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Effect of osmotic gradient on the physical properties of membrane lipids in liposomes
Chemistry and Physics of Lipids, 33 (1983) 81-85 Elsevier Scientific Publishers Ireland Ltd.
81
EFFECT OF OSMOTIC GRADIENT ON THE PHYSICAL PROPERTIES OF MEMBRANE LIPlDS IN LIPOSOMES
WITOLD K. SUREWICZ
Department of Biophysics, Institute of Biochemistry and Biophysics, University of Lodz, Banacha 12/16, 90-237 Lodz (Poland) Received August 27th, 1982
accepted October 21st, 1982
Osmotic swelling of multilameUar liposomes was shown to produce structural changes in liposomal membranes. Creation of transmembrane osmotic gradients results in an increased lateral mobility of membrane incorporated hydrophobic probe, pyrene. This is accompanied by some decrease in order parameter of membrane phospholipid polar head group regions. The perturbations are more marked in hydrophobic than in polar regions of the membranes. It is suggested that some functional changes in biomembranes arising upon osmotic swelling may be associated with structural alterations similar to those observed in liposomes.
Keywords: liposome; osmotic swelling; spin label; fluorescent probe; membrane structure.
Introduction Osmotic gradients are well known to induce swelling or shrinking of various cells and organelles [ 1--45]. This is accompanied by diverse functional perturbations at biomembrane level, such as changes in surface potential [ 1], increased leaking of metabolites [7] and changes in membrane transport o f various substances [ 3 , 8 - 1 0 ] . Although the nature of osmolarity gradient-induced functional perturbations in biomembranes is far from being clearly understood, the possibility that the osmotic swelling can affect cell physiology via changes in membrane fluidity has been recently raised [ 11 ]. Here, the effect of osmotic gradient on the physical state of membrane lipids in liposomes is examined in some detail. Changes in the structure of model phosphatidylcholine membranes are followed by electron spin resonance experiments using 5-doxylstearic acid as a spin label and by optical measurements using pyrene as a fluorescent probe. Multilamellar liposomes used in this study are known to be osmotic sensitive [11,12] and they provide a simple and convenient model for studying structural aspects of osmolarity gradient-induced perturbations in biological membranes. 0009-3084/83/$03.00 © 1983 Elsevier Scientific Publishers Ireland Ltd. Published and Printed in Ireland
82 Materials and methods
Chromatographically pure egg yolk phosphatidylcholine and dicetyl phosphate were obtained from Sigma Chemical Co. (St. Louis, MO, U.S.A.). Spin label 5doxylstearic acid was from Syva (Palo Alto, CA, U.S.A.). Pyrene and sucrose were purchased from Serva (D-6900 Heidelberg). Multilamellar liposomes were prepared by the method of Bangham et al. [12]. Stock solutions of phosphatidylcholine/dicetyl phosphate (molar ratio 95:5) containing pyrene (2.5 mol%) or spin label 5-doxylstearic acid (1 mol%) were made in chloroform. After evaporation of the solvent under argon Tris-HCl buffer (10 mM, pH 7.4) containing 1.5 M sucrose was added and the mixtures were shaken by means of vortex rotamixer for 5 min. The concentration of lipid was 0.6 mg/ml in the case of fluorescence-labeled liposomes and 50 mg/ml in the case of spin-labeled liposomes. In order to create osmotic gradients the above stock suspensions of liposomes were diluted three times in sucrose solutions of desired tonicity. After I h equilibration, when swelling was completed, spectroscopic measurements of labeled liposomes were performed at room temperature. Fluorescence spectra were recorded in a Jobin Yvon JY3 spectrofluorimeter. Electron spin resonance spectra were obtained with a SE/X-28 ESR spectrometer (Wroclaw Technical University) operating at 9.5 GHz. Results and discussion
Liposomes prepared in buffer containing 1.5 M sucrose were diluted with hypotonic sucrose solutions of desired tonicity. As expected from earlier reports [11,12], creation of such osmotic gradient resulted in liposome swelling. The swelling was followed by light scattering measurements and it was found to proceed within a few minutes. When a new equilibrium volume was reached it persisted for at least 12 h. Membrane structural changes associated with liposome swelling were studied using pyrene- and spin-labeling methods. Pyrene is a highly apolar molecule and, when added to liposome suspension, it is completely buried within the hydrophobic interior of the membrane [13-16].. The fluorescence emission spectrum of pyreneolabeled hposomes is shown in Fig. 1. The highly structured band (370-430 nm) corresponds to the emission of the excited monomer whereas the broad band (430-560 nm) corresponds to the emission of the excimer [13,17,18]. Pyrene excimers are formed by interaction of excited pyrene and the ground state pyrene. Excimer formation is a dynamic, diffusion controlled process. The rate of pyrene excimer formation in phospholipid bilayer is related to the lateral diffusion coefficient of the probe and it provides valuable information about the fluidity of the membrane [13,17-20]. It is reflected in fluorescence spectrum by lexe/Im ratio of the maximum fluorescence excimer (about 475 nm) and monomer (about 395 nm) intensities. At a
83 given concentration of pyrene in lipid bilayer the higher the Iexc/lm ratio the more probable excimer formation and, therefore, the more fluid the membrane environment of the probe. Figure 2 shows the effect of osmotic gradient-induced liposome swelling on the lexc/Im ratio. The creation of transmembrane osmotic gradient is reflected by an increase in excimer/monomer fluorescence ratio. This is indicative, as discussed above, of the increased fluidity of inner hydrocarbon regions in the liposomes. Concomitant with the above measurements, the relative intensities of the bands B and A in pyrene monomer fluorescence (Fig. 1) have been determined. The intensities of these vibronic bands have been shown to have a strong dependence on the solvent environment [21]. In the presence of polar solvents, there is a pronounced enhancement in the intensity of band A at the expense of other bands. As shown in Fig. 2 no significant changes in the polarity of the environment of pyrene molecules were detected upon liposome swelling in hypotonic solutions. This indicates that the increased fluidity in the hydrocarbon region of osmoticallyswollen liposomes is not accompanied by the increased water penetration into the interior of the bilayer [20]. Structural changes in liposomes have been studied also by means of spin-labeling technique. The spin probe used in this study, 5-doxylstearic acid, provides information about the fatty acid chains packing near the polar head group region of the bilayer [22]. Electron spin resonance spectrum of liposomes labeled with 5doxylstearic acid is shown in Fig. 3. This spectrum reflects the relatively high degree of order in the nitroxide reporter group intercalation region of the mem-
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OSMOTIC GRADIENT (M)
Fig. 1. Fluorescence spectrum of pyrene in phosphatidylcholine/dicetyl phosphate (95:5) liposomes. Label concentration 2.5 mol% with respect to lipid. Excitation wavelength: 340 rim. Fig. 2. Relationship between the osmotic gradient and intensity ratios in fluorescence spectrum
of pyrene-labeled multilameUar liposomes: Q, excimer/monomer fluorescence ratio, lexc/Im; o, intensity ratio of the B and A vibronic bands in pyrene monomer fluorescence,IB/I A. Osmotic gradients were calculated as sucrose concentration in the medium minus its concentration inside the liposomes.
84 brane. It may be analysed quantitatively using the order parameter, S, calculated according to the equation [23] : S =
All--At
a
Azz -- 1/2 (Axx + Ayy)
a
t
where Axx, Ayy, Azz are the components of hyperrme splitting tensor obtained from single crystal spectra [24], a = 1/3 (Axx + Ayy + Azz) and a ' = 1/3 (.4 FI + 2A1). The order parameter changes versus the osmotic gradient are shown in Fig. 4. In addition to the increased fluidity of inner hydrocarbon regions, osmotic swelling of liposomes is associated also with some decrease in molecular ordering near the phospholipid polar head group region. The changes in the molecular organization of membrane lipids in liposomes stated in this study are, in general, consistent with the recent results of Borochov and Borochov [11 ]. These authors have reported that osmotic swelling is associated with some decrease in the fluorescence anisotropy of membrane incorporated probe, 1,6-diphenyl-l,3,5-hexatriene. The application in the present study of membrane probes monitoring the organization of different regions of lipid bilayer allowed us to conclude that both hydrophobic and polar regions of the membrane are perturbed upon osmotic swelling. It should be emphasised, however, that
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Fig. 3. Electron spin resonance spectrum of 5-doxylstearic acid spin probe incorporated into phosphatidylcholine/dicetyl phosphate (95 : 5) liposomes. Fig. 4. The effect of osmotic gradient on order parameter change of 5-doxylstearic acid spinlabeled multilameUar liposomes . The mean order parameter for control liposomes was 0.615 ± 0.006. Osmotic gradients were calculated as sucrose concentration in the medium minus its concentration inside the liposomes.
85 structural changes in the hydrophobic interior of the membrane are much more pronounced than perturbations in more polar regions. To put it in context, the same change in order parameter of 5-doxylstearic acid-labeled liposomes as observed at osmotic gradient of 0.5 M is caused by an increase in temperature of about 4°C. The change in excimer/monomer ratio in pyrene fluorescence spectra observed at the same osmotic gradient is equivalent to the increase in temperature of approx. 14°C. Although great caution should be exercised when extrapolating results of model experiments to events at biomembranes, it may be suggested that some of functional changes in biological membranes arising upon osmotic swelling may be mediated by structural alterations similar to those observed in liposomes.
References 1 R.H. Racusen, A.M. Kinnesley and A.W. Galston, Science, 198 (1977) 405. 2 E.J.J. Van Zoelen, E.C.M. Van der Neut-Kok, J. De Gier and LL.M. Van Deenen, Biochim. Biophys. Acta, 394 (1975) 463. 3 M. Poznansky and A.K. Solomon, J. Membr. Biol., 10 (1972) 259. 4 C.T. Wangand P.S. Nobel, Biochim. Biophys. Acta, 241 (1971) 200. 5 I.B. Kirk and J.B. Hanson, Plant Physiol., 51 (1973) 357. 6 M. Kasai, T. Kanemasa and S. Fukomoto, J. Membr. Biol., 51 (1979) 311. 7 S.P. Burg, E.A. Burg and R. Marks, Plant Physiol., 39 (1964) 1.85. 8 R.A. Venom, Biochim. Biophys. Acta, 510 (1978) 378. 9 M.A. Hardy, Biochirn. Biophys. Acta, 552 (1979) 169. 10 J. Fischbarg, G.L. Hofer and R.A. Koatz, Biochim. Biophys. Acta, 603 (1980) 198. 11 A. Borochov and H. Borochov, Biochim. Biophys. Acta, 550 (1979) 546. 12 A.D. Bangham, J. De Gier and G.D. Greville, Chem. Phys. Lipids, 1 (1967) 225. 13 H.-J. GaUa and E. Sackmann, Bioehim. Biophys. Acta, 339 (1974) 103. 14 H.-J. Galla and E. Saekmann, BeE Bunsenges. Phys. Chem., 78 (1974) 949. 15 F. Podo and J.K. Blasie, Proc. Natl. Acad. Sci. U.S.A., 74 (1977) 1032. 16 J. Luisetti, H. Mohwald and H.-J. Galla, Bioehim. Biophys. Aeta, 552 (1979) 519. 17 A.K. Santar, H.J. Pownall, A.S. Hu and L.C. Smith, Biochemistry, 13 (1974) 2828. 18 J.M. Vanderkooi and J.B. Callis, Biochemistry, 13 (1974) 4000. 19 A. Waggoner, in: A. Martonosi (Ed.), The Enzymes of Biological Membranes, Vol. 1, Plenum Press, 1976, p. 111. 20 D.J.W. Barber and J.K. Thomas, Radiat. Res., 74 (1978) 51. 21 K. Kalyansandarm and J.K. Thomas, J. Am. Chem. Soc., 99 (1977) 2039. 22 S. Schreier-MuciUo, D. Marsh and I.C.P. Smith, Arch. Biochom. Biophys., 172 (1976) 1. 23 W.L. Hubbell and H.M. McConnell, J. Am. Chem. Soc., 93 (1971) 314. 24 P. Jost, L.J. Libertini, V.C. Hebert and O.H. Grfffith, J. Mol. Biol., 59 (1971) 77.
81
EFFECT OF OSMOTIC GRADIENT ON THE PHYSICAL PROPERTIES OF MEMBRANE LIPlDS IN LIPOSOMES
WITOLD K. SUREWICZ
Department of Biophysics, Institute of Biochemistry and Biophysics, University of Lodz, Banacha 12/16, 90-237 Lodz (Poland) Received August 27th, 1982
accepted October 21st, 1982
Osmotic swelling of multilameUar liposomes was shown to produce structural changes in liposomal membranes. Creation of transmembrane osmotic gradients results in an increased lateral mobility of membrane incorporated hydrophobic probe, pyrene. This is accompanied by some decrease in order parameter of membrane phospholipid polar head group regions. The perturbations are more marked in hydrophobic than in polar regions of the membranes. It is suggested that some functional changes in biomembranes arising upon osmotic swelling may be associated with structural alterations similar to those observed in liposomes.
Keywords: liposome; osmotic swelling; spin label; fluorescent probe; membrane structure.
Introduction Osmotic gradients are well known to induce swelling or shrinking of various cells and organelles [ 1--45]. This is accompanied by diverse functional perturbations at biomembrane level, such as changes in surface potential [ 1], increased leaking of metabolites [7] and changes in membrane transport o f various substances [ 3 , 8 - 1 0 ] . Although the nature of osmolarity gradient-induced functional perturbations in biomembranes is far from being clearly understood, the possibility that the osmotic swelling can affect cell physiology via changes in membrane fluidity has been recently raised [ 11 ]. Here, the effect of osmotic gradient on the physical state of membrane lipids in liposomes is examined in some detail. Changes in the structure of model phosphatidylcholine membranes are followed by electron spin resonance experiments using 5-doxylstearic acid as a spin label and by optical measurements using pyrene as a fluorescent probe. Multilamellar liposomes used in this study are known to be osmotic sensitive [11,12] and they provide a simple and convenient model for studying structural aspects of osmolarity gradient-induced perturbations in biological membranes. 0009-3084/83/$03.00 © 1983 Elsevier Scientific Publishers Ireland Ltd. Published and Printed in Ireland
82 Materials and methods
Chromatographically pure egg yolk phosphatidylcholine and dicetyl phosphate were obtained from Sigma Chemical Co. (St. Louis, MO, U.S.A.). Spin label 5doxylstearic acid was from Syva (Palo Alto, CA, U.S.A.). Pyrene and sucrose were purchased from Serva (D-6900 Heidelberg). Multilamellar liposomes were prepared by the method of Bangham et al. [12]. Stock solutions of phosphatidylcholine/dicetyl phosphate (molar ratio 95:5) containing pyrene (2.5 mol%) or spin label 5-doxylstearic acid (1 mol%) were made in chloroform. After evaporation of the solvent under argon Tris-HCl buffer (10 mM, pH 7.4) containing 1.5 M sucrose was added and the mixtures were shaken by means of vortex rotamixer for 5 min. The concentration of lipid was 0.6 mg/ml in the case of fluorescence-labeled liposomes and 50 mg/ml in the case of spin-labeled liposomes. In order to create osmotic gradients the above stock suspensions of liposomes were diluted three times in sucrose solutions of desired tonicity. After I h equilibration, when swelling was completed, spectroscopic measurements of labeled liposomes were performed at room temperature. Fluorescence spectra were recorded in a Jobin Yvon JY3 spectrofluorimeter. Electron spin resonance spectra were obtained with a SE/X-28 ESR spectrometer (Wroclaw Technical University) operating at 9.5 GHz. Results and discussion
Liposomes prepared in buffer containing 1.5 M sucrose were diluted with hypotonic sucrose solutions of desired tonicity. As expected from earlier reports [11,12], creation of such osmotic gradient resulted in liposome swelling. The swelling was followed by light scattering measurements and it was found to proceed within a few minutes. When a new equilibrium volume was reached it persisted for at least 12 h. Membrane structural changes associated with liposome swelling were studied using pyrene- and spin-labeling methods. Pyrene is a highly apolar molecule and, when added to liposome suspension, it is completely buried within the hydrophobic interior of the membrane [13-16].. The fluorescence emission spectrum of pyreneolabeled hposomes is shown in Fig. 1. The highly structured band (370-430 nm) corresponds to the emission of the excited monomer whereas the broad band (430-560 nm) corresponds to the emission of the excimer [13,17,18]. Pyrene excimers are formed by interaction of excited pyrene and the ground state pyrene. Excimer formation is a dynamic, diffusion controlled process. The rate of pyrene excimer formation in phospholipid bilayer is related to the lateral diffusion coefficient of the probe and it provides valuable information about the fluidity of the membrane [13,17-20]. It is reflected in fluorescence spectrum by lexe/Im ratio of the maximum fluorescence excimer (about 475 nm) and monomer (about 395 nm) intensities. At a
83 given concentration of pyrene in lipid bilayer the higher the Iexc/lm ratio the more probable excimer formation and, therefore, the more fluid the membrane environment of the probe. Figure 2 shows the effect of osmotic gradient-induced liposome swelling on the lexc/Im ratio. The creation of transmembrane osmotic gradient is reflected by an increase in excimer/monomer fluorescence ratio. This is indicative, as discussed above, of the increased fluidity of inner hydrocarbon regions in the liposomes. Concomitant with the above measurements, the relative intensities of the bands B and A in pyrene monomer fluorescence (Fig. 1) have been determined. The intensities of these vibronic bands have been shown to have a strong dependence on the solvent environment [21]. In the presence of polar solvents, there is a pronounced enhancement in the intensity of band A at the expense of other bands. As shown in Fig. 2 no significant changes in the polarity of the environment of pyrene molecules were detected upon liposome swelling in hypotonic solutions. This indicates that the increased fluidity in the hydrocarbon region of osmoticallyswollen liposomes is not accompanied by the increased water penetration into the interior of the bilayer [20]. Structural changes in liposomes have been studied also by means of spin-labeling technique. The spin probe used in this study, 5-doxylstearic acid, provides information about the fatty acid chains packing near the polar head group region of the bilayer [22]. Electron spin resonance spectrum of liposomes labeled with 5doxylstearic acid is shown in Fig. 3. This spectrum reflects the relatively high degree of order in the nitroxide reporter group intercalation region of the mem-
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390
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,¢70
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WAVELENGTH
081
/ ~
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-0,~
08
-72
OSMOTIC GRADIENT (M)
Fig. 1. Fluorescence spectrum of pyrene in phosphatidylcholine/dicetyl phosphate (95:5) liposomes. Label concentration 2.5 mol% with respect to lipid. Excitation wavelength: 340 rim. Fig. 2. Relationship between the osmotic gradient and intensity ratios in fluorescence spectrum
of pyrene-labeled multilameUar liposomes: Q, excimer/monomer fluorescence ratio, lexc/Im; o, intensity ratio of the B and A vibronic bands in pyrene monomer fluorescence,IB/I A. Osmotic gradients were calculated as sucrose concentration in the medium minus its concentration inside the liposomes.
84 brane. It may be analysed quantitatively using the order parameter, S, calculated according to the equation [23] : S =
All--At
a
Azz -- 1/2 (Axx + Ayy)
a
t
where Axx, Ayy, Azz are the components of hyperrme splitting tensor obtained from single crystal spectra [24], a = 1/3 (Axx + Ayy + Azz) and a ' = 1/3 (.4 FI + 2A1). The order parameter changes versus the osmotic gradient are shown in Fig. 4. In addition to the increased fluidity of inner hydrocarbon regions, osmotic swelling of liposomes is associated also with some decrease in molecular ordering near the phospholipid polar head group region. The changes in the molecular organization of membrane lipids in liposomes stated in this study are, in general, consistent with the recent results of Borochov and Borochov [11 ]. These authors have reported that osmotic swelling is associated with some decrease in the fluorescence anisotropy of membrane incorporated probe, 1,6-diphenyl-l,3,5-hexatriene. The application in the present study of membrane probes monitoring the organization of different regions of lipid bilayer allowed us to conclude that both hydrophobic and polar regions of the membrane are perturbed upon osmotic swelling. It should be emphasised, however, that
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OSMOTIC GRAOIENF (M)
Fig. 3. Electron spin resonance spectrum of 5-doxylstearic acid spin probe incorporated into phosphatidylcholine/dicetyl phosphate (95 : 5) liposomes. Fig. 4. The effect of osmotic gradient on order parameter change of 5-doxylstearic acid spinlabeled multilameUar liposomes . The mean order parameter for control liposomes was 0.615 ± 0.006. Osmotic gradients were calculated as sucrose concentration in the medium minus its concentration inside the liposomes.
85 structural changes in the hydrophobic interior of the membrane are much more pronounced than perturbations in more polar regions. To put it in context, the same change in order parameter of 5-doxylstearic acid-labeled liposomes as observed at osmotic gradient of 0.5 M is caused by an increase in temperature of about 4°C. The change in excimer/monomer ratio in pyrene fluorescence spectra observed at the same osmotic gradient is equivalent to the increase in temperature of approx. 14°C. Although great caution should be exercised when extrapolating results of model experiments to events at biomembranes, it may be suggested that some of functional changes in biological membranes arising upon osmotic swelling may be mediated by structural alterations similar to those observed in liposomes.
References 1 R.H. Racusen, A.M. Kinnesley and A.W. Galston, Science, 198 (1977) 405. 2 E.J.J. Van Zoelen, E.C.M. Van der Neut-Kok, J. De Gier and LL.M. Van Deenen, Biochim. Biophys. Acta, 394 (1975) 463. 3 M. Poznansky and A.K. Solomon, J. Membr. Biol., 10 (1972) 259. 4 C.T. Wangand P.S. Nobel, Biochim. Biophys. Acta, 241 (1971) 200. 5 I.B. Kirk and J.B. Hanson, Plant Physiol., 51 (1973) 357. 6 M. Kasai, T. Kanemasa and S. Fukomoto, J. Membr. Biol., 51 (1979) 311. 7 S.P. Burg, E.A. Burg and R. Marks, Plant Physiol., 39 (1964) 1.85. 8 R.A. Venom, Biochim. Biophys. Acta, 510 (1978) 378. 9 M.A. Hardy, Biochirn. Biophys. Acta, 552 (1979) 169. 10 J. Fischbarg, G.L. Hofer and R.A. Koatz, Biochim. Biophys. Acta, 603 (1980) 198. 11 A. Borochov and H. Borochov, Biochim. Biophys. Acta, 550 (1979) 546. 12 A.D. Bangham, J. De Gier and G.D. Greville, Chem. Phys. Lipids, 1 (1967) 225. 13 H.-J. GaUa and E. Sackmann, Bioehim. Biophys. Acta, 339 (1974) 103. 14 H.-J. Galla and E. Saekmann, BeE Bunsenges. Phys. Chem., 78 (1974) 949. 15 F. Podo and J.K. Blasie, Proc. Natl. Acad. Sci. U.S.A., 74 (1977) 1032. 16 J. Luisetti, H. Mohwald and H.-J. Galla, Bioehim. Biophys. Aeta, 552 (1979) 519. 17 A.K. Santar, H.J. Pownall, A.S. Hu and L.C. Smith, Biochemistry, 13 (1974) 2828. 18 J.M. Vanderkooi and J.B. Callis, Biochemistry, 13 (1974) 4000. 19 A. Waggoner, in: A. Martonosi (Ed.), The Enzymes of Biological Membranes, Vol. 1, Plenum Press, 1976, p. 111. 20 D.J.W. Barber and J.K. Thomas, Radiat. Res., 74 (1978) 51. 21 K. Kalyansandarm and J.K. Thomas, J. Am. Chem. Soc., 99 (1977) 2039. 22 S. Schreier-MuciUo, D. Marsh and I.C.P. Smith, Arch. Biochom. Biophys., 172 (1976) 1. 23 W.L. Hubbell and H.M. McConnell, J. Am. Chem. Soc., 93 (1971) 314. 24 P. Jost, L.J. Libertini, V.C. Hebert and O.H. Grfffith, J. Mol. Biol., 59 (1971) 77.