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Stabilization of liposome bilayers to freezing and thawing: Effects of cryoprotective agents and membrane proteins
CRYOBIOLOGY
17, 508-515
(1980)
Stabilization of Liposome of Cryoprotective
Bilayers to Freezing and Thawing: Agents and Membrane Proteins
G. STRAUSS Department
of Chemistry,
Douglass
College,
AND E. P. INGENITO’ Rutgers University, New Brunswick,
The nature of the protective action of glycerol and other cryoprotective agents (CPA) against freeze-thaw damage to biomaterials has been studied intensively for a long time. The accumulated evidence suggests that one or more of the following processes are responsible: (i) The formation of intracellular ice, found by microscopic observation (8). (ii) Osmotic dehydration of cells, resulting from the formation of ice outside, but not inside, the cell, as shown by observations of volume shrinkage of cells (6) and liposome model systems (11). This process depends on the water conductivity of the cell membrane and rate of cooling (4), and leads to a destructuve rise in salt concentration and denaturation of lipid-protein complexes (5). Hemolysis measurements on erythrocytes have shown that CPA such as glycerol and dimethyl sulfoxide (Me,SO) prevent intracellular ice formation and increase in salt concentration (6, 10). (iii) The integrity of the lipid membrane may suffer on freezing-thawing if a phase transition from a liquid-like (or liquid crystal) to a crystalline state occurs. This last process which has been studied in model systems (3), so far has received little attention in connection with cryoprotection of cells. In an effort to clarify the roles of these diverse processes and to better understand the behavior of the lipid membrane, as Received July 19, 1978; accepted May 27, 1980. L Present address: Medical Engineering and Medical Physics, Harvard-Massachusetts Institute of Technology Division of Health Sciences and Technology, Cambridge, Mass. 02139.
Copyright All rights
@ 1980 by Academic F’ress, Inc. of reproduction in any form reserved.
New Jersey 08903
such, at low temperatures, we investigated the loss in structural integrity of unilamellar lipid bilayer vesicles (liposomes) upon freezing and thawing. Liposomes are spherical or oblong vesicles, ca. 300 A in diameter. The walls, about 80 A thick, consist of lipid bilayers and enclose an aqueous phase. Lipid bilayers can serve as amphipathic solvents for other lipids, such as cholesterol, fatty acids, etc., and particularly for proteins which, in the case of living cells, constitute from 20 to 70% by weight of the lipid membrane. Liposomes can be prepared by suspending a phospholipid in an aqueous salt solution which then is sonicated. Altematively, an alcoholic solution of the lipid can be injected through a narrow hypodermic needle into a large (about 20-fold) volume of aqueous solution. These methods demonstrate that mechanical agitation or shear is sufficient to overcome the energy barrier in the transition in which a coarse suspension of the lipid (consisting of large multilamellar structures) is transformed into a suspension of small unilamellar vesicles. As we have found, freezing and thawing causes partial reversal of this transition. Unilamellar vesicles may then be restored once more by sonication. Addition of CPA, in contrast, causes dissolution of the phospholipid (at CPA concentrations beyond 20%), and irreversible destruction of the liposomes. Loss of membrane integrity was monitored (a) by change in turbidity, and (b) by loss in energy transfer between fluorescent probes incorporated into the bilayers. Turbidity changes, both positive and negative, were a sensitive, albeit only semiquantita-
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Effects
FREEZE-THAW
STABILITY
OF LIPOSOMES
509
cals) as energy donor, and chlorophyll a (highly purified, from Sigma Chemical Co.) as acceptor. Equal volumes of alcoholic solutions of PL (10 mg/ml), of the cyanine dye (1 mg/ml), and of chlorophyll (sufficient to give an optical density of about 0.1) were mixed. A 0.5ml volume of this solution was injected into 10 ml of an aqueous solution 0.16 M in KC1 and 0.05 M in Tris buffer, pH 7.5, containing up to 50% of glycerol or Me,SO. To prepare protein-containing liposomes, 75 mg of Cohn fraction VI serum glycoprotein (from U.S. Biochemical Corp.) was dissolved in 1.Oml water. Nine milliliters of ethanol and 75 mg PL were added, and the solution was agitated on a Vortex mixer until clear (20 min). This solution was then used in combination with the other stock solutions as before. To check how much of the protein present had become incorporated into the liposome membrane, the liposome suspensions were subjected to ultrafiltration under 20 lb/in.2 pressure through a Millipore PSJM Pellicon ultrafilter. These filters have a nominal molecular weight cutoff of 100,008 daltons, and thus retain liposomes (MW EXPERIMENTAL PROCEDURES about 2 x lo6 daltons) but pass flee serum Preparation of Liposomes glycoprotein (of the order of 50,000 dalThe phospholipid (PL) used for liposome tons). Absorption spectroscopy of the inipreparation either was lecithin from egg tial liposome suspensions and the filtrates yolk, obtained from Sigma Chemical Com- showed that 25% of the serum protein pany (Type IIE, commercial grade), and originally present had become bound, conpurified by chromatography on an alumina stituting 20% of the total membrane weight. This includes the protein trapped in the column, or DL-lecithin (1-octadec-9-enyl, 2-hexadecyl ether) obtained from Cal- internal aqueous phase of the liposomes, biochem and used without further purifi- which, however, amounts to only 0.03% of cation. Liposomes made from either of the membrane-bound protein. these PL showed comparable behavior. They were produced by the method of Bat- Freezing -Thawing Cycles zri and Korn (1) in which an ethanolic soluThree-milliliter volumes of the aqueous tion containing PL and the fluorescent liposome suspensions were placed into 13 probes is injected by a syringe through a 22- X loo-mm stoppered test tubes. Various gauge needle into a 20-fold volume of aque- cooling and warming rates were used, and ous solution. The fluorescent probes were the samples were held for 18 hr at bath temdioctadecyloxacarbocyanine-p -toluene peratures varying from -5 to -70°C. Samsulfonate (from Eastman Organic Chemi- ple temperatures were read on a remote tive, measure of liposome destruction. Energy transfer (discussed in a later section) measures the intermolecular distance of energy donor and acceptor molecules, which is small as long as the liposome membrane remains intact. Membrane destruction results in dispersion of these probe molecules over a vastly larger volume, and thus in loss of energy transfer. As reported previously (7), we found that addition of Triton X- 100, a nonionic detergent which is known to destroy liposomes, completely abolished energy transfer. This method, therefore, can be regarded as a measure of membrane integrity, in contrast to methods based on release of a trapped marker which respond not only to membrane destruction but also to creation of pores in an otherwise intact membrane. Loss of energy transfer was observed both when the lipid was dispersed by addition of CPA, and after a freeze-thaw cycle. In this latter case where the lipid became aggregated rather than dispersed, the loss in energy transfer can be ascribed to the fact that the probes became insolubie in the lipid below the transition temperature (3).
510
STRAUSS
AND
thermistor thermometer. For “slow” cooling (0.3Wmin) the sample tubes were surrounded by 50-mm-diameter tubes serving as air bath. For “fast” cooling (CC/min) the sample tubes were directly immersed in the cooling bath maintained at the desired final temperature. For temperatures from -5 to -30°C a refrigerated constant-temperature bath was used. An acetone-dry ice mixture in a Dewar flask was used to provide a -70°C bath. The latter was also used to cool samples at a very fast rate (750Wmin) by filling the suspensions into thin-walled 5-mm-diameter aluminum tubes which were directly immersed in the freezing mixture. “Slow” (0.3Wmin) and “fast” (5Wmin) warming rates were obtained by standing the sample tubes in air, at room temperature, or in a 25°C water bath, respectively.
INGENITO
the transfer quantum yield rJDA declines with the sixth power of the intermolecular distance. The energy transfer quantum yield +aA was found by taking fluorescence excitation spectra for the system under test, and for a control system produced either by destroying the liposomes by adding a detergent (Triton X-100), or by preparing liposomes containing only the acceptor, without donor. Both methods gave identical control spectra. As previously described in detail (7), &,* is given by
FD is the fluorescence intensity for excita-
tion at wavelength AD, the absorption maximum of the donor, and represents energy imparted to the acceptor both by energy transfer from the donor and directly (to the extent that the acceptor absorbs Turbidity and Fluorescence energy at A,,). FA, the fluorescence intenMeasurements sity for excitation at AA, the absorption A Hitachi-Perkin-Elmer MPF-3 fluo- maximum of the acceptor, represents direct rescence spectrophotometer was used. excitation of the acceptor, and serves as Turbidity was measured by reading scat- internal reference standard. The subscripts tered light intensity when both incident and T and 0 refer to the test and control system, emitted wavelengths were 750 nm where and K is a constant (related to absorbances there was no absorption or fluorescence by of the system). For the present work where any of the compounds present. Fluores- only ratios of +DA for different systems cence excitation spectra were obtained by were determined, K canceled out. The reading emission intensity at a fixed quantity F,, although theoretically a conwavelength while scanning through a range stant value, was measured separately for of excitation wavelengths. A corrected test and control systems to allow for inspectra accessory was used which compen- strumental variations. The ratio ( FDIFA)O sated for variation in lamp intensity and was highly reproducible. monochromator efficiency with waveRepresentative fluorescence excitation length. Data were corrected for attenua- spectra are shown in Fig. 1. The fluorestion of the incident and emitted beams cence emission of the cyanine dye (A,,, = while passing through the sample. 610 nm) used as energy donor overlaps well with an absorption region of chlorophyll a, Measurement of Energy Transfer the acceptor. The latter has a fluorescence Quantum Yields emission maximum at 680 nm. However, its Fluorescence induced in an energy ac- fluorescence was monitored at 730 nm ceptor by irradiation of an energy donor or where there is a subsidiary emission sensitizer occurs by electronic energy maximum, in order to avoid errors due to transfer. At donor-acceptor distances of 20 scattered light. These particular comto 100 A. tvnical of those within linosomes. Dounds were chosen because of their polar
FREEZE-THAW r
I
I
STABILITY
,
1\
450
I 550 650?mA AD Excitation Wavelength
FIG. 1. Fluorescence excitation spectra of chlorophyll a, the energy acceptor, monitored at 730 nm. Curve I, in presence of oxacarbocyanine as energy donor. Curve 2, in absence of energy donor. Curve 3, hypothetical case of 100% energy transfer. This curve corresponds to the additive absorption spectrum of energy donor and acceptor. The three curves have been normalized to each other at 660 nm.
head groups and long hydrocarbon tails which make them ideal solutes for the similarly constituted phospholipids. However, other donor-acceptor pairs, including entirely nonpolar compounds can be used. A useful donor-acceptor pair is p-terphenyl and 1,8-diphenyloctatetraene (7). The only requirement is that the probes should be practically insoluble in water and/or have little or no fluorescence in water.
511
OF LIPOSOMES
occurring. An increase in turbidity, and, in extreme cases, separation of a supernatant lipid phase gave evidence of aggregation of the lipid. Clarification of the initially slightly opalescent suspension was taken as evidence of dispersion of the lipid. Changes in turbidity are plotted in Fig. 2, as a function of CPA concentration. All samples in this series were cooled at 4”C/min, held at -27°C for 18 hr, then warmed at 5Wmin. In absence of protein, and without CPA, this freeze-thaw cycle resulted in a 30-fold increase in turbidity, evidently due to aggregation of the 300-A liposomes to much larger particles. This turbidity increase could be partly reversed by subsequent sonication. Addition of glycerol or Me,SO caused progressively smaller turbidity increases, showing that CPA are effective in preventing aggregation of the lipid upon freezing. In presence of glycoprotein, the initial turbidity was high, apparently due to the protein present in the aqueous phase. As already stated, only about 25% of the added 1
I
I
I a
IOC \
4
RESULTS
Two series of liposome suspensions, one without protein, and one with glycoprotein, were prepared in 0.16 M KCl-0.05 M Tris buffer (pH 7.9, in presence of various concentrations of glycerol or Me,SO, ranging up to 50%. Liposomes containing cytochrome c were also tested, with results similar to those with glycoprotein. Turbidity
Visual observation of turbidity changes gave a valuable clue of the basic changes
1 Oo
1 IO
30
50
I
“I. CPA
FIG. 2. Turbidity changes in liposomes as function of CPA concentration. (a) Without protein. (b) With glycoprotein. 0, Before freezing; 0, after cooling to -27”C, storage for 18 hr, and warming. Data are for glycerol as the CPA. Practically identical data obtained with Me,SO.
512
STRAUSS
AND
INGENITO
protein was incorporated into the lipid phase. Both glycerol and Me&SO reduced the turbidity at room temperature. Freezing- thawing of these protein-containing liposomes caused little or no change in turbidity, in absence or presence of CPA. These results must be interpreted with caution, however, since they show the combined effect of protein and lipid aggregation or dispersion. Moreover, the turbidity depends on both the amount of lipid undergoing change and the particle size.
series. The data were reproducible to within +5%. The findings may be summarized as follows: 1. At room temperature, protein-free liposomes had a lower &,A in the presence of CPA than in aqueous KCl, indicating a loss in membrane integrity. This effect was slight at up to 10% concentration of glycerol or Me,SO, but caused progressively larger liposome destruction at higher CPA concentrations. Energy transfer was almost completely abolished at Me,SO concentrations of 30% and over. Energy Transfer Efficiency 2. After freezing and thawing, proteinFigure 3 shows transfer efficiencies, +nA, free liposomes in KC1 suffered an approxiagain as function of CPA concentration, for mately 40% decrease in @on.Glycerol in the protein-free and glycoprotein-containing range of 5 to 20% reduced this loss to some liposomes. Transfer effrciences were mea- extent, and Me,SO in the range of 5 to 10% sured before and after subjecting the lipo- completely abolished it. The 4DA values somes to a freeze-thaw cycle. All values of were in fact slightly higher than for the KC1 4 are expressed relative to the sample in standard. 3. In protein-containing liposomes, the KD61,without CPA, before freezing. This destructive effect of CPA before freezing will be referred to as the “KC1 standard.” Separate standards were used for the was much less severe than in protein-free samples. This protective effect of the proprotein-free and the protein-containing tein was especially marked with high concentration of Me&SO. 4. After the freeze-thaw cycle, protein-containing liposomes either in KCl, or with glycerol or Me,SO, at up to 50%, showed little or no 1OSSeS in &,A. Effect of Storage Temperature Level and of Cooling and Warming Rates
t
I
0
IO
1
I
30
I
50
% CPA FIG. 3. Energy transfer yields, &,*, in liposomes as function of CPA concentration. Values relative to &A for liposomes without CPA, before freezing. (a) Without protein. (b) With glycoprotein. 0 and 0, With glycerol. 0 and n , with Me,SO. 0 and 0, Before freezing. 0 and w, After cooling and warming conditions as in Fig. 2.
The effect of holding liposomes at various temperature levels for 18 hr is shown in Fig. 4. When progressively lower storage temperatures were used, an increase in turbidity and a loss in energy transfer first appeared at -5°C. The turbidity increased roughly linearly with decreasing temperature, down to -27°C then became slightly less for storage at -70°C. Energy transfer, in contrast, decreased sharply between -5 and - ll”C, remained constant to -27”C, and increased slightly for -70°C. For both turbidity and energy transfer, the effect of changing the cooling and warming rates from 0.3 to 4”CYmin was
FREEZE-THAW
STABILITY
OF LIPOSOMES
513
ing force which acts on the dispersed PL molecules and bulk water, but not on the assembled membrane (13). / 2 20 .The destabilizing effect of CPA at room f /” temperature evidently depends on the abil2 IO ity of water-CPA mixtures to dissolve IA phospholipids in monomeric form, i.e., to I 0 O-A raise their critical micelle concentration. The greater destructiveness of Me&SO, compared to glycerol, may be due to hydrophobic interactions of its methyl groups with the phospholipid hydrocarbon chains. On freezing, in absence of CPA, both the hydration and the hydrophobic effect are suppressed, resulting in loss of membrane -25 -50 -75 0 +25 structure, a phase transition of the lipid, Storage Temperature, “C and ultimately in aggregation. CPA can FIG. 4. Effect of storage temperature for 18 hr, and of cooling and warming rate on freeze-damage in prevent such destabilization by (i) preliposomes. (a) Change in turbidity. (b) Change in venting loss of water of hydration, and posenergy transfer, &,*. A, Cooled and warmed at sibly (ii) lowering the liquid crystal-gel 0.3Wmin; 0, Cooled and warmed at 4”Cimin; 0, transition temperature. cooled at 750”Cimin and warmed at 4Wmin. Samples CPA at low temperature thus have both a contained no protein and no CPA. stabilizing and a destabilizing effect. This seems to be the reason why the protective minimal. For storage temperatures of -5 to action of CPA is maximal at an intermediate -27”C, the 0.3”C/min rate was marginally concentration. This maximum occurs near more damaging. At -7O”C, all cooling the threshold concentration for the onset of rates, including the very fast rate of membrane breakdown at room tempera75O”C/min produced the same degree of ture. Such an optimum CPA concentration damage. was also observed in the cryoprotection of red blood cells by Me&SO (at 20%) and DISCUSSION glycerol (at 11.S%) (10). The observed effects of CPA on proteinThe stabilizing effect of incorporated free liposomes can readily be accounted protein on membranes, in absence or presfor in terms of the liquid mosaic model ence of CPA, seems to arise from two effor bilayer membranes (12). Such mem- fects: (a) The hydrophilic areas of the probranes are stabilized by (a) hydration of tein protruding from the membrane attract the polar head groups of the phospho- water, and thus strengthen the hydration lipids (PL) and (b) the hydrophobic ef- layer on the membrane surface, (b) Hydrofect which prevents dispersion of the hy- phobic interaction of the nonpolar regions drocarbon chains of PL into the surrounding of the protein with the PL hydrocarbon aqueous phase. This latter effect arises chains serves to strengthen the membrane from the immobilization of water around structure. any molecularly dispersed hydrocarbon From the plot of energy transfer vs storgroup, with a consequent unfavorable de- age temperature (Fig. 4) it is very likely that crease in entropy. We may thus consider the sharp change between -5 and -11°C the bilayer membrane as being held to- arises from a phase transition of the lipid. gether by a repulsive thermodynamic driv- For egg lecithin, transition temperatures 30
514
STRAUSS
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between 0 and -15°C (depending on the lipid-water ratio) have been reported. The much more gradual change in turbidity with storage temperature suggests that the aggregation of the lipid is a secondary process, separate from, and following, the liquid crystal $ gel phase transition. It must be remembered, however, that the data show changes remaining after the samples have returned to room temperature and therefore are a measure of only that portion of change produced by cooling which was not reversed upon rewarming. The slightly lesser change seen after storage at -70°C as compared to -27°C may be due to failure of the system to attain equilibrium at this low temperature. The negligible effect of the cooling rate on the extent of bilayer destruction is in marked contrast with the behavior of various living cells (9) which show a minimum degree of destruction at certain optimum cooling and warming rates which are very different for different types of cells. This effect can account for the two-factor hypothesis of freeze damage; namely, internal freezing and damage by ice crystals (if cooled too fast), and external freezing, leading to deleterious dehydration and concentration of cell solutes (if cooled too slowly). Our finding of relative insensitivity of the bilayer itself to cooling rates is consistent with this hypothesis, which locates the cell-damaging events within the cytoplasm rather than in the cell membrane. SUMMARY
Lipid bilayer vesicles (liposomes) with and without glycoprotein incorporated into the membranes were tested for stability during freezing and thawing, in presence and absence of the cryoprotective agents (CPA) glycerol and dimethyl sulfoxide. Changes in turbidity and loss of energy transfer between fluorescent probes present in the bilayers were used to estimate membrane integrity. Freezing caused a 30 to 40% destruction
INGENITO
of protein-free liposomes, in absence of CPA. CPA at 10 to 20% concentration prevented such losses, but at higher concentrations destabilized liposomes even without freezing. Protein-containing liposomes suffered no loss on freezing in absence or presence of CPA at moderate concentrations. Lowering of the storage temperature of frozen samples within the range of -5 to - 27°C increased the freeze damage. Slower rates of cooling and warming caused a slightly greater loss. The results are interpreted in terms of the liquid mosaic model for lipid bilayers. CPA at higher concentrations destabilize bilayers by dissolving phospholipids. At moderate concentrations, however, they prevent the damaging effect of dehydration of the lipid on freezing. Proteins appear to stabilize bilayers by providing increased hydration at the membrane surface, and by additional hydrophobic binding in the membrane interior. ACKNOWLEDGMENTS
This work was supported by grants of the Charles and Johanna Busch Memorial Fund and the Rutgers Research Council. E.P.I. thanks Professor E. G. Cravalho and the U.R.O.P. at Massachusetts Institute of Technology for support. REFERENCES
1. Batzri, S., and Korn, E. D. Single bilayer liposomes prepared without sonication. Biochim. Biaphys. Acta 297, 1015- 1019 (1973). 2. Doebbler, G. F., and Rinfret, A. P. The influence of protective compounds and cooling and warming conditions on hemolysis of erythrocytes by freezing and thawing. Eiochim. Biophys. Acta 58, 449-458 (1962). 3. Lee, A. G. Fluorescence studies of chlorphyll a incorporated into lipid mixtures, and the interpretation of “phase” diagrams. Biochim. Biophys. Acra 413, 1 l-23
(1975).
4. Levitt, R. L., Cravalho, E. G., and Huggins, C. E. A membrane model describing the effect of temperature on the water conductivity of erythrocyte membranes at subzero temperatures. Cryobiology
13, 415-429
(1976).
5. Lovelock, J. E. The denaturation of lipid-protein
FREEZE-THAW
STABILITY
complexes as a cause of damage by freezing. Proc. Roy. SIX. Ser. B 147, 427-433 (1957).
6. Mazur, P. Causes of injury in frozen and thawed cells. Fed Proc. 24, S175- 182 (1965). 7. Mehreteab, A., and Strauss, G. Energy transfer and energy losses in bilayer membrane vesicles (liposomes). Phofochem. Photobiol. 27, 369-375 (1978). 8. Meryman, H. T. Physical limitations of the rapid freezing method. Proc. Roy. Sot. Ser. B 147, 452-459 (1957). 9. Miller, R. H., and Mazur, P. Survival of frozenthawed human red cells as a function of cooling
10. 11. 12. 13.
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13, and warming velocities. Cryobiology 404414 (1976). Pribor, D. B. Biological interactions between cell membranes and glycerol or DMSO. Cryobiology 12, 309-320 (1975). Siminovitch, D., and Chapman, D. Liposome bilayer model systems of freezing living cells. FEBS Lett. 16, 207-212 (1971). Singer, S. J., and Nicolson, G. L. The fluid mosaic model of the structure of cell membranes. Science 175, 720-731 (1972). Tanford, C. The hydrophobic effect and the organization of living matter. Science 200, 1012- 1018(1978).
17, 508-515
(1980)
Stabilization of Liposome of Cryoprotective
Bilayers to Freezing and Thawing: Agents and Membrane Proteins
G. STRAUSS Department
of Chemistry,
Douglass
College,
AND E. P. INGENITO’ Rutgers University, New Brunswick,
The nature of the protective action of glycerol and other cryoprotective agents (CPA) against freeze-thaw damage to biomaterials has been studied intensively for a long time. The accumulated evidence suggests that one or more of the following processes are responsible: (i) The formation of intracellular ice, found by microscopic observation (8). (ii) Osmotic dehydration of cells, resulting from the formation of ice outside, but not inside, the cell, as shown by observations of volume shrinkage of cells (6) and liposome model systems (11). This process depends on the water conductivity of the cell membrane and rate of cooling (4), and leads to a destructuve rise in salt concentration and denaturation of lipid-protein complexes (5). Hemolysis measurements on erythrocytes have shown that CPA such as glycerol and dimethyl sulfoxide (Me,SO) prevent intracellular ice formation and increase in salt concentration (6, 10). (iii) The integrity of the lipid membrane may suffer on freezing-thawing if a phase transition from a liquid-like (or liquid crystal) to a crystalline state occurs. This last process which has been studied in model systems (3), so far has received little attention in connection with cryoprotection of cells. In an effort to clarify the roles of these diverse processes and to better understand the behavior of the lipid membrane, as Received July 19, 1978; accepted May 27, 1980. L Present address: Medical Engineering and Medical Physics, Harvard-Massachusetts Institute of Technology Division of Health Sciences and Technology, Cambridge, Mass. 02139.
Copyright All rights
@ 1980 by Academic F’ress, Inc. of reproduction in any form reserved.
New Jersey 08903
such, at low temperatures, we investigated the loss in structural integrity of unilamellar lipid bilayer vesicles (liposomes) upon freezing and thawing. Liposomes are spherical or oblong vesicles, ca. 300 A in diameter. The walls, about 80 A thick, consist of lipid bilayers and enclose an aqueous phase. Lipid bilayers can serve as amphipathic solvents for other lipids, such as cholesterol, fatty acids, etc., and particularly for proteins which, in the case of living cells, constitute from 20 to 70% by weight of the lipid membrane. Liposomes can be prepared by suspending a phospholipid in an aqueous salt solution which then is sonicated. Altematively, an alcoholic solution of the lipid can be injected through a narrow hypodermic needle into a large (about 20-fold) volume of aqueous solution. These methods demonstrate that mechanical agitation or shear is sufficient to overcome the energy barrier in the transition in which a coarse suspension of the lipid (consisting of large multilamellar structures) is transformed into a suspension of small unilamellar vesicles. As we have found, freezing and thawing causes partial reversal of this transition. Unilamellar vesicles may then be restored once more by sonication. Addition of CPA, in contrast, causes dissolution of the phospholipid (at CPA concentrations beyond 20%), and irreversible destruction of the liposomes. Loss of membrane integrity was monitored (a) by change in turbidity, and (b) by loss in energy transfer between fluorescent probes incorporated into the bilayers. Turbidity changes, both positive and negative, were a sensitive, albeit only semiquantita-
508 OOl f -2240/80/050508-08$02.00/O
Effects
FREEZE-THAW
STABILITY
OF LIPOSOMES
509
cals) as energy donor, and chlorophyll a (highly purified, from Sigma Chemical Co.) as acceptor. Equal volumes of alcoholic solutions of PL (10 mg/ml), of the cyanine dye (1 mg/ml), and of chlorophyll (sufficient to give an optical density of about 0.1) were mixed. A 0.5ml volume of this solution was injected into 10 ml of an aqueous solution 0.16 M in KC1 and 0.05 M in Tris buffer, pH 7.5, containing up to 50% of glycerol or Me,SO. To prepare protein-containing liposomes, 75 mg of Cohn fraction VI serum glycoprotein (from U.S. Biochemical Corp.) was dissolved in 1.Oml water. Nine milliliters of ethanol and 75 mg PL were added, and the solution was agitated on a Vortex mixer until clear (20 min). This solution was then used in combination with the other stock solutions as before. To check how much of the protein present had become incorporated into the liposome membrane, the liposome suspensions were subjected to ultrafiltration under 20 lb/in.2 pressure through a Millipore PSJM Pellicon ultrafilter. These filters have a nominal molecular weight cutoff of 100,008 daltons, and thus retain liposomes (MW EXPERIMENTAL PROCEDURES about 2 x lo6 daltons) but pass flee serum Preparation of Liposomes glycoprotein (of the order of 50,000 dalThe phospholipid (PL) used for liposome tons). Absorption spectroscopy of the inipreparation either was lecithin from egg tial liposome suspensions and the filtrates yolk, obtained from Sigma Chemical Com- showed that 25% of the serum protein pany (Type IIE, commercial grade), and originally present had become bound, conpurified by chromatography on an alumina stituting 20% of the total membrane weight. This includes the protein trapped in the column, or DL-lecithin (1-octadec-9-enyl, 2-hexadecyl ether) obtained from Cal- internal aqueous phase of the liposomes, biochem and used without further purifi- which, however, amounts to only 0.03% of cation. Liposomes made from either of the membrane-bound protein. these PL showed comparable behavior. They were produced by the method of Bat- Freezing -Thawing Cycles zri and Korn (1) in which an ethanolic soluThree-milliliter volumes of the aqueous tion containing PL and the fluorescent liposome suspensions were placed into 13 probes is injected by a syringe through a 22- X loo-mm stoppered test tubes. Various gauge needle into a 20-fold volume of aque- cooling and warming rates were used, and ous solution. The fluorescent probes were the samples were held for 18 hr at bath temdioctadecyloxacarbocyanine-p -toluene peratures varying from -5 to -70°C. Samsulfonate (from Eastman Organic Chemi- ple temperatures were read on a remote tive, measure of liposome destruction. Energy transfer (discussed in a later section) measures the intermolecular distance of energy donor and acceptor molecules, which is small as long as the liposome membrane remains intact. Membrane destruction results in dispersion of these probe molecules over a vastly larger volume, and thus in loss of energy transfer. As reported previously (7), we found that addition of Triton X- 100, a nonionic detergent which is known to destroy liposomes, completely abolished energy transfer. This method, therefore, can be regarded as a measure of membrane integrity, in contrast to methods based on release of a trapped marker which respond not only to membrane destruction but also to creation of pores in an otherwise intact membrane. Loss of energy transfer was observed both when the lipid was dispersed by addition of CPA, and after a freeze-thaw cycle. In this latter case where the lipid became aggregated rather than dispersed, the loss in energy transfer can be ascribed to the fact that the probes became insolubie in the lipid below the transition temperature (3).
510
STRAUSS
AND
thermistor thermometer. For “slow” cooling (0.3Wmin) the sample tubes were surrounded by 50-mm-diameter tubes serving as air bath. For “fast” cooling (CC/min) the sample tubes were directly immersed in the cooling bath maintained at the desired final temperature. For temperatures from -5 to -30°C a refrigerated constant-temperature bath was used. An acetone-dry ice mixture in a Dewar flask was used to provide a -70°C bath. The latter was also used to cool samples at a very fast rate (750Wmin) by filling the suspensions into thin-walled 5-mm-diameter aluminum tubes which were directly immersed in the freezing mixture. “Slow” (0.3Wmin) and “fast” (5Wmin) warming rates were obtained by standing the sample tubes in air, at room temperature, or in a 25°C water bath, respectively.
INGENITO
the transfer quantum yield rJDA declines with the sixth power of the intermolecular distance. The energy transfer quantum yield +aA was found by taking fluorescence excitation spectra for the system under test, and for a control system produced either by destroying the liposomes by adding a detergent (Triton X-100), or by preparing liposomes containing only the acceptor, without donor. Both methods gave identical control spectra. As previously described in detail (7), &,* is given by
FD is the fluorescence intensity for excita-
tion at wavelength AD, the absorption maximum of the donor, and represents energy imparted to the acceptor both by energy transfer from the donor and directly (to the extent that the acceptor absorbs Turbidity and Fluorescence energy at A,,). FA, the fluorescence intenMeasurements sity for excitation at AA, the absorption A Hitachi-Perkin-Elmer MPF-3 fluo- maximum of the acceptor, represents direct rescence spectrophotometer was used. excitation of the acceptor, and serves as Turbidity was measured by reading scat- internal reference standard. The subscripts tered light intensity when both incident and T and 0 refer to the test and control system, emitted wavelengths were 750 nm where and K is a constant (related to absorbances there was no absorption or fluorescence by of the system). For the present work where any of the compounds present. Fluores- only ratios of +DA for different systems cence excitation spectra were obtained by were determined, K canceled out. The reading emission intensity at a fixed quantity F,, although theoretically a conwavelength while scanning through a range stant value, was measured separately for of excitation wavelengths. A corrected test and control systems to allow for inspectra accessory was used which compen- strumental variations. The ratio ( FDIFA)O sated for variation in lamp intensity and was highly reproducible. monochromator efficiency with waveRepresentative fluorescence excitation length. Data were corrected for attenua- spectra are shown in Fig. 1. The fluorestion of the incident and emitted beams cence emission of the cyanine dye (A,,, = while passing through the sample. 610 nm) used as energy donor overlaps well with an absorption region of chlorophyll a, Measurement of Energy Transfer the acceptor. The latter has a fluorescence Quantum Yields emission maximum at 680 nm. However, its Fluorescence induced in an energy ac- fluorescence was monitored at 730 nm ceptor by irradiation of an energy donor or where there is a subsidiary emission sensitizer occurs by electronic energy maximum, in order to avoid errors due to transfer. At donor-acceptor distances of 20 scattered light. These particular comto 100 A. tvnical of those within linosomes. Dounds were chosen because of their polar
FREEZE-THAW r
I
I
STABILITY
,
1\
450
I 550 650?mA AD Excitation Wavelength
FIG. 1. Fluorescence excitation spectra of chlorophyll a, the energy acceptor, monitored at 730 nm. Curve I, in presence of oxacarbocyanine as energy donor. Curve 2, in absence of energy donor. Curve 3, hypothetical case of 100% energy transfer. This curve corresponds to the additive absorption spectrum of energy donor and acceptor. The three curves have been normalized to each other at 660 nm.
head groups and long hydrocarbon tails which make them ideal solutes for the similarly constituted phospholipids. However, other donor-acceptor pairs, including entirely nonpolar compounds can be used. A useful donor-acceptor pair is p-terphenyl and 1,8-diphenyloctatetraene (7). The only requirement is that the probes should be practically insoluble in water and/or have little or no fluorescence in water.
511
OF LIPOSOMES
occurring. An increase in turbidity, and, in extreme cases, separation of a supernatant lipid phase gave evidence of aggregation of the lipid. Clarification of the initially slightly opalescent suspension was taken as evidence of dispersion of the lipid. Changes in turbidity are plotted in Fig. 2, as a function of CPA concentration. All samples in this series were cooled at 4”C/min, held at -27°C for 18 hr, then warmed at 5Wmin. In absence of protein, and without CPA, this freeze-thaw cycle resulted in a 30-fold increase in turbidity, evidently due to aggregation of the 300-A liposomes to much larger particles. This turbidity increase could be partly reversed by subsequent sonication. Addition of glycerol or Me,SO caused progressively smaller turbidity increases, showing that CPA are effective in preventing aggregation of the lipid upon freezing. In presence of glycoprotein, the initial turbidity was high, apparently due to the protein present in the aqueous phase. As already stated, only about 25% of the added 1
I
I
I a
IOC \
4
RESULTS
Two series of liposome suspensions, one without protein, and one with glycoprotein, were prepared in 0.16 M KCl-0.05 M Tris buffer (pH 7.9, in presence of various concentrations of glycerol or Me,SO, ranging up to 50%. Liposomes containing cytochrome c were also tested, with results similar to those with glycoprotein. Turbidity
Visual observation of turbidity changes gave a valuable clue of the basic changes
1 Oo
1 IO
30
50
I
“I. CPA
FIG. 2. Turbidity changes in liposomes as function of CPA concentration. (a) Without protein. (b) With glycoprotein. 0, Before freezing; 0, after cooling to -27”C, storage for 18 hr, and warming. Data are for glycerol as the CPA. Practically identical data obtained with Me,SO.
512
STRAUSS
AND
INGENITO
protein was incorporated into the lipid phase. Both glycerol and Me&SO reduced the turbidity at room temperature. Freezing- thawing of these protein-containing liposomes caused little or no change in turbidity, in absence or presence of CPA. These results must be interpreted with caution, however, since they show the combined effect of protein and lipid aggregation or dispersion. Moreover, the turbidity depends on both the amount of lipid undergoing change and the particle size.
series. The data were reproducible to within +5%. The findings may be summarized as follows: 1. At room temperature, protein-free liposomes had a lower &,A in the presence of CPA than in aqueous KCl, indicating a loss in membrane integrity. This effect was slight at up to 10% concentration of glycerol or Me,SO, but caused progressively larger liposome destruction at higher CPA concentrations. Energy transfer was almost completely abolished at Me,SO concentrations of 30% and over. Energy Transfer Efficiency 2. After freezing and thawing, proteinFigure 3 shows transfer efficiencies, +nA, free liposomes in KC1 suffered an approxiagain as function of CPA concentration, for mately 40% decrease in @on.Glycerol in the protein-free and glycoprotein-containing range of 5 to 20% reduced this loss to some liposomes. Transfer effrciences were mea- extent, and Me,SO in the range of 5 to 10% sured before and after subjecting the lipo- completely abolished it. The 4DA values somes to a freeze-thaw cycle. All values of were in fact slightly higher than for the KC1 4 are expressed relative to the sample in standard. 3. In protein-containing liposomes, the KD61,without CPA, before freezing. This destructive effect of CPA before freezing will be referred to as the “KC1 standard.” Separate standards were used for the was much less severe than in protein-free samples. This protective effect of the proprotein-free and the protein-containing tein was especially marked with high concentration of Me&SO. 4. After the freeze-thaw cycle, protein-containing liposomes either in KCl, or with glycerol or Me,SO, at up to 50%, showed little or no 1OSSeS in &,A. Effect of Storage Temperature Level and of Cooling and Warming Rates
t
I
0
IO
1
I
30
I
50
% CPA FIG. 3. Energy transfer yields, &,*, in liposomes as function of CPA concentration. Values relative to &A for liposomes without CPA, before freezing. (a) Without protein. (b) With glycoprotein. 0 and 0, With glycerol. 0 and n , with Me,SO. 0 and 0, Before freezing. 0 and w, After cooling and warming conditions as in Fig. 2.
The effect of holding liposomes at various temperature levels for 18 hr is shown in Fig. 4. When progressively lower storage temperatures were used, an increase in turbidity and a loss in energy transfer first appeared at -5°C. The turbidity increased roughly linearly with decreasing temperature, down to -27°C then became slightly less for storage at -70°C. Energy transfer, in contrast, decreased sharply between -5 and - ll”C, remained constant to -27”C, and increased slightly for -70°C. For both turbidity and energy transfer, the effect of changing the cooling and warming rates from 0.3 to 4”CYmin was
FREEZE-THAW
STABILITY
OF LIPOSOMES
513
ing force which acts on the dispersed PL molecules and bulk water, but not on the assembled membrane (13). / 2 20 .The destabilizing effect of CPA at room f /” temperature evidently depends on the abil2 IO ity of water-CPA mixtures to dissolve IA phospholipids in monomeric form, i.e., to I 0 O-A raise their critical micelle concentration. The greater destructiveness of Me&SO, compared to glycerol, may be due to hydrophobic interactions of its methyl groups with the phospholipid hydrocarbon chains. On freezing, in absence of CPA, both the hydration and the hydrophobic effect are suppressed, resulting in loss of membrane -25 -50 -75 0 +25 structure, a phase transition of the lipid, Storage Temperature, “C and ultimately in aggregation. CPA can FIG. 4. Effect of storage temperature for 18 hr, and of cooling and warming rate on freeze-damage in prevent such destabilization by (i) preliposomes. (a) Change in turbidity. (b) Change in venting loss of water of hydration, and posenergy transfer, &,*. A, Cooled and warmed at sibly (ii) lowering the liquid crystal-gel 0.3Wmin; 0, Cooled and warmed at 4”Cimin; 0, transition temperature. cooled at 750”Cimin and warmed at 4Wmin. Samples CPA at low temperature thus have both a contained no protein and no CPA. stabilizing and a destabilizing effect. This seems to be the reason why the protective minimal. For storage temperatures of -5 to action of CPA is maximal at an intermediate -27”C, the 0.3”C/min rate was marginally concentration. This maximum occurs near more damaging. At -7O”C, all cooling the threshold concentration for the onset of rates, including the very fast rate of membrane breakdown at room tempera75O”C/min produced the same degree of ture. Such an optimum CPA concentration damage. was also observed in the cryoprotection of red blood cells by Me&SO (at 20%) and DISCUSSION glycerol (at 11.S%) (10). The observed effects of CPA on proteinThe stabilizing effect of incorporated free liposomes can readily be accounted protein on membranes, in absence or presfor in terms of the liquid mosaic model ence of CPA, seems to arise from two effor bilayer membranes (12). Such mem- fects: (a) The hydrophilic areas of the probranes are stabilized by (a) hydration of tein protruding from the membrane attract the polar head groups of the phospho- water, and thus strengthen the hydration lipids (PL) and (b) the hydrophobic ef- layer on the membrane surface, (b) Hydrofect which prevents dispersion of the hy- phobic interaction of the nonpolar regions drocarbon chains of PL into the surrounding of the protein with the PL hydrocarbon aqueous phase. This latter effect arises chains serves to strengthen the membrane from the immobilization of water around structure. any molecularly dispersed hydrocarbon From the plot of energy transfer vs storgroup, with a consequent unfavorable de- age temperature (Fig. 4) it is very likely that crease in entropy. We may thus consider the sharp change between -5 and -11°C the bilayer membrane as being held to- arises from a phase transition of the lipid. gether by a repulsive thermodynamic driv- For egg lecithin, transition temperatures 30
514
STRAUSS
AND
between 0 and -15°C (depending on the lipid-water ratio) have been reported. The much more gradual change in turbidity with storage temperature suggests that the aggregation of the lipid is a secondary process, separate from, and following, the liquid crystal $ gel phase transition. It must be remembered, however, that the data show changes remaining after the samples have returned to room temperature and therefore are a measure of only that portion of change produced by cooling which was not reversed upon rewarming. The slightly lesser change seen after storage at -70°C as compared to -27°C may be due to failure of the system to attain equilibrium at this low temperature. The negligible effect of the cooling rate on the extent of bilayer destruction is in marked contrast with the behavior of various living cells (9) which show a minimum degree of destruction at certain optimum cooling and warming rates which are very different for different types of cells. This effect can account for the two-factor hypothesis of freeze damage; namely, internal freezing and damage by ice crystals (if cooled too fast), and external freezing, leading to deleterious dehydration and concentration of cell solutes (if cooled too slowly). Our finding of relative insensitivity of the bilayer itself to cooling rates is consistent with this hypothesis, which locates the cell-damaging events within the cytoplasm rather than in the cell membrane. SUMMARY
Lipid bilayer vesicles (liposomes) with and without glycoprotein incorporated into the membranes were tested for stability during freezing and thawing, in presence and absence of the cryoprotective agents (CPA) glycerol and dimethyl sulfoxide. Changes in turbidity and loss of energy transfer between fluorescent probes present in the bilayers were used to estimate membrane integrity. Freezing caused a 30 to 40% destruction
INGENITO
of protein-free liposomes, in absence of CPA. CPA at 10 to 20% concentration prevented such losses, but at higher concentrations destabilized liposomes even without freezing. Protein-containing liposomes suffered no loss on freezing in absence or presence of CPA at moderate concentrations. Lowering of the storage temperature of frozen samples within the range of -5 to - 27°C increased the freeze damage. Slower rates of cooling and warming caused a slightly greater loss. The results are interpreted in terms of the liquid mosaic model for lipid bilayers. CPA at higher concentrations destabilize bilayers by dissolving phospholipids. At moderate concentrations, however, they prevent the damaging effect of dehydration of the lipid on freezing. Proteins appear to stabilize bilayers by providing increased hydration at the membrane surface, and by additional hydrophobic binding in the membrane interior. ACKNOWLEDGMENTS
This work was supported by grants of the Charles and Johanna Busch Memorial Fund and the Rutgers Research Council. E.P.I. thanks Professor E. G. Cravalho and the U.R.O.P. at Massachusetts Institute of Technology for support. REFERENCES
1. Batzri, S., and Korn, E. D. Single bilayer liposomes prepared without sonication. Biochim. Biaphys. Acta 297, 1015- 1019 (1973). 2. Doebbler, G. F., and Rinfret, A. P. The influence of protective compounds and cooling and warming conditions on hemolysis of erythrocytes by freezing and thawing. Eiochim. Biophys. Acta 58, 449-458 (1962). 3. Lee, A. G. Fluorescence studies of chlorphyll a incorporated into lipid mixtures, and the interpretation of “phase” diagrams. Biochim. Biophys. Acra 413, 1 l-23
(1975).
4. Levitt, R. L., Cravalho, E. G., and Huggins, C. E. A membrane model describing the effect of temperature on the water conductivity of erythrocyte membranes at subzero temperatures. Cryobiology
13, 415-429
(1976).
5. Lovelock, J. E. The denaturation of lipid-protein
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STABILITY
complexes as a cause of damage by freezing. Proc. Roy. SIX. Ser. B 147, 427-433 (1957).
6. Mazur, P. Causes of injury in frozen and thawed cells. Fed Proc. 24, S175- 182 (1965). 7. Mehreteab, A., and Strauss, G. Energy transfer and energy losses in bilayer membrane vesicles (liposomes). Phofochem. Photobiol. 27, 369-375 (1978). 8. Meryman, H. T. Physical limitations of the rapid freezing method. Proc. Roy. Sot. Ser. B 147, 452-459 (1957). 9. Miller, R. H., and Mazur, P. Survival of frozenthawed human red cells as a function of cooling
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13, and warming velocities. Cryobiology 404414 (1976). Pribor, D. B. Biological interactions between cell membranes and glycerol or DMSO. Cryobiology 12, 309-320 (1975). Siminovitch, D., and Chapman, D. Liposome bilayer model systems of freezing living cells. FEBS Lett. 16, 207-212 (1971). Singer, S. J., and Nicolson, G. L. The fluid mosaic model of the structure of cell membranes. Science 175, 720-731 (1972). Tanford, C. The hydrophobic effect and the organization of living matter. Science 200, 1012- 1018(1978).