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Membrane dynamic alterations associated with viral transformation and reversion
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Experimental
MEMBRANE
DYNAMIC
VIRAL
Cell Research 118 (1979) 205-214
ALTERATIONS
TRANSFORMATION
ASSOCIATED
WITH
AND REVERSION
Decay of Fluorescence Emission and Anisotropy Studies of 3T3 Cells ABRAHAM
H. PAROLA,‘*
PHILLIPS W. ROBBINS* and ELKAN
R. BLOUT3
‘Departmentof Chemistry, Ben Gurion University of the Negev, Beer Sheva, Israel, Tenterfor Cancer Research, Massachusetts Institute of Technology, Cambridge, MA 02139, and 3Department of Biological Chemistry, Harvard Medical School, Boston, MA 02115, USA
SUMMARY Fluorescence lifetimes, anisotropies and rotational correlation time values of 1,6-diphenyl-1,3,5hexatriene (DPH) in membranes of normal, transformed, and revertant 3T3 cells were determined by nanosecond (nsec), photon counting spectrofluorimetry. No change in lifetime values with transformation or reversion is observed. Fluorescence anisotropy decay curves show at least two components; an initial relatively fast decay and a non-zero “plateau” level component. The observed changes in the average anisotropy values, which qualitatively follow steady-state fluorescence polarization values, is due primarily to changes in the non-zero “plateau” level component. The anisotropy decay curves suggest that the rotational motion of the probe is restricted to a limited angular range. The present results are compared with model membrane systems.
The result if malignant transformation of cells is an altered “social behavior” [14] most dramatically expressed by the absence or modification of contact inhibition of cell movement [5, 61 and density-dependent regulation of growth [7]. It is believed that changes in cell surface membranes are a major determinant in the transformation processes [g-l 11. The correlation between transformation and enhanced lectin-receptor mobility stresses the importance of understanding the dynamics of membrane proteins and the relationship of protein dynamics to membrane lipid (bilayer) fluidity [l, 3, 4, 12-141. The “fluidity” of membranes of transformed cells has been measured by a variety of physical methods, following either rotational or lateral motions, with both modes reflecting changes in membrane rigidity. Although Gaffney [15] found no sig-
nificant difference in membrane fluidity from spin-label electron spin resonance (ESR) studies of normal and Simian Virus 40 (SV40)-transformed 3T3 cells, we (using 1,6-diphenyl-1,3,5-hexatriene (DPH) as a probe) reported [ 131 significantly lower steady-state fluorescence polarization values (reflecting higher rotational relaxation time values) and lower calculated “microviscosities” for normal cells as compared with cells transformed either by polyoma virus or SV40. Lateral diffusion studies with pyrene also showed increased membrane lipid rigidity in Rous sarcoma virustransformed BKH-T-1 and BKH-T-2 cells as compared with their normal counterparts [ 161.A small decrease in “fluidity” was also observed with Rous sarcoma virus-trans* Address: Department of Chemistry, Ben Gurion University of the Negev, P.O. Box 653, Beer-Sheva 84120, Israel. Exp
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Parola, Robbins and Biotrt
formed chick embryo fibroblasts, which displayed a decreased unsaturation of membrane lipid acyl groups [17]. In contrast, fluorescence polarization studies with normal lymphocytes and lymphoma cells showed distinctly lower “microviscosity” in lipid membranes of the latter [ 181. In addition, opposing trends in membrane fluidity associated with cell transformations by 13Cnuclear magnetic resonance (NMR) [ 191 and our fluorescence measurements were reported [ 131.Moreover, no correlation between rotational mobility of Concanavalin A receptors [20] and lipid bilayer “fluidity” with transformation of either lymphocytes or 3T3 mouse cells were noted [13]. Heterogeneity in membrane lipids, as well as in the locus of the fluorescent probes could explain the differing results. In many NMR and ESR measurements directed toward elucidation of lateral diffusion constants, one follows the spectral line width rather than line shape analysis [21-241. Since line-width analysis is an averaging method, it is limited in its ability to detect heterogeneity in the system under investigation. Similarly, in steady-state fluorescence polarization studies, calculated “microviscosities” may not represent the actual microviscosity but a value intermediate among the various viscosities of the media present in the system, weighted according to the distribution of the fluorescent probe. We now report the application of timeresolved fluorescence anisotropy (and lifetime) measurements of intact normal and virus-transformed 3T3 cells. The kinetics of fluorescence anisotropy gives directly the functional form of the all-important autocorrelation function [25, 261. This function provides specific information about the heterogeneity and rotational modes of the probe in the anisotropic membrane interior.
Our results with the lipophilic fluorescent probe, DPH, are consistent with the hypothesis that there is a nearly homogeneous (95 %) distribution of the probe in the membrane and confirm and further extend our previous interpretation of the steady-state fluorescence polarization experiments [27]. When the results reported here are compared with studies of fluorescence anisotropy of DPH in model lipid bilayer systems at varying lipid composition [28], a better understanding of the anisotropic character of the probe’s rotation dynamics is obtained. Our results show an increase in apparent “viscosity” with cell transformation, and a decrease in viscosity associated with reversion of cells to the untransformed state. Yet, more significant is the change in structure in the lipid interior of the transformed cell membrane, expressed in the increased anisotropy values to which the decay curves of anisotropy with time level off. Increased lectin-receptor mobility and agglutinability with cell transformation must thus be regulated by another membrane associated system, e.g. microtubules and/or microfilaments [ 1, 113.
MATERIALS
AND METHODS
Cells The isolation, growth conditions, and characteristics of the cell lines involved in this study have been described oreviouslv in our steadv-state fluorescence polarizailon study [ 131. Mouse 3i’3d, 3T3Py6, Py6R, 3T3 BALB A-31, and SVT2 cells were obtained from Dr Thomas Benjamin, Department of Pathology, Harvard Medical School. 3T3Py6 [29] are polyoma virustransformed 3T3d cells [30]. Py6R1 [3l] is a revertant line isolated from 3T3Py6 by the fluorodeoxyuridine procedure [3 I]. SVT2 cells are SV40-transformed 3T3 A-3 1 cells.
DPH labeling procedure Exponentially growing cells harvested under mild identical conditions. were labeled with 2x 10m6M DPH dispersed in PBS according to our previously-described procedure [13]. Unlabeled control cells were
Membrane treated with a solution of tetrahydrofuran in PBS (1 : 1000) under the same conditions.
Decay offluorescence emission and anisotropy measurements The decay of both fluorescence emission and fluorescence anisotropy as an explicit function of time in the nanosecond (nsec) range [32-341 may be measured by the single photon counting technique [35]. This method allows, in principle, the determination of hydrodynamic parameters from a single accumulation at a given temperature. The sample is excited with very short pulses of polarized light. The intensity of the emission polarized parallel, 111(r),and perpendicular I,(r), to the polarized excitation light are then measured as a function of time. Fluorescence decay measurements were performed at 37-38°C and carried out on equipment kindly made available by Dr Lubert Stryer, following the general experimental and deconvolution procedure described by Yguerabide [36] and Yguerabide et al. [37]. Two major conditions are essential for successful measurements of time-resolved fluorescence on intact, turbid cell suspensions; viz. (1) proper correction for scatter and intrinsic fluorescence of non-DPH labeled cells and cell clumps; and (2) short duration for accumulation of reproducible data. The following modifications of the Yguerabide experimental procedures were found necessary: (a) automatically alternating accumulation of parallel and perpendicular components of the fluorescence emission for short intervals (10 set) was necessary to eliminate errors due to fluctuating lamp intensity and fluctuating numbers of cells in the light beam. The sectors of the multichannel analyzer addressed by the output of the time-to-amplitude converter were synchronically switched. (b) The time dependence of fluorescence emission could be calculated from the total fluorescence emission, s(r), defined by eq. (I) [36,37]:
dynamics and viral transformation
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measurement and at 5 min intervals. Accumulation times for labeled samples were 2&30 min (alternating parallel and perpendicular modes), resulting in 104-IO5 accumulated counts in the peak channel. The excitation light was filtered by a Coming 7-60 filter, and the emitted fluorescence was detected through a Coming 3-73 filter. Prior to performing the nsec photon counting measurements, DPH-labeled cells were examined by steady-state fluorescence polarization at 37°C.
RESULTS Total fluorescence emission as a function of time, S(t) (eq. l), depends on the rates of transitions which depopulate the lowest excited singlet state. For a homogeneous system it can be described by first-order kinet its (eq. 3): (3)
where S, is the initial emission intensity following a very short pulse of polarized light, t is the time in nsec, and r is the excited state lifetime; i.e., the inverse of the sum of the rates of all processes that depopulate the lowest excited singlet level of the chromophore. For more complex and heterogeneous systems, S(t) can be expressed as a sum of exponential terms [37]. S(r)=Z,1(r)+2z,(r) (1) Fig. 1 shows a representative curve of the while that of fluorescence anisotropy, A(t), from eq. nearly single exponential fluorescence de(2): cay of DPH embedded in 3T3d cells. Atqc-l,(r) A(t)= (2) tempts to improve the fit between the exSW perimental data and the theoretically comAn automated procedure [28] permitted the calculation puted double exponential decay curve indiof excited state lifetime (7) (calcu!ated by a least cated that as much as 95% of the intensity squares analysis), mean anisotropy (A), and a series of calculated convoluted anisotropy curves correspondis due to a single exponential decay mode ing to single rotational correlation times 7, were and only 5 % to a second mode. Moreover, plotted for comparison with the observed curves. The calculated curve with the same initial slope as the when compared to the decay curve of 8observed curve led to the here termed apparent rotaaniline-1-naphthalene sulfonic acid (ANS) tional correlation time, rpapD.The scattering background and intrinsic fluorescence of unlabeled cells, in ethanol (a homogeneous medium), DPH which was not larger than 5% of the fluorescence fluorescence decay curves of all cell lines emission of labeled cells, was subtracted. These procedures reduced the time required for data acquisition investigated showed similar or small deviaand increased the reliability of the resulting anisotropy tions from a calculated single exponential values. Cell suspensions were gently mixed 1 min before decay curve. Table 1 shows fluorescence Exp
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Pat-oh, Robbins und Blorrt
Table 1. Nunosecond cells ut 37°C
fluorescence
studies oj’ normul,
transformed
and re\lertunt
Cell line
7 (nsec)a
‘j
AZ6
pallp (nsec)Q,c
ht, (nsec)”
q (poise)+
3T3 A-3 I SVT2 3T3d 3T3Py6 PY6Rr
7.7to. I 7.8tO. I 8.0&O. 1 S.lrtO..l E.l+O.l
0. I I +0.01 0.14t0.01 0.12+0.01 0.14+0.01 0.10+0.01
0.07 0.12 0.10 0.12 0.08
6.0t 1.0 9.0f 1.5 5.8?0.3 9.3kO.3 4.8kO.8
2.5 3.1 2.8 3.7 2.7
1.0 I.6 1.0 1.5 0.8
3T3
0 Average values of three independent experiments. * Values corresponding to fig. 2. c The S.D.s are calculated on the basis of three independent experiments; for method of calculation see text and captions to the figures. d Rotational correlation time estimated by least-square fitting a line to the first 5 nsec of data expressed as log[A(r)-A,]. Since the psub values are not corrected for the finite width of the light pulse, they provide an estimate of relative correlation times (see text). p Taken from Fuchs et al. [l3].
lifetime values, 7, of DPH embedded in the membranes of normal A-31, SVT2-transformed, normal 3T3d, 3T3Py6 transformed and Py6R, revertant cells. Essentially no variation in DPH lifetime upon transformation (i.e. SVT2 vs A-3 1 and 3T3Py6 vs 3T3d
cells) or reversion (i.e. Py6R, vs 3T3d cells) is indicated. The decay of fluorescence anisotropy with time, A(t), expresses the change in orientation of the transition moment direction during the delay in emission from the first excited singlet state. For a very short light pulse, A(t) is at most slightly dependent on the excited state lifetime, since, by its definition, eq. (2), the division by S(t) eliminated the lifetime term. For the simplest case of a rigid sphere in an isotropic medium, A(t) decays exponentially (eq. 4): A(t)=A,,e-‘1”
time (nsec); ordinate: fluorescence intensity (counts). Total fluorescence emission decay for DPH in 3T3d cells at 37°C. The logarithm of fluorescence is plotted as a function of time. Solid line. -, Experimental data; ___, theoretical convoluted single exponential decay calculated for 7=8.0 nsec. The lamp profile is also shown. Data analysis is performed by the method of least squares.
Fig. 1. Abscissa:
E.rp Cdl
Rr,
118 fIY7Y)
(4)
where A0 is the initial value of the emission anisotropy [37] and cpis the rotational correlation time. The fluorescence anisotropy of the rod-shaped diphenylhexatriene molecule in an isotropic solvent should decay exponentially even though the molecule is not spherical because its longest wavelength transition moment is parallel to the long axis of the molecule. Kinetic determination of fluorescence depolarization would thus yield linear plots of In A vs time, the slope of which affords a value for cp directly; the slope is l/p and the intercept
Membrane
dynamics and viral transformation
20
209
so
Fig. 2. Abscissa:
time (nsec); ordinate: fluorescence emission anisotropy. Time dependence of the emission anisotropy of DPH, eq. (2), in membranes of 3T3 A-31 (...) and SVT2 (X x X) at 37°C (a) and in membranes of 3T3d (. . .) 3T3Py6 (X x X) and Py6Rr (000) at 37°C (b). The logarithm of the anisotropy is plotted as a function of time. The initial fast decay data points, coinciding
with the dashed line for 3T3 A-31 and with the solid line for SVT2 cell lines in (a) were visually compared with theoretically calculated convoluted plots of single exponential decay of anisotropy (detailed in [43]). Such calculated decay plots are included in (b), dashed lines. The scatter in points, particularly at the plateau region indicate the statistical counting errors where only a small number of photons are accumulated.
at time zero, is fnAO. Thus, all necessary information can be obtained from the same sample at physiological temperatures. Deviation from single exponential decay, eq. (4), may arise in various situations, e.g. restriction of the angular range of motion, or heterogeneity of the probe’s environment. In such cases A(t) will be a sum of exponentials,
two components; initially, a relatively fast one, within the first 12 (fig. 2a) and 10 (fig. 2b) nsec which then decays to a non-zero “plateau” value. The deviation of the anisotropy decay curves from those expected for a single component occurs between 12 and 20 nsec (fig. 2a) and between 10-15 nsec (fig. 2b) which corresponds to time intervals at which no deviation of total fluorescence emission (fig. 1) from a single exponential decay is observed; the deviation there (fig. 1) begins only after -25-30 nsec. The increased scatter in fluorescence emission anisotropy data points at longer time intervals, i.e. at the “plateau” region, arises from statistical variation in photon counting at a region where fluorescence emission is small. This scatter of the data points is due to the exponential decay of total fluorescence emission intensity (fig. l), which results in more than an order of magnitude
i.e. A(t)=&
Aie-“qf. i=l
Fig. 2 shows the typical time-dependent curves of the emission anisotropy of DPHlabeled A-3 1 and SVT2 cells (fig. 2a) and of DPH-labeled 3T3d, 3T3 Py6, and Py6R, cells (fig. 2 b). Characteristic of all cell lines studied here are the following features which are different from theoretically calculated single exponential decay of anisotropy curves: The decay curves exhibit at least
Exp Cell Res 118 (1979)
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Parola, Robbins and Blout
decrease in fluorescence intensity 20 nsec after the peak. Since accumuiation periods are limited by the vitality of the biological system, only 104-IO5counts were accumulated at the peak channel, resulting in low count number in the “plateau” region. Thus, obviously beyond 30 nsec, the scatter in data points is quite large and should be disregarded. Summarized in table 1 are values calculated for mean anisotropy, A, and the plateau value, AZ, corresponding to the slow motion (over 100nsec). Apparent rotational correlation times, &,,,, obtained by a comparison of the initial slope of the experimental data with the single exponential decay curves, are also included in table 1. These hpp values are used strictly for qualitative comparative purposes, and one should take into account the long-time limiting A2 anisotropy values. Thus, the nonzero values of A2 will cause the apparent initial slope of the anisotropy plot to be less than that for the fast component alone. Instead of fitting the data to a single exponential decay, eq. (4), it is helpful to fit the data to eq. (5), A(t)=A1e-“a+A2
(5)
The easiest way to estimate p is to fit the slope of the line to a function log (A(t)-A,). The (Sub values (subtracted rotational correlation time) derived from a weighted linear-least-squares analysis of the first 5 nsec of data (-20 data points) after subtraction of A2 are included in table 1. Consequently, rotational correlation times corrected for the difference in A2 values are shorter and range between 2.5-3.7 nsec. For both 3T3 A-31 and 3T3d cells, transformation with SV40 and polyoma viruses, respectively, resulted in an increase in A, AZ, cp,,,, and (Sub; the same trend was also reported in our steady-state fluorescence polarization Exp Cdl Res II8 (19791
study [ 131.While changes in (F~,,,,are similar in magnitude to changes in calculated “microviscosities” (50-60s increase with transformation and about 20% decrease with reversion), a smaller change (yet in the same direction) is indicated for (Pan,)values which are corrected for the differences in A 2 values. Yet, the good correlation between steady state fluorescence polarization results with papp values might be merely fortuitous. Moreover, even the qualitative correlation with ‘psu,,values should be cautiously regarded since the small differences in (P& VdUeS may not be that significant due to increased errors rising from convolution uncertainties. These observations suggest that the observed changes in A values associated with virus transformation are due not only to changes in membrane fluidity (which may be expressed in (Sub) but primarily to changes in the anisotropic nature of the lipid bilayer, which is expressed in AZ. Associated with transformation is loss of some degrees of freedom of rotation of the probe, which is essentially inhibited from completely depolarizing the fluorescence emission. Interestingly Py6R, revertant cells exhibit a small decrease in A, AZ, and papp; (P&, is not significantly smaller than that observed with 3T3d normal cells. DISCUSSION Steady-state fluorescence polarization values, P, depend on both the excited state lifetime, 7, and the rotational relaxation time, cp.Thus, DPH lifetime measurements are essential in order to establish whether differences in P-values observed between normal, virus-transformed, and revertant cells [ 131do actually reflect changes in rotational dynamics which may reflect differences in membrane fluidity. This will be the case if we assume no change in the limit-
Membrane dynamics and viral transformation
ing fluorescence polarization value, PO, to be associated with cell transformation. The similarity in lifetime values observed for 3T3 A-31 and SVT2 and among 3T3d, 3T3Py6, and Py6R, cells (table 1) indicate that changes in membrane “fluidity” and/ or anisotropy (rather than DPH lifetime) lead to the observed changes in P-values. These results further indicate that, upon transformation by either SV40 or polyoma virus, DPH molecules are exposed to similar dipolar interactions, suggesting no gross alteration in their immediate solvation shell. Any attempt to study lipid bilayer fluidity of plasma membranes in intact cells using lipophilic probes is complicated by the uncertainty in the location of the probe. A lipophilic probe, by its nature, cannot be anchored to the membrane-water interface, since if an ionic site is necessary for anchorage such an ionic moiety would prevent the probe’s penetration to the core of the lipid bilayer. Thus, if one is forced to use a lipophilic probe, its diffusion through the plasma membrane into intracellular membranes cannot be excluded a priori. While DPH penetration through the aqueous interior of the cells appears to be unlikely, due to its low solubility in water, one cannot exclude its partitioning into the endoplasmic reticulum linked to the plasma membrane, rather than confining itself only to the latter. Attempts were made to ascertain the cellular location of DPH through fluorescence microscopy. No obvious concentration of the dye in lipid droplets was observed. The experiments, however, were limited by rapid bleaching of the dye [38] under the conditions employed [49]. The fluorescence decay curves of DPH in each of the cell lines studied approximates a single exponential character (95%) (e.g. fig. 1). This may indicate a similar, relatively homogeneous distribution of DPH
211
molecules in the membranes of each of these cell lines. These observations are particularly striking since the non-complete single exponential decays may be due to some reversible bleaching [ 13, 181.This excited state isomerization may not be totally eliminated even in the photon counting mode, where short and weak illumination pulses are used. Deviations from monoexponential decay, ascribed to effects of solvent viscosity on cis-trans isomerization have been noted previously [39-41]. From a spectroscopic viewpoint, this is perhaps the single disadvantage in the use of DPH as a fluorescent probe, which otherwise is ideal for determination of fluorescence polarization of bilayer lipid membranes [4245]. Distinct double exponential fluorescence emission decay profiles were recently reported for DPH in dimytistoyl lecithine (DML) vesicles [42], yet single exponential decays were observed for DPH in multilamelar DML vesicles [45], in di(dehydrosterculayl)-phosphatidyl choline (PC) liposomes [28] and in DL-cr-dipalmitoylphosphatidylcholine (DPPC) liposomes [46]. In a different study, the fluorescence of 12(Panthroyl) stearic acid incorporated into red blood cell membranes also decays as single exponential [47]. The differences among the various systems, ranging from 87 to 95 and 100% “single” exponential decays, may be due to varying microheterogeneity of sites. This is more likely with small bilayer vesicles (such as those used in the Brand’s study [42] than with large multilamellar liposomes [45]. Thus, our cell system exhibits an intermediate situation, resembling more the multilamellar DML vesicles in terms of that microheterogeneity. However, again this may be due to cis-trans isomerization of the DPH, which may not be as clearly detectable in the present study where accumulation of data is restricted by
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the vitality of the biological system. It should be pointed out, however, that the lifetime of DPH in our study is similar to that reported in DML vesicles, i.e., S.O?O.l nsec for the 3T3d system and 7.7&O. 1 nsec for the 3T3 A-31 system vs 7.7 for single and 8.0 for double decay analysis in the artificial membranes. The fluorescence anisotropy decay profiles shown in fig. 2 are different from such plots for DPH in a homogeneous viscous medium, glycerol (A. Parola, unpublished results) or paraffm oil [39]. In the latter two systems, DPH decay of anisotropy exhibits a continuous decay with A2 approaching zero values. Yet similar characteristics to the present decay study were recently observed in three independent studies of artificial membranes: in PC liposomes with varying amounts of cholesterol particularly with higher cholesterol content [28], in DML vesicles, particularly at the temperature below the midpoint of the gel-liquid crystalline transition (at 37.2”C A2 did approach zero) [42], and in DPPC liposomes
[461. Heterogeneity in DPH molecular population, with a fraction of it highly immobilized resulting in the “plateau” region at long times, and the rest free to rotate leading to the initial fast decay region (fig. 2) [45] may adequately describe the situation in small, single-walled vesicles [42], particularly since a double exponential decay of total fluorescence, characteristic of two different lifetimes, is observed in addition to the nonsingle exponential decay of A(t) vs time plots. However, due to the close to single exponential character of the fluorescence emission decay (fig. 1) and particularly since the anisotropy decay curves reach their plateau value about 10 nsec before the deviation from single exponential decay of total fluorescence emission is noted (though Exp Cd Rr., 118 (1979)
we cannot totally exclude such heterogeneity) an alternative model may be more plausible. It is based on the anisotropic character of the rotational motion of each DPH molecule, free to reorient only over a limited angular range. Such a model is consistent with the interpretation of the data for DPH observed with artificial membranes [28] and spin resonance experiments done with spin labels in lipid bilayers [48]. Such motion may be expected of DPH molecules because of their elongated shape and the highly anisotropic environment inside the membrane. Rotation of the chromophore about the long axis of the molecule cannot result in any depolarization because it does not lead to reorientation of the transition moment. However, a very rapid motion over a limited angular range, corresponding to a restricted “wobble” about the long axis of the molecule, could contribute to the initial decrease from A0 (0.33 for DPH in glycerol at O’C) to the initially observed anisotropy values (see fig. 2). Such a “wobble” motion seems to occur in liposomes, too, [28, 461. Veatch & Stryer have carried out time-resolved fluorescence studies of DPH in liposomes. They observed fluorescence anisotropy decay curves in this model system very similar to those shown in fig. 2, with the magnitude of A2 increasing with increasing cholesterol mole fraction. They concluded that the addition of cholesterol to phosphatidylcholine liposomes renders the environment highly non-isotropic. In steady-state fluorescence polarization experiment of complex membrane systems, the observed polarization is an average value. Therefore, the “microviscosity” values calculated from Perrin’s equation (invoking a number of gross assumptions [13]) may not represent the actual “microviscosity” but a value intermediate between the various viscosities in the media. Moreover, the
Membrane
interpretation of steady state data in terms of “microviscosity” assumes that equivalent A(t) curves would be observed for equivalent viscosities of reference (oil solution) and membrane systems and thereby implicitly that the same viscosity concept applies in both media. Yet comparison with time-resolved fluorescence anisotropy me urements, which is not an averaging / method, reveals qualitatively similar trends in the results: i.e. A, AZ, and (Sub increase with transformation and decrease with reversion, as “microviscosity” values do too [13]. Thus, despite the fact that estimated values of viscosity in a cone were found to be an order of magnitude smaller than the value of “microviscosity” estimated from steady state emission anisotropies [46], our observations qualitatively do confirm our previous steady-state measurements and are consistent with a relatively homogeneous distribution of the probe. Still, the term “microviscosities” (obtained from steadystate studies) should be used very cautiously; the quotation marks do mean to indicate the over-simplification in this expression, since they represent both fluidity and major structural effects. A comparison with model membranes further clarifies these combined fluidity and structural factors. Thus, while similar, non-single exponential decay curves could be obtained with PC vesicles containing varying amounts of cholesterol, these curves seem to differ from our cell systems in that the changes in A2 were not associated with changes in &b [28]. Yet in the cell lines studied here both A2 and &b vary with transformation. Thus, while changes in cholesterol levels in the different cell lines could presumably be the cause for the major difference observed inA, (rendering the membrane more anisotropic with higher cholesterol levels), other factors (lipid and protein composition) could lead to
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the relatively small differences observed in $4ub values, which may represent actual relative fluidity differences. However, even in our system, the observed small differences in $&ubmay be within experimental uncertainties due to convolution problems. A question often raised is whether the new properties detected in virus-transformed cells are due to the presence of the virus in the cells. We found similar trends in membrane fluidity and anisotropy with a BUdR dependent cell line derived from a malignant Syrian hamster melanoma line [49]. With this system, “transformation” can be reversibly controlled by the presence or absence of the small BUdR moiety. Fluorescence anisotropy decay curves for BUdR “transformed” cells showed similar profiles to those observed for SV40 and polyoma “transformed’ characteristics independent of the actual cause of “transformation”. In conclusion, this study shows that the careful design of experiments using two techniques-steady-state and nsec spectrofluorimetry-allows more confidence in the interpretation of the dynamic behavior of biological membranes. Experiments aimed at detecting phase transitions and fusion activation energies from plots of In r) vs l/T [13] may be most adequately performed with steady-state spectrofluorimetry. On the other hand, fluorescence lifetime information, rotational correlation times, and information about membrane anisotropy can be obtained only from the time-resolved spectrofluorimetry. For the cell system studied here, the present work indicates that increased fluorescence polarization values observed with viral transformation are primarily due to anisotropy changes in the membrane lipid bilayer and thus may indicate only a relatively small change in its microviscosity. Exp Cell Res 118 (1979)
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We thank Dr Thomas Benjamin of the Department of Pathology at Harvard Medical School for providing us with the cell lines. The technical assistance of MS Lin Chen is gratefully acknowledged. We are very much indebted to Professor Lubert Stryer, Standford University, for the use of his photon counting equipment. We thank Professor William Veatch and Professor Samuel Latt, Harvard Medical School, for helpful discussions. This work was supported, in part, by NIH Grants AM07300 and 5-Rpl-CA14143-12 and the NC1 Grants CA14142 and CA14051. A. H. P. was a recipient of an NIH Postdoctoral Fellowship. A preliminary report on this work was presented at the Meeting of the Israel Biochemical Society, April I, 1976.Beersheva, Israel.
REFERENCES I. Nicolson, G L, Biochim biophys acta 458 (1976) I. 2. - - Ibid 457 (1976) 57. 3. Singer, S J & Nicolson, G L, Science 175 (1972) 720. 4. Rapin, A M C & Burger, M M, Adv cancer res 20 (1974) I. 5. Abercrombie, M, Cells and tissues in culture (ed E N Willmer) vol. 1, p. 177. Academic Press, New York (1%5). Harris, A, Cell communication (ed R P Cox) p. 147. John Wiley & Sons, New York (1974). Halley, R W, Nature 258 (1975) 487. Oserof, A R, Robbins, P W & Burger, M M, Ann rev biochem 42 (1973) 647. Robbins, J C & Nicolson, G L, Cancel-: a comprehensive treatise (ed F F Becker) vol. 14, p. 3. Plenum Press, New York (1975). 10. Hynes, R 0, Biochim biophys acta 458 (1976) 73. II. Edelman, G M, Science 192(1976) 218. 12. Lis, H & Sharon, N, Ann rev biochem 43 (1973) 541. 13. Fuchs, P. Parola, A. Robbins, P W & Blout, E R, Proc natl acad sci US 72 (1975) 3351. 14. Edidin, M, Ann rev biophys bioeng 3 (1974) 179. 15, Gaffney, B J, Proc natl acad sci US 72 (1975) 664. 16. Edwards, H E, Thomas, J K, Buleson, G E & Kupla, C F, Biochim biophys acta 448 (1976) 45I. 17. Yau, T M, Buckman, T, Hale, A & Weber, M J, Biochemistry 15 (1976) 3212. 18. Shinitzky, M & Inbar, M, J mol biol85 (1974) 603. 19. Nicolau, U, Dietrich, W, Steiner, M, Steiner, S & Melnick, J L, Biochim biophys acta 382(1975) 31I. 20. Inbar, M, Shinitzky. M & Sachs. L. J mol biol 81 (1973) 245. 21 Davis, D G, Biochem biophys res commun 49 (1972) 1492. 33 McFarland, B G, Chem phys lipids 8 (1972) 303. I_.
23. Lee. A G, Birdsall. N J M & Metcalf. J c‘. Biochemistry 12 (1973) 1650. 24. Komberg, R & McConnell. H M, Proc natl acad sci US 68 (1971) 2564. 25. Jones, R E, Ph.D. thesis. Stanford University (1970). 26. b’dllach, D, J them phys 47 (1967) 5258. 27. Parola, A, Robbins, P W & Blout, E R. Israel j med sci 12 (1976) 1362. 28. Veatch, W & Stryer, L, J mol biol 117(1977) 1109. 29. Benjamin, T L & Berger, M M, Proc natl acad sci US 67 (1970) 929. 30. Todaro, G J & Green, H, J cell biol 17(1963) 299. 31. Benjamin, T L & Goldman, E, Cold Spring Harbor symp quant biol 39 (1974) 41. 32. Jablonski, A, 2 Naturforsch 16a (196I) I. 33. Fayet, M & Wahl, P, Biochim biophys acta 181 (1969) 373. 34. Wahl, Ph, Biochemical fluorescence concepts (ed R F Chen & H Edelhoch) vol. I ~ p. I. Marcel Dekker Inc., New York (1975). 35. Lami, H, Pfeffer, G & Laustriat. G. J phys 27 (1966) 398. 36. Yguerabide, J, Methods in enzymology (ed C H W Hirs & S N Timasheff) vol. 26, p. 498, part C. Academic Press, New York, London (1972). 37. Yguerabide, J, Epstein, H F & Stryer, L, J mol biol 51 (1970) 573. 38. Berlin, R D & Fera. J P. Proc natl acad sci US 74 (1977) 1972. 39. Cehelnik, E D, Cundall, R B, Lockwood, .l R & Palmer, T F, J them sot Faraday trans 2.70 (1974) 244. 40. Cehelnik, E D, Cundall, R B, Lockwood, .I R & Palmer, T F, J phys them 70 (1975) 1369. 41. Birks, J B, Chem phys lett 38 (1976) 437. 42. Chen, L A, Dale, R E, Roth, S & Brand, L, J biol them 252 (1977) 2163. 43. Andrich, M P & Vanderkooi. J M, Biochemistry I5 (1976) 1257. 44. Stubbs, G W, Litman, B J & Barenholz, Y, Biochemistry 15 (1976) 2766. 45. Lentz. B R. Barenholz. Y & Thomnson. T E. Biochemistry 15 (1976) 4521. 46. Kawato. S. Kinosita. K. Jr & Ikerami, A. Biochemistry 16 (1977) 2319. 47. Vanderkooi, 3, Fischkoff, S, Chance, B &Cooper. R A, Biochemistry 13 (1974) 1589. 48. Israelachvili, J, Sjiisten, J, Erickson, L E G, EhrStrom, M, Gdslund, A & Ehrenberg, A, Biochim biophys acta 382 (1975) 125. 49. Rosenthal, S L, Parola, A H, Blout, E R & Davidson, R L, Exp cell res 112(1978) 419. Received April 14, 1978 Revised version received July 24, 1978 Accepted August 3, 1978
Experimental
MEMBRANE
DYNAMIC
VIRAL
Cell Research 118 (1979) 205-214
ALTERATIONS
TRANSFORMATION
ASSOCIATED
WITH
AND REVERSION
Decay of Fluorescence Emission and Anisotropy Studies of 3T3 Cells ABRAHAM
H. PAROLA,‘*
PHILLIPS W. ROBBINS* and ELKAN
R. BLOUT3
‘Departmentof Chemistry, Ben Gurion University of the Negev, Beer Sheva, Israel, Tenterfor Cancer Research, Massachusetts Institute of Technology, Cambridge, MA 02139, and 3Department of Biological Chemistry, Harvard Medical School, Boston, MA 02115, USA
SUMMARY Fluorescence lifetimes, anisotropies and rotational correlation time values of 1,6-diphenyl-1,3,5hexatriene (DPH) in membranes of normal, transformed, and revertant 3T3 cells were determined by nanosecond (nsec), photon counting spectrofluorimetry. No change in lifetime values with transformation or reversion is observed. Fluorescence anisotropy decay curves show at least two components; an initial relatively fast decay and a non-zero “plateau” level component. The observed changes in the average anisotropy values, which qualitatively follow steady-state fluorescence polarization values, is due primarily to changes in the non-zero “plateau” level component. The anisotropy decay curves suggest that the rotational motion of the probe is restricted to a limited angular range. The present results are compared with model membrane systems.
The result if malignant transformation of cells is an altered “social behavior” [14] most dramatically expressed by the absence or modification of contact inhibition of cell movement [5, 61 and density-dependent regulation of growth [7]. It is believed that changes in cell surface membranes are a major determinant in the transformation processes [g-l 11. The correlation between transformation and enhanced lectin-receptor mobility stresses the importance of understanding the dynamics of membrane proteins and the relationship of protein dynamics to membrane lipid (bilayer) fluidity [l, 3, 4, 12-141. The “fluidity” of membranes of transformed cells has been measured by a variety of physical methods, following either rotational or lateral motions, with both modes reflecting changes in membrane rigidity. Although Gaffney [15] found no sig-
nificant difference in membrane fluidity from spin-label electron spin resonance (ESR) studies of normal and Simian Virus 40 (SV40)-transformed 3T3 cells, we (using 1,6-diphenyl-1,3,5-hexatriene (DPH) as a probe) reported [ 131 significantly lower steady-state fluorescence polarization values (reflecting higher rotational relaxation time values) and lower calculated “microviscosities” for normal cells as compared with cells transformed either by polyoma virus or SV40. Lateral diffusion studies with pyrene also showed increased membrane lipid rigidity in Rous sarcoma virustransformed BKH-T-1 and BKH-T-2 cells as compared with their normal counterparts [ 161.A small decrease in “fluidity” was also observed with Rous sarcoma virus-trans* Address: Department of Chemistry, Ben Gurion University of the Negev, P.O. Box 653, Beer-Sheva 84120, Israel. Exp
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Parola, Robbins and Biotrt
formed chick embryo fibroblasts, which displayed a decreased unsaturation of membrane lipid acyl groups [17]. In contrast, fluorescence polarization studies with normal lymphocytes and lymphoma cells showed distinctly lower “microviscosity” in lipid membranes of the latter [ 181. In addition, opposing trends in membrane fluidity associated with cell transformations by 13Cnuclear magnetic resonance (NMR) [ 191 and our fluorescence measurements were reported [ 131.Moreover, no correlation between rotational mobility of Concanavalin A receptors [20] and lipid bilayer “fluidity” with transformation of either lymphocytes or 3T3 mouse cells were noted [13]. Heterogeneity in membrane lipids, as well as in the locus of the fluorescent probes could explain the differing results. In many NMR and ESR measurements directed toward elucidation of lateral diffusion constants, one follows the spectral line width rather than line shape analysis [21-241. Since line-width analysis is an averaging method, it is limited in its ability to detect heterogeneity in the system under investigation. Similarly, in steady-state fluorescence polarization studies, calculated “microviscosities” may not represent the actual microviscosity but a value intermediate among the various viscosities of the media present in the system, weighted according to the distribution of the fluorescent probe. We now report the application of timeresolved fluorescence anisotropy (and lifetime) measurements of intact normal and virus-transformed 3T3 cells. The kinetics of fluorescence anisotropy gives directly the functional form of the all-important autocorrelation function [25, 261. This function provides specific information about the heterogeneity and rotational modes of the probe in the anisotropic membrane interior.
Our results with the lipophilic fluorescent probe, DPH, are consistent with the hypothesis that there is a nearly homogeneous (95 %) distribution of the probe in the membrane and confirm and further extend our previous interpretation of the steady-state fluorescence polarization experiments [27]. When the results reported here are compared with studies of fluorescence anisotropy of DPH in model lipid bilayer systems at varying lipid composition [28], a better understanding of the anisotropic character of the probe’s rotation dynamics is obtained. Our results show an increase in apparent “viscosity” with cell transformation, and a decrease in viscosity associated with reversion of cells to the untransformed state. Yet, more significant is the change in structure in the lipid interior of the transformed cell membrane, expressed in the increased anisotropy values to which the decay curves of anisotropy with time level off. Increased lectin-receptor mobility and agglutinability with cell transformation must thus be regulated by another membrane associated system, e.g. microtubules and/or microfilaments [ 1, 113.
MATERIALS
AND METHODS
Cells The isolation, growth conditions, and characteristics of the cell lines involved in this study have been described oreviouslv in our steadv-state fluorescence polarizailon study [ 131. Mouse 3i’3d, 3T3Py6, Py6R, 3T3 BALB A-31, and SVT2 cells were obtained from Dr Thomas Benjamin, Department of Pathology, Harvard Medical School. 3T3Py6 [29] are polyoma virustransformed 3T3d cells [30]. Py6R1 [3l] is a revertant line isolated from 3T3Py6 by the fluorodeoxyuridine procedure [3 I]. SVT2 cells are SV40-transformed 3T3 A-3 1 cells.
DPH labeling procedure Exponentially growing cells harvested under mild identical conditions. were labeled with 2x 10m6M DPH dispersed in PBS according to our previously-described procedure [13]. Unlabeled control cells were
Membrane treated with a solution of tetrahydrofuran in PBS (1 : 1000) under the same conditions.
Decay offluorescence emission and anisotropy measurements The decay of both fluorescence emission and fluorescence anisotropy as an explicit function of time in the nanosecond (nsec) range [32-341 may be measured by the single photon counting technique [35]. This method allows, in principle, the determination of hydrodynamic parameters from a single accumulation at a given temperature. The sample is excited with very short pulses of polarized light. The intensity of the emission polarized parallel, 111(r),and perpendicular I,(r), to the polarized excitation light are then measured as a function of time. Fluorescence decay measurements were performed at 37-38°C and carried out on equipment kindly made available by Dr Lubert Stryer, following the general experimental and deconvolution procedure described by Yguerabide [36] and Yguerabide et al. [37]. Two major conditions are essential for successful measurements of time-resolved fluorescence on intact, turbid cell suspensions; viz. (1) proper correction for scatter and intrinsic fluorescence of non-DPH labeled cells and cell clumps; and (2) short duration for accumulation of reproducible data. The following modifications of the Yguerabide experimental procedures were found necessary: (a) automatically alternating accumulation of parallel and perpendicular components of the fluorescence emission for short intervals (10 set) was necessary to eliminate errors due to fluctuating lamp intensity and fluctuating numbers of cells in the light beam. The sectors of the multichannel analyzer addressed by the output of the time-to-amplitude converter were synchronically switched. (b) The time dependence of fluorescence emission could be calculated from the total fluorescence emission, s(r), defined by eq. (I) [36,37]:
dynamics and viral transformation
207
measurement and at 5 min intervals. Accumulation times for labeled samples were 2&30 min (alternating parallel and perpendicular modes), resulting in 104-IO5 accumulated counts in the peak channel. The excitation light was filtered by a Coming 7-60 filter, and the emitted fluorescence was detected through a Coming 3-73 filter. Prior to performing the nsec photon counting measurements, DPH-labeled cells were examined by steady-state fluorescence polarization at 37°C.
RESULTS Total fluorescence emission as a function of time, S(t) (eq. l), depends on the rates of transitions which depopulate the lowest excited singlet state. For a homogeneous system it can be described by first-order kinet its (eq. 3): (3)
where S, is the initial emission intensity following a very short pulse of polarized light, t is the time in nsec, and r is the excited state lifetime; i.e., the inverse of the sum of the rates of all processes that depopulate the lowest excited singlet level of the chromophore. For more complex and heterogeneous systems, S(t) can be expressed as a sum of exponential terms [37]. S(r)=Z,1(r)+2z,(r) (1) Fig. 1 shows a representative curve of the while that of fluorescence anisotropy, A(t), from eq. nearly single exponential fluorescence de(2): cay of DPH embedded in 3T3d cells. Atqc-l,(r) A(t)= (2) tempts to improve the fit between the exSW perimental data and the theoretically comAn automated procedure [28] permitted the calculation puted double exponential decay curve indiof excited state lifetime (7) (calcu!ated by a least cated that as much as 95% of the intensity squares analysis), mean anisotropy (A), and a series of calculated convoluted anisotropy curves correspondis due to a single exponential decay mode ing to single rotational correlation times 7, were and only 5 % to a second mode. Moreover, plotted for comparison with the observed curves. The calculated curve with the same initial slope as the when compared to the decay curve of 8observed curve led to the here termed apparent rotaaniline-1-naphthalene sulfonic acid (ANS) tional correlation time, rpapD.The scattering background and intrinsic fluorescence of unlabeled cells, in ethanol (a homogeneous medium), DPH which was not larger than 5% of the fluorescence fluorescence decay curves of all cell lines emission of labeled cells, was subtracted. These procedures reduced the time required for data acquisition investigated showed similar or small deviaand increased the reliability of the resulting anisotropy tions from a calculated single exponential values. Cell suspensions were gently mixed 1 min before decay curve. Table 1 shows fluorescence Exp
Cd
Res
I18 (1979)
208
Pat-oh, Robbins und Blorrt
Table 1. Nunosecond cells ut 37°C
fluorescence
studies oj’ normul,
transformed
and re\lertunt
Cell line
7 (nsec)a
‘j
AZ6
pallp (nsec)Q,c
ht, (nsec)”
q (poise)+
3T3 A-3 I SVT2 3T3d 3T3Py6 PY6Rr
7.7to. I 7.8tO. I 8.0&O. 1 S.lrtO..l E.l+O.l
0. I I +0.01 0.14t0.01 0.12+0.01 0.14+0.01 0.10+0.01
0.07 0.12 0.10 0.12 0.08
6.0t 1.0 9.0f 1.5 5.8?0.3 9.3kO.3 4.8kO.8
2.5 3.1 2.8 3.7 2.7
1.0 I.6 1.0 1.5 0.8
3T3
0 Average values of three independent experiments. * Values corresponding to fig. 2. c The S.D.s are calculated on the basis of three independent experiments; for method of calculation see text and captions to the figures. d Rotational correlation time estimated by least-square fitting a line to the first 5 nsec of data expressed as log[A(r)-A,]. Since the psub values are not corrected for the finite width of the light pulse, they provide an estimate of relative correlation times (see text). p Taken from Fuchs et al. [l3].
lifetime values, 7, of DPH embedded in the membranes of normal A-31, SVT2-transformed, normal 3T3d, 3T3Py6 transformed and Py6R, revertant cells. Essentially no variation in DPH lifetime upon transformation (i.e. SVT2 vs A-3 1 and 3T3Py6 vs 3T3d
cells) or reversion (i.e. Py6R, vs 3T3d cells) is indicated. The decay of fluorescence anisotropy with time, A(t), expresses the change in orientation of the transition moment direction during the delay in emission from the first excited singlet state. For a very short light pulse, A(t) is at most slightly dependent on the excited state lifetime, since, by its definition, eq. (2), the division by S(t) eliminated the lifetime term. For the simplest case of a rigid sphere in an isotropic medium, A(t) decays exponentially (eq. 4): A(t)=A,,e-‘1”
time (nsec); ordinate: fluorescence intensity (counts). Total fluorescence emission decay for DPH in 3T3d cells at 37°C. The logarithm of fluorescence is plotted as a function of time. Solid line. -, Experimental data; ___, theoretical convoluted single exponential decay calculated for 7=8.0 nsec. The lamp profile is also shown. Data analysis is performed by the method of least squares.
Fig. 1. Abscissa:
E.rp Cdl
Rr,
118 fIY7Y)
(4)
where A0 is the initial value of the emission anisotropy [37] and cpis the rotational correlation time. The fluorescence anisotropy of the rod-shaped diphenylhexatriene molecule in an isotropic solvent should decay exponentially even though the molecule is not spherical because its longest wavelength transition moment is parallel to the long axis of the molecule. Kinetic determination of fluorescence depolarization would thus yield linear plots of In A vs time, the slope of which affords a value for cp directly; the slope is l/p and the intercept
Membrane
dynamics and viral transformation
20
209
so
Fig. 2. Abscissa:
time (nsec); ordinate: fluorescence emission anisotropy. Time dependence of the emission anisotropy of DPH, eq. (2), in membranes of 3T3 A-31 (...) and SVT2 (X x X) at 37°C (a) and in membranes of 3T3d (. . .) 3T3Py6 (X x X) and Py6Rr (000) at 37°C (b). The logarithm of the anisotropy is plotted as a function of time. The initial fast decay data points, coinciding
with the dashed line for 3T3 A-31 and with the solid line for SVT2 cell lines in (a) were visually compared with theoretically calculated convoluted plots of single exponential decay of anisotropy (detailed in [43]). Such calculated decay plots are included in (b), dashed lines. The scatter in points, particularly at the plateau region indicate the statistical counting errors where only a small number of photons are accumulated.
at time zero, is fnAO. Thus, all necessary information can be obtained from the same sample at physiological temperatures. Deviation from single exponential decay, eq. (4), may arise in various situations, e.g. restriction of the angular range of motion, or heterogeneity of the probe’s environment. In such cases A(t) will be a sum of exponentials,
two components; initially, a relatively fast one, within the first 12 (fig. 2a) and 10 (fig. 2b) nsec which then decays to a non-zero “plateau” value. The deviation of the anisotropy decay curves from those expected for a single component occurs between 12 and 20 nsec (fig. 2a) and between 10-15 nsec (fig. 2b) which corresponds to time intervals at which no deviation of total fluorescence emission (fig. 1) from a single exponential decay is observed; the deviation there (fig. 1) begins only after -25-30 nsec. The increased scatter in fluorescence emission anisotropy data points at longer time intervals, i.e. at the “plateau” region, arises from statistical variation in photon counting at a region where fluorescence emission is small. This scatter of the data points is due to the exponential decay of total fluorescence emission intensity (fig. l), which results in more than an order of magnitude
i.e. A(t)=&
Aie-“qf. i=l
Fig. 2 shows the typical time-dependent curves of the emission anisotropy of DPHlabeled A-3 1 and SVT2 cells (fig. 2a) and of DPH-labeled 3T3d, 3T3 Py6, and Py6R, cells (fig. 2 b). Characteristic of all cell lines studied here are the following features which are different from theoretically calculated single exponential decay of anisotropy curves: The decay curves exhibit at least
Exp Cell Res 118 (1979)
210
Parola, Robbins and Blout
decrease in fluorescence intensity 20 nsec after the peak. Since accumuiation periods are limited by the vitality of the biological system, only 104-IO5counts were accumulated at the peak channel, resulting in low count number in the “plateau” region. Thus, obviously beyond 30 nsec, the scatter in data points is quite large and should be disregarded. Summarized in table 1 are values calculated for mean anisotropy, A, and the plateau value, AZ, corresponding to the slow motion (over 100nsec). Apparent rotational correlation times, &,,,, obtained by a comparison of the initial slope of the experimental data with the single exponential decay curves, are also included in table 1. These hpp values are used strictly for qualitative comparative purposes, and one should take into account the long-time limiting A2 anisotropy values. Thus, the nonzero values of A2 will cause the apparent initial slope of the anisotropy plot to be less than that for the fast component alone. Instead of fitting the data to a single exponential decay, eq. (4), it is helpful to fit the data to eq. (5), A(t)=A1e-“a+A2
(5)
The easiest way to estimate p is to fit the slope of the line to a function log (A(t)-A,). The (Sub values (subtracted rotational correlation time) derived from a weighted linear-least-squares analysis of the first 5 nsec of data (-20 data points) after subtraction of A2 are included in table 1. Consequently, rotational correlation times corrected for the difference in A2 values are shorter and range between 2.5-3.7 nsec. For both 3T3 A-31 and 3T3d cells, transformation with SV40 and polyoma viruses, respectively, resulted in an increase in A, AZ, cp,,,, and (Sub; the same trend was also reported in our steady-state fluorescence polarization Exp Cdl Res II8 (19791
study [ 131.While changes in (F~,,,,are similar in magnitude to changes in calculated “microviscosities” (50-60s increase with transformation and about 20% decrease with reversion), a smaller change (yet in the same direction) is indicated for (Pan,)values which are corrected for the differences in A 2 values. Yet, the good correlation between steady state fluorescence polarization results with papp values might be merely fortuitous. Moreover, even the qualitative correlation with ‘psu,,values should be cautiously regarded since the small differences in (P& VdUeS may not be that significant due to increased errors rising from convolution uncertainties. These observations suggest that the observed changes in A values associated with virus transformation are due not only to changes in membrane fluidity (which may be expressed in (Sub) but primarily to changes in the anisotropic nature of the lipid bilayer, which is expressed in AZ. Associated with transformation is loss of some degrees of freedom of rotation of the probe, which is essentially inhibited from completely depolarizing the fluorescence emission. Interestingly Py6R, revertant cells exhibit a small decrease in A, AZ, and papp; (P&, is not significantly smaller than that observed with 3T3d normal cells. DISCUSSION Steady-state fluorescence polarization values, P, depend on both the excited state lifetime, 7, and the rotational relaxation time, cp.Thus, DPH lifetime measurements are essential in order to establish whether differences in P-values observed between normal, virus-transformed, and revertant cells [ 131do actually reflect changes in rotational dynamics which may reflect differences in membrane fluidity. This will be the case if we assume no change in the limit-
Membrane dynamics and viral transformation
ing fluorescence polarization value, PO, to be associated with cell transformation. The similarity in lifetime values observed for 3T3 A-31 and SVT2 and among 3T3d, 3T3Py6, and Py6R, cells (table 1) indicate that changes in membrane “fluidity” and/ or anisotropy (rather than DPH lifetime) lead to the observed changes in P-values. These results further indicate that, upon transformation by either SV40 or polyoma virus, DPH molecules are exposed to similar dipolar interactions, suggesting no gross alteration in their immediate solvation shell. Any attempt to study lipid bilayer fluidity of plasma membranes in intact cells using lipophilic probes is complicated by the uncertainty in the location of the probe. A lipophilic probe, by its nature, cannot be anchored to the membrane-water interface, since if an ionic site is necessary for anchorage such an ionic moiety would prevent the probe’s penetration to the core of the lipid bilayer. Thus, if one is forced to use a lipophilic probe, its diffusion through the plasma membrane into intracellular membranes cannot be excluded a priori. While DPH penetration through the aqueous interior of the cells appears to be unlikely, due to its low solubility in water, one cannot exclude its partitioning into the endoplasmic reticulum linked to the plasma membrane, rather than confining itself only to the latter. Attempts were made to ascertain the cellular location of DPH through fluorescence microscopy. No obvious concentration of the dye in lipid droplets was observed. The experiments, however, were limited by rapid bleaching of the dye [38] under the conditions employed [49]. The fluorescence decay curves of DPH in each of the cell lines studied approximates a single exponential character (95%) (e.g. fig. 1). This may indicate a similar, relatively homogeneous distribution of DPH
211
molecules in the membranes of each of these cell lines. These observations are particularly striking since the non-complete single exponential decays may be due to some reversible bleaching [ 13, 181.This excited state isomerization may not be totally eliminated even in the photon counting mode, where short and weak illumination pulses are used. Deviations from monoexponential decay, ascribed to effects of solvent viscosity on cis-trans isomerization have been noted previously [39-41]. From a spectroscopic viewpoint, this is perhaps the single disadvantage in the use of DPH as a fluorescent probe, which otherwise is ideal for determination of fluorescence polarization of bilayer lipid membranes [4245]. Distinct double exponential fluorescence emission decay profiles were recently reported for DPH in dimytistoyl lecithine (DML) vesicles [42], yet single exponential decays were observed for DPH in multilamelar DML vesicles [45], in di(dehydrosterculayl)-phosphatidyl choline (PC) liposomes [28] and in DL-cr-dipalmitoylphosphatidylcholine (DPPC) liposomes [46]. In a different study, the fluorescence of 12(Panthroyl) stearic acid incorporated into red blood cell membranes also decays as single exponential [47]. The differences among the various systems, ranging from 87 to 95 and 100% “single” exponential decays, may be due to varying microheterogeneity of sites. This is more likely with small bilayer vesicles (such as those used in the Brand’s study [42] than with large multilamellar liposomes [45]. Thus, our cell system exhibits an intermediate situation, resembling more the multilamellar DML vesicles in terms of that microheterogeneity. However, again this may be due to cis-trans isomerization of the DPH, which may not be as clearly detectable in the present study where accumulation of data is restricted by
212
Parola, Robbins and Blout
the vitality of the biological system. It should be pointed out, however, that the lifetime of DPH in our study is similar to that reported in DML vesicles, i.e., S.O?O.l nsec for the 3T3d system and 7.7&O. 1 nsec for the 3T3 A-31 system vs 7.7 for single and 8.0 for double decay analysis in the artificial membranes. The fluorescence anisotropy decay profiles shown in fig. 2 are different from such plots for DPH in a homogeneous viscous medium, glycerol (A. Parola, unpublished results) or paraffm oil [39]. In the latter two systems, DPH decay of anisotropy exhibits a continuous decay with A2 approaching zero values. Yet similar characteristics to the present decay study were recently observed in three independent studies of artificial membranes: in PC liposomes with varying amounts of cholesterol particularly with higher cholesterol content [28], in DML vesicles, particularly at the temperature below the midpoint of the gel-liquid crystalline transition (at 37.2”C A2 did approach zero) [42], and in DPPC liposomes
[461. Heterogeneity in DPH molecular population, with a fraction of it highly immobilized resulting in the “plateau” region at long times, and the rest free to rotate leading to the initial fast decay region (fig. 2) [45] may adequately describe the situation in small, single-walled vesicles [42], particularly since a double exponential decay of total fluorescence, characteristic of two different lifetimes, is observed in addition to the nonsingle exponential decay of A(t) vs time plots. However, due to the close to single exponential character of the fluorescence emission decay (fig. 1) and particularly since the anisotropy decay curves reach their plateau value about 10 nsec before the deviation from single exponential decay of total fluorescence emission is noted (though Exp Cd Rr., 118 (1979)
we cannot totally exclude such heterogeneity) an alternative model may be more plausible. It is based on the anisotropic character of the rotational motion of each DPH molecule, free to reorient only over a limited angular range. Such a model is consistent with the interpretation of the data for DPH observed with artificial membranes [28] and spin resonance experiments done with spin labels in lipid bilayers [48]. Such motion may be expected of DPH molecules because of their elongated shape and the highly anisotropic environment inside the membrane. Rotation of the chromophore about the long axis of the molecule cannot result in any depolarization because it does not lead to reorientation of the transition moment. However, a very rapid motion over a limited angular range, corresponding to a restricted “wobble” about the long axis of the molecule, could contribute to the initial decrease from A0 (0.33 for DPH in glycerol at O’C) to the initially observed anisotropy values (see fig. 2). Such a “wobble” motion seems to occur in liposomes, too, [28, 461. Veatch & Stryer have carried out time-resolved fluorescence studies of DPH in liposomes. They observed fluorescence anisotropy decay curves in this model system very similar to those shown in fig. 2, with the magnitude of A2 increasing with increasing cholesterol mole fraction. They concluded that the addition of cholesterol to phosphatidylcholine liposomes renders the environment highly non-isotropic. In steady-state fluorescence polarization experiment of complex membrane systems, the observed polarization is an average value. Therefore, the “microviscosity” values calculated from Perrin’s equation (invoking a number of gross assumptions [13]) may not represent the actual “microviscosity” but a value intermediate between the various viscosities in the media. Moreover, the
Membrane
interpretation of steady state data in terms of “microviscosity” assumes that equivalent A(t) curves would be observed for equivalent viscosities of reference (oil solution) and membrane systems and thereby implicitly that the same viscosity concept applies in both media. Yet comparison with time-resolved fluorescence anisotropy me urements, which is not an averaging / method, reveals qualitatively similar trends in the results: i.e. A, AZ, and (Sub increase with transformation and decrease with reversion, as “microviscosity” values do too [13]. Thus, despite the fact that estimated values of viscosity in a cone were found to be an order of magnitude smaller than the value of “microviscosity” estimated from steady state emission anisotropies [46], our observations qualitatively do confirm our previous steady-state measurements and are consistent with a relatively homogeneous distribution of the probe. Still, the term “microviscosities” (obtained from steadystate studies) should be used very cautiously; the quotation marks do mean to indicate the over-simplification in this expression, since they represent both fluidity and major structural effects. A comparison with model membranes further clarifies these combined fluidity and structural factors. Thus, while similar, non-single exponential decay curves could be obtained with PC vesicles containing varying amounts of cholesterol, these curves seem to differ from our cell systems in that the changes in A2 were not associated with changes in &b [28]. Yet in the cell lines studied here both A2 and &b vary with transformation. Thus, while changes in cholesterol levels in the different cell lines could presumably be the cause for the major difference observed inA, (rendering the membrane more anisotropic with higher cholesterol levels), other factors (lipid and protein composition) could lead to
dynamics and viral transformation
213
the relatively small differences observed in $4ub values, which may represent actual relative fluidity differences. However, even in our system, the observed small differences in $&ubmay be within experimental uncertainties due to convolution problems. A question often raised is whether the new properties detected in virus-transformed cells are due to the presence of the virus in the cells. We found similar trends in membrane fluidity and anisotropy with a BUdR dependent cell line derived from a malignant Syrian hamster melanoma line [49]. With this system, “transformation” can be reversibly controlled by the presence or absence of the small BUdR moiety. Fluorescence anisotropy decay curves for BUdR “transformed” cells showed similar profiles to those observed for SV40 and polyoma “transformed’ characteristics independent of the actual cause of “transformation”. In conclusion, this study shows that the careful design of experiments using two techniques-steady-state and nsec spectrofluorimetry-allows more confidence in the interpretation of the dynamic behavior of biological membranes. Experiments aimed at detecting phase transitions and fusion activation energies from plots of In r) vs l/T [13] may be most adequately performed with steady-state spectrofluorimetry. On the other hand, fluorescence lifetime information, rotational correlation times, and information about membrane anisotropy can be obtained only from the time-resolved spectrofluorimetry. For the cell system studied here, the present work indicates that increased fluorescence polarization values observed with viral transformation are primarily due to anisotropy changes in the membrane lipid bilayer and thus may indicate only a relatively small change in its microviscosity. Exp Cell Res 118 (1979)
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Parola, Robbins and Blout
We thank Dr Thomas Benjamin of the Department of Pathology at Harvard Medical School for providing us with the cell lines. The technical assistance of MS Lin Chen is gratefully acknowledged. We are very much indebted to Professor Lubert Stryer, Standford University, for the use of his photon counting equipment. We thank Professor William Veatch and Professor Samuel Latt, Harvard Medical School, for helpful discussions. This work was supported, in part, by NIH Grants AM07300 and 5-Rpl-CA14143-12 and the NC1 Grants CA14142 and CA14051. A. H. P. was a recipient of an NIH Postdoctoral Fellowship. A preliminary report on this work was presented at the Meeting of the Israel Biochemical Society, April I, 1976.Beersheva, Israel.
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