due to other factors than the presence of various degrees of water.
The amount of cis double bonds in the acyl chains of phospholipids is known to have dramatic effects on the order and dynamics of the membrane. How the trans double bond that is present in the sphingoid base of most natural SMs affects the membrane dynamics has not been that extensively studied. Here we addressed this question by studying the dynamics of phospholipid-bound DPH in 16:0-SM and 16:0-DHSM vesicles. Previous studies have indicated that DHSMs pack more tightly, and have stronger intermolecular interactions than SMs (Talbott et al., 2000; Kuikka et al., 2001). Therefore, it was not surprising that DPH-DHSM in 16:0-DHSM membranes was less mobile than the DPH-SM in 16:0-SM membranes, as seen from the anisotropy decays of the reporter molecules. DPPC, like 16:0-DHSM, is fully saturated, but still DPPC membranes were not as ordered as 16:0-DHSM membranes, and had Tm:s 7 degrees lower than the acyl-chain matched 16:0-DHSM. This could be due to tighter lateral packing of 16:0-DHSM than of DPPC, but also to differences in hydrogen bonding in the two membrane types.
The water exposure of dansyl-phosphatidylethanolamine in the membrane-water interface
To evaluate the role of water in the membrane-water interface, dansyl-PE was included into the phospholipid vesicles. The dansyl moiety of dansyl-PE has been shown to localize within the headgroup region of the bilayer (Asuncion-Punzalan et al., 1998). Hence, the results contain structural information of the phosphocholine headgroups in DPPC, 16:0-SM, and 16:0-DHSM. The same method was used by Saito and co-workers to study how the inclusion of egg yolk SM and DPPC alters the membrane surface structure of egg yolk phosphatidylcholine-triolein emulsions (Saito et al., 2000). They showed that the dansyl moiety was less accessible to water when SM was included in the emulsions than when DPPC was included. They concluded that this probably was an effect of the inter- and intramolecular interactions that are believed to be formed by SMs but not by phosphatidylcholines. Similar conclusions were made by Jendrasiak and Smith (2001), who studied the hydration of SM and phosphatidylcholine monolayers. In our work we found that the degree to which the dansyl group was exposed to water in the three systems studied increased in the order: 16:0-SM < DPPC < 16:0-DHSM. We also found that dansyl-PE had a shorter fluorescence lifetime in DPPC vesicles than in the SM vesicles, which is in agreement with a previous report (Saito et al., 2000). The mechanism for this remains unclear. However, it emphasizes the differences in interfacial properties between SM and PC membranes.
Interfacial differences as revealed by Laurdan emission spectra
In monolayers studies, it has previously been shown that DHSMs had a 100 mV lower surface potential than acyl-chain matched SMs (Kuikka et al., 2001). Data from NMR spectroscopy again indicates that DHSMs and SMs interact differently with water molecules (Talbott et al., 2000). To obtain more information of the properties of the headgroup region of bilayers composed of DHSMs, SMs, and phosphatidylcholines, we measured emission spectra of Laurdan in the different phospholipid environments. The amino-naphtalene-carbonyl fluorescent moiety of Laurdan renders it particularly sensitive to the polarity of the environment (Weber and Farris, 1979; Massey and Pownall, 1998; Krasnowska et al., 1998). In phospholipid membranes, Laurdan monitors relevant differences in the polarity and the different phase states (Zeng and Chong, 1991; Rottenberg, 1992; Parasassi et al., 1994, 1997).
Excitation and emission spectra of Laurdan in DPPC and different SM membranes have already been presented in a number of studies (Bagatolli et al., 1997, 1998, 1999; Massey, 2001). The SMs that have been used in these studies have been natural SMs like egg yolk or bovine brain SM, which contain SMs with several different acyl-chain lengths. In the present study, only D-erythro-N-16:0-DHSM or D-erythro-N-16:0-SM has been used, resulting in a more homogenous system from which the signal is measured. In the gel phase, Laurdan had broader emission spectra in 16:0-SM and 16:0-DHSM membranes than in DPPC membranes. This could indicate a higher degree of heterogeneity in the headgroup region of the SM bilayers than in DPPC bilayers (cf. (Bagatolli et al., 1997)), but it could also derive from excited state reaction (Lakowicz, 1999). At present we cannot determine which explanation is more plausible. The matter is further complicated by the fact that SM and DHSM are both hydrogen donors and acceptors, whereas DPPC and Laurdan are hydrogen acceptors. Thus, hydrogen bonds between Laurdan and SM or DHSM are possible, but not between DPPC and Laurdan. Compared to previously published emission spectra of Laurdan in SM membranes (Bagatolli et al., 1999; Massey, 2001), the spectra we present are broader and more red shifted. However, the cited studies were performed at lower temperatures (23 and 25°C, respectively) compared to the present study. We also recorded emission spectra at 23°C that were more blue shifted and not as broad as at 36°C (results not shown). In the liquid-crystalline phase, the emission spectra of Laurdan were similar in the SM bilayers, whereas it was a little broader in the blue end in DPPC bilayers. This trend has also been shown in other studies (Bagatolli et al., 1999; Massey, 2001).
Membrane partitioning of Prodan
Prodan differs from Laurdan by having a shorter acyl chain (three carbon atoms) compared to Laurdan (12 carbon atoms). Based on this difference, Laurdan and Prodan detect membrane properties at different depths, e.g., the phospholipid polar headgroup pretransition is observable only with Prodan (Krasnowska et al., 1998). In the gel phase, Prodan had a blue-shifted emission spectra in the SM compared to DPPC vesicles, indicating that the probe had a less polar surrounding in SM membranes than in DPPC membranes. Massey showed similar spectra in a report where the Prodan emission in bovine brain SM and DPPC bilayers was compared (Massey, 2001). The emission spectra of Prodan from a DPPC membrane in the liquid-crystalline phase were, as in the gel phase, red shifted in relation to those recorded from comparable SM membranes.
The emission spectra of Prodan are very dependent on the degree of probe partitioning between solvent and the host membrane. It was observed, based on both steady-state and time-resolved measurements, that Prodan partitioned differently between water and the different membranes studied. Prodan partitioning into the highly ordered 16:0-DHSM membranes was less extensive than into e.g., 16:0-SM. This finding is actually expected, because we have shown that detergent partitioning into DHSMs is significantly hindered as compared to the less-ordered SMs (Ollila and Slotte, 2002).
An interesting observation on the fluorescence properties of Prodan was that the fluorescence lifetimes were shorter in SM and DHSM bilayers than in DPPC membranes, although the emission spectra in SM and DHSM membranes were blue shifted compared to the DPPC membranes. Normally a blue shift of the emission spectrum is expected to result in longer lifetimes. It is unclear what caused the unusual behavior of Prodan in SM and DHSM membranes, but we see it as an indication of interfacial differences between PC and SM membranes, similar to those discussed earlier concerning Laurdan emission spectra.
In conclusion, this study with acyl-chain defined phospholipids has clearly shown that DHSMs are more ordered than their counterparts containing the trans 4 double bond in the sphingoid long-chain base. DHSM membranes also resist partitioning of amphiphilic molecules to a greater extent than the SM counterparts (this study and Ollila and Slotte, 2002). The interpretation of the interfacial data was not, however, straightforward. The complications were due to the partially unknown position of the probes in the different membranes, and the possible hydrogen bonds between probes and the SMs. However, the interfacial properties of SM or DHSM membranes differed clearly from the interfacial properties of DPPC membranes, most likely due differences in the hydrogen-bonding properties of the SMs and the glycerophospholipids. The differences between properties of DHSM and SM membranes were not as clear, especially in the liquid-crystalline phase. However, both Laurdan and Prodan reported small differences in the gel phase.
Because of the ordering properties of DHSM in membranes, further studies are warranted to examine the functional role of this particular sphingomyelin species, especially with respect to its possible participation in the formation of raft structures in cell membranes.
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FOOTNOTES |
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Submitted on May 3, 2002; accepted for publication September 26, 2002.
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REFERENCES |
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