Proc. Natl. Acad. Sci. USA Vol. 74, No. 10, pp. 4458-4461 October 1977
Cell Biology
Microviscosity modulation during the cell cycle of neuroblastoma cells (membrane fluidity/cell surface/lipid layer/fluorescence polarization/synchronized cells)
SIEGFRIED W. DE LAAT*t, PAUL T. VAN DER SAAG*, AND MEIR SHINITZKYt *
Hubrecht Laboratory, International Embryological Institute, Utrecht, The Netherlands; and * Department of Membrane Research, The Weizmann Institute of
Science, Rehovot, Israel
Communicated by H. A. Scheraga, July 11, 1977
ABSTRACT Microviscosity (ij) of the cell membrane lipid layer was determined in synchronized C1300 mouse neuroblastoma cells (clone Neuro-2A) by fluorescence polarization of 1,6-diphenyl-1,3,5-hexatriene. Measurements were performed directly with cells in situ on a glass substratum. The determined v value was maximal in mitosis, decreased markedly in the GI phase, remained constant at a low level during the S phase, and increased a ain during the G2 phase. These findings imply a direct role of the cell membrane fluidity in regulation of the cell cycle. Accumulated evidence has implicated the cell surface as a primary site for the control of the cell cycle, differentiation, and transformation (1-3). A variety of cell membrane propertiessuch as the expression of receptor sites and antigens, transport processes, electrical properties, enzymic activities, and morphology-are implemented in the cell-cycle events. It has been suggested that at least some of the observed changes during the cell cycle are due to the sequential insertion of new membrane components (3). The fluidity of the lipid matrix is probably one of the most important factors that regulate the dynamic features of the cell membrane, in particular the lipid-protein interactions. It not only controls the lateral and rotational mobility of membrane proteins (4), but also modulates their degree of exposure (5). In addition, membrane-associated microtubules and microfilaments might exert similar influences (6). Furthermore, it has been demonstrated that membrane-bound enzymes require and select out a specific lipid annulus to become functional. Examples of such enzymes are (Na+,K+)-ATPase (7, 8) and adenylate cyclase (9), both of which are supposed to be involved in growth control (10, 11). Fluorescence polarization measurements, using 1,6-diphenyl-1,3,5-hexatriene as a lipid probe, have shown significant differences in microviscosity between normal and transformed cells (12, 13), suggesting that the microviscosity of the membrane lipid layer is involved in the regulatory mechanisms of cell proliferation. This suggestion implies that the microviscosity of the cell membrane lipid layer is modulated during the cell cycle. In the present study we show that the microviscosity, as measured by DPH-fluorescence polarization of synchronized neuroblastoma cells, changes markedly during the cell cycle, reaching a maximum in mitosis and a minimum during S phase.
modified Eagle's medium without bicarbonate, but with 25 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (Hepes) buffer, supplemented with 10% fetal calf serum (DMEMHepes). The serum was obtained from Flow Laboratories, Irvine, Scotland. Synchronization of Cells. The cells were synchronized by selective detachment of metaphase cells upon shaking of the culture flask, without the addition of any mitotic inhibitor. In each experiment cells from the stock culture were replated in 150-cm2 tissue culture flasks (Costar, Cambridge, MA), 15 X 106 cells in 20 ml of DMEM-Hepes per flask. From these flasks mitotic cells were harvested the next day by shaking (2-3 min), either manually or on a gyratory shaker at 400 rpm, after reduction of the medium volume to 5 ml. The first shake-off was discarded. The cells obtained by subsequent shakes (30-to45-min intervals) were replated. For [3H]thymidine incorporation studies, cells were replated on sterile cover slips (1.5 cm in diameter) in Repli dishes (Sterilin, Teddington, England) at 105 cells in 1 ml of DMEM-Hepes per cover slip and incubated for 30 min at 370 in the presence of [3H]thymidine at 5 MiCi/ml (52 Ci/mmol; Radiochemical Centre, Amersham, England). After incubation the cells on the cover slips were washed twice in ice-cold Dulbecco's phosphate-buffered saline (PBS), treated with 10% trichloroacetic acid for 30 min, and washed twice with 5% trichloroacetic acid and then with 1:1 (vol/vol) ethanol/ether. The cover slips were air dried and transferred to scintillation vials containing 10 ml of Instagel (Packard Instrument Co., Downers Grove, IL). Radioactivity was determined in a Packard model 2450 Tri-Carb liquid scintillation counter. For time-lapse cinematographic analysis mitotic cells were replated at an equivalent density in 25-cm2 Costar tissue culture flasks. The films obtained were used for mitotic index determinations. Fluorescence Labeling of Cells. The fluorescent hydrocarbon 1,6-diphenyl-1,3,5-hexatriene (DPH) was used as a probe for determining the fluidity properties of the hydrocarbon region of the cell membrane lipid layer (15, 16). The labeling dispersion of 2-6 ,gM DPH in PBS was prepared by appropriate dilution of 2 mM DPH (Koch-Light Laboratories, Colnbrook, England) in tetrahydrofuran with vigorously stirred PBS. Stirring was continued under N2 bubbling for 30 min at room temperature for evaporation of the tetrahydrofuran. For fluorescence polarization measurements of cells in situ attached to the glass substratum, 5 X 104 cells were grown in Sterilin Repli dishes on rectangular cover slips (0.9 X 1.8 cm). The ad-
MATERIALS AND METHODS Cells. C1300 mouse neuroblastoma cells, clone Neuro-2A (14), were obtained from the American Type Culture Collection, Rockville, MD. The cells were grown in Dulbecco's
Abbreviations: DPH, 1,6-diphenyl-1,3,5-hexatriene; Hepes, N-2hydroxyethylpiperazine-N'-2-ethanesulfonic acid; DMEM-Hepes, Dulbecco's modified Eagle's medium without bicarbonate but with 25 mM Hepes and 10% fetal calf serum; PBS, phosphate-buffered saline. t To whom reprint requests should be addressed.
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U. S. C. §1734 solely to indicate this fact.
4458
Proc. Nati. Acad. Sci. USA 74 (1977)
Cell Biology: de Laat et al. M
O- 5 -
Fluorescence detection and readout
S
M G2
G1
,,
x
E
aO 4 -
c
365 nm Polarizer
CD
.0
0
E
0.
150 =a) U
FIG. 1. Instrumental setup used for measurements of fluorescence polarization of cells in monolayer on a thin glass substratum.
s'I
V1 0-
herent cells were washed twice with PBS and incubated at 370 for 30-45 min in the labeling solution. The cover slips were then washed twice with PBS and placed in 1 X 1-cm quartz cuvettes
(see below). Mitotic cells were centrifuged directly after collection, washed with PBS, and incubated in suspension at 370 for 30 min in the labeling solution. The cells were then washed with PBS and resuspended in an appropriate volume of PBS. For fluorescence polarization measurements in suspensions of exponentially growing cells, the cells were grown at densities equivalent to those used for measurements in situ, for 24 hr in 75-cm2 tissue culture flasks. The flasks were washed twice with PBS and incubated for 30 min in the labeling solution. Two different procedures were used for removal of the labeled cells from the substratum. (i) The flasks were washed twice with PBS, flushed with 0.25% trypsin in PBS, and incubated upside-down for 5 min at 37°. Trypsinization was stopped by adding 10 ml of DMEM-Hepes. The cells were removed, centrifuged, washed with PBS, and resuspended in an appropriate volume of PBS. (ii) The flasks were washed twice with Ca2+and Mg2+-free PBS and incubated for 5 min at 370 in 5 ml of Ca2+- and Mg2+-free PBS containing 2.5 mM EDTA. The cells were removed, centrifuged, and resuspended in an appropriate volume of PBS. For all fluorescence measurements of cells in suspension, 0.5 to 1.0 X 106 cells per ml of PBS were used. Fluorescence Polarization Measurements. Fluorescence polarization and intensity were measured with a specially designed instrument employing the mercury lamp and the cuvette holder from a Zeiss PMQ-2 fluorescence spectrophotometer. The cuvette holder can contain four cuvettes simultaneously, which can be determined sequentially. For excitation the Hg 365 nm band, which was passed through a turnable sheet polarizer, was used. The fluorescence intensity was measured after the light had passed through a turnable sheet analyzer and an aqueous 2 M NaNO2 solution, used as a cut-off filter for wavelengths below 390 nm. The temperature was controlled by thermoelectric elements (Cambion model 3959-01, Cambridge Thermoionic Corp., Cambridge, MA) and held to +0. 10 over the range of 0-40°. Temperature was measured continuously by a diode immersed in the sample. The cuvette identification number, temperature, fluorescence intensity with the relative position of the polarizer and analyzer (parallel, Ill, or perpendicular, II) were recorded on punch tape for subsequent computer calculations. In this way the temperature dependence of the fluorescence polarization over the range of 0-40° could be measured in four independent samples within 15 min. For measurements of cells in situ on a glass substratum, cover slips were positioned in a cuvette filled with PBS, as shown in Fig. 1. In measurements of cell suspensions, the cells were gently mixed before each reading. In all cases corrections for stray light and intrinsic fluorescence were made by subtracting the values for Ill and _1 of unlabeled samples from those of identical but labeled samples. The corrections for cover slips were below 20% and for suspensions, below 3%.
4459
t
0
2
4 8 6 10 Hours after shake-off
FIG. 2. Synchronization of Neuro-2A cells: [3H]thymidine incorporation (0) and % cell number (0). Mean values of three independent samples are given for the [3H]thymidine incorporation. The number of cells was determined from time-lapse films.
Determination of Microviscosity. The fluorescence anisotropy, r, and the total fluorescence intensity, F, were calculated from the corrected fluorescence intensities III and I according to Eq. 1 F~
; F
=Ill+ 2I1
[1]
Microviscosity (-) is an operational term that translates the resistance of the environments to rotation of the probe molecule into macroscopic units (poise, = 0.1 Pa-sec). Because the membrane structure is anisotropic the obtained value is a weight average of the subpopulations. Microviscosities were derived as described previously (12, 15, 16) by a method based on the Perrin equation (Eq. 2) for rotational depolarization of a nonspherical fluorophore: - = 1 + C(r)-r wi
[2]
in which rO and r are the limiting and measured fluorescence anisotropies, T is the absolute temperature, and r is the excited state lifetime, which was calculated from a second-order polynomial regression line of the dependence of F on temperature (15). C(r) is a molecular shape parameter having a precalibrated value for each value of r. From the determined r, T, and r values and the corresponding C(r) the microviscosity was derived. The temperature dependence of v was expressed as log v versus l/T. A linear regression analysis was made to determine the flow activation energy, AE, and to detect possible phase transitions (15, 16). All computations and plots were made with a Wang 2200-B system (Wang Laboratories, Tewksbury,
MA). Fluorescence Microscopy. A Zeiss Universal microscope was used for fluorescence microscopy. Fluorescence micrographs of DPH-labeled cells were taken with epi-illumination at 365 nm and emission above 410 nm. To overcome the rapid bleaching of DPH-labeled cells, micrographs were taken with a highly sensitive recording film (Kodak 2475), which was specially processed to increase its sensitivity to about 50
DIN.
RESULTS The Neuro-2A cells were chosen for this study because they can be easily synchronized without the use of metabolic or mitotic inhibitors. The mitotic shake-off method, under the employed
Proc. Natl. Acad. Sci. USA 74 (1977)
Cell Biology: de Laat et al.
4460 -
0
0.20
1.0
Table 1. Microviscosity of the membrane lipid layer of exponentially growing Neuro-2A cells in PBS, mean values ± SEM
a S
0.
Cell system
Microviscosity at 370, poise
On glass substratum After EDTA detachment After trypsin detachment
2.1 ± 0.1 2.9 ± 0.1 3.1 + 0.1
C0
*0c
xCu
0.15
E
U.
a)
cJC
a)
upC)cn
To avoid possible alterations in cell membrane by detach-
0
ment treatments, as with trypsin or EDTA, we have developed
01
0.10
0
10
20
30
40 50 Incubation time, min
60
FIG. 3. Fluorescence anisotropy and relative fluorescence intensity (F/Fmax) during the uptake of DPH by exponentially growing Neuro-2A cells.
conditions, resulted in a high degree of synchronization. The yield of mitotic cells per shake-off was 2-4% of the total cell number. Fig. 2 illustrates the degree of synchronization, as determined by [3H]thymidine incorporation and time-lapse cinematography. The mean duration of the cell cycle was 9.5 hr in the synchronized cultures, which was not significantly different from that observed in the exponentially growing cultures. The duration of the S phase was about 6 hr. while the GI and G2 phases lasted about 1.5 hr and 1 hr, respectively. Labeling of intact cells with fluorescent hydrocarbons like DPH or perylene may be only initially confined to the cell surface membrane. To check for any possible contribution of DPH incorporated into intracellular compartments, the fluorescence anisotropy and the total fluorescence intensity were determined simultaneously during the labeling period. Fig. 3 gives an example of such a measurement. No significant alterations in the r value were observed during the uptake of DPH, indicating that DPH is uniformly dissolved into one fluid compartment. Taking into account that the plasma membrane is the first compartment DPH molecules encounter during their uptake, the contribution of intracellular DPH was thus assumed to be negligible. Furthermore, in the fluorescence microscope, DPH-labeled neuroblastoma cells (Fig. 4) displayed an evenly glowing periphery with minor intracellular fluorescence, in contrast to reported data on 3T3 cells (17).
a simple method by which fluorescence polarization can be measured with the cells attached to a thin glass substratum (see Fig. 1). Table 1 shows the differences in microviscosity obtained for cells attached to substratum and for cells in suspension, after trypsin or EDTA treatment. Clearly, the latter treatments both lead to an overall increase in microviscosity, trypsin to a larger extent than EDTA. Four independent synchronization experiments were carried out in triplicate for each time. The determined microviscosity parameters of the synchronized cells are summarized in Figs. 5 and 6, and Table 2. As stated before, the measurements were made on cells attached to a glass substratum, with the exception of the first point (t = 0 hr) in Fig. 5. These mitotic cells were measured in suspension, immediately after shake-off, because the reattachment period lasted about 15 min. As shown, the microviscosity appears to be maximal during mitosis, and at the GI phase it decreases rapidly, reaching about a half-maximal value at the onset of DNA synthesis. The microviscosity remained constant at this low level during the S phase and increased again during the G2 phase. The determined X value always reached a lower maximum during the second mitosis as compared to the first (Fig. 5). This was presumably due to partial loss of synchrony with time (see Fig. 2), because the first cells started to divide again at 8 hr after the shake-off, but all cells were divided only after 11 hr. Consequently, at any time between 8 and 11 hr after shake-off only a fraction of the cells were in mitosis and the apparent microviscosity had an intermediate value between the maximum of mitotic cells and the minimum of S-phase cells. Plots of log I versus 1/T for cells in different phases of the cell cycle gave linear relations (see Fig. 6) with no signs of phase transitions. The flow activation enerS
M 3.5 -i
3.0
-
2.5
-
2.0
-
M
G6
G2
a),
.0
F
Igr
FIG. 4. Micrographs of Neuro-2A cells after 90-min labeling with DPH showing the label uniformly distributed over the cell surface with minor intracellular fluorescence; bars equal 10 tm. (A) Phase contrast micrograph; (B) fluorescence micrographs of the same cells. After 30-min labeling, when fluorescence measurements were made, the intracellular fluorescence was largely missing. However, the overall fluorescence was too weak to be photographed.
-~
"
1.5 -r 0
I-4
2
4
-~
6
-f
-
8
10
12
Hours after shake-off FIG. 5. Microviscosity (ij) at 370 as a function of time during the cell cycle of Neuro-2A cells. Mean + SEM is indicated. All measurements were made on cells in situ on a glass substratum, with the ex-
ception of the freshly selected mitotic cells (t measured in suspension.
=0
hr), which were
Cell Biology: de Laat et al.
Proc. Nati. Acad. Sci. USA 74 (1977)
4461
Table 2. Measured fluorescence anisotropy (r), and corresponding microviscosity (61), and flow activation energy (AE) of the membrane lipid layer at different stages of the cell cycle
10
Stage of cell cycle M Early G1 Late G1 S
5
gr
250
370 r 0.198 0.181 0.166 0.150
v7 3.5 2.9 2.2 1.9
r
0.225 0.214 0.192 0.178
40 v 4.7 4.2 3.2 2.7
r
i
0.269 0.262 0.241 0.232
8.7 8.4 6.3 5.2
AE 4.8 5.4 5.4 5.2
v is expressed in poise; AE is expressed in kcal/mol (1 cal = 4.184 J).
I 3.6 3.4 3.5 (1/T) X 10-3, K-l FIG. 6. Temperature dependence of microviscosity (ii) in Neuro-2A cells at different stages of the cell cycle, plotted on a logarithmic scale versus 1T. Mitosis (0), early G1 phase (0), late G1 phase (*), and S phase (*).
3.2
3.3
gies (AE) obtained from the slopes of these lines are given in Table 2.
DISCUSSION The microviscosity of the cell membrane lipid layer varies by a factor of two during the cell cycle of neuroblastoma cells. It is maximal in mitosis, minimal during the S phase, and shows a rapid decrease and increase in the Gi and G2 phase, respectively. This confirms earlier suggestions that the cell membrane of mitotic cells is more rigid than that of interphase cells (18, 19).
Previously, differences in microviscosity have been associated with growth characteristics. Resting lymphocytes were found to have a higher microviscosity than proliferating lymphoblasts (20) and lymphoma cells (12), while normal 3T3 cells showed a lower microviscosity than virus-transformed 3T3 cells (13). In view of the described cell cycle dependency of the microviscosity, these differences, as measured in nonsynchronized cell populations, could at least partially be a reflection of differences in the relative distribution of the cells through the cell cycle. It seems likely that the rate of lipid insertion into the cell surface is maximal during mitosis and the GI phase; for references see ref. 3. Accordingly, our results indicate a rapid alteration of the lipid composition of the cell membrane during the Gi phase (16). An excess insertion of lipids during this phase could also explain the described surface bleb formation (21) and decrease in the density of intramembranous particles in the GI phase (22). A great number of publications have led to the conclusion that surface antigens and (lectin-) receptor sites exhibit their lowest degree of expression or accessibility during the S phase; for references see ref 3. In view of the reported relationship between the microviscosity and the displacements of proteins perpendicular to the plane of the membrane (5), the possibility should be considered that the variation during the cell cycle of the expression of membrane receptor sites originates from alterations in the fluidity of the membrane lipid layer, as described here. Cyclic nucleotide metabolism (23) and ion metabolism (24) have been inferred to be involved in growth regulation. In both cases the activities of membrane-bound enzymes, i.e., adenylate
cyclase and (Na+, K+)-ATPase have been shown to be dependent on the membrane lipid composition and fluidity (7-9). Assuming that the observed correlation of the cell cycle with membrane microviscosity is a general phenomenon, it can be speculated that the membrane exerts its influence on the regulation of the cell cycle by modulating the activities of such. enzymes through changes in the lipid fluidity. We thank Mr. J. H. Beeker and Mr. P. Hogeweg for excellent technical support in constructing the fluorescence polarization apparatus; Mrs. Manuela M. Marques da Silva Pimenta Guarda, Miss Alie Feyen, and Mr. M. A. da Silva Guarda for their skillful technical assistance; and Mr. L. Boom for the photography. This work was supported by a travel grant (to S.W.deL.) of the Netherlands Organization for the Advancement of Pure Research.
1. Nicolson, G. L. (1976) Biochim. Biophys. Acta 457,57-108. 2. Nicolson, G. L. (1976) Biochim. Biophys. Acta 458, 1-72. 3. Bluemink, J. G. & de Laat, S. W. (1977) in Cell Surface Reviews, eds. Post, G. & Nicolson, G. L. (North-Holland Publishing Co., Amsterdam), Vol. 4, in press. 4. Edidin, M. (1974) Annu. Rev. Biophys. Bioeng. 3, 179-201. 5. Borochov, H. & Shinitzky, M. (1976) Proc. Nati. Acad. Sci. USA
73,4526-4530. 6. Schlessinger, J., Elson, E. L., Webb, W. W., Yahara, I., Rutishauser, U. & Edelman, G. M. (1977) Proc. Natl. Acad. Sci. USA 74, 1110-1114. 7. Kimelberg, H. K. & Papahadjopoulos, D. (1972) Biochim. Biophys. Acta 282, 277-292. 8. Kimelberg, H. K. (1975) Biochim. Biophys. Acta 413, 143156. 9. Orly, J. & Schramm, M. (1975) Proc. Natl. Acad. Sci. USA 72, 3433-3437. 10. Anderson, W. B., Russell, T. R., Carchman, R. A. & Pastan, I. (1973) Proc. Natl. Acad. Sci. USA 70,3802-3805. 11. Graham, F. M., Summer, M. C. B., Curtis, D. H. & Pasternak, C. A. (1973) Nature 246, 291-295. 12. Shinitzky, M. & Inbar, M. (1974) J. Mol. Biol. 85,603-615. 13. Fuchs, P., Parola, A., Robbins, P. W. & Blout, E. R. (1975) Proc. Natl. Acad. Sci. USA 72,3351-3354. 14. Klebe, R. J. & Ruddle, F. H. (1969) J. Cell Biol. 43, 69A. 15. Shinitzky, M. & Barenholz, Y. (1974) J. Biol. Chem. 249, 2652-2657. 16. Shinitzky, M. & Inbar, M. (1976) Biochim. Biophys. Acta 433, 133-147. 17. Berlin, R. D. & Fera, J. P. (1977) Proc. Natl. Acad. Sci. USA 74, 1072-1076. 18. Furcht, T. & Scott, R. E. (1974) Exp. Cell Res. 88, 311-318. 19. Garrido, J. (1975) Exp. Cell Res. 94, 159-175. 20. Inbar, M. & Shinitzky, M. (1975) Eur. J. Immunol. 5, 166170. 21. Porter, K., Prescott, D. & Frey, J. (1973) J. Cell Biol. 57, 815836. 22. Scott, R. E., Carter, R. L. & Kidwell, W. R. (1971) Nature New Biol. 233, 219-220. 23. Burger, M. M., Bombik, B. M., Breckenridge, B. M. L. & Sheppard, J. R. (1972) Nature New Biol. 239, 161-163. 24. Cone, C. D., Jr. (1974) Ann. N.Y. Acad. Sci. 238,420-435.
Cell Biology
Microviscosity modulation during the cell cycle of neuroblastoma cells (membrane fluidity/cell surface/lipid layer/fluorescence polarization/synchronized cells)
SIEGFRIED W. DE LAAT*t, PAUL T. VAN DER SAAG*, AND MEIR SHINITZKYt *
Hubrecht Laboratory, International Embryological Institute, Utrecht, The Netherlands; and * Department of Membrane Research, The Weizmann Institute of
Science, Rehovot, Israel
Communicated by H. A. Scheraga, July 11, 1977
ABSTRACT Microviscosity (ij) of the cell membrane lipid layer was determined in synchronized C1300 mouse neuroblastoma cells (clone Neuro-2A) by fluorescence polarization of 1,6-diphenyl-1,3,5-hexatriene. Measurements were performed directly with cells in situ on a glass substratum. The determined v value was maximal in mitosis, decreased markedly in the GI phase, remained constant at a low level during the S phase, and increased a ain during the G2 phase. These findings imply a direct role of the cell membrane fluidity in regulation of the cell cycle. Accumulated evidence has implicated the cell surface as a primary site for the control of the cell cycle, differentiation, and transformation (1-3). A variety of cell membrane propertiessuch as the expression of receptor sites and antigens, transport processes, electrical properties, enzymic activities, and morphology-are implemented in the cell-cycle events. It has been suggested that at least some of the observed changes during the cell cycle are due to the sequential insertion of new membrane components (3). The fluidity of the lipid matrix is probably one of the most important factors that regulate the dynamic features of the cell membrane, in particular the lipid-protein interactions. It not only controls the lateral and rotational mobility of membrane proteins (4), but also modulates their degree of exposure (5). In addition, membrane-associated microtubules and microfilaments might exert similar influences (6). Furthermore, it has been demonstrated that membrane-bound enzymes require and select out a specific lipid annulus to become functional. Examples of such enzymes are (Na+,K+)-ATPase (7, 8) and adenylate cyclase (9), both of which are supposed to be involved in growth control (10, 11). Fluorescence polarization measurements, using 1,6-diphenyl-1,3,5-hexatriene as a lipid probe, have shown significant differences in microviscosity between normal and transformed cells (12, 13), suggesting that the microviscosity of the membrane lipid layer is involved in the regulatory mechanisms of cell proliferation. This suggestion implies that the microviscosity of the cell membrane lipid layer is modulated during the cell cycle. In the present study we show that the microviscosity, as measured by DPH-fluorescence polarization of synchronized neuroblastoma cells, changes markedly during the cell cycle, reaching a maximum in mitosis and a minimum during S phase.
modified Eagle's medium without bicarbonate, but with 25 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (Hepes) buffer, supplemented with 10% fetal calf serum (DMEMHepes). The serum was obtained from Flow Laboratories, Irvine, Scotland. Synchronization of Cells. The cells were synchronized by selective detachment of metaphase cells upon shaking of the culture flask, without the addition of any mitotic inhibitor. In each experiment cells from the stock culture were replated in 150-cm2 tissue culture flasks (Costar, Cambridge, MA), 15 X 106 cells in 20 ml of DMEM-Hepes per flask. From these flasks mitotic cells were harvested the next day by shaking (2-3 min), either manually or on a gyratory shaker at 400 rpm, after reduction of the medium volume to 5 ml. The first shake-off was discarded. The cells obtained by subsequent shakes (30-to45-min intervals) were replated. For [3H]thymidine incorporation studies, cells were replated on sterile cover slips (1.5 cm in diameter) in Repli dishes (Sterilin, Teddington, England) at 105 cells in 1 ml of DMEM-Hepes per cover slip and incubated for 30 min at 370 in the presence of [3H]thymidine at 5 MiCi/ml (52 Ci/mmol; Radiochemical Centre, Amersham, England). After incubation the cells on the cover slips were washed twice in ice-cold Dulbecco's phosphate-buffered saline (PBS), treated with 10% trichloroacetic acid for 30 min, and washed twice with 5% trichloroacetic acid and then with 1:1 (vol/vol) ethanol/ether. The cover slips were air dried and transferred to scintillation vials containing 10 ml of Instagel (Packard Instrument Co., Downers Grove, IL). Radioactivity was determined in a Packard model 2450 Tri-Carb liquid scintillation counter. For time-lapse cinematographic analysis mitotic cells were replated at an equivalent density in 25-cm2 Costar tissue culture flasks. The films obtained were used for mitotic index determinations. Fluorescence Labeling of Cells. The fluorescent hydrocarbon 1,6-diphenyl-1,3,5-hexatriene (DPH) was used as a probe for determining the fluidity properties of the hydrocarbon region of the cell membrane lipid layer (15, 16). The labeling dispersion of 2-6 ,gM DPH in PBS was prepared by appropriate dilution of 2 mM DPH (Koch-Light Laboratories, Colnbrook, England) in tetrahydrofuran with vigorously stirred PBS. Stirring was continued under N2 bubbling for 30 min at room temperature for evaporation of the tetrahydrofuran. For fluorescence polarization measurements of cells in situ attached to the glass substratum, 5 X 104 cells were grown in Sterilin Repli dishes on rectangular cover slips (0.9 X 1.8 cm). The ad-
MATERIALS AND METHODS Cells. C1300 mouse neuroblastoma cells, clone Neuro-2A (14), were obtained from the American Type Culture Collection, Rockville, MD. The cells were grown in Dulbecco's
Abbreviations: DPH, 1,6-diphenyl-1,3,5-hexatriene; Hepes, N-2hydroxyethylpiperazine-N'-2-ethanesulfonic acid; DMEM-Hepes, Dulbecco's modified Eagle's medium without bicarbonate but with 25 mM Hepes and 10% fetal calf serum; PBS, phosphate-buffered saline. t To whom reprint requests should be addressed.
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U. S. C. §1734 solely to indicate this fact.
4458
Proc. Nati. Acad. Sci. USA 74 (1977)
Cell Biology: de Laat et al. M
O- 5 -
Fluorescence detection and readout
S
M G2
G1
,,
x
E
aO 4 -
c
365 nm Polarizer
CD
.0
0
E
0.
150 =a) U
FIG. 1. Instrumental setup used for measurements of fluorescence polarization of cells in monolayer on a thin glass substratum.
s'I
V1 0-
herent cells were washed twice with PBS and incubated at 370 for 30-45 min in the labeling solution. The cover slips were then washed twice with PBS and placed in 1 X 1-cm quartz cuvettes
(see below). Mitotic cells were centrifuged directly after collection, washed with PBS, and incubated in suspension at 370 for 30 min in the labeling solution. The cells were then washed with PBS and resuspended in an appropriate volume of PBS. For fluorescence polarization measurements in suspensions of exponentially growing cells, the cells were grown at densities equivalent to those used for measurements in situ, for 24 hr in 75-cm2 tissue culture flasks. The flasks were washed twice with PBS and incubated for 30 min in the labeling solution. Two different procedures were used for removal of the labeled cells from the substratum. (i) The flasks were washed twice with PBS, flushed with 0.25% trypsin in PBS, and incubated upside-down for 5 min at 37°. Trypsinization was stopped by adding 10 ml of DMEM-Hepes. The cells were removed, centrifuged, washed with PBS, and resuspended in an appropriate volume of PBS. (ii) The flasks were washed twice with Ca2+and Mg2+-free PBS and incubated for 5 min at 370 in 5 ml of Ca2+- and Mg2+-free PBS containing 2.5 mM EDTA. The cells were removed, centrifuged, and resuspended in an appropriate volume of PBS. For all fluorescence measurements of cells in suspension, 0.5 to 1.0 X 106 cells per ml of PBS were used. Fluorescence Polarization Measurements. Fluorescence polarization and intensity were measured with a specially designed instrument employing the mercury lamp and the cuvette holder from a Zeiss PMQ-2 fluorescence spectrophotometer. The cuvette holder can contain four cuvettes simultaneously, which can be determined sequentially. For excitation the Hg 365 nm band, which was passed through a turnable sheet polarizer, was used. The fluorescence intensity was measured after the light had passed through a turnable sheet analyzer and an aqueous 2 M NaNO2 solution, used as a cut-off filter for wavelengths below 390 nm. The temperature was controlled by thermoelectric elements (Cambion model 3959-01, Cambridge Thermoionic Corp., Cambridge, MA) and held to +0. 10 over the range of 0-40°. Temperature was measured continuously by a diode immersed in the sample. The cuvette identification number, temperature, fluorescence intensity with the relative position of the polarizer and analyzer (parallel, Ill, or perpendicular, II) were recorded on punch tape for subsequent computer calculations. In this way the temperature dependence of the fluorescence polarization over the range of 0-40° could be measured in four independent samples within 15 min. For measurements of cells in situ on a glass substratum, cover slips were positioned in a cuvette filled with PBS, as shown in Fig. 1. In measurements of cell suspensions, the cells were gently mixed before each reading. In all cases corrections for stray light and intrinsic fluorescence were made by subtracting the values for Ill and _1 of unlabeled samples from those of identical but labeled samples. The corrections for cover slips were below 20% and for suspensions, below 3%.
4459
t
0
2
4 8 6 10 Hours after shake-off
FIG. 2. Synchronization of Neuro-2A cells: [3H]thymidine incorporation (0) and % cell number (0). Mean values of three independent samples are given for the [3H]thymidine incorporation. The number of cells was determined from time-lapse films.
Determination of Microviscosity. The fluorescence anisotropy, r, and the total fluorescence intensity, F, were calculated from the corrected fluorescence intensities III and I according to Eq. 1 F~
; F
=Ill+ 2I1
[1]
Microviscosity (-) is an operational term that translates the resistance of the environments to rotation of the probe molecule into macroscopic units (poise, = 0.1 Pa-sec). Because the membrane structure is anisotropic the obtained value is a weight average of the subpopulations. Microviscosities were derived as described previously (12, 15, 16) by a method based on the Perrin equation (Eq. 2) for rotational depolarization of a nonspherical fluorophore: - = 1 + C(r)-r wi
[2]
in which rO and r are the limiting and measured fluorescence anisotropies, T is the absolute temperature, and r is the excited state lifetime, which was calculated from a second-order polynomial regression line of the dependence of F on temperature (15). C(r) is a molecular shape parameter having a precalibrated value for each value of r. From the determined r, T, and r values and the corresponding C(r) the microviscosity was derived. The temperature dependence of v was expressed as log v versus l/T. A linear regression analysis was made to determine the flow activation energy, AE, and to detect possible phase transitions (15, 16). All computations and plots were made with a Wang 2200-B system (Wang Laboratories, Tewksbury,
MA). Fluorescence Microscopy. A Zeiss Universal microscope was used for fluorescence microscopy. Fluorescence micrographs of DPH-labeled cells were taken with epi-illumination at 365 nm and emission above 410 nm. To overcome the rapid bleaching of DPH-labeled cells, micrographs were taken with a highly sensitive recording film (Kodak 2475), which was specially processed to increase its sensitivity to about 50
DIN.
RESULTS The Neuro-2A cells were chosen for this study because they can be easily synchronized without the use of metabolic or mitotic inhibitors. The mitotic shake-off method, under the employed
Proc. Natl. Acad. Sci. USA 74 (1977)
Cell Biology: de Laat et al.
4460 -
0
0.20
1.0
Table 1. Microviscosity of the membrane lipid layer of exponentially growing Neuro-2A cells in PBS, mean values ± SEM
a S
0.
Cell system
Microviscosity at 370, poise
On glass substratum After EDTA detachment After trypsin detachment
2.1 ± 0.1 2.9 ± 0.1 3.1 + 0.1
C0
*0c
xCu
0.15
E
U.
a)
cJC
a)
upC)cn
To avoid possible alterations in cell membrane by detach-
0
ment treatments, as with trypsin or EDTA, we have developed
01
0.10
0
10
20
30
40 50 Incubation time, min
60
FIG. 3. Fluorescence anisotropy and relative fluorescence intensity (F/Fmax) during the uptake of DPH by exponentially growing Neuro-2A cells.
conditions, resulted in a high degree of synchronization. The yield of mitotic cells per shake-off was 2-4% of the total cell number. Fig. 2 illustrates the degree of synchronization, as determined by [3H]thymidine incorporation and time-lapse cinematography. The mean duration of the cell cycle was 9.5 hr in the synchronized cultures, which was not significantly different from that observed in the exponentially growing cultures. The duration of the S phase was about 6 hr. while the GI and G2 phases lasted about 1.5 hr and 1 hr, respectively. Labeling of intact cells with fluorescent hydrocarbons like DPH or perylene may be only initially confined to the cell surface membrane. To check for any possible contribution of DPH incorporated into intracellular compartments, the fluorescence anisotropy and the total fluorescence intensity were determined simultaneously during the labeling period. Fig. 3 gives an example of such a measurement. No significant alterations in the r value were observed during the uptake of DPH, indicating that DPH is uniformly dissolved into one fluid compartment. Taking into account that the plasma membrane is the first compartment DPH molecules encounter during their uptake, the contribution of intracellular DPH was thus assumed to be negligible. Furthermore, in the fluorescence microscope, DPH-labeled neuroblastoma cells (Fig. 4) displayed an evenly glowing periphery with minor intracellular fluorescence, in contrast to reported data on 3T3 cells (17).
a simple method by which fluorescence polarization can be measured with the cells attached to a thin glass substratum (see Fig. 1). Table 1 shows the differences in microviscosity obtained for cells attached to substratum and for cells in suspension, after trypsin or EDTA treatment. Clearly, the latter treatments both lead to an overall increase in microviscosity, trypsin to a larger extent than EDTA. Four independent synchronization experiments were carried out in triplicate for each time. The determined microviscosity parameters of the synchronized cells are summarized in Figs. 5 and 6, and Table 2. As stated before, the measurements were made on cells attached to a glass substratum, with the exception of the first point (t = 0 hr) in Fig. 5. These mitotic cells were measured in suspension, immediately after shake-off, because the reattachment period lasted about 15 min. As shown, the microviscosity appears to be maximal during mitosis, and at the GI phase it decreases rapidly, reaching about a half-maximal value at the onset of DNA synthesis. The microviscosity remained constant at this low level during the S phase and increased again during the G2 phase. The determined X value always reached a lower maximum during the second mitosis as compared to the first (Fig. 5). This was presumably due to partial loss of synchrony with time (see Fig. 2), because the first cells started to divide again at 8 hr after the shake-off, but all cells were divided only after 11 hr. Consequently, at any time between 8 and 11 hr after shake-off only a fraction of the cells were in mitosis and the apparent microviscosity had an intermediate value between the maximum of mitotic cells and the minimum of S-phase cells. Plots of log I versus 1/T for cells in different phases of the cell cycle gave linear relations (see Fig. 6) with no signs of phase transitions. The flow activation enerS
M 3.5 -i
3.0
-
2.5
-
2.0
-
M
G6
G2
a),
.0
F
Igr
FIG. 4. Micrographs of Neuro-2A cells after 90-min labeling with DPH showing the label uniformly distributed over the cell surface with minor intracellular fluorescence; bars equal 10 tm. (A) Phase contrast micrograph; (B) fluorescence micrographs of the same cells. After 30-min labeling, when fluorescence measurements were made, the intracellular fluorescence was largely missing. However, the overall fluorescence was too weak to be photographed.
-~
"
1.5 -r 0
I-4
2
4
-~
6
-f
-
8
10
12
Hours after shake-off FIG. 5. Microviscosity (ij) at 370 as a function of time during the cell cycle of Neuro-2A cells. Mean + SEM is indicated. All measurements were made on cells in situ on a glass substratum, with the ex-
ception of the freshly selected mitotic cells (t measured in suspension.
=0
hr), which were
Cell Biology: de Laat et al.
Proc. Nati. Acad. Sci. USA 74 (1977)
4461
Table 2. Measured fluorescence anisotropy (r), and corresponding microviscosity (61), and flow activation energy (AE) of the membrane lipid layer at different stages of the cell cycle
10
Stage of cell cycle M Early G1 Late G1 S
5
gr
250
370 r 0.198 0.181 0.166 0.150
v7 3.5 2.9 2.2 1.9
r
0.225 0.214 0.192 0.178
40 v 4.7 4.2 3.2 2.7
r
i
0.269 0.262 0.241 0.232
8.7 8.4 6.3 5.2
AE 4.8 5.4 5.4 5.2
v is expressed in poise; AE is expressed in kcal/mol (1 cal = 4.184 J).
I 3.6 3.4 3.5 (1/T) X 10-3, K-l FIG. 6. Temperature dependence of microviscosity (ii) in Neuro-2A cells at different stages of the cell cycle, plotted on a logarithmic scale versus 1T. Mitosis (0), early G1 phase (0), late G1 phase (*), and S phase (*).
3.2
3.3
gies (AE) obtained from the slopes of these lines are given in Table 2.
DISCUSSION The microviscosity of the cell membrane lipid layer varies by a factor of two during the cell cycle of neuroblastoma cells. It is maximal in mitosis, minimal during the S phase, and shows a rapid decrease and increase in the Gi and G2 phase, respectively. This confirms earlier suggestions that the cell membrane of mitotic cells is more rigid than that of interphase cells (18, 19).
Previously, differences in microviscosity have been associated with growth characteristics. Resting lymphocytes were found to have a higher microviscosity than proliferating lymphoblasts (20) and lymphoma cells (12), while normal 3T3 cells showed a lower microviscosity than virus-transformed 3T3 cells (13). In view of the described cell cycle dependency of the microviscosity, these differences, as measured in nonsynchronized cell populations, could at least partially be a reflection of differences in the relative distribution of the cells through the cell cycle. It seems likely that the rate of lipid insertion into the cell surface is maximal during mitosis and the GI phase; for references see ref. 3. Accordingly, our results indicate a rapid alteration of the lipid composition of the cell membrane during the Gi phase (16). An excess insertion of lipids during this phase could also explain the described surface bleb formation (21) and decrease in the density of intramembranous particles in the GI phase (22). A great number of publications have led to the conclusion that surface antigens and (lectin-) receptor sites exhibit their lowest degree of expression or accessibility during the S phase; for references see ref 3. In view of the reported relationship between the microviscosity and the displacements of proteins perpendicular to the plane of the membrane (5), the possibility should be considered that the variation during the cell cycle of the expression of membrane receptor sites originates from alterations in the fluidity of the membrane lipid layer, as described here. Cyclic nucleotide metabolism (23) and ion metabolism (24) have been inferred to be involved in growth regulation. In both cases the activities of membrane-bound enzymes, i.e., adenylate
cyclase and (Na+, K+)-ATPase have been shown to be dependent on the membrane lipid composition and fluidity (7-9). Assuming that the observed correlation of the cell cycle with membrane microviscosity is a general phenomenon, it can be speculated that the membrane exerts its influence on the regulation of the cell cycle by modulating the activities of such. enzymes through changes in the lipid fluidity. We thank Mr. J. H. Beeker and Mr. P. Hogeweg for excellent technical support in constructing the fluorescence polarization apparatus; Mrs. Manuela M. Marques da Silva Pimenta Guarda, Miss Alie Feyen, and Mr. M. A. da Silva Guarda for their skillful technical assistance; and Mr. L. Boom for the photography. This work was supported by a travel grant (to S.W.deL.) of the Netherlands Organization for the Advancement of Pure Research.
1. Nicolson, G. L. (1976) Biochim. Biophys. Acta 457,57-108. 2. Nicolson, G. L. (1976) Biochim. Biophys. Acta 458, 1-72. 3. Bluemink, J. G. & de Laat, S. W. (1977) in Cell Surface Reviews, eds. Post, G. & Nicolson, G. L. (North-Holland Publishing Co., Amsterdam), Vol. 4, in press. 4. Edidin, M. (1974) Annu. Rev. Biophys. Bioeng. 3, 179-201. 5. Borochov, H. & Shinitzky, M. (1976) Proc. Nati. Acad. Sci. USA
73,4526-4530. 6. Schlessinger, J., Elson, E. L., Webb, W. W., Yahara, I., Rutishauser, U. & Edelman, G. M. (1977) Proc. Natl. Acad. Sci. USA 74, 1110-1114. 7. Kimelberg, H. K. & Papahadjopoulos, D. (1972) Biochim. Biophys. Acta 282, 277-292. 8. Kimelberg, H. K. (1975) Biochim. Biophys. Acta 413, 143156. 9. Orly, J. & Schramm, M. (1975) Proc. Natl. Acad. Sci. USA 72, 3433-3437. 10. Anderson, W. B., Russell, T. R., Carchman, R. A. & Pastan, I. (1973) Proc. Natl. Acad. Sci. USA 70,3802-3805. 11. Graham, F. M., Summer, M. C. B., Curtis, D. H. & Pasternak, C. A. (1973) Nature 246, 291-295. 12. Shinitzky, M. & Inbar, M. (1974) J. Mol. Biol. 85,603-615. 13. Fuchs, P., Parola, A., Robbins, P. W. & Blout, E. R. (1975) Proc. Natl. Acad. Sci. USA 72,3351-3354. 14. Klebe, R. J. & Ruddle, F. H. (1969) J. Cell Biol. 43, 69A. 15. Shinitzky, M. & Barenholz, Y. (1974) J. Biol. Chem. 249, 2652-2657. 16. Shinitzky, M. & Inbar, M. (1976) Biochim. Biophys. Acta 433, 133-147. 17. Berlin, R. D. & Fera, J. P. (1977) Proc. Natl. Acad. Sci. USA 74, 1072-1076. 18. Furcht, T. & Scott, R. E. (1974) Exp. Cell Res. 88, 311-318. 19. Garrido, J. (1975) Exp. Cell Res. 94, 159-175. 20. Inbar, M. & Shinitzky, M. (1975) Eur. J. Immunol. 5, 166170. 21. Porter, K., Prescott, D. & Frey, J. (1973) J. Cell Biol. 57, 815836. 22. Scott, R. E., Carter, R. L. & Kidwell, W. R. (1971) Nature New Biol. 233, 219-220. 23. Burger, M. M., Bombik, B. M., Breckenridge, B. M. L. & Sheppard, J. R. (1972) Nature New Biol. 239, 161-163. 24. Cone, C. D., Jr. (1974) Ann. N.Y. Acad. Sci. 238,420-435.
