In the Laboratory
Fluorescence Polarization as a Tool to Pinpoint Vesicle Thermal Phase Transitions
W
Gary A. Baker Department of Chemistry, University at Buffalo, The State University of New York, Buffalo, NY 14260 Thomas A. Betts Department of Physical Sciences, Kutztown University of Pennsylvania, Kutztown, PA 19530 Siddharth Pandey* Department of Chemistry, New Mexico Institute of Mining and Technology, Socorro, NM 87801;
[email protected] Background The dichotomy between hydrophilic (“water-loving”) and oleophilic (“lipid-loving”) species provides molecular motivation for the self-assembly of an important class of interrelated states of matter that have become very pervasive in our modern technology (1). The building block for such assemblies (e.g., monolayers, micelles, lyotropic liquid crystals, microemulsions, bilayers, and vesicles) is the amphipathic lipid, which contains spatially distinct hydrophilic and oleophilic segments. After its introduction to the scientific community in 1965 by A. D. Bangham, who proposed it as a useful model of the cell membrane (2), one such assembly, the liposome, rapidly changed in status from an intellectual curiosity or laboratory plaything to an indispensable tool for the industrialist and biotechnologist alike. Besides utility in product formulation, liposome technology is being developed in areas of diagnosis, drug and transfection agent delivery, in vivo immunomodulation, and energy storage/photon harvesting (3, 4). While broadly defined as a lipid vesicle enclosing an aqueous space, more technically, the term liposome refers to a vesicle formed exclusively from purified phospholipids. Phosphatidylcholines (PC), the most abundant lipid class in mammalian membranes, are critical constituents of human lung surfactant, serum lipoproteins, and bile; they are also the most widely employed phospholipid for model membrane studies (5, 6 ). Saturated diacyl PCs with fatty acyl chains 15–22 carbons long are characterized by a rich polymorphism (7 ). Over a broad range of temperature, these cylindricalshaped lipids undergo many phase transitions. After the socalled pretransition has taken place, upon further heating, the rippled gel phase, Pβ′, undergoes a highly cooperative melting transition, which involves the collapse of the crystal lattice to form a lamellar liquid crystalline Lα phase in which the hydrocarbon chains assume a disordered or gauche conformation. Although the magnitude of this main transition temperature (Tm) is influenced by the chemistry of the lipid, the hydration level, and the dissolved solute, the gel to liquidcrystalline transition is sharp and well defined. The fundamental principles of fluorescence polarization (FP), a measure of the time-averaged rotational motion of a sampled population of fluorescent molecules, were fully developed by Perrin (8) in a series of papers beginning in 1926. Later, pioneering work by Weber (9) ushered in an
1100
area of biophysical luminescence by clever application of these techniques to biological systems. To fully appreciate the elegant simplicity of the FP, a brief overview is in order. In FP measurements, vertically polarized radiation selectively photo-excites those molecules whose absorption transition moments have a significant vector component in the plane of the electric vector of the exciting light. As a result, in the absence of energy migration effects, the excited molecules will emit radiation that is vertically polarized to a degree inversely related to the amount of Brownian motion occurring in the time interval between light absorption and emission (i.e., the excited-state lifetime of the molecule). In practice, the state of polarization of the fluorescence is conveniently characterized by the fluorescence anisotropy (r) defined as
F F r= ⊥ F F⊥
–1 (1)
+2
where F and F⊥ are the parallel and perpendicular fluorescence intensity components, respectively, when the sample is excited with vertically polarized light. To correct for the selective transmission of one orthogonal component through the collection/detection optics, an instrument-specific correction factor G is used, which is the ratio of the sensitivities of the detection system for the vertically and horizontally polarized light. The G factor is easily measured using horizontally polarized excitation and is equal to FHV/FHH, where FHV and FHH are the vertically and horizontally polarized fluorescence intensities, respectively. When G is known, the ratio F/F⊥ can be calculated:
F F⊥
F = 1 × VV G F VH
(2)
where F VV and F VH are the fluorescence intensity components respectively parallel and perpendicular to the electric vector of the vertically polarized incident light. With constant excited-state lifetime in the absence of energy transfer or multiple scattering events, decreased local viscosity or hydrodynamic volume of rotation manifests itself in a subsequently decreased average fluorescence anisotropy. The
Journal of Chemical Education • Vol. 78 No. 8 August 2001 • JChemEd.chem.wisc.edu
In the Laboratory L-α-Dimyristoylphosphatidylcholine
broad application of FP to study biomolecular interactions (antigen–antibody, ligand–receptor, protein–nucleic acid) lies in this potentiality toward interrogating molecular-level dynamics (10–12). We were surprised to learn how few were the contributions to this Journal in the area of artificial membranes (13), and we identified but a single laboratory report that deals directly with liposomes (14). Similarly, although fluorescence polarization is unique in its capacity to gauge molecular mobility, association, and solvation while being simultaneously well suited to an undergraduate curriculum, it too has received scant attention in this Journal to date (15, 16 ). To help to ameliorate this situation, we coupled these two technologies in the formulation of an inexpensive, microscale laboratory exercise equally suited to biochemistry, instrumental analysis, or biophysics. In this laboratory experiment, students determine the Pβ′to-Lα transition temperature (Tm) of L-α-dimyristoylphosphatidylcholine (DMPC) small unilamellar vesicles (SUVs, composed of a single bilayer) by measuring the temperaturedependent depolarization of a membrane probe (BODIPY-C12) sequestered within the membrane (Fig. 1). DMPC was selected as the system of study because it is a frequently studied lipid and because it undergoes a phase transition at a convenient temperature (22 °C < Tm < 25 °C). From the polarization data, a sigmoidal melting curve is generated whereupon Tm is determined by nonlinear least squares fitting or direct differentiation.
(DMPC, 99+%) was obtained from Sigma Chemical Company (St. Louis, MO) and used as received. Water used in this study was quartz bi-distilled and deionized.
Vesicle Preparation Fluorescently labeled DMPC SUVs are prepared beforehand by the instructor as detailed in the online lab documentation.W Fluorescence Polarization Measurements Steady-state fluorescence polarization was measured using a SLM-AMINCO model 8100 spectrofluorometer (Spectronic Instruments) operating in ratiometric mode to eliminate source intensity fluctuations. Excitation was provided at 488 nm (8 nm bandwidth) by a Xe arc lamp. Emission was monitored using a 515-nm long-pass filter, and linear polarization selection was provided by calcite GT linear polarizers (Oriel, Stratford, CT). All measurements were made in disposable 10-mm path-length methacrylate disposable cuvettes (Fisher Scientific, Pittsburgh, PA). Temperaturedependent polarization data were acquired following slow heating of samples using a water-circulating thermostatted cuvette holder with continuous feedback to an external solidstate relay (Omega Engineering, Stamford, CT) connected to a circulating bath (Lauda Model RLS-6). To generate “melting” curves, the temperature was raised at a rate of 1–2 °C/min beginning at 0 °C. Data analyses were performed using TableCurve 3.0 or SigmaPlot 3.2 software packages (Jandel Scientific, San Rafael, CA).
Materials and Methods
Materials 4,4-Difluoro-5-methyl-4-bora-3a,4a-diaza-s-indacene-3dodecanoic acid (BODIPY-C12) was purchased from Molecular Probes (Eugene, OR) and used without further purification.
Hazards There are no hazards or special precautions associated with the use of these materials and procedures.
O O O
O
N
O P O O
DMPC
+
O−
O N H3C
F
B
N OH
F
BODIPY-C12 SUV
Figure 1. Diagram depicting the bilayer structure of small unilamellar vesicles (SUVs). Also shown are the molecular structures for the constituent lipid DMPC and the membrane probe molecule BODIPY-C12.
JChemEd.chem.wisc.edu • Vol. 78 No. 8 August 2001 • Journal of Chemical Education
1101
In the Laboratory
0.6
2
0
0
10
20
30
40
Temperature / ˚C 0.4
λEX
0.2
0.0 450
550
600
650
dr/dT
0.15
Tm
−0.02 −0.04 −0.06 10
15
20
25
30
Temperature / ˚C
0.10
0.05
0.00 500
0.02 0.00
4
Steady-State Anisotropy (r )
Lifetime / ns
0.8
Fluorescence Intensity
0.20
6
1.0
0
Wavelength / nm
10
20
30
40
50
Temperature / ˚C
Figure 2. Fluorescence excitation and emission spectra for BODIPYC12 labeled DMPC SUVs (the vertical bar denotes the excitation wavelength: λex = 488 nm, ∆λ = 8 nm). Excited-state fluorescence lifetimes as a function of temperature are provided in the inset.
Figure 3. Steady-state emission anisotropies as a function of temperature for BODIPY-C12-labeled DMPC SUVs. Error bars represent 1 SD; n = 30. The inset shows the first derivative polarization spectrum identifying Tm as the global minimum in dr/dT.
Results and Discussion
versus T or d2r /dT 2 versus T, respectively) of the original spectrum. Upon differentiation, maxima or minima result where the original spectrum exhibits an inflection point and zero crossings result where the original spectrum exhibits maxima or minima (18). In the inset of Figure 3, the first derivative of the “melting curve” r (T ) is shown where the minimum at 23.5 °C clearly identifies Tm.
Normalized fluorescence excitation and emission spectra for BODIPY-C12 embedded in DMPC SUVs are shown in Figure 2. The shaded region under the emission profile denotes the relative fluorescence envelope passed by our filter into the polarization/detection channel of our fluorimeter. As expected for BODIPY derivatives that are known to be relatively insensitive to environmental conditions (17), the excited-state lifetime for DMPC membrane-bound BODIPY-C12 remains essentially constant at 5.53 ± 0.17 ns from 3 to 38 °C (Fig. 2 inset). This is a desirable feature of polarization probes because the experimental values of r relate directly to the reorientational rate of the probe (12). Figure 3 provides representative data for the thermally induced depolarization of fluorescence for BODIPY-C12 associated with DMPC SUVs. At low temperatures, the anisotropy is high (r ≈ 0.16 at 0 °C), indicating limited probe reorientation during its excited-state lifetime. Upon the sharp thermotropic phase transition near 23 °C, however, the gel phase is dramatically transformed into the more mobile liquidcrystalline phase, permitting a greater degree of molecular reorientation with a corresponding reduction in steady-state anisotropy (r ≈ 0.03 at 30 °C). Since the polarization data are clearly sigmoidal, we instructed our students to use the following four-parameter logistic expression to model their data:
r=
a 1 + exp b T – c
+d
(3)
Here, a is the range in anisotropy, b is a slope coefficient, c is the T at the maximal rate of change (i.e., Tm), and d is the minimum anisotropy value. The solid line in Figure 3 is the nonlinear least squares fit of the polarization data to eq 2 (r 2 = .983). The recovered value for Tm (23.0 °C) compares well with the reported value of 23.6 ± 1.5 °C (7). A common way of obtaining greater spectral detail thereby enhancing particular features of a spectrum (e.g., r versus T ) is to display the first or second derivative (dr /dT 1102
Summary Students determine the gel-to-liquid transition temperature (Tm) in DMPC model bilayer systems based on static FP of a lipid-tailed fluorescent marker bound therein. First, students are exposed to both organized media and emission polarization phenomena. Then they apply these fundamentals to readily and accurately (±1–2 °C) determine Tm for a neat DMPC vesicular system. W
Supplemental Material
A data entry form for students and notes for the instructor on SUV preparation and equipment needs are available in this issue of JCE Online. Some additional tips for the instructor are also provided. Literature Cited 1. Ball, P. Life’s Matrix: A Biography of Water; Farrar Straus & Giroux: New York, 2000. 2. Bangham, A. D.; Standish, M. M.; Watkins, J. C. J. Mol. Biol. 1965, 13, 238. 3. Liposomes: A Practical Approach; New, R. R. C., Ed.; Oxford University Press: Oxford, 1990; p 63. 4. Rosoff, M. Vesicles; Dekker: New York, 1996. 5. Fendler, J. H. Membrane Mimetic Chemistry; Wiley Interscience: New York, 1982. 6. Ti Tien, H.; Ottova-Leitmannova, A. Membrane Biophysics; Elsevier: New York, 2000
Journal of Chemical Education • Vol. 78 No. 8 August 2001 • JChemEd.chem.wisc.edu
In the Laboratory 7. 8. 9. 10.
Koynova, R.; Caffrey, M. Biochim. Biophys. Acta 1997, 1376, 91. Perrin, F. J. Phys. Radium 1926, 7, 390. Weber, G. Adv. Protein Chem. 1953, 8, 415. Cantor, C. R.; Schimmel, P. R. Biophysical Chemistry: Part II, Techniques for the Study of Biological Structure and Function; Freeman: New York, 1980; pp 454–465. 11. Jameson, D. M.; Sawyer, W. H. Methods Enzymol. 1995, 246, 283–300. 12. Lakowicz, J. R. Principles of Fluorescence Spectroscopy, 2nd ed.; Kluwer: New York, 1999; Chapter 10.
13. Kohlwein, S. D. J. Chem. Educ. 1992, 69, 3. 14. Jakubowski, H. V.; Penas, M.; Saunders, K. J. Chem. Educ. 1994, 71, 347. 15. Carper, M. A.; Carper, W. R. J. Chem. Educ. 1968, 45, 662. 16. Bigger, S. W.; Craig, R. A.; Ghiggino, K. P.; Scheirs, J. J. Chem. Educ. 1993, 70, A234. 17. Haugland, R. P. Handbook of Fluorescent Probes and Research Chemicals, 7th ed. [CD-ROM]; Molecular Probes: Eugene, OR, 1999. 18. Green, G. L.; O’Haver, T. C. Anal. Chem. 1974, 46, 2191.
JChemEd.chem.wisc.edu • Vol. 78 No. 8 August 2001 • Journal of Chemical Education
1103
