ARCHIVES
OF BIOCHEMISTRY
AND
BIOPHYSICS
Vol. 196, No. 2, September, pp. 424-429, 1979
The Effects of Cell Proliferation and Fluidity of Hepatocyte SHIRLEY Department
of Biochemistry, Los
on the Lipid Composition Plasma Membranes’
CHENG
AND DANIEL
University
of Soutbxzrn
Angeles, California
Califwia, 90&V
LEVY2 School
of Medicine,
Received February 28, 1979; revised April 1, 19’79 The fluorescence probe, 1,6-diphenyl-1,3,5-hexatriene, has been used to investigate the effects of controlled and uncontrolled growth on the dynamic properties of the lipid regions of hepatocyte plasma membranes. DPH was incubated with plasma membranes derived from quiescent and regenerating liver and Morris hepatoma 7777, and the resulting systems were studied by fluorescence polarization spectroscopy. Membranes from the rapidly growing hepatoma exhibited a significantly lower fluorescence polarization than observed in quiescent liver, suggesting the presence of a more fluid membrane lipid domain. Membranes from regenerating liver exhibited a time-dependent increase in membrane fluidity, reaching a maximum 12 h after growth stimulation. A close correspondence between membrane fluidity and the cholesterol-phospholipid ratio was also observed where a decrease in this ratio resulted in a more fluid lipid matrix. These results suggest that cell cycling, as observed in regenerating liver and Morris hepatoma 7777, results in significant increases in membrane fluidity, a property which may play an important regulatory role in various cell functions.
Recent studies have demonstrated that the cell surface plays an important role in the control of cell growth and transformation (l-3). Properties such as transport, cell surface components, hormone binding, enzymatic activities, lipid composition, cellcell interactions, and growth regulation have been shown to be altered in transformed or malignant cell systems (4). Many of the dynamic features of membranes may be determined by the fluidity of the lipid bilayer through its effect on the mobility and exposure of membrane proteins (5, 6) and on membrane-associated enzymatic activities (7, 8). In an effort to investigate the effects of controlled and uncontrolled cell proliferation on the dynamic properties of the lipid regions of hepatocyte plasma membranes, we have utilized the technique of * This investigation was supported by Grant PCM 76-23852 from the National Science Foundation and Grant CA 14089 from the National Institutes of Health. ’ To whom correspondence should be addressed. 0003-9861/79/100424-06$02.00/O Copyright 0 1979 by Academic Press, All rights
of reproduction
Inc. in any form reserved.
fluorescence polarization with the lipid probe 1,6-diphenyl-1,3,5-hexatriene (DPH).” This technique has been used extensively to define fluidity parameters in a wide variety of systems (9). Biological events such as transformation (lo- IS), growth (13), cell cycle (14), and differentiation (15) have been shown to be accompanied by alterations in the fluidity of the lipid matrix. In this study we report the use of DPH to evaluate the dynamic properties of plasma membranes derived from normal, regenerating, and malignant hepatocytes. EXPERIMENTAL
PROCEDURES
Isolation of plasma membranes. Hepatocyte plasma membranes were isolated from the livers of male Sprague-Dawley rats fed ad libitum, using a sucrose density gradient procedure according to the method of Neville (16), as modified by Ray (17), or by the aqueous two-phase polymer system as developed by Lesko
3 Abbreviations atriene. 424
used: DPH, 1,6-Diphenyl-1,3,5-hex-
EFFECTS
OF GROWTH ON PLASMA
et al. (18). Morris hepatoma 7’777, which is a rapidly growing tumor, was transplanted intramuscularly every 3 weeks in male Buffalo rats. The tumors were harvested and the neoplastic tissue carefully excised and the plasma membranes isolated, essentially according to procedures previously described (19). Regenerating liver was obtained following partial (70%) hepatectomy under ether anesthesia, as previously reported (ZO), and the plasma membranes purified utilizing an aqueous two-phase polymer system (18). The yield and purity of the membrane fractions were assessed by measuring the protein content by the method of Lowry et al. (21), as modified by Hartree (22), and the activities of marker enzymes for plasma membranes (5’-nucleotidase (18)), endoplasmic reticulum (glucose 6-phosphatase (23)), and mitochondria (succinate dehydrogenase
(241). Fluorescence labeling of plasma membranes. A solution of DPH (2 x 10m3M) in tetrahydrofuran was diluted lOOO-fold by injection into a vigorously stirred solution of phosphate-buffered saline, affording a stable dispersion which was essentially void of fluorescence. The fluorescence probe was used to determine the fluidity properties of the lipid regions of the plasma membranes (25, 26). The plasma membrane fractions in 50 mM Tris-HCl, pH 7.5, (100 pg protein/ml) were mixed with an equal volume of the DPH dispersion and incubated for 1 h at 25”C, followed by the fluorescence polarization measurements. Fluorescence microscopy. The cellular distribution of DPH in the hepatocyte system was analyzed using a Zeiss Universal fluorescence microscope. Fluorescence measurements. Fluorescence polarization and intensity were measured on a Perkin-Elmer Model MPF-4 spectrofluorometer equipped with a polarization attachment. The temperature in the cuvettes was controlled with a thermostated circulating water pump and measured with a thermistor probe. The fluorescence polarization value, P, was calculated from Eq. [l] (27),
MEMBRANE
FLUIDITY
425
- = 1 + C(r) 2, r II
r0
where r and r0 are the measured and limiting anisotropies; T, the absolute temperature; 7, the excited state lifetime; C(r), a molecular shape parameter; and r), the viscosity of the medium. This value does not consider the anisotropy of the membrane matrix or the possible heterogeneity of DPH binding sites and thus is a weighted average of all populations of the probe (28). The flow activation energy (AE) is calculated as previously described (26). Excited state lifetimes (7) are estimated from the measured relative fluorescence intensities assuming a r0 value of 11.4 ns (25). Lipid analysis. Purified plasma membranes (1 mg protein) suspended in water (200 ~1) were added to 4 ml of chloroform/methanol (l/l, v/v) and the lipids extracted at 4’C for 24 h. An additional portion of chloroform (2 ml) was added and the extract filtered through Whatman No. 1 filter paper which was then washed twice with a small volume of chloroform/ methanol (2/l, v/v). The extract was separated into two phases by the addition of 0.29% NaCl, according to the method of Folch et al. (29), and the lower phase collected after standing at 4°C for 18 h. The extract was evaporated to dryness and the cholesterol and phospholipid determined as previously described (30-32). Chemicals. 1,6-Dipenyl-1,3,5-hexatriene was purchased from Sigma Chemical Company. Other reagents were all of analytical grade. RESULTS
Plasma Membranes from Resting and Regenerating Liver and Morris Hepatoma 7777
Plasma membranes were prepared from resting and regenerating liver and from Morris hepatoma 7777. An analysis of the appropriate enzymatic markers (Table I) indicates that all preparations were highly enp = II, - 11 111 riched in 5’-nucleotidase activity and subI,, + 1, stantially free of contamination from mitowhere I,, and I, are the fluorescence intensities polarchondria and endoplasmic reticulum.
ized parallel and perpendicular, respectively, to the direction of polarization of the excitation beam. The intensity of light obtained from unlabeled control membrane suspensions due to light scattering was negligible in these experiments. Excitation was affected at 365 nm and the emission was measured at 426 nm. A 390-nm cutoff filter was used for the emission channel and 5-nm slit widths were used for both excitation and emission channels. The average microviscosity (7) of the domains occupied by DPH in the different membrane systems was derived as previously described (26) using the Perrin equation (Eq. [2])
Fluorescence Polarization Studies of Hepatocyte Plasma Membranes
Examination of DPH-treated intact hepatocytes by fluorescence microscopy indicated that the fluorescence probe was rapidly taken up into the intracellular compartments (data not shown) and thus studies on whole cells would afford data which would be a weighted average, reflecting the solubility in surface as well as intracellular hy-
426
CHENG AND LEVY TABLE
I
ENZYMATICANALYSISOF LIVERPLASMAMEMBRANES"
Resting liver Homogenate Plasma membrane Regenerating liver
5’-Nucleotidase (pmoi/30 mini mg protein)
Succinate dehydrogenase (~molihl mg protein)
Glucose 6phosphatase (pmoY30 min/ mg protein)
1.40 2 0.7 38.0 2 9.0
1.20 k 0.3 0.20 2 0.1
1.80 ” 0.8 0.40 k 0.3
1.10 c 0.5 25.0 f 6.5
0.75 t 0.2 0.14 + 0.1
3.90 + 1.3 2.20 -+ 0.8
0.90 2 0.4 23.0 5 4.8
0.73 + 0.2 0.29 k 0.2
2.25 + 0.7 1.40 k 0.4
3.20 2 1.2 56.0 2 9.0
0.89 2 0.2 0.18 k 0.1
2.00 + 0.6 0.85 k 0.3
(1 day) Homogenate Plasma membrane Regenerating liver (3 days) Homogenate Plasma membrane Morris hepatoma 7777 Homogenate Plasma membrane
0 The values reported are the means + SE. All analyses were carried out on at least five different preparations.
drophobic structures. Similar observations in several cell systems have been reported (11, 33, 34). Measurements were thus carried out on isolated plasma membranes which were incubated with an aqueous dispersion of DPH. After 60 min, a maximum fluorescence intensity was reached. As shown in Table II, the values obtained for the polarization and microviscosity of the membranes derived from the hepatoma are significantly lower than the values for the normal resting liver. This increase in lipid fluidity upon transformation has been reported in several systems (lo- 12). A plot of IogP versus l/T for these two systems (Fig. TABLE
1) gave straight lines indicating the absence of a phase transition over this temperature range. The flow activation energies (AE) which were obtained as described by Shinitzky et aZ. (26) are shown in Table II. Plasma membranes obtained from hepatocytes at various times after partial hepatectomy were incubated with DPH and the fluorescence polarization measured for each system. The results of this study are shown in Fig. 2. After 4 h the P value indicated a dramatic increase in membrane fluidity reaching a maximum after 12 h. Subsequent membrane samples indicated a gradual return to P values exhibited by resting liver, II
FLUORESCENCE POLARIZATION (P), MICROVISCOSITY ($, ANDFLOWACTIVATION ENERGY (AE) OFNORMALANDTRANSFORMED LIVERPLASMAMEMBRANES" Plasma membrane
P
(poiseT) 25°C)
AE (kcal/mol)
Hepatocyte Morris hepatoma 7777
0.224 + 0.004 0.188 2 0.004
1.20 0.86
4.9 5.7
a The values reported are the mean + SE. All determinations tions of each membrane system.
were carried out on four different prepara-
EFFECTS
OF GROWTH ON PLASMA
%+eeei+ HIT) x10'
,"K-'
FIG. 1. Temperature dependence of fluorescence polarization (P) for resting liver (0) and Morris hepatoma 7777 (0). Measurements were performed with plasma membranes (50 pg/ml) labeled with 2 x 10e6 M DPH. Each experiment was carried out at least five times. The standard error for the reported values was less than 5%.
reaching this plateau after 72 h, when the growth rate of the regenerating liver was substantially reduced. Analysis of Membrane Cholesterol and Phospholipid
MEMBRANE
FLUIDITY
427
namic properties of plasma membranes. These studies suggest that various functional properties of the membrane systems may be regulated by the nature of the lipid environment. The objective of this study was to investigate the effect of rapid controlled and uncontrolled growth on the fluidity of hepatocyte plasma membranes and to relate this property to lipid composition. A comparison of resting hepatocytes with Morris hepatoma 7777, which is a rapidly growing tumor, demonstrated that there was a significant decrease in the membrane microviscosity of the malignant cell system. This decrease was associated with an increase in the flow activation energy, L\E, which indicated a reduction in the degree of order in the lipid domain (26). The effect of controlled or nonmalignant rapid growth on plasma membrane dynamics was evaluated as a function of time in regenerating liver. Resection of 70% of the liver results in a wave of cellular proliferation in the residual parenchyma (351, the maximum of which is at a level of the proliferative activity of malignant cells. DNA synthesis reaches a maximum between 22-24 h. In contrast to malignant
An analysis of cholesterol and phospholipid constituents in the hepatocyte membrane systems is shown in Table III. A close correspondence is observed between the cholesterol-phospholipid ratio and the po0.2407 larization values (Table II, Fig. 2), where a decrease in the cholesterol-phospholipid 0. ratio is accompanied by a decrease in the polarization values, suggesting an increase in lipid fluidity. In the hepatoma system, 0. an increase in the amount of cholesterol as P well as phospholipid is observed when compared to normal liver, however, the result0. ing ratio of these components is decreased. Regenerating liver, 12 h after partial hepa0. tectomy, showed a 45% decrease in the cholesterol-phospholipid ratio which was accompanied by a 40% decrease in the polar0. ization value. As opposed to the hepatoma system, the content of phospholipid remains DAYS AFTER PARTIAL HEPATECTOMY essentially the same as observed in quiescent liver, while the content of cholesterol FIG. 2. Fluorescence polarization as a function of time after partial hepatectomy. The arrow indicates undergoes a time-dependent modulation. DISCUSSION
Fluorescence polarization techniques with DPH have been used to investigate the dy-
the polarization value obtained from quiescent liver. Measurements were performed with plasma membranes (50 pg/ml) labeled with 2 x 10eB M DPH. The values reported are the average of triplicate determinations with a standard error of less than 5%.
428
CHENG AND LEVY TABLE PHOSPHOLIPID RESTING
AND CHOLESTEROL AND REGENERATING
Plasma membrane Hepatocytes Morris hepatoma 7777 Regenerating liver (days after partial hepatectomy) 0.17 0.5 1.0 2.0 3.0 5.0
III
ANALYSIS OF PLASMA MEMBRANES DERIVED FROM LIVER AND FROM MORRIS HEPATOMA 7777a
Fmol Cholesterol mg protein
pmol Phospholipid/ mg protein
CIP
0.262 2 0.02 0.365 + 0.02
0.350 + 0.01 0.680 2 0.02
0.75 0.54
0.165 0.155 0.163 0.220 0.259 0.270
0.377 0.375 0.380 0.370 0.355 0.357
0.44 0.41 0.43 0.59 0.73 0.76
5 + 2 t f +
0.04 0.02 0.03 0.01 0.02 0.01
a The values reported are the mean t SE. All determinations preparations of each membrane system.
growth, cellular proliferation subsides after liver weight is restored and the stationary state of the organ is reestablished. The fluorescence studies (Fig. 2) on this system suggest a dramatic alteration in membrane dynamics accompanying increased cellular proliferation. As the liver returns to the quiescent state a corresponding change in membrane fluidity is observed. The concept that the fluidity of the membrane lipid domains is in part determined by the ratio of cholesterol to phospholipid (26,36) is supported by the results of these studies (Table III). A rapid and dramatic decrease in the cholesterol content of plasma membranes from regenerating liver suggests a decrease in the insertion and/or increase in the turnover of this component during the period of rapid cellular proliferation. As the liver returns to the quiescent state, the content of cholesterol is reestablished which is reflected in the fluidity measurements. Whether a more fluid lipid matrix is necessary for rapid cellular proliferation remains to be determined. The modulation of membrane cholesterol levels may be thus utilized to regulate the fluidity of membranes under various physiological conditions. Many functional differences between the membranes of rapidly proliferating and quiescent cells have been reported. Significant increases in the transport of 2-deoxy-
+ f + + 2 +
0.03 0.01 0.01 0.01 0.02 0.02
were carried out on at least three different
glucose, uridine, and phosphate have been demonstrated (37-39) in normal and transformed cell systems which are undergoing rapid cellular proliferation. Membrane-bound enzymes such as adenylate cyclase and (Na+,K+)-ATPase, which may play an important role in growth regulation through the modulation of cyclic nucleotide and ion metabolism, have been shown to be affected by membrane lipid composition and fluidity (7, 8). Studies with regenerating liver (40) have indicated that amino acid transport is elevated in the proliferating cells. The results of this study strongly suggest that controlled (regenerating liver) and uncontrolled (Morris hepatoma 7777) cellular proliferation is associated with a significant increase in membrane lipid fluidity. This alteration may regulate various cellular functions such as transport, cell cycling, and differentiation which depend on the dynamics of the cell surface. REFERENCES 1. BURGER, M. M. (1973) Fed. Proc. 32, 91-101. 2. NICOLSON, G. L. (1976) Biochim. Biophys. Acla 457, 57-108. 3. NICOLSON, G. L., POSTE, G., AND JI,‘T. H. (1977) in Cell Surface Reviews (Poste, G., and Nicolson, G. L., eds.), Vol. 3, pp. l-73, Elsevier, Amsterdam.
EFFECTS
OF GROWTH
ON PLASMA
4. NICOLSON, G. L. (1976) Biochim. Biophys. Acta 458, l-72. 5. EDIDIN, M. (1974) Annu. Rev. Biophys. Bioeng. 3, 179-201. 6. BOROCHOV, H., AND SHINITZKY, M. (1976) Proc. Nat. Acad. Sci. USA 73, 4526-4530. 7. KIMELBERG, H. K. (1975) Biochim. Biophys. Acta 413, 143-156. 8. ORLY, J., AND SCHRAMM, M. (1975) Proc. Nat. Acad. Sci. USA 72, 3433-3437. 9. SHINITZKY, M., AND BARENHOLZ, Y. (1978) Biochim. Biophys. Acta 515, 367-394. 10. SHINITZKY, M., AND INBAR, M. (1974) J. Mol. Biol. 85, 603-615. 11. VAN BLITTERSWIZK, W. J., EMMELOT, P., HILKMANN, H. R. M., OOMEN-MEULEMANS, E. P. M., AND INBAR, M. (1977) Biochim. Biophys. Acta 467, 309-320. 12. BOROCHOV, H., ZAHLER, P., WILBRANDT, W., AND SHINITZKY, M. (1977) Biochim. Biophys. Acta 470, 382-388. 13. COLLARD, J. G., DEWILDT, A., OOMEN-MEULEMANS, E. P. M., SMEEKENS, J., EMMELOT, P., AND INBAR, M. (1977)FEBSLett. 77,173-178. 14. DE LAAT, S. W., VAN DER SAAG, P. T., AND SHINITZKY, M. (1977) Proc. Nat. Acad. Sci. USA 74, 4458-4461. 15. KAWASAKI, Y., WAKAYAMA, N., KOIKE, T., KAWAI, M., AND AMANO, T. (1978) Biochim. Biophys. Acta 509, 440-449. 16. NEVILLE, D. M. (1960) J. Biophys. &o&m. Cytol. 8, 413-422. 17. RAY, T. K. (1970) Biochim. Biophys. Acta 196, 1-9. 18. LESKO, L., DONLON, M., MARINETTI, G. V., AND HARE, G. D. (1973) Biochim. Biophys. Acta 311, 173-17s. 19. BACHMANN, W., HARMS, E., HASSELS, B., HENNINGER, H., AND REUTTER, W. (1977) Biothem. J. 166, 455-462. 20. HIGGENS, G. M., AND ANDERSON, R. M. (1931) Arch. Patkol. 12, 186-202. 21. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J. (1951) J. Biol. Chem. 193, 265-275.
MEMBRANE
FLUIDITY
429
22. HARTREE, E. F. (1972) Anal. Biochem. 48, 422427. 23. ARSON, N., AND TOUSTER, 0. (1974) in Methods in Enzymology (Fleischer, S., and Packer, L., eds.), Vol. 31, P. A, pp. 90-102, Academic Press, New York. 24. KING, T. E. (1967) in Methods in Enzymology (Estabrook, R. W., and Pullman, M. E., eds.), Vol. 10, pp. 322-331, Academic Press, New York. 25. SHINITZKY, M., AND BARENHOLZ, Y. (1974) J. Biol. Chem. 249, 2652-2657. 26. SHINITZKY, M., AND INBAR, M. (1976) Biochim. Biophys. Acta 433, 133-149. 27. WEBER, G. (1953) Advan. Protein Chem. 8,415459. 28. CHEN, L. A., DALE, R. E., ROTH, S., AND BRAND, L. (1977) J. Biol. Chem. 252, 2163-2169. 29. FOLCH, J., LEES, M., AND SLOANE-STANLEY, G. H. (1957) J. Biol. Chem. 226, 497-509. 30. BROWN, H. H., ZLATKIS, A., ZAK, B., AND BOYLE, A., JR. (1954) Anal. Chem. 26, 397399. 31. COURCHAINE, A. J., MILLER, W. H., AND STEIN, D. B., JR. (1959) Clin. Chem. 5, 609-614. 32. BARTLETT, G. R. (1959)J. Biol. Chem. 234,466468. 33. JOHNSON, S. M., AND NICOLAU, C. (1977) Bio&em. Biophys. Res. Commun. 76, 869-874. 34. PAGANO, R. E., OZATO, K., AND RUYSSCHAERT, J-M. (1977) Biochim. Biophys. Acta 465, 661666. 35. BUCHER, N. L. R. (1967) N. Engl. J. Med. 277, 686-696. 36. COOPER, R. A., LESLIE, M. H., FISCHKOFF, S., SHINITZKY, M., AND SHATTIL, S. J. (1978) Biochemistry 17, 327-331. 37. KLETZIEN, R. F., AND PERDUE, J. F. (1974) J. Biol. Chem. 249, 3383-3387. 38. CUNNINGHAM, D. D., AND PARDEE, A. B. (1969) Proc. Nat. Acad. Sci. USA 64, 1049-1056. 39. ROZENGURT, E., AND DE ASUA, L. J. (1973) Proc. Nat. Acad. Sci. USA 70, 3609-3612. 40. WALKER, P. K., AND WHITFIELD, J. F. (1978) Proc. Nat. Acad. Sci. USA 75, 1394-1398.
OF BIOCHEMISTRY
AND
BIOPHYSICS
Vol. 196, No. 2, September, pp. 424-429, 1979
The Effects of Cell Proliferation and Fluidity of Hepatocyte SHIRLEY Department
of Biochemistry, Los
on the Lipid Composition Plasma Membranes’
CHENG
AND DANIEL
University
of Soutbxzrn
Angeles, California
Califwia, 90&V
LEVY2 School
of Medicine,
Received February 28, 1979; revised April 1, 19’79 The fluorescence probe, 1,6-diphenyl-1,3,5-hexatriene, has been used to investigate the effects of controlled and uncontrolled growth on the dynamic properties of the lipid regions of hepatocyte plasma membranes. DPH was incubated with plasma membranes derived from quiescent and regenerating liver and Morris hepatoma 7777, and the resulting systems were studied by fluorescence polarization spectroscopy. Membranes from the rapidly growing hepatoma exhibited a significantly lower fluorescence polarization than observed in quiescent liver, suggesting the presence of a more fluid membrane lipid domain. Membranes from regenerating liver exhibited a time-dependent increase in membrane fluidity, reaching a maximum 12 h after growth stimulation. A close correspondence between membrane fluidity and the cholesterol-phospholipid ratio was also observed where a decrease in this ratio resulted in a more fluid lipid matrix. These results suggest that cell cycling, as observed in regenerating liver and Morris hepatoma 7777, results in significant increases in membrane fluidity, a property which may play an important regulatory role in various cell functions.
Recent studies have demonstrated that the cell surface plays an important role in the control of cell growth and transformation (l-3). Properties such as transport, cell surface components, hormone binding, enzymatic activities, lipid composition, cellcell interactions, and growth regulation have been shown to be altered in transformed or malignant cell systems (4). Many of the dynamic features of membranes may be determined by the fluidity of the lipid bilayer through its effect on the mobility and exposure of membrane proteins (5, 6) and on membrane-associated enzymatic activities (7, 8). In an effort to investigate the effects of controlled and uncontrolled cell proliferation on the dynamic properties of the lipid regions of hepatocyte plasma membranes, we have utilized the technique of * This investigation was supported by Grant PCM 76-23852 from the National Science Foundation and Grant CA 14089 from the National Institutes of Health. ’ To whom correspondence should be addressed. 0003-9861/79/100424-06$02.00/O Copyright 0 1979 by Academic Press, All rights
of reproduction
Inc. in any form reserved.
fluorescence polarization with the lipid probe 1,6-diphenyl-1,3,5-hexatriene (DPH).” This technique has been used extensively to define fluidity parameters in a wide variety of systems (9). Biological events such as transformation (lo- IS), growth (13), cell cycle (14), and differentiation (15) have been shown to be accompanied by alterations in the fluidity of the lipid matrix. In this study we report the use of DPH to evaluate the dynamic properties of plasma membranes derived from normal, regenerating, and malignant hepatocytes. EXPERIMENTAL
PROCEDURES
Isolation of plasma membranes. Hepatocyte plasma membranes were isolated from the livers of male Sprague-Dawley rats fed ad libitum, using a sucrose density gradient procedure according to the method of Neville (16), as modified by Ray (17), or by the aqueous two-phase polymer system as developed by Lesko
3 Abbreviations atriene. 424
used: DPH, 1,6-Diphenyl-1,3,5-hex-
EFFECTS
OF GROWTH ON PLASMA
et al. (18). Morris hepatoma 7’777, which is a rapidly growing tumor, was transplanted intramuscularly every 3 weeks in male Buffalo rats. The tumors were harvested and the neoplastic tissue carefully excised and the plasma membranes isolated, essentially according to procedures previously described (19). Regenerating liver was obtained following partial (70%) hepatectomy under ether anesthesia, as previously reported (ZO), and the plasma membranes purified utilizing an aqueous two-phase polymer system (18). The yield and purity of the membrane fractions were assessed by measuring the protein content by the method of Lowry et al. (21), as modified by Hartree (22), and the activities of marker enzymes for plasma membranes (5’-nucleotidase (18)), endoplasmic reticulum (glucose 6-phosphatase (23)), and mitochondria (succinate dehydrogenase
(241). Fluorescence labeling of plasma membranes. A solution of DPH (2 x 10m3M) in tetrahydrofuran was diluted lOOO-fold by injection into a vigorously stirred solution of phosphate-buffered saline, affording a stable dispersion which was essentially void of fluorescence. The fluorescence probe was used to determine the fluidity properties of the lipid regions of the plasma membranes (25, 26). The plasma membrane fractions in 50 mM Tris-HCl, pH 7.5, (100 pg protein/ml) were mixed with an equal volume of the DPH dispersion and incubated for 1 h at 25”C, followed by the fluorescence polarization measurements. Fluorescence microscopy. The cellular distribution of DPH in the hepatocyte system was analyzed using a Zeiss Universal fluorescence microscope. Fluorescence measurements. Fluorescence polarization and intensity were measured on a Perkin-Elmer Model MPF-4 spectrofluorometer equipped with a polarization attachment. The temperature in the cuvettes was controlled with a thermostated circulating water pump and measured with a thermistor probe. The fluorescence polarization value, P, was calculated from Eq. [l] (27),
MEMBRANE
FLUIDITY
425
- = 1 + C(r) 2, r II
r0
where r and r0 are the measured and limiting anisotropies; T, the absolute temperature; 7, the excited state lifetime; C(r), a molecular shape parameter; and r), the viscosity of the medium. This value does not consider the anisotropy of the membrane matrix or the possible heterogeneity of DPH binding sites and thus is a weighted average of all populations of the probe (28). The flow activation energy (AE) is calculated as previously described (26). Excited state lifetimes (7) are estimated from the measured relative fluorescence intensities assuming a r0 value of 11.4 ns (25). Lipid analysis. Purified plasma membranes (1 mg protein) suspended in water (200 ~1) were added to 4 ml of chloroform/methanol (l/l, v/v) and the lipids extracted at 4’C for 24 h. An additional portion of chloroform (2 ml) was added and the extract filtered through Whatman No. 1 filter paper which was then washed twice with a small volume of chloroform/ methanol (2/l, v/v). The extract was separated into two phases by the addition of 0.29% NaCl, according to the method of Folch et al. (29), and the lower phase collected after standing at 4°C for 18 h. The extract was evaporated to dryness and the cholesterol and phospholipid determined as previously described (30-32). Chemicals. 1,6-Dipenyl-1,3,5-hexatriene was purchased from Sigma Chemical Company. Other reagents were all of analytical grade. RESULTS
Plasma Membranes from Resting and Regenerating Liver and Morris Hepatoma 7777
Plasma membranes were prepared from resting and regenerating liver and from Morris hepatoma 7777. An analysis of the appropriate enzymatic markers (Table I) indicates that all preparations were highly enp = II, - 11 111 riched in 5’-nucleotidase activity and subI,, + 1, stantially free of contamination from mitowhere I,, and I, are the fluorescence intensities polarchondria and endoplasmic reticulum.
ized parallel and perpendicular, respectively, to the direction of polarization of the excitation beam. The intensity of light obtained from unlabeled control membrane suspensions due to light scattering was negligible in these experiments. Excitation was affected at 365 nm and the emission was measured at 426 nm. A 390-nm cutoff filter was used for the emission channel and 5-nm slit widths were used for both excitation and emission channels. The average microviscosity (7) of the domains occupied by DPH in the different membrane systems was derived as previously described (26) using the Perrin equation (Eq. [2])
Fluorescence Polarization Studies of Hepatocyte Plasma Membranes
Examination of DPH-treated intact hepatocytes by fluorescence microscopy indicated that the fluorescence probe was rapidly taken up into the intracellular compartments (data not shown) and thus studies on whole cells would afford data which would be a weighted average, reflecting the solubility in surface as well as intracellular hy-
426
CHENG AND LEVY TABLE
I
ENZYMATICANALYSISOF LIVERPLASMAMEMBRANES"
Resting liver Homogenate Plasma membrane Regenerating liver
5’-Nucleotidase (pmoi/30 mini mg protein)
Succinate dehydrogenase (~molihl mg protein)
Glucose 6phosphatase (pmoY30 min/ mg protein)
1.40 2 0.7 38.0 2 9.0
1.20 k 0.3 0.20 2 0.1
1.80 ” 0.8 0.40 k 0.3
1.10 c 0.5 25.0 f 6.5
0.75 t 0.2 0.14 + 0.1
3.90 + 1.3 2.20 -+ 0.8
0.90 2 0.4 23.0 5 4.8
0.73 + 0.2 0.29 k 0.2
2.25 + 0.7 1.40 k 0.4
3.20 2 1.2 56.0 2 9.0
0.89 2 0.2 0.18 k 0.1
2.00 + 0.6 0.85 k 0.3
(1 day) Homogenate Plasma membrane Regenerating liver (3 days) Homogenate Plasma membrane Morris hepatoma 7777 Homogenate Plasma membrane
0 The values reported are the means + SE. All analyses were carried out on at least five different preparations.
drophobic structures. Similar observations in several cell systems have been reported (11, 33, 34). Measurements were thus carried out on isolated plasma membranes which were incubated with an aqueous dispersion of DPH. After 60 min, a maximum fluorescence intensity was reached. As shown in Table II, the values obtained for the polarization and microviscosity of the membranes derived from the hepatoma are significantly lower than the values for the normal resting liver. This increase in lipid fluidity upon transformation has been reported in several systems (lo- 12). A plot of IogP versus l/T for these two systems (Fig. TABLE
1) gave straight lines indicating the absence of a phase transition over this temperature range. The flow activation energies (AE) which were obtained as described by Shinitzky et aZ. (26) are shown in Table II. Plasma membranes obtained from hepatocytes at various times after partial hepatectomy were incubated with DPH and the fluorescence polarization measured for each system. The results of this study are shown in Fig. 2. After 4 h the P value indicated a dramatic increase in membrane fluidity reaching a maximum after 12 h. Subsequent membrane samples indicated a gradual return to P values exhibited by resting liver, II
FLUORESCENCE POLARIZATION (P), MICROVISCOSITY ($, ANDFLOWACTIVATION ENERGY (AE) OFNORMALANDTRANSFORMED LIVERPLASMAMEMBRANES" Plasma membrane
P
(poiseT) 25°C)
AE (kcal/mol)
Hepatocyte Morris hepatoma 7777
0.224 + 0.004 0.188 2 0.004
1.20 0.86
4.9 5.7
a The values reported are the mean + SE. All determinations tions of each membrane system.
were carried out on four different prepara-
EFFECTS
OF GROWTH ON PLASMA
%+eeei+ HIT) x10'
,"K-'
FIG. 1. Temperature dependence of fluorescence polarization (P) for resting liver (0) and Morris hepatoma 7777 (0). Measurements were performed with plasma membranes (50 pg/ml) labeled with 2 x 10e6 M DPH. Each experiment was carried out at least five times. The standard error for the reported values was less than 5%.
reaching this plateau after 72 h, when the growth rate of the regenerating liver was substantially reduced. Analysis of Membrane Cholesterol and Phospholipid
MEMBRANE
FLUIDITY
427
namic properties of plasma membranes. These studies suggest that various functional properties of the membrane systems may be regulated by the nature of the lipid environment. The objective of this study was to investigate the effect of rapid controlled and uncontrolled growth on the fluidity of hepatocyte plasma membranes and to relate this property to lipid composition. A comparison of resting hepatocytes with Morris hepatoma 7777, which is a rapidly growing tumor, demonstrated that there was a significant decrease in the membrane microviscosity of the malignant cell system. This decrease was associated with an increase in the flow activation energy, L\E, which indicated a reduction in the degree of order in the lipid domain (26). The effect of controlled or nonmalignant rapid growth on plasma membrane dynamics was evaluated as a function of time in regenerating liver. Resection of 70% of the liver results in a wave of cellular proliferation in the residual parenchyma (351, the maximum of which is at a level of the proliferative activity of malignant cells. DNA synthesis reaches a maximum between 22-24 h. In contrast to malignant
An analysis of cholesterol and phospholipid constituents in the hepatocyte membrane systems is shown in Table III. A close correspondence is observed between the cholesterol-phospholipid ratio and the po0.2407 larization values (Table II, Fig. 2), where a decrease in the cholesterol-phospholipid 0. ratio is accompanied by a decrease in the polarization values, suggesting an increase in lipid fluidity. In the hepatoma system, 0. an increase in the amount of cholesterol as P well as phospholipid is observed when compared to normal liver, however, the result0. ing ratio of these components is decreased. Regenerating liver, 12 h after partial hepa0. tectomy, showed a 45% decrease in the cholesterol-phospholipid ratio which was accompanied by a 40% decrease in the polar0. ization value. As opposed to the hepatoma system, the content of phospholipid remains DAYS AFTER PARTIAL HEPATECTOMY essentially the same as observed in quiescent liver, while the content of cholesterol FIG. 2. Fluorescence polarization as a function of time after partial hepatectomy. The arrow indicates undergoes a time-dependent modulation. DISCUSSION
Fluorescence polarization techniques with DPH have been used to investigate the dy-
the polarization value obtained from quiescent liver. Measurements were performed with plasma membranes (50 pg/ml) labeled with 2 x 10eB M DPH. The values reported are the average of triplicate determinations with a standard error of less than 5%.
428
CHENG AND LEVY TABLE PHOSPHOLIPID RESTING
AND CHOLESTEROL AND REGENERATING
Plasma membrane Hepatocytes Morris hepatoma 7777 Regenerating liver (days after partial hepatectomy) 0.17 0.5 1.0 2.0 3.0 5.0
III
ANALYSIS OF PLASMA MEMBRANES DERIVED FROM LIVER AND FROM MORRIS HEPATOMA 7777a
Fmol Cholesterol mg protein
pmol Phospholipid/ mg protein
CIP
0.262 2 0.02 0.365 + 0.02
0.350 + 0.01 0.680 2 0.02
0.75 0.54
0.165 0.155 0.163 0.220 0.259 0.270
0.377 0.375 0.380 0.370 0.355 0.357
0.44 0.41 0.43 0.59 0.73 0.76
5 + 2 t f +
0.04 0.02 0.03 0.01 0.02 0.01
a The values reported are the mean t SE. All determinations preparations of each membrane system.
growth, cellular proliferation subsides after liver weight is restored and the stationary state of the organ is reestablished. The fluorescence studies (Fig. 2) on this system suggest a dramatic alteration in membrane dynamics accompanying increased cellular proliferation. As the liver returns to the quiescent state a corresponding change in membrane fluidity is observed. The concept that the fluidity of the membrane lipid domains is in part determined by the ratio of cholesterol to phospholipid (26,36) is supported by the results of these studies (Table III). A rapid and dramatic decrease in the cholesterol content of plasma membranes from regenerating liver suggests a decrease in the insertion and/or increase in the turnover of this component during the period of rapid cellular proliferation. As the liver returns to the quiescent state, the content of cholesterol is reestablished which is reflected in the fluidity measurements. Whether a more fluid lipid matrix is necessary for rapid cellular proliferation remains to be determined. The modulation of membrane cholesterol levels may be thus utilized to regulate the fluidity of membranes under various physiological conditions. Many functional differences between the membranes of rapidly proliferating and quiescent cells have been reported. Significant increases in the transport of 2-deoxy-
+ f + + 2 +
0.03 0.01 0.01 0.01 0.02 0.02
were carried out on at least three different
glucose, uridine, and phosphate have been demonstrated (37-39) in normal and transformed cell systems which are undergoing rapid cellular proliferation. Membrane-bound enzymes such as adenylate cyclase and (Na+,K+)-ATPase, which may play an important role in growth regulation through the modulation of cyclic nucleotide and ion metabolism, have been shown to be affected by membrane lipid composition and fluidity (7, 8). Studies with regenerating liver (40) have indicated that amino acid transport is elevated in the proliferating cells. The results of this study strongly suggest that controlled (regenerating liver) and uncontrolled (Morris hepatoma 7777) cellular proliferation is associated with a significant increase in membrane lipid fluidity. This alteration may regulate various cellular functions such as transport, cell cycling, and differentiation which depend on the dynamics of the cell surface. REFERENCES 1. BURGER, M. M. (1973) Fed. Proc. 32, 91-101. 2. NICOLSON, G. L. (1976) Biochim. Biophys. Acla 457, 57-108. 3. NICOLSON, G. L., POSTE, G., AND JI,‘T. H. (1977) in Cell Surface Reviews (Poste, G., and Nicolson, G. L., eds.), Vol. 3, pp. l-73, Elsevier, Amsterdam.
EFFECTS
OF GROWTH
ON PLASMA
4. NICOLSON, G. L. (1976) Biochim. Biophys. Acta 458, l-72. 5. EDIDIN, M. (1974) Annu. Rev. Biophys. Bioeng. 3, 179-201. 6. BOROCHOV, H., AND SHINITZKY, M. (1976) Proc. Nat. Acad. Sci. USA 73, 4526-4530. 7. KIMELBERG, H. K. (1975) Biochim. Biophys. Acta 413, 143-156. 8. ORLY, J., AND SCHRAMM, M. (1975) Proc. Nat. Acad. Sci. USA 72, 3433-3437. 9. SHINITZKY, M., AND BARENHOLZ, Y. (1978) Biochim. Biophys. Acta 515, 367-394. 10. SHINITZKY, M., AND INBAR, M. (1974) J. Mol. Biol. 85, 603-615. 11. VAN BLITTERSWIZK, W. J., EMMELOT, P., HILKMANN, H. R. M., OOMEN-MEULEMANS, E. P. M., AND INBAR, M. (1977) Biochim. Biophys. Acta 467, 309-320. 12. BOROCHOV, H., ZAHLER, P., WILBRANDT, W., AND SHINITZKY, M. (1977) Biochim. Biophys. Acta 470, 382-388. 13. COLLARD, J. G., DEWILDT, A., OOMEN-MEULEMANS, E. P. M., SMEEKENS, J., EMMELOT, P., AND INBAR, M. (1977)FEBSLett. 77,173-178. 14. DE LAAT, S. W., VAN DER SAAG, P. T., AND SHINITZKY, M. (1977) Proc. Nat. Acad. Sci. USA 74, 4458-4461. 15. KAWASAKI, Y., WAKAYAMA, N., KOIKE, T., KAWAI, M., AND AMANO, T. (1978) Biochim. Biophys. Acta 509, 440-449. 16. NEVILLE, D. M. (1960) J. Biophys. &o&m. Cytol. 8, 413-422. 17. RAY, T. K. (1970) Biochim. Biophys. Acta 196, 1-9. 18. LESKO, L., DONLON, M., MARINETTI, G. V., AND HARE, G. D. (1973) Biochim. Biophys. Acta 311, 173-17s. 19. BACHMANN, W., HARMS, E., HASSELS, B., HENNINGER, H., AND REUTTER, W. (1977) Biothem. J. 166, 455-462. 20. HIGGENS, G. M., AND ANDERSON, R. M. (1931) Arch. Patkol. 12, 186-202. 21. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J. (1951) J. Biol. Chem. 193, 265-275.
MEMBRANE
FLUIDITY
429
22. HARTREE, E. F. (1972) Anal. Biochem. 48, 422427. 23. ARSON, N., AND TOUSTER, 0. (1974) in Methods in Enzymology (Fleischer, S., and Packer, L., eds.), Vol. 31, P. A, pp. 90-102, Academic Press, New York. 24. KING, T. E. (1967) in Methods in Enzymology (Estabrook, R. W., and Pullman, M. E., eds.), Vol. 10, pp. 322-331, Academic Press, New York. 25. SHINITZKY, M., AND BARENHOLZ, Y. (1974) J. Biol. Chem. 249, 2652-2657. 26. SHINITZKY, M., AND INBAR, M. (1976) Biochim. Biophys. Acta 433, 133-149. 27. WEBER, G. (1953) Advan. Protein Chem. 8,415459. 28. CHEN, L. A., DALE, R. E., ROTH, S., AND BRAND, L. (1977) J. Biol. Chem. 252, 2163-2169. 29. FOLCH, J., LEES, M., AND SLOANE-STANLEY, G. H. (1957) J. Biol. Chem. 226, 497-509. 30. BROWN, H. H., ZLATKIS, A., ZAK, B., AND BOYLE, A., JR. (1954) Anal. Chem. 26, 397399. 31. COURCHAINE, A. J., MILLER, W. H., AND STEIN, D. B., JR. (1959) Clin. Chem. 5, 609-614. 32. BARTLETT, G. R. (1959)J. Biol. Chem. 234,466468. 33. JOHNSON, S. M., AND NICOLAU, C. (1977) Bio&em. Biophys. Res. Commun. 76, 869-874. 34. PAGANO, R. E., OZATO, K., AND RUYSSCHAERT, J-M. (1977) Biochim. Biophys. Acta 465, 661666. 35. BUCHER, N. L. R. (1967) N. Engl. J. Med. 277, 686-696. 36. COOPER, R. A., LESLIE, M. H., FISCHKOFF, S., SHINITZKY, M., AND SHATTIL, S. J. (1978) Biochemistry 17, 327-331. 37. KLETZIEN, R. F., AND PERDUE, J. F. (1974) J. Biol. Chem. 249, 3383-3387. 38. CUNNINGHAM, D. D., AND PARDEE, A. B. (1969) Proc. Nat. Acad. Sci. USA 64, 1049-1056. 39. ROZENGURT, E., AND DE ASUA, L. J. (1973) Proc. Nat. Acad. Sci. USA 70, 3609-3612. 40. WALKER, P. K., AND WHITFIELD, J. F. (1978) Proc. Nat. Acad. Sci. USA 75, 1394-1398.
