Detection of Sphingomyelin Clusters by Raman Spectroscopy Koichiro Shirota,1 Kiyoshi Yagi,2 Takehiko Inaba,1 Pai-Chi Li,2 Michio Murata,3,4 Yuji Sugita,2 and Toshihide Kobayashi1,5,* 1 Lipid Biology Laboratory and 2Theoretical Molecular Science Laboratory, RIKEN, Saitama, Japan; 3Department of Chemistry, Graduate School of Science, Osaka University, Osaka, Japan; 4Lipid Active Structure Project, Japan Science and Technology Agency, ERATO, Osaka, Japan; and 5UMR 7213 CNRS, University of Strasbourg, Illkirch, France
ABSTRACT Sphingomyelin (SM) is a major sphingolipid in mammalian cells that forms specific lipid domains in combination with cholesterol (Chol). Using molecular-dynamics simulation and density functional theory calculation, we identified a characteristic Raman band of SM at ~1643 cm1 as amide I of the SM cluster. Experimental results indicate that this band is sensitive to the hydration of SM and the presence of Chol. We showed that this amide I Raman band can be utilized to examine the membrane distribution of SM. Similarly to SM, ceramide phosphoethanolamine (CerPE) exhibited an amide I Raman band in almost the same region, although CerPE lacks three methyl groups in the phosphocholine moiety of SM. In contrast to SM, the amide I band of CerPE was not affected by Chol, suggesting the importance of the methyl groups of SM in the SM-Chol interaction.
INTRODUCTION Sphingomyelin (SM) is a major sphingolipid in mammalian cells that is involved in membrane signal transduction through the formation of a second messenger (ceramide), as well as through the formation of specific lipid domains (1). The formation of SM-rich domains is affected by the surrounding lipids. Naturally occurring SM is enriched with saturated fatty acids and thus exhibits a high liquid crystalline-gel phase transition temperature (Tc). In other words, SM is rigid at physiological temperature. When egg SM (Tc ¼ ~41 C) is mixed with a fluid lipid such as 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), whose Tc is 22 C (2), egg SM and DOPC are phase separated, resulting in the formation of SM-rich domains. In contrast, egg SM is well mixed with 1,2-dipalmitoyl-sn-glycero-3phosphocholine (DPPC) (Tc ¼ 41 C). Fluorescence microscopy is often employed to study the formation of lipid domains in model membranes, using giant unilamellar vesicles labeled with a fluorescent marker that partitions to specific membrane domains (3). Fluorescence spectroscopy is also used to examine the partitioning of fluorescent dye (4). Fluorescence-based techniques require exogenous fluorescent dye to be present in the membrane, and thus the effect
Submitted February 22, 2016, and accepted for publication July 22, 2016. *Correspondence: [email protected]
or [email protected]
Editor: Arne Gericke. http://dx.doi.org/10.1016/j.bpj.2016.07.035
of the dye on the membrane cannot be excluded. Label-free methods are also employed to study the formation of membrane domains. Differential scanning calorimetry is frequently used to measure phase separation (5). However, because this approach measures the difference in Tc of two lipids, it cannot be applied to study the miscibility of two lipids with the same Tc, such as egg SM and DPPC. Atomic force microscopy is also a powerful tool for studying the miscibility of lipids (6). Because atomic force microscopy measures the height difference of two lipids, it is difficult to use this method to analyze the miscibility of two lipids of similar sizes without a special setup (7). Infrared (IR) and Raman spectroscopy provides information about the structure and dynamics of lipid molecules. Recently, Raman spectroscopy was used to visualize lipid domains containing SM analogs (8). However, Raman spectroscopy has not been applied to study lipid domains composed of naturally occurring SM. The structure of SM is similar to that of the mammalian major phospholipid, phosphatidylcholine (PC). The primary characteristics of SM are the interfacial hydroxyl and amide residues of the molecule. These residues are capable of donating and accepting hydrogen bonds, whereas the carbonyl group of the N-acyl chain and the headgroup phosphodiester moiety may act as a hydrogen bond acceptor. By contrast, PC has only hydrogen bond acceptors at the interface. Based on a molecular-dynamics (MD) study, Mombelli et al. (9) claimed that the sphingosine OH group is mainly
Ó 2016 Biophysical Society.
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MATERIALS AND METHODS
involved in intramolecular hydrogen bonds of SM, in contrast to the almost exclusive intermolecular hydrogen bonds formed by the amide NH moiety. Thus, the amide vibration mode can provide information to distinguish between intra- and intermolecular hydrogen bonds. Fourier transform IR spectroscopy indicates that the addition of cholesterol (Chol) has a strong effect on the amide I band of SM (10,11). In dried SM, the amide I band is split into two peaks. In the presence of Chol, the amide I band is still asymmetric, although the splitting is unresolved (11). It is speculated that the addition of SM-SM intermolecular interactions and the SM-Chol interaction increases the heterogeneity of the environment surrounding the amide group. Raman spectroscopy has been applied to elucidate the membrane properties of SM (12,13). However, the assignment of the Raman spectrum corresponding to intra- and intermolecular hydrogen bonds has not been established. In this study, we assigned the Raman peak responsible for the intermolecular hydrogen bonds of SM by means of Raman spectrum measurements and theoretical calculations. Our results indicate that the intermolecular hydrogen bonds of SM are sensitive to hydration, Chol, and the membrane distribution of SM. Our results also show that, in contrast to SM, the SM analog ceramide phosphoethanolamine (CerPE), which lacks three methyl groups in the phosphocholine moiety of SM, does not form a hydrogen bond with Chol, suggesting the importance of the methyl group of SM in the SM-Chol interaction.
Materials We purchased SM (egg, chicken; egg SM; >80% of the amide-linked fatty acid of egg SM is palmitic acid according to the manufacturer), DPPC, DOPC, N-lauroyl-D-erythro-sphingosylphosphorylcholine (lauroyl SM), N-stearoyl-D-erythro-sphingosylphosphorylcholine (stearoyl SM), and N-palmitoyl-d31-D-erythro-sphingosylphosphorylcholine (d31 SM) from Avanti Polar Lipids (Alabaster, AL); Chol from Sigma (St. Louis, MO); and N-Acyl-sphingosylphosphorylethanolamine (CerPE) from Matreya LLC (Pleasant Gap, PA). 10 -13C-N-stearoyl-SM was prepared as described previously (14). Chloroform and methanol were purchased from Nacalai Tesque (Tokyo, Japan). Deuterium oxide (D2O) was purchased from Acros Organics (Morris, NJ). Glass-bottomed dishes (3911-035) were obtained from AGC Techno Glass (Tokyo, Japan). Fig. 1 summarizes the structures of lipids employed in this study.
Preparation of lipid samples Lipids were dissolved in chloroform, except that CerPE was dissolved in chloroform/methanol (2:1) and aliquoted to glass test tubes to prepare a lipid mixture. Solvent was evaporated under nitrogen flow and then in vacuo for 2 h. MilliQ water (Millipore, Billerica, MA) was added to the resultant lipid films (2 mM total lipids) and the test tubes were warmed up to 60 C and vigorously mixed until the solution turned turbid. The liposome solutions were centrifuged for 25 min at 16,000 g. After removal of the supernatant, the concentrated suspensions were used for measurements. For the D2O experiment, D2O was used instead of H2O. D2O was added to the lipid film and the same procedure was followed. After centrifugation, the floated lipid suspensions were collected. Fig. S1 in the Supporting Material shows a dark-field microscope image of the egg SM suspensions employed in this study. The suspensions exhibit different sizes and shapes, and seem to be multilamellar as deduced from
O P O ONH H
O P O ONH H
egg SM O
O P O ONH H
H OH D DD DD DD DD DD DD D
C stearoyl SM O
O P O O-
O O P O O-
O O H
O P O ONH H
FIGURE 1 Chemical structure of the lipids used in this study.
D DD DD DD DD DD DD D O
d31 palmitoyl SM
O O O H
O P O O-
H H H H
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O P O ONH H
Detection of Sphingomyelin Clusters the strong light scattering. Dynamic light scattering of the suspension solutions performed with a Zetasizer Nano S (Malvern Instruments, Malvern, UK) resulted in a high polydispersity (polydispersity index ¼ 1–0.9) with a variety of liposome sizes, including a diameter greater than the detection limit of the apparatus (3 mm), consistent with the microscope image.
Raman data acquisition Raman spectra were obtained with a laboratory-made laser Raman microscope based on an inverted microscope (IX71; Olympus, Tokyo, Japan). This microscope employed a 532 nm laser (Sapphire SF-532; Coherent, Santa Clara, CA) for excitation and an oil-immersion objective lens (UPLSAPO 60XO, NA ¼ 1.35; Olympus). Through the objective lens, the excitation laser beam was focused on the sample. Then, the backscattered Raman radiation was collected with the same objective lens and passed through sharp-edge filters to block Rayleigh scattering and focused on the entrance slit of a spectrograph (SP2300; Princeton Instruments, Acton, MA). The signal light was dispersed by a 1200 lines/mm grating and finally detected by a thermoelectrically cooled charge-couple device (Pixsus 400BR-eXcelon; Princeton Instruments, Trenton, NJ) as a Raman spectrum. The sample suspension (2–10 mL) was deposited on a 35 mm glass-bottomed dish and placed on the sample stage of the microscope. All spectra were obtained at room temperature (23 C). The laser power at the sample and the exposure time were typically 10 mW and 60 s, respectively. All measurements were recorded with the charge-couple device software WinSpec (Princeton Instruments) and processed with MATLAB (The MathWorks, Natick, MA).
Simulation Details of the computational procedure have been given elsewhere (15). An MD simulation of the SM bilayer was conducted using an all-atom force field for SM (16). After equilibration of a system consisting of 128 SM and 5012 water molecules, the trajectory was propagated for 150 ns with constant temperature (296 K) and pressure (1 bar). The temperature was controlled using the Langevin thermostat with a coupling time of 1 ps1. A semi-isotropic pressure coupling scheme was used, and the pressure was controlled using the Nose´-Hoover Langevin piston method (17–19) with the oscillation period time and the damping time set to 0.05 ps and 25 fs, respectively. A periodic boundary condition was used in all directions. The electrostatic interaction was treated using the smooth particlemesh Ewald scheme (20,21). The nonbonding interaction was decreased ˚ and 12 A ˚ by employing a switching function. The to zero between 10 A neighbor list was updated every 20 fs. The bond length involving hydrogen atoms was restrained by the SHAKE method (22) and the time step for integration was set to 2 fs. The MD calculation was performed using NAMD (23). We confirmed that the thickness and the area per lipid of the SM bilayer were kept stable during the simulation, and that the mean values ˚ and 47.5 A ˚ 2, respectively) were in good agreement with the exper(43.5 A ˚ and 55 A ˚ 2) (24). imental values (42.2 A From the last 100 ns trajectory, we first searched for a cluster of SM and water molecules formed by hydrogen bonds. Then, we classified the clusters according to type (e.g., SM þ water, SM þ two waters, or SM dimer). For a cluster type with an existence weight greater than 1%, we selected a representative structure by using the k-means clustering algorithm, and performed a density function theory (DFT) calculation using the B3LYP exchange-correlation functionals (25,26) and mixed basis sets of 6-31G(d,p) and 6-31þþG(d,p) (27,28). The latter basis set with diffuse functions was used for atoms involved in the hydrogen bonds (i.e., hydrogen, nitrogen, and oxygen atoms of the amino group and water). The combined basis set is denoted 6-31(þþ)G(d,p) in the following text. All of the DFT calculations were carried out using Gaussian 09 (29). Although the MD simulation based on the force field treated all atoms of the SM molecule explicitly, the DFT calculation was computationally much more intensive. Therefore,
we used a fragment of the SM molecule in the DFT calculation, removing the headgroup and most of the acyl chains, and retaining only the atoms in the vicinity of the amide group and the C-C double bond that were essential for this study. The size of the fragment was carefully validated in a preliminary calculation (see Fig. S2 and Table S1, as well as data in (15)). The geometry was optimized starting from the MD snapshot of the clusters. Then, the harmonic frequency and the Raman activity were calculated for each cluster. The frequencies were corrected by
ucorr ¼ f ðu þ aÞ;
where a ¼ 16 and 3 cm1, and f ¼ 0.976 and 0.981 for the C-C and C-O double-bond stretching modes, respectively. The shift parameter (a) accounts for the size of the fragment and basis set, and the scaling factor (f) incorporates the anharmonic effects. The scaling factors were derived from an anharmonic vibrational structure calculation on the SM monomer. The final Raman spectrum was obtained by a weight average of the spectrum of each cluster. The convoluted spectrum was obtained by using the Lorentz function with a full width at half-maximum of 15 cm1.
RESULTS AND DISCUSSION Hydration-dependent alteration of the Raman spectra of SM Fig. 2 A shows the Raman spectra of egg SM suspensions at 23 C. The bottom and top spectra were measured 1 h and 25 h, respectively, after sample deposition on a glassbottomed dish. The two spectra are almost identical except for the peak at 1643 cm1 (arrow). A spectrum similar to that of the 25 h sample was obtained previously (30). However, the origin of the 1643 cm1 band has not been well characterized. Tentative assignments of the main Raman bands of egg SM (after 1 h) are listed in Table 1. Fig. 2 B shows the changes in the Raman spectra of egg SM in the region of 1600–1720 cm1 from 1 to 48 h after sample deposition. A shoulder that was seen in the spectrum 1 h after sample deposition gradually evolved into a sharp band at ~1643 cm1. Because the sample was left on the glassbottomed dish throughout the measurement process, this change is most likely due to water evaporation. To confirm the influence of water, we added 2 mL of water to the same volume of 1-day-dried suspension (100 mM egg SM) and recorded the temporal changes in the Raman spectrum (Fig. 2 C). The band at 1643 cm1 that existed in the 1-day-dried sample became the shoulder within 15 min after the addition of water and reappeared as the band within 30 min as the water evaporated. Water evaporation in our experiment was confirmed by the OH stretching band at ~3400 cm1 (Fig. S3). The band at ~1643 cm1 did not depend on the excitation laser intensity, as shown in Fig. S4. Thus, the Raman analysis of egg SM revealed that its spectrum has a hydration-sensitive band at ~1643 cm1. In addition to the 1643 cm1 band, the 830 and 850 cm1 bands also correlated with the water content, as shown in Fig. S5. The molecular origin of these bands has not yet been characterized. In this study, we primarily examined the 1643 cm1 band.
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1h 6h 25 h 48 h
dry t = 0 min t = 15 min t = 30 min
Raman Shift (cm-1)
Raman Shift (cm-1)
Raman Shift (cm-1)
E a. Lauroyl SM
b. Egg SM / D2O
c. D31 palmitoyl SM
d. 13C stearoyl SM
e. Stearoyl SM
f. Egg SM
Raman Shift (cm-1)
TABLE 1 Assignment of the Main Raman Bands for Egg SM Shown in Fig. 2 A
1676 1643 1445 1301 1135 1097 1067 961 896 881 769 721
Raman Shift (cm-1)
The origin of the Raman band of SM at ~1643 cm1 is not established. Levin et al. (12) pointed out that the broad feature centered at ~1644 cm1 reflects the amide I band associated with amide linkage between the acyl chain and amino group of sphingosine base. In contrast, Lamba et al. (13) claimed that a weak feature at ~1644 cm1 was the HOH deformation mode of water molecules involved in the bilayer hydrogen-bonding network. In Fig. 2, D and E,
Raman shift (cm1)
FIGURE 2 Time variations of the Raman spectrum of egg SM suspension and a spectral comparison of different SM suspensions. All samples were deposited on the glass-bottomed dish and measured at 23 C. (A) Raman spectra collected 1 h (bottom) and 25 h after sample deposition (top). (B) Changes in Raman spectra of egg SM in the region between 1600 and 1720 cm1. Lipid suspensions were deposited on a glass-bottomed dish from 1 h to 48 h at 23 C (blue, 1 h; red, 6 h; green, 25 h; black, 48 h). Measurements were made on 10 mL of a 40 mM lipid suspension. (C) Changes in Raman spectra in the region between 1600 and 1720 cm1 after addition of water (2 mL) to a 1-day-dried lipid suspension: before addition of water (red), elapsed time t ¼ 0 (blue), t ¼ 15 min (green), and t ¼ 30 min (black). Measurements were made on 2 mL of a 100 mM lipid suspension. (D) Raman spectra of (a) lauroyl SM, (b) egg SM prepared with D2O, (c) d31 palmitoyl SM, (d) 13C stearoyl SM, (e) stearoyl SM, and (f) egg SM. (E) Raman spectra of the 1560–1720 cm1 region for (a) lauroyl SM, (b) egg SM with D2O, (c) d31 palmitoyl SM, (d) 13C stearoyl SM, (e) stearoyl SM, and (f) egg SM. All spectra in (E) and (F) were recorded after the same time interval from sample deposition.
Assignments C¼C stretching (12,13) Amide I band (12) CH2 deformation (12,30,43) CH2 twist (13,30) C-C stretching (trans) (30,44,45) C-C stretching (gauche) (30,44,45) C-C stretching (trans) (30,44,45) CN asymetric stretching (46) acyl C1-C2 streching (trans) (47) acyl C1-C2 streching (gauche) þ choline deformation (47) O-P-O symmetric streching (45) C-N symmetric stretching of the choline group (46,48)
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the Raman spectra of different SM samples are compared. All of these spectra were recorded after the same time interval from sample deposition. Measuring the Raman spectrum of egg SM in D2O instead of H2O did not alter the 1643 cm1 band, indicating that the HOH deformation mode of water is not responsible for this band. SMs with different fatty acid compositions (lauroyl (C12:0) SM, stearoyl (C18:0) SM, and egg SM (mainly C16:0)) exhibited the 1643 cm1 band. This observation supports the idea that the acyl chain length does not affect the orientation and position of NH- and OH- groups at the lipid-water interface, and consequently the formation of intermolecular hydrogen bonds and the assignment of the 1643 cm1 band. D31-palmitoyl SM did not alter the 1643 cm1 band, indicating that the fatty acid residue of SM is not involved in this band. Raman spectra were measured at 23 C. At this temperature, egg SM and stearoyl SM are in the gel phase, whereas lauroyl SM is in the liquid crystalline phase (31). Fig. 2 E indicates that the 1643 cm1 band was not affected by the phase state of SM. In contrast to stearoyl SM, 10 -13C-Nstearoyl-SM exhibited a Raman shift of the 1643 cm1 band to 1603 cm1. The wavenumber ratio of the 13C¼O and 12C¼O bands (1603/1643 ¼ 0.976) is consistent with the value calculated from reduced masses (0.977), and the
Detection of Sphingomyelin Clusters
To further investigate the physical origin of this band, we conducted MD simulations and DFT calculations for the SM bilayer to obtain the weight of SM clusters in an isothermal-isobaric (NPT) ensemble formed by the intermolecular hydrogen bonds of the amide group. We then calculated the harmonic frequency and Raman activity for each cluster at the level of B3LYP/6-31(þþ)G(d,p) and corrected the frequencies according to Eq. 1. We note that both the level of the DFT calculation and the correction scheme were carefully examined to obtain the frequency with quantitative accuracy. For example, the C¼O stretching frequency was obtained as 1774 cm1 for a single SM fragment at the B3LYP/6-31G(d,p) level. Adding the diffuse functions lowered the frequency by 30 cm1, and the correction mainly for the anharmonicity further lowered the frequency by 40 cm1. Consequently, the scheme used here yielded a C¼O stretching frequency of 1706 cm1 for a single SM fragment. See Table S1 and (15) for more details. Fig. 3 A plots the calculated Raman shift of the C¼O stretching mode for isolated SM and SM clusters of the amide groups terminated by water molecules (denoted w(SM)nw). The C¼O stretching frequency is strongly dependent on the cluster type. The formation of hydrogen bonds between the amide group and water molecules induces a sizable red shift of the C¼O stretching by 36 cm1. Furthermore, the extension of the hydrogen bond network (i.e., the amide chain) exhibits a red shift of 9.6 and 5.3 cm1 when the value of n is varied from 1 to 2 and from 2 to 3. The hydrogen bond of the amide groups gives rise to a band at ~1650 cm1, as shown in Fig. 3 B. The band is mainly composed of the C¼O stretching modes of wSMw and w(SM)2w, although that of the larger cluster is hardly visible due to its small probability. The C¼C stretching mode gives a rather strong band at 1685 cm1. The calculated spectrum is in good agreement with the observed spectrum 1 h after sample deposition (Fig. 2 B, 1 h, blue). Therefore, the two bands observed at 1672 and 1642 cm1 are assigned to the C¼C stretching modes and the C¼O stretching modes of the hydrogen-bonded amide group, respectively. The C¼O stretching mode in the amide group predominantly contributes to the amide I band (33). Consequently, these results support the findings obtained using Raman spectroscopy. In essence, the characteristic shift in the position of the amide I band observed in the Raman spectra of SM reflects the degree of intermolecular hydrogen bonding of SM.
C=O 1700 1680 1660 1640 SM wSMw w(SM)2w w(SM)3w
Cluster type B
Simulation of the Raman band of SM at ~1643 cm1
Raman shift (cm-1)
isotope shift was reported as ~40 cm1 (32). This strongly supports the idea that 1643 cm1 band reflects the amide I band associated with amide linkage between the acyl chain and amino group of sphingosine base.
Convoluted C=C str SM dimer SM monomer
Experiment 1 h dried sample
Raman shift (cm-1) FIGURE 3 (A) The Raman shift of the C¼O stretching modes of w(SM)nw (n ¼ 1–3) clusters and isolated SM was calculated at the level of B3LYP with mixed 6-31G(d,p) and 6-31þþG(d,p) basis sets, using the correction scheme in Eq. 1. The Raman band that exhibits the highest Raman activity is shown for n ¼ 2 and 3. (B) Comparison of theoretical and experimental Raman spectra. The experimental spectrum is the same curve shown in Fig. 2 B (1 h, blue).
Chol inhibits hydrogen bonding between SM molecules A preferential interaction between SM and Chol was reported previously (34). This preference is caused not only by the interaction between the steroid ring of Chol and the saturated acyl chains of the SM but also by the hydrogen bond between the 3-hydroxyl group of Chol and the amide group of SM (34). A previous study using Fourier transform IR showed that the amide I band at ~1650 cm1 in dried egg SM was altered by the presence of Chol (35). We measured the effect of adding Chol to egg SM on the Raman band at ~1643 cm1 (Fig. 4). The presence of Chol abolished the band at 1643 cm1 even in the mixture of SM/Chol ¼ 1:0.2. It is noteworthy that above this concentration of Chol, the sharp gel-to-liquid crystalline transition of
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Chol-rich area suddenly forms a network that extends over the entire SM bilayer. These results suggest that ~20% Chol is sufficient to interact with whole SM molecules in the membrane, and interaction with Chol disrupts the intermolecular hydrogen bonding of SM molecules.
SM/Chol 1/0 1/0.1 1/0.2 1/0.5 1/1
The amide I Raman band reflects the presence of SM clusters in the membrane
We next asked whether the characteristic 1643 cm1 band of SM derived from intermolecular hydrogen bonding could be utilized to study the membrane distribution of SM. Egg SM has a gel-to-liquid crystalline phase transition temperature (Tc) at ~41 C and mixes well with DPPC (Tc ¼ 41 C), but not with DOPC (Tc ¼ 22 C) (2). Thus, in an egg SM/DOPC mixture, the lipids are phase separated and SM is present in the form of clusters (38). By contrast, palmitoylsphingomyelin is dispersed in DPPC (39). Fig. 5 A shows the Raman spectra in the 600–1800 cm1 region for egg SM, egg SM/DOPC (1:1), and egg SM/ DPPC, DOPC, and DPPC. The 1741 cm1 band in the DOPC and DPPC spectra was assigned to the C¼O stretching mode of the saturated aliphatic ester group, and the 1659 cm1 band in the DOPC was assigned to the cis C¼C stretching mode in the hydrocarbon chains. The
Raman Shift (cm ) FIGURE 4 Raman spectra in the region of 1600–1720 cm1 for egg SM (red) and four mixtures of egg SM/Chol (1:0.1 (blue), 1:0.2 (black dotted), 1:0.5 (black dashed), and 1:1 (black solid)). The Raman spectrum was collected 25 h after sample deposition at 23 C.
SM is no longer detectable in differential scanning calorimetry (36). Through Monte Carlo calculations using interaction energies deduced from calorimetric results, Snyder and Freire (37) previously showed that at ~20% Chol, the A
1700 1680 1660 1640 1620 -1
Raman Shift (cm )
Raman Shift (cm )
D SM/DOPC trans CC amide I
SM/DPPC trans CC amide I
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FIGURE 5 Raman spectra of various lipid suspensions. (A) Raman spectra of egg SM, egg SM/DOPC (1:1), egg SM/DPPC (1:1), DOPC, and DPPC. (B) Raman spectra of the 1620– 1720 cm1 region for egg SM (red), egg SM/ DOPC (1:1) (blue), and egg SM/DPPC (1:1) (black). (C) Concentration dependence of the relative intensities of the trans C¼C stretching band (circle) and the amide I band (square) for a 1:1 mixture of egg SM and DOPC. (D) Concentration dependence of the relative intensities of the trans C¼C stretching band (circle) and the amide I band (square) for a 1:1 mixture of egg SM and DPPC. The Raman spectrum was collected 25 h after sample deposition at 23 C.
Detection of Sphingomyelin Clusters
amide I region of egg SM and the egg SM/DOPC (1:1) mixture is enlarged in Fig. 5 B, whereas its existence is unclear in the egg SM/DPPC (1:1) mixture. As shown in Fig. 2 B, egg SM shows the amide I (C¼O stretching) band at 1644 cm1. The shoulder of amide I clearly exists in the egg SM/DOPC mixture, whereas its existence is unclear in the egg SM/DPPC mixture. The relative intensities of the egg SM trans C¼C stretching band at ~1675 cm1 and the amide I band at ~1643 cm1 for egg SM/DOPC and egg SM/DPPC with different SM/PC ratios are shown in Fig. 5, C and D, respectively. Whereas the intensity of the trans C¼C stretching band decreased linearly with a decreasing SM/PC ratio in both egg SM/DOPC and egg SM/DPPC suspensions, the decrease of the amide I band was much less sensitive to the SM concentration in egg SM/DOPC suspensions. By contrast, the addition of DPPC dramatically reduced the intensity of the amide I band of egg SM. These results indicate that the characteristic amide I Raman band reflects the clustering of SM. The amide I band of CerPE is insensitive to Chol The sphingolipid composition differs among different species. Although SM is the major sphingolipid in mammalian cells, the major sphingolipid in Drosophila is CerPE. In CerPE, the phosphocholine residue of SM is replaced with phosphoethanolamine, resulting in a stronger intermolecular interaction compared with SM, as indicated by its higher phase transition temperature (40). CerPE shows almost the same Raman spectrum in the region of 1620–1700 cm1 as SM, as shown in Fig. 6. We next asked whether the amide I band of CerPE at ~1641 cm1 is sensitive to the addition of other lipids. Similarly to SM, a CerPE/DOPC (1:1) mixture exhibited a shoulder at ~1641 cm1, which was abolished in CerPE/DPPC (1:1).
These results indicate that CerPE forms clusters in DOPC and is dispersed in DPPC. However, in contrast to egg SM/Chol, the CerPE/Chol (1:1) membrane clearly shows an amide I band. Fig. 6 shows the immiscibility of CerPE and Chol. This result is consistent with a previous observation that CerPE failed to form sterol-rich domains (40). A previous MD simulation (41) indicated that CerPE has more hydrogen bonds with Chol than with SM. However, the overall decrease in the number of hydrogen bonds is compensated for by an increasing number of charge pairs between SM and Chol, which leads to an increase in the total number of intermolecular interactions. In addition, the small headgroup of CerPE results in a strong CerPE-CerPE interaction that excludes Chol. Furthermore, the small headgroup of CerPE results in less effective shielding of the sterol molecule from unfavorable interactions with water. Drosophila contains very low levels of sterols compared with mammalian cells (42). Our results suggest that the sphingolipid domains in Drosophila and mammalian cells have different lipid compositions and physical properties. In conclusion, we identified the Raman amide I band at ~1643 cm1 of SM, which reflects the intermolecular hydrogen bond of SM clusters. The amide I band is sensitive to the hydration of lipids and the presence of Chol. We showed that this Raman band can be utilized as a simple marker to examine the membrane distribution of SM in model systems. However, the application of this method in cell systems may be hindered by the amide I band of proteins. We also showed that the interaction of SM and Chol is dependent on the methyl groups of the phosphocholine moiety of the molecule. SUPPORTING MATERIAL Six figures and one table are available at http://www.biophysj.org/biophysj/ supplemental/S0006-3495(16)30615-4.
AUTHOR CONTRIBUTIONS T.K. conceived and coordinated the project. K.S. performed Raman spectroscopy and data analysis. T.I. prepared lipids for measurement. K.Y., P-C.L., and Y.S. performed the simulation. M.M. provided the 10 -13C-Nstearoyl-SM. K.S., K.Y., Y.S., and T.K. wrote the manuscript.
ACKNOWLEDGMENTS We are grateful to Prof. Hiroshi Takahashi of Gunma University for his valuable comments.
FIGURE 6 Raman spectra in the region of 1620–1700 cm1 for CerPE (red) and three mixtures of CerPE/DOPC (1:1) (blue), CerPE /DPPC (1:1) (green), and CerPE/Chol (1:1) (black). The Raman spectrum was collected 25 h after sample deposition at 23 C.
This work was supported by the Integrated Lipidology Program of RIKEN, Grants-in Aid for Scientific Research (25293015 to T.K. and 26620141 to K.S.) from the Japan Society for the Promotion of Science (JSPS), the Center of Innovation Program of the Japan Science and Technology Agency (JST), and the Naito Foundation. The computational resources of the HPCI system were provided by the University of Nagoya through the HPCI System Research Project (Project ID: hp140105), the RIKEN Integrated Cluster of Clusters (RICC), and MEXT SPIRE Supercomputational Life Science (SCLS).
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