Biophysical and structural characterization of 1H-NMR-detectable mobile lipid domains in NIH-3T3 fibroblasts

Biochimica et Biophysica Acta 1438 (1999) 329^348 www.elsevier.com/locate/bba

Biophysical and structural characterization of 1H-NMR-detectable mobile lipid domains in NIH-3T3 ¢broblasts Amalia Ferretti a , Arno Knijn a , Egidio Iorio a , Simonetta Pulciani b , Massimo Giambenedetti a , Agnese Molinari c , Stefania Meschini c , Annarita Stringaro c , Annarica Calcabrini c , Isabel Freitas d , Roberto Strom e , Giuseppe Arancia c , Franca Podo a; * a

d e

Laboratory of Cell Biology, Istituto Superiore di Sanita©, Viale Regina Elena, 299, 00161 Rome, Italy b Laboratory of Virology, Istituto Superiore di Sanita©, 00161 Rome, Italy c Laboratory of Ultrastructures, Istituto Superiore di Sanita©, 00161 Rome, Italy Department of Animal Biology and C.N.R. Center for Histochemistry, University of Pavia, 27100 Pavia, Italy Department of Cellular Biotechnology and Hematology, University of Rome `La Sapienza', 00185 Rome, Italy Received 21 January 1999; received in revised form 8 April 1999; accepted 26 April 1999

Abstract Nature and subcellular localization of 1 H-NMR-detectable mobile lipid domains (ML) were investigated by NMR, Nile red fluorescence and electron microscopy, in NIH-3T3 fibroblasts and their H-ras transformants (3T3ras ) transfected with a high number of oncogene copies. Substantial ML levels (ratio of (CH2 )n /CH3 peak areas R = 1.56 þ 0.33) were associated in untransformed fibroblasts with both (a) intramembrane amorphous lipid vesicles, about 60 nm in diameter, distinct from caveolae; and (b) cytoplasmic, osmiophilic lipid bodies surrounded by own membrane, endowed of intramembrane particles. 2D NMR maps demonstrated that ML comprised both mono- and polyunsaturated fatty chains. Lower ML signals were detected in 3T3ras (R = 0.76 þ 0.37), under various conditions of cell growth. Very few (if any) lipid bodies and vesicles were detected in the cytoplasmic or membrane compartments of 3T3ras cells with R 6 0.4, while only intramembrane lipid vesicles were associated with moderate R values. Involvement of phosphatidylcholine hydrolysis in ML generation was demonstrated by selective inhibition of endogenous phospholipase C (PC-plc) or by exposure to bacterial PC-plc. This study indicates that: (1) both cytoplasmic lipid bodies and membrane vesicles (possibly in mutual dynamic exchange) may contribute (although to a different extent) to ML signals ; and (2) high levels of ras-transfection either inhibit ML formation or facilitate their extrusion from the cell. ß 1999 Elsevier Science B.V. All rights reserved. Keywords: Lipid body; Fibroblast; Ras-transformation; NMR; Electron microscopy; Fluorescence microscopy

Abbreviations: CE, cholesteryl esters; COSY, correlation spectroscopy; DG, diacylglycerol; DMEM, Dulbecco's modi¢ed Eagle's medium; FCS, fetal calf serum; FFA, free fatty acid; HBSS, Hank's balanced salt solution; IMP, intramembrane particle; ML, mobile lipid ; NMR, nuclear magnetic resonance; PBS, phosphate-bu¡ered saline; PC, phosphatidylcholine; PF, protoplasmic fracture (face); SPF, S-phase fraction; TEM, transmission electron microscopy; TG, triacylglycerol * Corresponding author. Fax: +39-6-4938-7144; E-mail: [email protected] 1388-1981 / 99 / $ ^ see front matter ß 1999 Elsevier Science B.V. All rights reserved. PII: S 1 3 8 8 - 1 9 8 1 ( 9 9 ) 0 0 0 7 1 - 2

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1. Introduction Still limited knowledge is available on speci¢c alterations occurring in actively proliferating cells at the level of metabolism, turnover and subcellular compartmentalization of lipids, in spite of increasing indications on the involvement of these components in cell physiology regulation [1] and also in tumor growth and invasiveness [2]. Novel perspectives were recently opened in this ¢eld by nuclear magnetic resonance (NMR) spectroscopy, whose non-invasive nature, combined with high chemical speci¢city, allows detection of intracellular pools of neutral lipids and phospholipid metabolites in intact cells and tissues and measurements of their alterations during tumor development and progression [3,4]. In particular, 1 H-NMR-visible mobile lipid domains (ML) are reported as a peculiar feature of malignant cells in vitro and in vivo [5]. ML signals also dominate the 1 H-NMR spectra of activated lymphocytes [6,7], cultured embryo cells [8], and also cells undergoing apoptosis [9]. The diagnostic signi¢cance of ML signals detected by ex vivo 1 HNMR spectroscopy and spectroscopic imaging methods has even been claimed for a number of invasive human tumors, including uterine cervix, thyroid and colon cancer [3,5]. The nature, subcellular localization and biological function of ML are still under investigation. Alternative hypotheses propose that narrow NMR lipid signals may arise from triacylglycerols (TGs) and/or cholesteryl esters (CE), organized in isotropically tumbling lipoprotein-like domains incorporated in the plasma membrane bilayer [10^12] or from cytoplasmic lipid bodies surrounded by phospholipid membranes [13,14]. Histochemical studies also demonstrated that neutral lipids are particularly abundant in perinecrotic cells and in necrotic areas of both experimental and human tumors ex vivo [15], while a positive correlation has been observed between NMR-visible ML signals and necrosis in high-grade human astrocytomas [16]. In an attempt to isolate the speci¢c contribution(s) given by oncogene-induced cell transformation to the generation of ML domains, and to further investigate the chemical nature and intracellular localization of these lipid structures, One-dimensional (1D) and two-dimensional (2D) 1 H-NMR experiments

were recently undertaken in our laboratory [17] on mouse embryo NIH-3T3 ¢broblasts and their oncogene-transformed cell variants, obtained by stable human H-ras transfection [18]. The capability of ras oncogenes to induce neoplastic transformation has been demonstrated in a number of cell systems in vitro [19]. In fact, ras oncogenes, mostly activated by point mutations at one or two main sites of their coding sequences and ¢rst identi¢ed as transforming genes of RNA tumor viruses, have also been recognized to play a role in human cancer [19,20]. Our previous NMR experiments have given the unexpected result that a clone of ras-transformed, in vivo tumorigenic ¢broblasts was characterized by signi¢cantly lower NMR-visible ML levels with respect to its untransformed, non-tumorigenic parental cell line [17]. This ¢nding, which clearly represents an exception to the general assumption that malignancy is associated with elevated ML contents, reinforced the interest of utilizing this particular cell model to investigate in more detail the molecular and biological mechanisms underlying the formation of NMRvisible lipid structures in eukaryotic cells. The present work reports the results of combined NMR, biochemical, cytochemical and ultrastructural studies carried out on ML domains in NIH-3T3 ¢broblasts and their H-ras-transformed cell variant, under di¡erent conditions of cell growth or mitogenic stimulation. Moreover, involvement of phosphatidylcholine (PC) hydrolysis in ML formation was assessed either by selectively inhibiting PC-speci¢c phospholipase C (PC-plc) in intact cells or by adding to them exogenous, bacterial PC-plc. The subcellular localization of NMR-visible ML structures was investigated by means of static and £ow cyto£uorimetric analyses of cells stained with a £uorescent lipid probe (Nile red), as well as by transmission electron microscopy (TEM) of either ultrathin sections or replicas of freeze-fractured samples. 2. Materials and methods 2.1. Chemicals Tricyclodecan-9-yl-xantogenate (D609) was purchased from Calbiochem (La Jolla, CA, USA) and Nile red from Kodak Laboratory Chemicals (New

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York, NY, USA). Puri¢ed bacterial phosphatidylcholine-speci¢c phospholipase C (EC 3.1.4.3) from Bacillus cereus (contamination by sphingomyelinase lower than 0.05%) was supplied in ammonium sulfate solution by Boerhinger Mannheim, Germany. Dioleoyl-phosphatidylcholine, cholesteryl arachidonate, Q-O-alkyl-L-acetyl- and Q-O-octadecyl-L-acetyl-phosphatidylcholine (used as standards for NMR peak assignments) were purchased from Sigma (St. Louis, MO, USA). 2.2. Cells Stabilized mouse embryo NIH-3T3 ¢broblasts, clone A6109 (3T3) and their transformed line induced by transfection with human H-ras oncogene (mutation G12V), clone 109.3.2 (3T3ras ) possessing a high number of oncogene copies (20^40), were grown under sterile conditions in 75 cm2 £asks at 37³C, in a 5% CO2 ^95% air incubator, in Dulbecco's modi¢ed Eagle's medium (DMEM) supplemented with 10% fetal calf serum (FCS) and prepared as previously described [18]. Stock cultures were split every 3^4 days at ratios of 1:10 or 1:20 and maintained without medium change. Fresh stocks were revived every 2 months (i.e. after about 10 sub-cultures) in order to avoid possible metabolic alterations associated with high passage number [21]. Cells were harvested in phosphate-bu¡ered saline (PBS) containing 1 mM EDTA. Tumorigenicity was tested by s.c. injection of either 106 or 107 cells into newborn mice [18]. All animals injected with 3T3ras cells developed solid tumors and died within 15 days. Mice injected with untransformed ¢broblasts were instead found to be tumor free up to 25 days after injection of 107 cells. For NMR, £uorescence and electron microscopy analyses, ¢broblasts were equally seeded at a density of 1.0U104 cells/cm2 and cultured in DMEM supplemented with 10% FCS; the medium was changed at 48 h and cells harvested at 72 h (late log phase). When speci¢ed, di¡erent seeding densities were adopted (either 0.5 or 2.0U104 cells/cm2 ) in order to collect cells at di¡erent growth phases (log phase and stationary (or con£uent) phase, respectively, at 72 h). Cell quiescence was induced by 24 h FCS-deprivation on ¢broblasts, which had been grown for 20 h in

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DMEM supplemented with 10% FCS (seeding density 2.0U104 cells/cm2 ). After quiescence, cells were again exposed to 10% FCS and analyzed at di¡erent times of re-stimulation (0, 1, 2 and 24 h). For experiments on PC-plc inhibition, cells were grown for 48 h in DMEM-FCS, and then cultured for the subsequent 24 h in the presence of tricyclodecan-9-yl-xantogenate (D609, 10 Wg/ml), a speci¢c PC-plc inhibitor [22]. Under these conditions, no inhibition of cell growth was induced on either cell line, as assessed by viable cell counting and £ow cytometry on propidium iodide-stained cells (L. Lenti and F. d'Agostino, personal communication). Bacterial PC-plc (from B. cereus, 0.2 U/ml) was added exogenously to the cells, harvested at the late log phase and incubated for 30 min in PBS (at a cell density of about 17U106 cells/ml). Flow cytometry of cells stained with the £uorescent DNA probe propidium iodide demonstrated that at quiescence (t = 0) the cell cycle distribution was very similar in both 3T3 and 3T3ras ¢broblasts, the (G0 +G1 ) and the S phase fraction (SPF) being 63^70% and 20^25%, respectively, in both cell types. Upon cell re-stimulation, SPF remained unaltered for at least 2 h and then evolved at 24 h to about 40% in 3T3 and 55% in 3T3ras (C. Ramoni, L. Dupuis et al., personal communication). 2.3. NMR analyses Intact cells were counted, washed three times in saline solution, centrifuged at 2700Ug and resuspended in 700 Wl of PBS (60% D2 O) before transfer to a 5 mm NMR tube (40^80U106 cells). The percentage of viable cells, determined by Trypan blue exclusion test, ranged between 80 and 95%, both before and after NMR analyses. 1 H-NMR experiments were performed on a Bruker AMX 400WB spectrometer (magnetic ¢eld 9.4 T). Analyses on intact cells were carried out at 25³C, using the 1D version of a sequence of homonuclear correlation via dipolar coupling and nuclear Overhauser e¡ect (NOESYPR 1D), included in the Bruker program library. The spectra were obtained in 17 min (320 scans), using 60³ pulses preceded by 1.0 s presaturation for water signal suppression (acquisition time 2.0 s, spectral width 4032 Hz, 16 K data points). Only the spectra in Fig. 5A were per-

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formed on a Gemini Varian 200 MHz spectrometer (magnetic ¢eld 4.7 T). Trimethylsilyl-2,2,3,3-d4 -propionate (10 mM in D2 O) was used as external chemical shift standard, contained in a 1-mm diameter capillary inserted in the NMR tube. Lipid extracts, prepared as previously described [17], were analyzed by applying a single pulse 1D sequence at equilibrium (90³ pulses and 5.0 s delay time), accumulating 128 scans. Two-dimensional 1 H-NMR experiments were performed on intact cells and extracts, using a 2D homonuclear shift correlation sequence (COSY). The COSY spectra of intact cells (or their extracts) were acquired with 80 (or 64) transients; measurements consisted of 256 (or 128) time domain points respectively in t1 and 512 in t2 , acquisition time 64 ms, with a spectral width in both dimensions of 4032 Hz (increment in t1 : vt1 = 0.248 ms). The repetition time was 1.0 s for intact cells or 2.5 s for extracts; continuous-wave irradiation was used to presaturate the water signal. The data were zero-¢lled in t1 and multiplied in both directions by a sine-bell or squared sine-bell window prior to 2D Fourier transformation to obtain 512U512 spectra; spectral processing consisted in t1 ridge removal and diagonal symmetrization. On-dimensional spectra of intact cells acquired before and after 2D experiments demonstrated that no metabolic changes occurred during the overall measurement time. All types of 1D and 2D NMR spectra were repeated on standard compounds for veri¢cation of signal assignments, which were further con¢rmed by literature reports [23^26]. Spectra of extracts were zero-¢lled by doubling the number of data points (to 32 K) and line-broadened by 0.2 Hz prior to Fourier transformation and subsequently quantitated by integration using WINNMR Bruker software (Germany). The latter allowed baseline correction by application of a cubic splines function through appropriate points. Signal processing and data analysis of the spectra of intact cells were performed directly in the time domain using the AMARES algorithm [27] within the Magnetic Resonance User Interface (MRUI) software package [28], i.e. by estimation of model function parameters through non-linear least-squares minimization. The model chosen to estimate the free

induction decays was a sum of exponentially damped sinusoids (i.e. Lorentzian lineshapes in the frequency domain) with the following parameters: amplitude (proportional to concentration and/or mobility of protons), phase, linewidth (1/T2 ), frequency (chemical shift). Since the signals decayed at a relatively fast rate, only the ¢rst 2048 data points were included in the quanti¢cation. The residual water signal was eliminated, ¢tting it with 10 exponentials by Hankel Lanczos Singular Value Decomposition program [29]; no line broadening or other signal processing was applied before quantitative analysis. As general prior knowledge, so-called `soft constraints' were used to constrain model parameters within logical bounds in order to maximize the accuracy and robustness of the model function. Furthermore, the zero-order phases of all signals were supposed to be equal, the spectrometer pre-scan-delay was set at 0 and the linewidths of the resonances at 1.59, 1.68 and 2.25 ppm were kept equal, to guarantee stability. In the region from 1.10 to 1.50 ppm, the spectra of intact cells are characterized by an extensive overlap of resonances. By modeling this region with four exponentials, it was possible to quantify the signal exclusively arising from the ^(CH2 )n ^ protons of the mobile lipids' fatty chains (resonating at 1.29 ppm) with one exponential, while the contributions from overlapping neighboring peaks (such as lactate, alanine, threonine, etc.) were ¢tted by the model with the other three exponentials. Thus the ratio R of the peak areas of ^(CH2 )n ^ with respect to ^CH3 could be calculated with a higher precision. 2.4. Chemical analyses Lipid analysis by gas chromatography and enzymatic determinations of triacylglycerol and free fatty acids (FFA) concentration, were performed as previously described [17]. 2.5. Fluorescence microscopy For £uorescence microscopy observations, cells grown on glass coverslips (10 mm in diameter) were air-dried and stained with Nile red [15,30]. To this end, a stock solution of the dye in acetone (0.5 mg/ml) was diluted 1:200 in glycerol (75% by vol), the solution thoroughly degassed and an aliquot

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added directly to the cell preparation. The sample was incubated for at least 5 min before £uorescence microscopy analyses, carried out with a Nikon Microphot-SA (Nikon, Tokyo, Japan). The Nile red, yellow-gold £uorescence was observed with the ¢lter set for £uorescein and namely: 450^500 nm bandpass excitation ¢lter; 510-nm centered dichroic mirror; and 528 nm long-pass barrier ¢lter. The stained cells were photographed in black and white on Kodak Tri-X pan ¢lm (400 ASA). 2.6. Flow cytometry The cells, harvested with EDTA and trypsin, were resuspended in Hanks' balanced salt solution (HBSS) and stained with 100 ng/ml Nile red at 37³C for 15 min. After washing in PBS, the samples were resuspended in the same bu¡er and immediately analyzed with a FACScan £ow cytometer (Becton Dickinson, Mountain View, CA) equipped with a 15-mW, 488-nm air-cooled argon ion laser. Fluorescence emission was analyzed by employing laser excitation wavelength of 488 nm and emission wavelength of 585 nm (yellow £uorescence). Data were collected and analyzed on a Hewlett-Packard model 310 computer interfaced with the FACScan equipment. At least a minimum of 10 000 cells was counted per analysis. The log of the £uorescence intensity was plotted on the x-axis and the number of cells possessing a given intensity on the y-axis. 2.7. TEM of ultrathin sections Cells were ¢xed with 2.5% glutaraldehyde in 0.2 M cacodylate bu¡er (pH 7.4) for 2 h at room temperature, washed and post¢xed with 1% OsO4 in the same bu¡er for 1 h at room temperature. Cells were then dehydrated in an alcohol gradient and embedded in epoxy resin (Agar 100 resin, Agar Scienti¢c, Stansted, UK) by routine procedures. Ultrathin sections, obtained with LKB Ultrotome Nova ultramicrotome, were stained with uranyl acetate and lead citrate and examined with a Zeiss EM10C transmission electron microscope at 60 kV. 2.8. Freeze-fracture electron microscopy Cells were ¢xed with 2.5% glutaraldehyde in the

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culture medium. After 20 min of ¢xation, cells were detached mechanically and centrifuged for 10 min at 120Ug. After two washings in HBSS, cells were resuspended in the same medium containing 25% glycerol and incubated for 20 min at room temperature. The suspension was then centrifuged at 170Ug for 15 min and the pellet was put on carriers and quickly frozen in Freon 22 partially solidi¢ed by cooling with liquid nitrogen. The mounted carriers were then transferred into a Balzers BAF 300 freeze-etch unit, cleaved at 3100³C at a pressure of 2^4U1037 mbar, shadowed with 2.5 nm of platinum^carbon and replicated with 20 nm carbon deposits. Platinum^carbon evaporation (at an angle of 45³) and carbon evaporation (at an angle of 90³) were performed using electron beam guns. The thickness of the deposit was evaluated by means of a quartz crystal thin ¢lm monitor. Cells were digested for 2 h from the replica by Chlorox. The replicas were mounted on naked 300-mesh grids and examined with a Zeiss EM10C electron microscope at 60 kV. For each sample, several replicas were obtained and examined. 3. Results 3.1. 1 H-NMR signals of ML in 3T3 ¢broblasts and their H-ras transformants Con¢rming previous results [17], untransformed 3T3 ¢broblasts, grown to the late log phase of growth, exhibited more intense mobile lipid signals from fatty chains' chemical groups, than the rastransformed cell clone, grown under similar conditions (examples in Fig. 1). A more accurate determination of the ratio R between the ^(CH2 )n ^ and the ^ CH3 peak areas, currently used as an indicator of mobile lipid NMR visibility, could be performed (Table 1). This parameter is also dependent upon average ML fatty chains' length and unsaturation level. Quantitative spectral analyses showed that the ^CHNCH^ to ^CH3 peak ratio was signi¢cantly lower in 3T3ras than in 3T3 ¢broblasts (Table 1), indicating a lower degree of unsaturation of NMRvisible fatty chains in the transformed clone. This evidence was further substantiated by the fact that the signal at 2.04 ppm (at least partly due to NCH^ CH2 ^CH2 ^ groups) was also decreased in the ras-

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transformed cells. Since a higher degree of fatty chain unsaturation implicates a lower intensity of the ^(CH2 )n ^ peak, the di¡erence in unsaturation between 3T3 and 3T3ras further accentuates the difference in ML visibility between the two cell lines. On the basis of cross-peaks of correlation between adjacent chemical groups, 2D COSY spectra of intact cells allowed the identi¢cation of several other signals, some of which due to mobile lipids, including speci¢c unsaturated fatty acid chains. In the example shown in Fig. 2A (2D NMR map of intact 3T3 cells), it is possible to observe the cross-correlations respectively arising from the linolenic acid NCH^CH2 ^ CH3 group (L, 0.9/2.1 ppm) and the OOC^CH2 ^ CH2 ^CH2 ^CHNCH^ group of arachidonic acid (M, 1.7/2.1 ppm). Moreover, the cross-peaks respectively due to NCH^CH2 ^CH2 ^ (C, 2.0/5.4 ppm) and NCH^CH2 ^CHN (D, 2.8/5.4 ppm) groups of monoand poly-unsaturated fatty chains can be clearly identi¢ed (Fig. 2A). Further peak assignments, either to lipids or to other compounds, are reported in the legend of Fig. 2. In comparison with 3T3, 3T3ras cells showed lower or barely detectable cross-peaks from unsaturated chains (example in Fig. 2B) with substantially decreased intensities of C and D cross-peaks. Furthermore, the speci¢c cross-correlation arising from arachidonic acid (M) was always below detection level. Among the other cross-peaks detected in 2D COSY maps of intact untransformed 3T3 cells, of particular interest are those indicated by K and GE in Fig. 2A, respectively arising from the ^O^CH2 ^ CH2 ^ (1.6/3.5 ppm) and ^O^CH2 ^CH^O^ (3.5/5.1 ppm) groups of ether lipids. The resonances of these

Fig. 1. Examples of 1D 1 H-NMR spectra on intact NIH-3T3 ¢broblasts (top, R = 2.19) and their ras-transformants (bottom, R = 0.38) with assignment of the major mobile lipid signals. The resonance attributed to OOC^CH2 ^CH2 ^ (1.59 ppm) may also comprise contributions from ^O^CH2 ^CH2 ^ (either ether-linked or free), OOC^CH2 ^CH2 ^CH2 ^CHNCH^ (e.g. arachidonic acid) and cholesterol C25 methine, as better resolved in 2D spectra (Fig. 2). The peak attributed to NCH^CH2 ^CH2 ^ (2.04 ppm) may include contributions from NCH^CH2 ^CH3 (linolenic acid), glutamate and a still unknown signal (having a cross-correlation with a signal at 3.42 ppm, as shown in Fig. 2A). The resonance attributed to OOC^CH2 ^CH2 includes contributions from valine and an unknown compound. The residual water peak (4.8 ppm) has been removed from the spectra using Hankel Lanczos singular value decomposition as described in Section 2.

groups could not be detected in 1D spectra of intact cells, due to their relatively low intensity and heavy overlap with neighboring resonances, but were clearly resolved in 1D spectra of 3T3 lipid extracts, together with a sharp triplet at about 3.5 ppm, due to the ^CH2 ^OH group of free fatty alcohols (data not shown). On the contrary, K and GE cross-peaks were

Table 1 Peak area ratios of di¡erent ML signals to the ^CH3 resonance, in intact untransformed NIH-3T3 ¢broblasts (3T3) and their H-ras transformants (3T3ras )a NMR signalb

N (ppm)

3T3 (n = 8)

3T3ras (n = 9)

Pc

^(CH2 )n ^ OOC^CH2 ^CH2 NCH^CH2 ^CH2 ^ OOC^CH2 ^CH2 ^ ^CHNCH^

1.29 1.59 2.04 2.25 5.32

1.56 þ 0.33 0.17 þ 0.05 0.95 þ 0.24 0.31 þ 0.07 0.08 þ 0.02

0.76 þ 0.37 0.12 þ 0.04 0.74 þ 0.15 0.32 þ 0.08 0.02 þ 0.02

6 0.0005 6 0.025 6 0.06 6 0.90 6 0.0005

a

Cells harvested at the late log phase of growth. Some resonances include additional contributions from other signals, as indicated in the legend of Fig. 1. c Statistical signi¢cance of di¡erences (two-tailed Student's t-test). b

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Fig. 2. Typical examples of 2D COSY maps of intact NIH-3T3 ¢broblasts (A) re-stimulated by FCS for 2 h, after 24 h of serum deprivation (R = 1.56) and their ras-transformants (B) harvested at the stationary phase of growth (seeding density 2.0U104 cells/cm2 ; R = 1.00). For further details see Section 2. Cross-peak assignments : lipid fatty acyl chains, A (0.9/1.3 ppm), ^CH2 ^CH2 ^CH3 ; B (1.3/ 2.1 ppm), NCH^CH2 ^CH2 ^; C (2.0/5.4 ppm), NCH^CH2 ^CH2 ^; D (2.8/5.4 ppm) NCH^CH2 ^CHN; E (1.3/1.6 ppm), OOC^CH2 ^ CH2 ^CH2 ^CH2 ^; F (1.6/2.3 ppm) OOC^CH2 ^CH2 ^CH2 ^; L (0.9/2.1 ppm) NCH^CH2 ^CH3 (linolenic); M (1.7/2.1 ppm) OOC^CH2 ^ CH2 ^CH2 ^CHNCH^ (arachidonic); glycerol backbone, GPP (4.1/4.3 ppm), C(1)HHP^C(2)H^ and C(3)HHP^C(2)H (triglycerides) ; G (4.2/5.3 ppm), C(1)HHP^C(2)H^ (triglycerides, diglycerides, phospholipids); ether lipids (glyceryl ether diesters), K (1.6/3.5 ppm), O^ CH2 ^CH2 ^; GE (3.5/5.1 ppm), O^CH2 ^CH^O; cholesterol moiety, Z (0.9/1.5 ppm), methyl ^C(26)H3 and ^C(27)H3 and methine ^C(25)H^; aqueous soluble cytoplasmic metabolites, 1, alanine; 2, aspartic acid; 3, glutamic acid/glutamine/glutathione; 4, myo-inositol; 5, leucine; 6, lysine and polyamine; 7, threonine; 8, valine; 9, taurine; 10, choline; 11, ethanolamine; 12, glycerophosphocholine; 13, lactate; 14, phosphocholine; 15, phosphoethanolamine.

below detection in 2D COSY maps of intact 3T3ras cells, although the corresponding resonances were present in total cell lipid extracts. As for TGs, the intensity of the glycerol backbone cross-peak G (4.2/5.3 ppm) was relatively weak in 3T3 and even weaker (if not absent) in 3T3ras cells. On the other hand, the cross-peak Z (0.9/1.5 ppm), attributed to cholesterol, was also equally present in the 2D maps of both 3T3 and 3T3ras cells, whereas the cross-peak at 2.2/4.6 ppm, characteristic of esteri¢ed cholesterol, was not visible in either cell line. 3.2. E¡ects of cell culture conditions on ML signals Recent evidence demonstrated higher NMR-visible mobile lipid levels in murine L ¢broblasts harvested at con£uence with respect to those collected in the log phase of growth [24]. 1 H-NMR experiments were therefore carried out to compare the ML spectral features of 3T3 and 3T3ras cells cultured at di¡erent

seeding densities, and then harvested in di¡erent phases of cell growth, respectively log phase, late log phase and stationary (or con£uent) phase. Analyses of two independent experiments showed a slight (if any) di¡erence in ML signal intensity in 3T3 cells at either log phase (R = 1.23), late log phase (R = 1.32) or stationary phase (R = 1.39). Parallel experiments con¢rmed lower R ratios in 3T3ras cells, with no substantial alteration on going from early (R = 0.95) to late (R = 0.86) exponential phase of growth, or stationary phase (R = 0.96). In order to further assess the e¡ects of di¡erent conditions of cell growth on ML levels, both 3T3 and 3T3ras ¢broblasts (cultured under normal conditions for 24 h) were brought to quiescence and their 1 H-NMR spectra analyzed at di¡erent times of restimulation by FCS. 1 H-NMR experiments showed that in untransformed 3T3 ¢broblasts (Fig. 3, left panel) the R ratio, already as high as 1.47 in quiescent cells (t = 0)

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Fig. 3. One-dimensional 1 H-NMR spectra of NIH-3T3 ¢broblasts (left panel) and their ras-transformants (right panel) at di¡erent times of FCS-induced cell re-stimulation after quiescence (for further details see Section 2). Closely similar results were obtained in two independent series of experiments.

remained constant at early stages of FCS re-stimulation (up to 2 h, R = 1.56) and slightly increased at 24 h (R = 1.80). A much lower NMR visibility was consistently found for mobile lipids detected in rastransformed cells (Fig. 3, right panel), although the R ratio underwent a 2^3-fold increase from 0^2 h (R = 0.25) to 24 h (R = 0.68), after FCS re-addition. 3.3. E¡ects of PC-speci¢c plc on ML signals It has recently been proposed [6,7] that activation

of the PC cycle [31] may contribute to the generation of NMR-visible lipids in both thymic and splenic lymphocytes stimulated by phorbol myristate acetate and ionomycin. Furthermore, in a previous work, we demonstrated that PC hydrolysis mediated by a PC-speci¢c plc enzyme actually occurs at substantially higher rates in homogenates of both 3T3 (0.66 þ 0.14 nmol/106 cells/h) and 3T3ras (0.38 þ 0.10 nmol/106 cells/h) cells than in mature, non-proliferative mammalian cells [32]. This evidence points to a possible role of the PC cycle in the generation of

Fig. 4. E¡ect of D609 (speci¢c PC-plc inhibitor) on NMR-visibility of mobile lipids in intact NIH-3T3 and 3T3ras ¢broblasts at late log phase. The R value decreased from 1.76 to 1.44 in the former and from 0.99 to 0.69 in the latter cells. The bottom traces represent the di¡erence between the two respective upper and middle spectra. For further details see Section 2.

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NMR-detectable lipids in these cells. In order to verify this hypothesis, 1 H-NMR spectra were obtained in both untransformed and transformed intact ¢broblasts, following incubation with D609, a speci¢c PC-plc inhibitor (Fig. 4). Quantitative analyses with AMARES on two independent series of experiments, showed that the R ratio decreased by 20 þ 10% in both 3T3 and 3T3ras cells exposed to D609, as compared to the respective control preparations. An even more direct evidence on the involvement of PC hydrolysis in ML formation was obtained by incubating the cells in the presence of bacterial PCplc. 1 H-NMR measurements demonstrated a substantial R increase in intact 3T3 cells (from 1.78 to

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2.43), while in 3T3ras , this ratio was even more markedly enhanced (from 0.56 to 1.30), reaching levels typically observed in 3T3 cells (Fig. 5A). A second experiment on 3T3ras cells exposed to bacterial PCplc con¢rmed this behavior (R increasing from 1.09 to 1.71). Therefore, treatment of 3T3ras cells with exogenous PC-plc apparently stimulated the formation of NMR-visible ML domains, making their spectra pro¢les similar to those of untransformed 3T3 cells, not only with respect to the ^(CH2 )n ^ signal, but also regarding the relative intensity of the ^ CHNCH^ resonance (as con¢rmed by quantitative analyses). This body of evidence indicates that at least partial contributions to the production of NMR-detectable mobile lipids in 3T3 and 3T3ras ¢broblasts are associated with the activity of PC-plc, with consequent production of diacylglycerol (DG) and phosphorylcholine. Since high DG levels are not compatible with plasma membrane integrity and cell viability, an excess of these compounds is likely converted into other neutral lipids, particularly TG and FFA, besides providing precursors to phospholipid re-synthesis. These considerations suggested the interest of performing chemical analyses on TG and FFA contents in the two cell lines. 3.4. Chemical characterization of total cell lipid components

Fig. 5. Induction of alterations on ML domains by 3T3ras cell exposure to bacterial PC-plc (from B. cereus). (A) 1D 1 H-NMR spectra of 3T3ras cells incubated for 30 min either in the presence (bottom trace; R = 1.30) or in the absence (upper trace; R = 0.56) of the exogenous enzyme. (B) Flow cytometry histograms of untreated (999) and treated (c c c) cells, stained with Nile red. The mean value of £uorescence intensity increased by a factor 2.3 upon exposure to the enzyme.

Lipid analyses were performed by: (1) chemical assays on whole cell extracts in organic phase (chloroform/methanol) to measure the respective total concentrations of TG and FFA; and (2) gas chromatography on total lipid extracts, to determine composition and levels of unsaturation of fatty chains. In substantial agreement with preliminary measurements already reported [17], chemical assays performed on total lipid extracts of 3T3 ¢broblasts (after cell harvest at the late log phase and NMR analysis) resulted in an average TG level of 1.1 þ 0.5 nmol/106 cells (n = 7), associated with a rather substantial amount of FFA (15.2 þ 4.7 nmol/ 106 cells). The period of cell packing in the NMR tube was short enough not to induce any substantial change in TG and FFA concentration levels, as demonstrated by comparison with the results of analyses

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on freshly collected ¢broblasts. The total TG (1.9 þ 1.0 nmol/106 cells) and FFA (8.6 þ 1.5 nmol/ 106 cells) levels were not signi¢cantly di¡erent in ras-transformed ¢broblasts, whether extracted immediately after cell harvest or after NMR analysis. No correlation was found between NMR-visible ML levels (R value) in intact cells and the concentration levels of TG and FFA in the extracts of the two cell lines, within the respective ranges of variability. Preliminary analyses also showed similar CE contents (V0.3 nmol/106 cells) in both transformed and untransformed cells. Gas chromatography analysis allowed the determination in both 3T3 and 3T3ras cells of total lipid (including phospholipid) fatty chains' composition. The predominant fatty chain residues were 16:0 (palmitic), 18:0 (stearic) and 18:1 (mostly oleic (18:1(9)), but also vaccenic 18:1(11)). The percent distribution of [16:0]:[18:0]:[18:1] were (30 þ 3):(21 þ 4):(32 þ 7) for 3T3 and (23 þ 4):(22 þ 4):(37 þ 6) for 3T3ras , independently from particular conditions of growth phase or cell stimulation. The other mono-unsaturated chains, 16:1 (palmitoleic) and 14:1 (myristoleic) occurred with percent distributions (respectively, 9 þ 1% and below 1%) which were similar in both cell lines. Poly-unsaturated chains (18:2 (linoleic), 18:3 (linolenic) and arachidonic (20:4)) represented a limited percentage of lipids in both 3T3 and 3T3ras cells. The percent distribution of fatty chains in saturated (S), mono-unsaturated (M) and poly-unsaturated (P) classes, measured under the di¡erent conTable 2 Percent distribution of saturated (S), mono-unsaturated (M) and poly-unsaturated (P) and average lengths of fatty chains in total lipid extracts of untransformed NIH-3T3 cells (3T3) and their H-ras transformed clone (3T3ras ) Class of saturation and chain lengths

3T3 (n = 6)

3T3ras (n = 6)

S M P LCH2 a LFA b

56.7 þ 8.3 38.5 þ 10.1 5.8 þ 3.1 11.26 þ 0.12 17.12 þ 0.40

51.3 þ 6.9 47.0 þ 6.9 1.7 þ 2.2 11.17 þ 0.29 17.20 þ 0.08

Di¡erences were not statistically signi¢cant (P s 0.1 by Student's t-test). a LCH2 , average length of saturated (CH2 )n segments. b LFA , average total length of fatty chains (from both phospholipids and lipid components).

Fig. 6. Fluorescence microscopy of Nile red-stained NIH-3T3 ¢broblasts (A; R = 2.19) and their ras-transformants (B; R = 0.38). The inset (3U) in A shows a string of £uorescence particles at the surface level of 3T3 cells. Scale bar: 10 Wm. (C) Flow cytometry histograms of a di¡erent pair of 3T3 and 3T3ras cell preparations (3T3, R = 1.40, 999; 3T3ras , R = 0.76, c c c).

ditions of cell growth described in the previous sections, are shown in Table 2. No signi¢cant correlation was found between the R values of ML domains measured in intact cells and the P or (M+P)

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contents, nor with the average total length (LFA ) or the average length of the saturated ^(CH2 )n ^ segments (LCH2 ) of fatty chains in total cell extracts. 3.5. Cytochemical characterization of lipid domains in 3T3 and 3T3ras cells Fluorescence microscopy examinations of Nile red-stained ¢broblasts demonstrated the presence of a large number of globular lipid structures inside the cytoplasm of virtually all the examined 3T3 cell preparations (example in Fig. 6A, R = 2.19). The diameter of these £uorescent bodies ranged from the resolution limit (0.1^0.2 Wm) to about 1 Wm. Moreover, in some cells, a string of £uorescent spots was visible at the cell surface level (inset of Fig. 6A). Conversely, no cytoplasmic lipid droplets were detected and only a weak and di¡use £uorescence could be observed (Fig. 6B) upon Nile red staining of ras-

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transformed cells characterized by a low R value (0.38). The di¡erent expression of Nile red-positive structures between normal and transformed ¢broblasts was con¢rmed by £ow cytometry analysis. Fig. 6C shows an example of £uorescence intensity distribution obtained from either 3T3 (R = 1.40) or 3T3ras (R = 0.76) cells. In the former, the signal at 585 nm (corresponding to the staining of neutral lipids) appeared to be signi¢cantly more intense (1.8U) than that produced by the transformed cells. However, when 3T3ras cells were exposed to exogenous PC-plc, which resulted in a striking increase (vR = 0.74 units) in NMR-visible ML (Fig. 5A), the £uorescence intensity (obtained upon Nile red staining) also substantially increased (2.3U; Fig. 5B). Accordingly, £uorescence microscopy showed the formation of numerous intracytoplasmic lipid bodies (data not shown).

Fig. 7. TEM analyses of ultrathin sections of NIH-3T3 ¢broblasts (R = 2.19; A^C) and their ras-transformants (R = 0.38; D,E) stained with osmium tetraoxide. Scale bar: 1 Wm.

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Fig. 8. TEM analyses of freeze-fractured NIH-3T3 ¢broblasts (R = 2.19). (A)Protoplasmic fracture face (PF) of plasma membrane. Cross-fractured microvilli (full arrowheads) and numerous protein intramembrane particles (IMPs) are visible. Several spherical smooth structures (arrows), protruding from the surface of the fracture face, are also detectable. The inset (4U) shows a high magni¢cation of a spherical smooth structure thrusting out of the membrane bilayer. Empty arrowheads point to plasma membrane invaginations attributed to caveolae. (B) Cytoplasm of a cross-fractured cell. Several globular bodies enveloped by a membrane with distinct IMPs are present. The inset (2.5U) shows a high magni¢cation of a cross-fractured lipid body with a smooth content, distinguishable from the surrounding rough cytoplasmic matrix. Scale bar: 1 Wm.

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Fig. 9. TEM analyses of freeze-fractured 3T3ras ¢broblasts. (A) PF face of the plasma membrane of a cell preparation with low ML content (R = 0.38). Several IMPs randomly distributed are detectable. (B) Cytoplasm of a cross-fractured cell (R = 0.38). Numerous cytoplasmic organelles, such as mitochondria (arrowheads) and Golgi apparatus (arrows), are visible. On the left of the picture, a portion of the nuclear envelope can be detected. (C) PF of the plasma membrane of a 3T3ras cell with moderate ML content (R = 0.56) showing the presence of globular lipid bodies. In the same cell preparation, no intracytoplasmic lipid bodies could be detected (D). Scale bar: 1 Wm.

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3.6. Ultrastructural characterization of lipid domains in 3T3 and 3T3ras cells The observation by TEM of ultrathin sections allowed detection of a number of electrondense globular bodies in the cytoplasm of 3T3 cells (Fig. 7A). Due to their strong osmiophilic nature, these structures could be identi¢ed as lipid bodies containing high amounts of unsaturated fatty acids. Quite often these particles appeared to be arranged in a grapelike shape with fusion processes taking place among adjacent granules (Fig. 7B). The maximum diameter of the individual granules was about 1.5 Wm. In some sections, lipid bodies appeared well delimited and surrounded by a membrane-like layer (Fig. 7C). These osmiophilic bodies were not visible (Fig. 7D), even at high magni¢cation (Fig. 7E), in the cytoplasm of most of 3T3ras cells possessing a low R ratio of 0.38 (processed exactly as 3T3 cells). In order to further investigate ML intracellular localization, 3T3 and 3T3ras cells were analyzed by freeze-fracture electron microscopy. In freeze-fractured cells, the fracture passes either through the hydrophobic plane of the plasma membrane or across the cytoplasm. In the former case, the molecular ultrastructural organization of both the protoplasmic (PF) and exoplasmic (EF) fracture faces of the plasma membrane could be examined; in the latter case, the intracytoplasmic components could be visualized. Fig. 8A shows a large portion of PF in the plasma membrane of a 3T3 cell. Some crossfractured microvilli (full arrowheads) and numerous protein intramembrane particles (IMPs), randomly distributed, were well visible. In addition, a number of spherical smooth structures with an average diameter of about 60 nm (subtracting the platinum-carbon coating thickness) appeared to protrude from the surface of the fracture face (arrows). These structures, intercalated within the plasma membrane, very likely correspond to lipid vesicles, that assume a globular shape and segregate from the phospholipid bilayer as well as from membrane proteins. The inset of Fig. 8A shows one of these structures surrounded by numerous IMPs, observed at high magni¢cation. Quantitative estimation indicated that the overall volume of these lipid vesicles did not exceed the value of about 0.2 Wm3 /cell. Another kind of component with about the same size as the before-mentioned

smooth spherical vesicles, appears as invaginations in the fractured plane (empty arrowheads). These invaginations, given their shape and size (about 60 nm) are most probably caveolae [33,34]. The di¡erent disposition of platinum coating allows easy discrimination between the globular lipid structures protruding from the fracture and the concave caveolar invaginations. In the cytoplasm of cross-fractured 3T3 cells, a variety of intracellular organelles could be detected (Fig. 8B). Among these, several globular bodies enveloped by a membrane with evident IMPs were present. In cross-fractured globular bodies, it was possible to con¢rm the presence of an enveloping membrane and to establish their amorphous content, as better demonstrated at high magni¢cation (inset of Fig. 8B). These observations strongly suggest the lipid nature of their content. The total volume of these cytoplasmic lipid bodies was estimated to be 4^8 Wm3 per cell. In freeze-fractured 3T3ras cells possessing low ML contents (R = 0.38) only very few (if any) 60 nm lipid particles and caveolae were detected in the plasma membrane. These cells also lacked intracytoplasmic spherical lipid bodies. Fig. 9A shows the PF face of the plasma membrane of a ras-transformed cell with randomly distributed IMPs, while Fig. 9B shows a cross-fractured 3T3ras cell with numerous cytoplasmic organelles, but apparently no spherical lipid bodies. On the other hand, in 3T3ras cells possessing moderate ML contents (R = 0.56) lipid vesicles were observed only at the membrane level but, again, no lipid bodies were detected in the cytoplasm (Fig. 9C,D). When transformed cells were exposed to exogenous PC-plc, elevation in the R value was associated with the massive appearance of cytoplasmic lipid bodies detected by TEM of either ultrathin sections or freeze-fractured cells (data not shown). 4. Discussion Although high resolution 1 H-NMR spectroscopy has been applied to intact cells for over 15 years, no de¢nite explanation is as yet available on the nature, subcellular localization and biological function of mobile neutral lipid structures responsible for the characteristic elevation of methylene, methyl and

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methine peaks frequently occurring in the spectra of malignant cells and activated lymphocytes [12^14, 35]. In order to give rise to these relatively narrow NMR signals, the corresponding lipid domains should be endowed with a su¤ciently high level of isotropic mobility [36] and cannot therefore be simply aligned in a lipid bilayer. NMR-visible ML domains are generally assumed to be associated with cell transformation and positively correlated with tumor malignancy grade. However, the 1 H-NMR analyses carried out in this study demonstrated lower NMR visibility of ML domains in intact, in vivo tumorigenic ras-transformed ¢broblasts than in the parental, non-tumorigenic cell line, at di¡erent phases of cell growth, as well as during cell stimulation from quiescence. Cell transformation cannot, therefore, be assumed to necessarily bring about an increase of ML domains. 4.1. Chemical nature of ML in 3T3 and 3T3ras ¢broblasts Limited information is available on biochemical contents of NMR-visible ML domains in intact cells, although it is generally accepted that TG (in exchange with both exogenous and endogenous free fatty acids) and CE are the prevailing components [5]. Chemical assays on total cell extracts demonstrated that, in spite of their di¡erent ML levels, 3T3 and 3T3ras cells are characterized by very similar TG and CE contents and only a slight, if any, di¡erence in FFA concentration. These results indicate that, besides the overall capability of the cell to synthesize and store TG and CE, other factors (including subcellular lipid compartmentalization) may modulate the NMR-visibility of ML domains in intact cells. It is also interesting to point out that, in spite of their high R values, 3T3 cells are not particularly rich in TG. The overall TG concentration of about 1 nmol/106 cells, measured in 3T3 cell extracts, is in fact similar to that found in cells constitutively possessing rather low lipid contents (e.g. liver endothelial cells), and about 50 times lower than that of cells committed to metabolize and store high amounts of lipids (e.g. parenchymal liver cells) [37]. These considerations suggest that the amount of

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lipids in the mobile fraction is only a very small percentage of the total cell lipids. While any information on the speci¢c nature and selective aggregation of lipids contributing to NMRvisible ML domains is inevitably lost in chemical analyses of total cell extracts, 1D and 2D NMR spectra provide unique, non-invasive insight on the chemical composition of these domains, as they occur in living intact cells. Spectral analyses con¢rmed the presence of TG in NMR-visible lipid domains, as indicated by the appearance in 2D spectra of 3T3 cells of a distinct glycerol backbone cross-peak G, whose intensity decreased (or even vanished) in the transformed cell variants. A substantial di¡erence between 3T3 and 3T3ras was also a signi¢cantly higher ^CHNCH^ peak area in 1D spectra of the former cells, associated in the 2D maps with higher intensities of the C and D cross peaks, respectively, due to mono- and polyunsaturated fatty chains (including linolenic and arachidonic acid). Since no signi¢cant changes were found in the average degree of unsaturation of fatty chains in the total extracts of the two cell lines, it is reasonable to conclude that the fatty acyl composition of ML domains may substantially di¡er from the rest of the cell. Another peculiar feature of ML domains in intact 3T3 cells was a speci¢c enrichment in alkyl ether chains, identi¢ed by the 2D cross-peak K. The latter, not observed in 2D COSY maps of intact 3T3ras ¢broblasts, was, however, detected in the total cells extracts of both cell lines. 4.2. Dependence of ML upon mitogenic cell stimulation and PC-plc hydrolysis ML signals in quiescent 3T3 ¢broblasts exhibited R values of about 1.5, i.e. very close to the levels measured at the late log phase. This ¢nding is in general agreement with experiments by Holmes et al. [38] on activated B- and T-lymphocytes, where the spectral lipid pro¢les remained practically unaltered upon cell block in G1 . In quiescent 3T3ras ¢broblasts, instead, ML signals fell to a very low level (RW0.2), i.e. well below the average content exhibited by these cells at the late log phase. The ML levels respectively measured in the spectra of quiescent 3T3 and 3T3ras ¢broblasts remained un-

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changed in the ¢rst 2 h of cell re-stimulation by FCS (when the SPF was maintained at about 20^25% for both cell lines); at later stages (24 h; SPF about 40% for 3T3 and 55% for 3T3ras ) R increased by about 0.3 units in 3T3 (vR/R = 0.2) and about 0.5 units in 3T3ras (vR/R = 2.5), the vR/vSPF ratio being about 0.015 in both cell lines. These results suggest that the production of ML domains may, at least partly depend, in both cell lines, upon similar biochemical mechanisms, and their modulations during the cell cycle. The hypothesis has been proposed that in activated T- and B-cells, the development of narrow NMR-detectable ML signals, occurs in G1 [35,38], a cell phase characterized by high rates of membrane phospholipid turnover and elevated levels of both phospholipid precursors and derivatives [39]. Moreover, the fact that PC-plc, similarly to a growth factor, is required through G1 for maximal mitogenic activity [40] further supports a causal relationship between PC turnover and G1 progression [41]. 1 HNMR analyses on stimulated lymphocytes exposed to D609, a speci¢c PC-plc inhibitor, allowed Veale et al. to con¢rm a relevant contribution of the PC cycle to ML formation [6,7]. NMR analyses performed in this study demonstrated that, similarly to lymphocytes [6,7], speci¢c inhibition of the endogenous PC-plc enzyme by D609 is also associated with some decrease (20 þ 10%) in the ML contents of both 3T3 and 3T3ras ¢broblasts. Furthermore, exposure to exogenous PC-plc induced R increases of about 0.7 units in both cell lines. With this increment, the lipid pro¢le of intact 3T3ras ¢broblasts became very similar to that usually exhibited by untransformed cells. These results demonstrate a substantial contribution of either endogenous or exogenous PC-plc to ML formation in both transformed and untransformed ¢broblasts, and suggest a role of DGs as possible ML constituents and/or precursors of ML components. In this respect, it is interesting to note that exposure of Rat-1 ¢broblasts to exogenous PC-plc was reported to produce in 30 min a 6-fold raise in DG levels, which remained elevated for at least 6 h, with no sign of conversion of DG into phosphatidate [42]. The hypothesis can, therefore, be suggested that a long-lasting excess of DGs might participate in the formation of mobile lipid microdomains in the plas-

ma membrane, which may then migrate into discrete pools [43]. Taken together, these results support the view that PC-plc-triggered PC cycle activation and DG production may at least partly contribute to generate mobile lipid domains in the plasma membrane of both 3T3 and 3T3ras cells. Solely on this basis, however, ras-transformed cell variants, for which constitutive activation of PC-plc [44] and higher DG levels [45] have been reported with respect to the parental cells, should be characterized by higher, rather than lower ML levels. It should, however, be noted that other phospholipases of the PC-cycle, such as plA2 and PC-pld might interfere with ML formation, especially in 3T3ras , where the activity of these enzymes is reported to be enhanced with respect to the untransformed cells [46,47]. The relative contributions given to ML production by these di¡erent pathways of PC hydrolysis can at the moment be hardly predicted. 4.3. Subcellular localization of ML domains As far as the localization of ML domains is concerned, two di¡erent models have been proposed in the literature to interpret the appearance of NMRvisible lipid signals in intact cells: (a) lipoprotein-like micro-domains with a core of neutral lipid intercalated in the plasma membrane bilayer [10,12]; and (b) lipid droplets localized in the cytoplasm [14]. Arguments in favor of the membrane micro-domain hypothesis were based upon: (a) experiments with paramagnetic probes [11]; (b) comparison of 1 H-NMR spectra of cells and their ghosts [11] ^ although it may be di¤cult to avoid contamination of plasma membrane preparations by lipid droplets [13]; and (c) lack of signi¢cant di¡erence in cytoplasmic lipid droplet content of CHO cell lines exhibiting di¡erent ML signal intensities [10]. However, it has to be said that no direct evidence has so far been provided on the presence of isotropic lipid microdomains in the membrane of cells exhibiting elevated NMR lipid signals. On the other hand, a correlation between ML signal intensity and cytoplasmic lipid bodies has been proposed on the basis of NMR studies on myeloma cells grown in the presence of oleic acid [13] and measurements of the average root mean square dis-

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placement of the lipid acyl ^(CH2 )n ^ signal at 1.29 ppm through di¡usion experiments in vivo in rat brain glioma [14]. Cytoplasmic lipid bodies, detected by TEM analysis of osmium-stained cells or by £uorescent microscopy of cells incubated with speci¢c £uorophores, were reported in a variety of cells either committed to metabolism and storage of neutral lipids [37,48^ 51] or involved in in£ammatory processes [52], as well as in malignant cells [14,15,53]. These lipid bodies, often (but not always) surrounded by a limiting membrane [14,37,48,54], are considered to be essentially made of TG and CE and may in some cells (such as macrophages) be enriched in arachidonic acid [52]. These bodies may also contain proteins and enzymes possessing esterase, peroxidase, lipase and also acyltransferase activities, which play signi¢cant roles in lipid metabolism, storage and release of cellular material to other subcellular structures (such as lysosomes). Cytochemical and ultrastructural analyses performed in this study on 3T3 ¢broblasts provide the ¢rst direct evidence on association of NMR-visible ML with the presence of both cytoplasmic lipid bodies and lipid vesicles located at the plasma membrane level. This evidence, together with the contribution given to ML formation by PC-plc, an enzyme acting at the plasma membrane level, strongly supports the possible coexistence of the two alternative models so far proposed for subcellular ML location. Our results indicate that membrane lipid vesicles and cytoplasmic lipid domains contribute to a di¡erent extent to ML signals. In fact, a di¡erence in the R value of about 1.6 units between 3T3 and 3T3ras ¢broblasts was associated with a massive accumulation of cytoplasmic lipid bodies (for a total volume of 4^8 Wm3 per cell) in the former but not in the latter cells. On the other hand, an increase in the total volume of membrane lipid vesicles of less than 0.2 Wm3 was associated with an R increase of 0.2 units in 3T3ras cells. The di¡erent contribution given by these two types of lipid domains to ML signals is likely due to their di¡erent chemical composition and/or fatty acyl chains' mobility. Regarding cytoplasmic lipid bodies, £uorescence microscopy analyses on Nile red-stained 3T3 cells (late log phase) demonstrated in fact the presence of bright yellow £uorescent lipid bodies, mostly with-

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in the cytoplasm, a small portion being located at (or near to) the plasma membrane. A similar peri-membrane distribution of £uorescent lipid bodies has been reported by Freitas et al. in Nile red-stained Yoshida hepatoma and Ehrlich carcinoma grown in the ascites form in rodents, especially when the tumor cell density in the peritoneal exudate was very high [15]. TEM analyses con¢rmed the presence of numerous electron dense lipid droplets in the cytoplasm of osmium-stained 3T3 (but not 3T3ras ) cells. The high electron density of these particles is in agreement with the substantial ML fatty chains' unsaturation level detected by 1D and 2D NMR in intact 3T3 ¢broblasts. Particularly, the presence in ML domains of polyunsaturated fatty chains (identi¢ed by speci¢c cross-peaks in 2D COSY spectra of intact 3T3 cells) seems to further substantiate the view, proposed by Dvorak et al. [52], that lipid bodies may act as speci¢c carriers of arachidonic acid. Moreover, ultrastructural analyses (Fig. 8B) gave a clear-cut demonstration that the cytoplasmic lipid bodies detected in 3T3 cells are not only surrounded by a membrane, but are also endowed of intramembrane particles. Regarding plasma membrane, freeze-fracture TEM examinations provided the ¢rst direct evidence on the association of high ML signals with the presence of amorphous lipid vesicles (about 60 nm in diameter) spanning the membrane bilayer of 3T3 cells. In transformed ¢broblasts, the number of these vesicles was instead much lower and practically vanished at R 6 0.4. Particles of such a large diameter, well exceeding the phospholipid bilayer thickness, likely involve stabilizing interactions with molecular components located in the cortical cytoplasmic region and may be related to the £uorescent spots detected at the plasma membrane level in Nile red-stained 3T3 cells (Fig. 6). These smooth lipid aggregates are morphologically well distinct from caveolae (Fig. 8). Caveolae, i.e. caveolin-coated lipid microdomains enriched in cholesterol and glycosphingolipids [55] and involved in signal transduction [56], have already been described in NIH-3T3 ¢broblasts, and their levels have been shown to be reduced by an oncogenic stimulus [34,57]. In our experiments (Figs. 8 and 9) the abundance of caveolae-like invaginations paralleled that of the smooth spherical lipid vesicles in the

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plasma membrane, suggesting that this correlation might not be coincidental. The question ¢nally arises on the mechanisms whereby the H-ras oncogene product, p21ras , interferes with the accumulation of ML domains in 3T3ras cells, in both membrane and cytoplasmic compartments. Following a previous hypothesis [17], it seems reasonable to attribute the reduced ML levels observed in this heavily transfected cell line to the extensive morphogenetic and architectural alterations induced by the mutated oncogene product, p21ras , on both cytoskeleton organization [58,59] and membrane structure [46]. In fact, formation and maintenance of cytoplasmic lipid bodies require close interactions of these aggregates with regular networks of cytoskeleton components [48^52], which are, however, strongly perturbed upon ras-induced cytoskeleton remodeling (e.g. disappearance of polymerized actin stress ¢bers, subcortical accumulation of F-actin near peripheral ru¥ing membranes, re-organization of actin-binding proteins, large reduction of vimentin ¢laments and alterations of microtubules [58,59]). Moreover, membrane ru¥ing and £uidphase pynocytosis described in transformed ¢broblasts [46] likely result into a reduced capability of the 3T3ras cell membrane to accommodate substantial amounts of the 60-nm lipid vesicles detected in untransformed ¢broblasts. 5. Conclusions This study demonstrates that: (a) both intra-membrane lipid vesicles and cytoplasmic lipid bodies (likely maintaining a mutual dynamic exchange of lipid components) may contribute, although to a different extent, to ML signals in NMR spectra of intact murine embryo-derived NIH-3T3 ¢broblasts; (b) phosphatidylcholine hydrolysis by neutral active speci¢c phospholipase C is involved in ML generation; (c) ML domains are enriched in unsaturated fatty chains; and (d) ras-transformation is associated with a reduced capability of ¢broblasts to accumulate ML domains, especially in the cytoplasmic compartment.

Acknowledgements We acknowledge partial ¢nancial support by CNR PF ACRO and from the Italian Ministry for Scienti¢c and Technological Research (MURST-Progetti di Ricerca di Interesse Nazionale, 1996 and 1997). Our thanks go to Dr. V. Bertone (Department of Animal Biology, University of Pavia) for information and collaboration concerning lipid £uorochromization; to Dr. A. Cantafora (Department of Metabolism and Pathological Biochemistry, Istituto Superiore di Sanita©, Rome) for scienti¢c discussions and the use of gas chromatography facilities in his laboratory; to Mr. M. Giannini for excellent assistance in ensuring top level performance of NMR equipment. References [1] A.H. Futerman, R. Ghidoni, G. van Meer, EMBO Workshop Report `Lipids: Regulatory Functions in Membrane Tra¤c and Cell Development', EMBO J. 17 (1998) 6772^ 6775. [2] F.A. Ho¡man, Metabolic changes in malignancy, in: L.A. Liotta (Ed.), In£uence of Tumor Development on the Host, Kluwer Academic, Dordrecht, 1989, pp. 18^27. [3] C.E. Mountford, C.L. Lena, W.B. Mackinnon, P. Russell, The use of proton MR in cancer pathology, in: G.A. Webbs (Ed.), Annual Reports on NMR Spectroscopy, Vol. 27, Academic Press, New York, 1993, pp. 173^215. [4] F. Podo, J.D. de Certaines (Eds.), Proceedings of the Special Symposium on `Lipid Metabolism and Function in Cancer. Signi¢cance of Magnetic Resonance Spectroscopy (MRS) Measurements in Relation to Biochemical Processes and Cellular Control', Anticancer Res. 16 (1996) 1305^1594. [5] C.E. Mountford, W.B. Mackinnon, P. Russell, A. Rutter, E.J. Delikatny, Human cancers detected by proton MRS and chemical shift imaging ex vivo, Anticancer Res. 16 (1996) 1521^1532. [6] M.F. Veale, A.J. Dingley, G.F. King, N.J.C. King, 1 H NMR visible neutral lipids in activated T lymphocytes: relationship to phosphatidylcholine cycle, Biochim. Biophys. Acta 1303 (1996) 215^221. [7] M.F. Veale, N.J. Roberts, G.F. King, N.J.C. King, The generation of 1 H-NMR-detectable mobile lipid in stimulated lymphocytes: relationship to cellular activation, the cell cycle, and phosphatidylcholine-speci¢c phospholipase C, Biochem. Biophys. Res. Commun. 239 (1997) 868^874. [8] G.L. May, L.C. Wright, K.T. Holmes, P.G. Williams, I.C.P. Smith, P.E. Wright, R.M. Fox, C.E. Mountford, Assignment of methylene proton resonances in NMR spectra of embryonic and transformed cells to plasma membrane triglyceride, J. Biol. Chem. 261 (1986) 3048^3053.

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