Investigation of Model Cell Membranes with Raman Spectroscopy: A

In the Laboratory

Investigation of Model Cell Membranes with Raman Spectroscopy

W

A Biochemistry Laboratory Experiment Norman C. Craig,* William H. Fuchsman, and Nanette N. Lacuesta Department of Chemistry, Oberlin College, Oberlin, OH 44074-1083; *[email protected]

A renaissance in Raman spectroscopy has occurred in the last decade. New developments make Raman spectroscopy more feasible for the study of biochemical systems and for applications in instructional laboratories. This article reports new developments in Raman spectroscopy and describes the use of Raman spectroscopy in the undergraduate biochemistry laboratory for the investigation of temperature-dependent changes in the hydrocarbon interiors of bilayers prepared by dispersing phospholipids in water. Because phospholipids are the dominant matrix material of cell membranes, such dispersions are models of cell membranes and are examples of molecular self-assembly. The experiments described here are outgrowths of research done in Levin’s laboratory at the National Institutes of Health (1, 2) and by other workers in the field (3). An alternative way of studying the effect of temperature on phospholipid dispersions is with differential scanning calorimetry (DSC). Recently, Ohline and coworkers described a suite of undergraduate experiments of this type (4). Their paper provides a fuller discussion of the biophysics of model membranes than is supplied here. Before turning to a more detailed description of the biochemical system of interest, as well as a description of the experimental arrangements, we review two important developments in instrumentation that minimize or eliminate unwanted fluorescence in Raman spectroscopy. We also recall the essentials of the Raman effect. New Developments in Instrumentation for Raman Spectroscopy The introduction of Fourier transform (FT) Raman spectroscopy, led by Chase at Dupont (5), has eliminated interfering electronic fluorescence. The exciting laser light comes from a Nd:Yglass1 laser producing light in the near-infrared (IR) region at 1064 nm. An FT-IR spectrometer detects the Raman-shifted light. Excitation with light of 1064 nm avoids the problem of fluorescence as a result of electronic transitions, which often accompanies the investigation of biologically interesting samples, especially those from natural sources, when visible light excitation is used.2 Because the intensity of the Raman effect decreases with the inverse fourth power of the wavelength, the use of near-infrared light excitation produces a Raman signal that is about 1兾16 as intense as the signal produced by visible light excitation with the green line at 514.5 nm of an argon-ion laser. The light processing efficiency of the FT-IR spectrometer helps compensate for the reduced intensity of the Raman effect with 1064-nm excitation. Levin and Lewis describe the application of FT-Raman spectroscopy to studies of phospholipids (6). 1282

The other important development in Raman spectroscopy involves dispersive spectroscopy and a red laser. Using a krypton-ion laser with a wavelength of 752 nm in the deep red or a solid-state red laser producing a comparable wavelength avoids most fluorescence excitation and yet gives a larger Raman intensity than is obtained with neodyniumbased near-infrared lasers. A simple monochromator contains a holographic transmission grating, and the detector is a charge-coupled device, CCD (7 ). The CCD, a multichannel device, is the electronic equivalent of a photographic film but with much greater convenience and processing flexibility. Both of the new types of Raman techniques take advantage of holographic notch filters that efficiently reject the intense, unshifted light from the exciting laser without appreciably decreasing the wanted Raman signals. Although the experiment described here uses the FT method, the dispersive method with red laser excitation is an attractive alternative. Essentials of Raman Spectroscopy Like IR spectroscopy the Raman effect reports on the stretching and bending of chemical bonds, but the Raman effect is caused by a roundabout process involving two separate photons in an inelastic scattering process. The Raman scattering, observed at 90⬚ or 180⬚ to the direction of the exciting laser beam, is inelastic because energy states in the molecular material change during the Raman event. Thus, the Raman effect is quite different from the direct one-photon absorption process observed in the IR region, and the Raman effect depends on different molecular characteristics for its occurrence. For an IR transition to occur, the vibrational motion must be accompanied by a change in dipole moment. For a Raman transition to occur, the molecular vibration must be accompanied by a change in polarizability of the electron cloud within the molecule. Polarizability reflects the tightness with which the electron cloud is held by the molecule and can change in size and direction as a vibration occurs. The difference in the mechanisms by which IR absorption takes place and the Raman effect occurs results in these two kinds of vibrational spectroscopies being nearly complementary in intensities. Thus, bond vibrations that contribute strong intensities to IR spectra are typically weak contributors to Raman spectra and vice versa. Vibrations of polar bonds, such as O⫺H, N⫺H, C⫺O, C⫺F, P⫺O, or C⫽O bonds, usually give strong IR bands and weak Raman bands. Vibrations of nonpolar bonds, such as C⫺H, C⫺C, or C⫽C bonds, and of bonds involving larger atoms such as sulfur or chlorine with loosely held electrons usually give strong Raman bands and weaker IR bands.

Journal of Chemical Education • Vol. 80 No. 11 November 2003 • JChemEd.chem.wisc.edu

In the Laboratory

Rayleigh scattering

Raman anti-Stokes

Raman Spectroscopy and Biochemical Systems With the fluorescence problem of Raman spectroscopy largely eliminated by excitation with long-wavelength light, the use of water, a natural choice for biochemical systems, as a solvent for Raman spectroscopy is secured. Because of the vibrations of its polar O⫺H bonds, water is a strong absorber of light in the mid-IR region, and water systems are difficult to study by IR spectroscopy. In contrast, water is a relatively weak Raman scatterer, and thus water is a good solvent for Raman spectroscopy. A typical phospholipid component of cell membranes is dipalmitoylphosphatidylcholine (DPPC). The structure of this molecule is shown in Figure 2. The long hydrocarbon chains are the hydrophobic part of the molecule. The rest of the molecule is the hydrophilic polar head group. The bilayers form the shells of vesicles, called liposomes, by self-assembly

IR absorption

A

H3C

CH3 CH3

N+

B

CH2

water

H2C

virtual state

O–

O

P O O H2 CH2 C O CH

h νin > h νout'

C H2C

−+

−+

−+

−+

−+

−+

−+

Raman Stokes

simple example of this effect, we consider the cis and trans isomers of 1,2-dichloroethylene (8). For the cis isomer, a band for the symmetric stretching of the C⫺Cl bonds appears in the Raman spectrum with good intensity at 711 cm᎑1. For the trans isomer, the band for symmetric C⫺Cl stretching appears strongly in the Raman spectrum at 654 cm᎑1. Because of the center of symmetry in the trans-1,2-dichloroethylene molecule, a transition for antisymmetric C⫺Cl stretching has no accompanying polarizability change and thus is not allowed in the Raman spectrum. For the cis isomer, though allowed, the band for antisymmetric C⫺Cl stretching is very weak in the Raman spectrum. Thus, relative intensities of the two bands in the C⫺Cl stretching region of a Raman spectrum of a mixture of cis and trans 1,2-dichloroethylene would tell the relative quantities of the two isomers. As a second example, the bands due principally to C⫺Cl stretching in the Raman spectrum of 1,2dichloroethane tell how the dynamic composition of gauche and trans rotamers changes with temperature (8).

−+

Whereas IR spectra result from the interaction of a continuous spectral range of IR photons of different frequencies with molecular material, Raman spectra are excited by a single-frequency laser line. A whole spectrum of Raman transitions, typically 3500–100 cm᎑1, is produced by the action on the sample of a single frequency of light. The roundabout two-photon process and the accompanying “virtual state” of the Raman effect are illustrated for a single net vibrational excitation on the far left of the energy diagram in Figure 1. This net vibrational excitation is called Raman Stokes. The single exciting photon is the same for each Raman transition, but the scattered photons have characteristic energies for the different net vibrational excitations. The most likely process in the Raman experiment is, however, Rayleigh scattering, as shown second from the left in Figure 1. In Rayleigh scattering, which is elastic, the scattered photon has the same frequency as the incident photon. As indicated in Figure 1, Rayleigh scattering is about 103 more likely than Raman scattering. (A holographic filter removes the intense Rayleigh line of the exciting light from the scattered light.) The incident photon may also induce net vibrational deexcitation in a process called Raman anti-Stokes. This type of Raman process, which is shown third on the energy diagram in Figure 1, depends on prior excitation of vibrational modes. The antiStokes bands are weaker than the Stokes bands because the populations of excited vibrational modes are very small at room temperature except for very low-frequency modes. For comparison with the Raman effect, a direct, singlephoton, IR transition is shown on the right side of Figure 1. Absorption of a photon in the infrared region is about 108 more probable than inelastic scattering due to the Raman effect. Raman spectra, typically as the Stokes part only, are presented as the difference between the frequency of the exciting line and the Raman shifted light and thus appear on a frequency scale comparable to IR spectra. Intensities (and frequencies) of bands in vibrational spectra are sensitive to the conformations of molecules. As a

O O

C

O

CH2 H2C

h νin = h νout

CH2

H2C CH2 H2C

CH2

H2C CH2 H2C

h νin < h νout''

CH2

H2C CH2 H2C

CH2

H2C CH2 H2C

CH2

−+

−+

h νIR

CH2

H2C

water

CH2 H2C

ground state

−+

CH2 H2C

−+

H2C

vib. state

CH2

H3C H3C

rel. intensity

~10ⴚ8

~10ⴚ5

ⴚ8 < ~ 10

1

Figure 1. Basics of photon–molecule interactions in Raman spectroscopy and infrared spectroscopy.

Figure 2. (A) Structure of DPPC and (B) a schematic cross-section of a phospholipid bilayer.

JChemEd.chem.wisc.edu • Vol. 80 No. 11 November 2003 • Journal of Chemical Education

1283

In the Laboratory

in water. As shown to the right in Figure 2, the hydrocarbon chains of the two layers point inward and interact favorably with each other. The polar head groups interact strongly with themselves and with water molecules on the inside and outside of each shell. The shells are thought to surround one another in water-spaced layers in loose onion-skin fashion. The dominant Raman signal comes from the nonpolar hydrocarbon interiors of the bilayers. The polar bonds in the head groups contribute little to the Raman spectrum. The strongest Raman effect for phospholipids comes from the C⫺H stretching region in the vicinity of 2900 cm᎑1, as shown in Figure 3 for a DPPC dispersion at 15 ⬚C.3 The relative intensities of various features in the C⫺H stretching region are sensitive to the conformation of the hydrocarbon chains. Frequencies also depend on conformation, but these frequency shifts will not be used in this experiment. The Raman spectrum of the C⫺H stretching region of a dispersion of DPPC at 15 ⬚C is shown in Figure 4A while the same sample at ∼65 ⬚C is shown in Figure 4B. The spectra shown in Figure 4 are accumulations of 200 scans at a resolution of 8 cm᎑1. The signal-to-noise ratio for a single scan is too low for a useful determination of the intensity ratios. The scaled sum of 200 scans, accumulated in about 5 min, has a signalto-noise improvement of about 14 (= √200). Watching the spectra improve on the computer screen with successive scans, students see how signal accumulation enhances signal-to-noise ratios. A second region containing conformationally sensitive intensity ratios is the C⫺C stretching region around 1100 cm᎑1, for which three bands are seen in Figure 3. However, the peaks in this region are weaker than the peaks in the C⫺H stretching region and would require the accumulation of many more spectra for adequate signal-to-noise ratios. An example of the spectral contribution from the polar head groups is the weak band owing to the carbonyl stretching mode at about 1735 cm᎑1 in Figure 3. At low temperatures the phospholipid dispersions are gellike. The interior hydrocarbon chains are largely in the uniform zigzag, trans arrangement characteristic of solid paraffins. At higher temperatures the phospholipid dispersions are more fluid or cream-like. They are regarded as liquid crystalline at higher temperatures, and the hydrocarbon interiors are largely in random coils as in the liquid phase of the pure hydrocarbon. As the temperature rises, the transition in the dispersion from the gel phase at lower temperature to the liquid-crystalline phase at higher temperature is relatively sharp (4, 9). During the transition, the largely trans configurations of the “frozen” hydrocarbon chains increasingly become gauche configurations of the “melted” hydrocarbon chains. Consequently, intensities (and frequencies) of various C⫺H stretching modes change during the transition. Although the hydrocarbon interior of the bilayer changes structure during the transition, the overall bilayer structure is retained. The transition temperature rises with an increase in length of the hydrocarbon chain of the phospholipid. Thus, it is a good experimental design to have different student pairs in a laboratory section experiment with dispersions of different phospholipids. Although the H2O of a phospholipid dispersion does not make a large contribution to the Raman spectrum, the tail of an H2O absorption band overlaps the 1064-nm line of the exciting laser. In addition, an H2O overtone absorption 1284

Figure 3. Spectrum of a DPPC dispersion in D2O at 15 ⬚C: 400 scans at 8 cm-1 resolution and white light corrected.

A

B

Figure 4. C⫺H stretching region of a DPPC dispersion in D2O: (A) at ~15 °C; (B) at ~65 °C; 200 scans at 8 cm-1 resolution.


In the Laboratory

Sample Preparation Dispersions are made by vortexing a small test tube containing 0.03 g of phospholipid6 and 60 µL of D2O. Capillaries are standard hard glass melting point capillaries (1.5–1.8-mm diameter, 50-mm length). The warmed dispersion is transferred with a plastic syringe into a capillary in the creamy, liquid-crystalline state and compacted in the capillary by centrifuging. Heated test tubes are used to prepare dispersions, to warm syringes prior to transferring dispersions into a capillary, and to warm capillaries prior to centrifuging.

200 scans at 8 cm᎑1 resolution, which gives adequate resolution for these liquid-phase spectra and a reasonable signalto-noise ratio. Omnic and Excel software packages are used on a Compaq Deskpro computer. Other Raman spectrometers may be employed for this experiment. Many observations have been made with conventional, dispersive spectrometers and argon-ion excitation (2). Dedicated FT-Raman spectrometers are available from several instrument manufacturers. These instruments are less expensive than the versatile IR-Raman instrument used here. The least expensive option is undergoing rapid commercial development. For example, Ocean Optics supplies a system with a solid-state, 500-W red laser (785 nm), a monochromator, a notch filter, and a CCD detector for about $15,000. This system has a high-frequency cutoff of 2700 cm᎑1 and a 15-cm᎑1 resolution. Whereas the Ocean Optics instrument is not useful for the C⫺H stretching region, as described here, this instrument is likely to be useful for a comparable study of the C⫺C stretching region. For the temperature study of the phospholipid dispersions, a variable-temperature sample holder is needed for which the temperature can be set quickly below and above room temperature. Because samples are contained in melting point capillaries, a small thermoelectric cooling and heating element works well. When the current flows in one direction through the thermoelectric device, the temperature rises on one face and drops on the other face. When the current flows in the opposite direction, the first plate has the lower temperature. The design of the sample holder is shown in Figure 5. The thermoelectric cooler element is sandwiched between the aluminum block, which holds the sample, and a finned heat sink made of aluminum.7 A small fan is attached to the heat sink unit. Thermoconductive grease containing zinc oxide provides good thermal contact between the thermoelectric cooling element and the aluminum block and the heat sink. The aluminum sample holder has a hole drilled vertically to fit capillaries snugly and a wedge-shaped window to allow collection of the scattered light over a reasonable angular range. For monitoring the temperature of the sample-holder block, a diode temperature sensor is embedded in the block. Because some local heating of the dispersion in the capillary occurs, we found it necessary to use a

Instrumentation For the experiments described here, the Raman method is excitation with a 1064-nm Nd:YVO4 laser, InGaAs (indium gallium arsenide) detection, and Fourier transform analysis of the scattered light. The particular instrument is a Nicolet Magna 760 infrared bench with a CaF2 beamsplitter and an associated Raman Accessory Module. The Raman module is set for the high optical efficiency of 180⬚ observation of Raman scattering. The laser is focused on the sample capillary, and the power is about 0.5 W at the sample. Aligning samples of phospholipid dispersions in the FT-Raman instrument is impractical because of their weak Raman scattering. We optimize the position of a capillary containing polycrystalline diphenylacetylene (Aldrich), a strong Raman scatterer, in the sample holder used for the study of phospholipid dispersions. The red beam of the helium–neon laser coming from the IR module makes this alignment particularly easy with the Nicolet system. Spectra are recorded with

Figure 5. Sample holder and thermoelectric cooler and heater. The aluminum block is 3.4 x 3.4 x 1.0 cm.

band in the IR overlaps the C⫺H stretching region of a phospholipid in the Raman-shifted spectrum. Because of this band, H2O absorbs the scattered Raman photons and weakens the observed spectrum in this region. Thus, if H2O is used in the experiment, the strength of the Raman effect is weakened, and the sample warms owing to absorption of laser light. A good solution to this problem is using D2O (6). The spectra in Figures 3 and 4 are dispersions in D2O.4,5 Experimental Considerations Sample preparation in a melting point capillary is done as a supplement to another experiment in a prior laboratory session, and the sample, sealed with Parafilm, is stored at room temperature. A week later, a three-hour laboratory session is devoted to recording and analyzing the spectra. Different student pairs work with diacylphosphatidylcholines based on fatty acids with chains of C14, C16, C18, C20, and C22, and the data are shared. Of course, phospholipids with an odd number of carbon atoms in the acyl chains may be studied, but samples of the biologically uncommon, odd-numbered species are more expensive than samples of the phospholipids with even-numbered acyl chains. In exploratory experiments temperature profiles were run for the C15 and C17 phospholipids. The laboratory writeup of the experiment, including details of the experimental methods but excluding the introduction, is supplied the Supplemental Material.W Specifics of apparatus used in sample preparation and of the construction of the variable temperature sample holder are given in the laboratory writeup.


1285

In the Laboratory 1.0 0.9

C16

C14

C20

C18

0.8

C22

0.7

Ishoulder Ipeak

tiny thermocouple in the dispersion to measure the local temperature. For insulation, the aluminum block and thermoelectric cooler are clad in fiberboard. An adjustable, constant current supply is needed for the thermoelectric cooler. The size of the current determines the magnitude of cooling or heating of the sample holder. With a current of 2.0 A in one direction the temperature was about 4 °C, and with 2.0 A in the opposite direction the temperature was about 100 ⬚C. The diode sensor is used in setting approximate temperatures. The reading of the copper–constantan thermocouple with its reference junction in an ice bath is used to compute the actual temperature from the measured emf. Sensitivity to at least 0.01 mV is needed.

0.6 0.5 0.4 0.3

0.0

Experiment In a laboratory session of three hours, a pair of students can make observations at about 12 temperatures. Typically, the students start with an observation at room temperature, move to a temperature below room temperature, unless the transition temperature for their phospholipid is expected to be well above room temperature, and then raise the temperature in increments of about 7 ⬚C. This survey gives a good determination of the plateau regions below and above the transition temperature and an approximate determination of the region of the transition. The students then make measurements with careful temperature control to map out the sharp transition region. Hazards The laser light from a Nd:Yglass laser is doubly hazardous. Not only does this IR light cause eye damage, but its invisibility makes its beam unnoticed by the human eye. Thus, the Raman module, as is true of the Nicolet Raman Accessory, must have secure interlocks that cause blocking of the laser light whenever the hatch cover of the Raman module is open. Leads for the power supply, its fan, and the temperature sensors must pass out of the Raman module. Special care must be exercised to be sure no laser light gets out of this port. In contrast to the near-IR laser light, the light from the helium–neon laser beam used to track the movement of the interferometer in the IR module and to aid in alignment in the Raman module is not hazardous unless it is viewed directly down the beam. Results Intensities are found with the aid of the Omnic software. Two expansions of the spectra in the C⫺H stretching region of a DPPC dispersion that are used to measure relative intensities of two temperature-sensitive features are shown in Figure 4. The upper spectrum shows the dispersion at ∼15 ⬚C and the lower spectrum shows the dispersion at ∼65 ⬚C. The two features are the shoulder at about 2930 and the peak at about 2885 cm᎑1. A local baseline spanning the C⫺H stretching region is found with the software and used in the measurement of the intensities at peak maxima. As can be seen in Figure 4, the shoulder and the peak change in frequency during the transition. For DPPC the peak moves from ∼2882 to ∼2892 cm᎑1 as the temperature rises. The higher 1286

0

10

20

30

40

50

60

70

80

90

100

Temperature / °C Figure 6. Temperature profiles of a series of diacylphosphatidylcholines dispersed in D2O.

frequency shoulder moves from ∼2933 to ∼2927 cm᎑1. These shifts in frequency are ignored in the analysis. The ratio Ishoulder兾Ipeak is plotted versus temperature with the aid of Excel software. Students also use the Omnic software to plot two representative spectra, such as the spectrum shown in Figure 3, one from the low-temperature plateau region and one from the high-temperature plateau region. The temperature profiles measured by students for a series of diacylphosphatidylcholines where the fatty acid moiety ranges from the C14 chain of myristic acid to the C22 chain of behenic acid are shown in Figure 6. These are all saturated acids. The transition temperature rises as the length of the fatty acid chain increases. Local bumps in the temperature profiles reflect noise in the intensity measurements and are not significant. The student results for transition temperatures, which are midpoints in the transition regions, for the even-numbered C14 – C22 acyl chains are compared with published values in Table 1. The transition temperatures observed by students in the Raman experiment are consistently somewhat lower than the reported values. This difference undoubtedly reflects some local warming by the laser beam, not reflected in the temperature measured by the thermocouple, which is located just outside the point of incidence of the laser beam. Table 1 includes observed transition temperatures for two phospholipids with odd-numbered acyl chains, C15 and C17. Future Investigations Phospholipid systems other than ones containing only saturated fatty acid moieties have been studied by the Raman method (6). For the biochemistry laboratory experiment, we have not experimented with phospholipids containing unsaturated fatty acid components because of concern that deterioration might occur during storage periods. Another direction for elaboration of this experiment is to work with dispersions of phospholipids with different head groups such as ones containing ethanolamine. We have obtained a satis-


In the Laboratory

Table 1. Transition Temperatures for a Series of Diacylphosphatidylcholines Acyl Chain

Chain Length

Obs. Transition Temp./⬚C

Dimyristoyl

C14

22.5

23.9a, 23.5b, 24.0c

Dipentadecanoyl

C15

31.0

34.7a, 34.0b

Dipalmitoyl

C16

40.5

41.4a, 41.4b, 41.5c

Diheptadecanoyl

C17

47.5

49.8a, 48.2b

Distearoyl

C18

53.0

55.3a, 55.1b, 55.0c

Diarachidoyl

C20

61.5

66.4a, 64.5b

Dibehenoyl

C22

70.0

74.0b

a

Reference 10.

b

Reference 11.

Lit. Transition Temp./⬚C

c

Reference 4.

factory temperature profile of dipalmitoylphosphatidylethanolamine. Yet another direction is to study phospholipid dispersions to which materials such as cholesterol have been added. Cholesterol, a component of cell membranes, has the effect of broadening the transition region (3, 4). The preparation of dispersions of phospholipids with cholesterol addends takes more effort than the sample preparation described here. The phospholipid and cholesterol are dissolved together in chloroform and the mixture is freeze-dried (4). Then, the mixture of the phospholipid and cholesterol can be properly dispersed in water. As described by Ohline and coworkers, studies of the effect of temperature on phospholipid dispersions can also be done by DSC in the undergraduate laboratory (4). The authors describe experiments with phospholipids with double bonds in the chains, with mixtures of phospholipids, and with the cholesterol addend. We have considered having some students use the DSC method while others use the Raman method. We have yet to experiment with this design. Acknowledgments We are grateful to a number of people who assisted in developing this experiment. The impetus for the experiment for the undergraduate laboratory traces to research done by NCC with conventional Raman methods and visible light excitation in Ira Levin’s laboratory during a leave of absence. Recently, Ira Levin made a number of valuable contributions to the laboratory experiment described here. Bruce Chase and Paul Carey were also helpful. William Mohler, Oberlin College electronics technician, designed and constructed the electronics module for the diode temperature sensor. William Marton, Oberlin College machinist, constructed the sample holder. When Jeannine Chan used this experiment in the biochemistry laboratory, she resolved some problems with the sample preparation technique. Many Oberlin College students in three semesters of the biochemistry laboratory did this experiment with enthusiasm and discernment. Their data are reported in Figures 3– 5 and in Table 1 of this paper. The purchase of the Nicolet spectrometer was supported by an NSF ILI grant, DUE9850474, and matching money was supplied by Henry Z. Friedlander to Oberlin College. NNL was supported by a grant to Oberlin College made by the Howard Hughes Medical Institute.

W

Supplemental Material

The laboratory writeup of the experiment, including details of the experimental methods, specifics of apparatus used in sample preparation and of the construction of the variable temperature sample holder, is available in this issue of JCE Online. Notes 1. Lasers are often based on Nd:YAG glasses, where AG stands for aluminum garnet. However, the laser used in the Nicolet instrument is a Nd:YVO4 laser, which has a vanadate matrix. Thus, in general, we refer to the matrix in this type of laser as a “glass.” The newer Nd:YVO4 lasers operate with simple air cooling of the red diode pump laser. 2. Under the best of conditions the Raman effect is very weak. Fluorescence is many orders of magnitude more efficient. Thus, a trace impurity that fluoresces gives reemitted, broadband light that competes in intensity with the Raman bands. Typically, the noise level also increases in association with fluorescence. 3. A white light correction, which compensates for the variation in response of the instrument over the spectral range, has been applied to the spectrum in Figure 3 (5). 4. In Figure 3 the two broad and overlapped bands due to O⫺D stretching in D2O are in the 2500-cm᎑1 region. Because of the changes in hydrogen bonding with temperature the relative intensities of these two bands are also temperature sensitive. This difference has been used to assess temperature in dispersions prepared in D2O (6). 5. The spectra in Figure 4 have not been treated with a white light correction to account for changes in instrument response over the spectral range. Local intensity comparisons without a white light correction are satisfactory. 6. Samples of phospholipids were purchased from SigmaAldrich. 7. A sketch of the construction of the sample holder is supplied with dimensions in the experiment available in the Supplemental Material. W The diode temperature sensor and the construction of the thermocouple are also described there.

Literature Cited 1. Levin, I. W. Adv. Infrared Raman Spectrosc. 1984, 11, 1. 2. Craig, N. C.; Bryant, G. T.; Levin. I. W. Biochem. 1987, 26, 2449.


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In the Laboratory 3. Lippert, J. L.; Peticolas, W. L. Proc. Natl. Acad. Sci. USA 1971, 68, 1572. 4. Ohline, S. M.; Campbell, M. L.; Turnbull, M. T.; Kohler, S. J. J. Chem. Educ. 2001, 78, 1251. 5. Fourier Transform Raman Spectroscopy: from Concept to Experiment; Chase, D. B., Rabolt, J. F., Eds.; Academic Press: San Diego, CA, 1994. 6. Levin, I. W.; Lewis, E. N. In Fourier Transform Raman Spectroscopy: from Concept to Experiment; Chase, D. B., Rabolt, J. F., Eds.; Academic Press: San Diego, CA, 1994; Chapter 6.

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7. Dong, J.; Dinakarpandian, D.; Cary, P. R. Appl. Spectrosc. 1998, 52, 1117. 8. Shimanouchi, T. Tables of Molecular Vibrational Frequencies: Consolidated Volume I; NSRDS-NBS 39; U. S. Government Printing Office: Washington, DC, 1972. 9. Kirchhoff, W. H.; Levin, I. W. J. Res. Natl. Bur. Stds. 1987, 92, 113. 10. Lewis, R. N. A. H.; Mak, N.; McElhaney, R. N. Biochemistry 1987, 26, 6118. 11. Marsh, D. CRC Handbook of Lipid Bilayers; CRC Press, Inc.: Boca Raton, FL, 1990; Table 136.


Investigation of Model Cell Membranes with Raman Spectroscopy: A

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