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Cell Biology International (2005) 29, 33–39 (Printed in Great Britain)
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Analysis of 1H NMR-detectable mobile lipid domains for assessment of apoptosis induced by inhibitors of DNA synthesis and replication
V.M. Mikhailenko*, A.A. Philchenkov and M.P. Zavelevich
R.E. Kavetsky Institute of Experimental Pathology, Oncology and Radiobiology, National Academy of Sciences of Ukraine, 45, Vasilkivska Street, Kyiv 03022, Ukraine

Abstract

Cell membrane rearrangements coincident with apoptosis may contribute to the increase in the ratio of methylene (CH2 at 1.3ppm) to methyl (CH3 at 0.9ppm) resonance signal intensity as observed by proton nuclear magnetic resonance (1H NMR).

We studied CH2 and CH3 resonances in cultured cell lines treated with etoposide and fludarabine or bioflavonoid quercetin. Etoposide treatment (10μM, 18h) resulted in 3.3 fold increase of the CH2/CH3 signal intensity ratio and 6.4 fold decrease in choline signal of MT4 cells. Incubation of Namalwa cells with fludarabine (3μM, 72h) increased the CH2/CH3 signal intensity ratio by 2.4 fold and choline resonance intensity was unchanged. Quercetin treatment (30μM, 1.5 month) increased CH2/CH3 ratio by 2.1 fold. Necrotic cell death upon ethanol (20%) or DMSO (30%) treatment did not change the CH2/CH3 signal intensity ratio. 1H NMR-based study of mobile lipid domains is sensitive for detection of early engagement into apoptosis.

Keywords: Apoptosis, 1H NMR, Mobile lipid domains.

*Corresponding author.

1 Introduction

Apoptotic cell death is coincident with changes in the cell membrane such as altered lipid packing of the lipid bilayer, membrane blebbing, decreased membrane microviscosity and a loss of membrane asymmetry with surface exposure of phosphatidylserine (Schlegel et al., 1993). Those changes were attributed to the increase in the ratio of the methylene (CH2) resonance (at 1.3ppm) to the methyl (CH3) resonance (at 0.9ppm) signal intensity as observed by proton nuclear magnetic resonance spectroscopy (1H NMR) (Blankenberg et al., 1996). These resonances are also associated with malignancy and detected in many cancer cells and in cancer tissue (Le Moyec et al., 1992; Barba et al., 1999; Hakumaki et al., 1999). All these findings make 1H NMR spectroscopy as a very useful tool for noninvasive assessment of cell proliferation, malignancy and apoptosis.

Cellular origin of these resonances is related to lipid turnover and cell membrane structure. NMR signals from CH2 and CH3 groups originate mainly from mobile fatty acyl chains of tissue triacylglycerides with lesser contributions from free fatty acids and cholesteryl esters. Considering physical basis of NMR, lipid resonances must arise from the isotropically tumbling molecules, with sufficient molecular mobility (Hakumaki and Kauppinen, 2000). Currently, two main models have been suggested to explain the presence of NMR-detectable lipids in cells. Mountford and Wright (1988) considered triacylglycerides in globular plasma membrane microdomains (22–28nm in diameter), similar to the serum low-density lipoproteins or malignancy-associated lipoproteins as a source for 1H NMR signals in cells. This model was supported by the data from electron microphotograph showing microdomains of 60nm in diameter within the plasma membrane of tumor cells (Ferretti et al., 1999; Rosi et al., 1999). Another possible source of lipid resonances in cells is related to intracellular lipid bodies, either adjacent to the plasma membrane or within the cytoplasm (Callies et al., 1993; Barba et al., 1999; Remy et al., 1997). A lipid body formation involves endoplasmic reticulum membrane budding, initiated by bilayer cleavage and is facilitated by biosynthetic enzymes. Lipid bodies containing neutral lipids such as triacylglycerides, cholesteryl esters, diacylglycerides or phospholipids range from 0.1 to 50μm in diameter (Murphy and Vance, 1999).

1H NMR-visible mobile lipid domains (MLD) are reported as a peculiar feature of malignant cells in vitro and in vivo (Mountford et al., 1996) and also cells undergoing apoptosis (Blankenberg et al., 1997). 1H NMR spectral changes have been shown to be associated with apoptotic cell death in a variety of cell lines. The onset of apoptosis is associated with at least a 2 fold rise in the signal intensity of the membrane lipid methylene resonance (Blankenberg et al., 1996). The signal intensity of CH3 resonance practically does not change during apoptosis and can be used as an internal standard. The ratio of CH2/CH3 signal intensity is directly proportional to the percentage of apoptotic cells in vitro. There is also a direct temporal relationship between an increase in the CH2/CH3 signal intensity ratio and the onset of apoptosis as detected by nuclear morphologic analysis, fluorescein-annexin V flow cytometry, and DNA gel electrophoresis. Thin-layer chromatography confirmed that a dynamic and/or compositional change of the plasma membrane, rather than increase in lipase activity or fatty acid production, appears to account for the increase in the CH2/CH3 signal intensity ratio during apoptosis (Blankenberg et al., 1997).

In several NMR studies the models of cell death caused in vitro by chemotherapeutic drugs varying in mechanisms of apoptosis induction have been explored (Blankenberg et al., 1996). Nevertheless, it has not yet been known whether induction of apoptosis by different stimuli results in similar alterations in cell membrane structure revealed by NMR technique and to what extent the changes in cell membranes run in parallel with other manifestations of apoptosis.

Since the majority of anticancer drugs directly or indirectly alter DNA replication and synthesis it is worthwhile assessing by NMR technique the development of apoptosis induced by chemotherapeutic drugs of various classes affecting DNA. Apoptosis induced by these drugs is mediated by Fas-independent pathways that are far from being understood (Eischen et al., 1997). DNA topoisomerase inhibitors as well as metabolic inhibitors of DNA synthesis represent two different groups of widely used potent cytotoxic drugs known as inducers of the apoptosis in cancer cells of different genesis.

The cytotoxic and apoptotic effects of DNA topoisomerase II inhibitors seem to be associated with the stabilization of DNA/DNA-topoisomerase complexes finally resulting in DNA breaks (Stanulla et al., 1997). Nucleoside analogs are structurally and metabolically related agents affecting the structural integrity of DNA after incorporation during replication or DNA excision repair synthesis. In general, the intrinsic pathway of apoptosis triggered by DNA topoisomerase II inhibitors as well as by nucleotide analogs results in mitochondrial-dependent activation of effector caspases, which leads to the characteristic apoptotic phenotype. Nevertheless, the mechanisms of apoptosis as well as its kinetics are different for the drugs of these two groups. In particular, in contrast to early induction of apoptosis by DNA topoisomerase II inhibitors, some metabolic DNA inhibitors resulted in delayed apoptosis realized not earlier than in 72h (Robertson et al., 1993).

Earlier we have studied the effects of the etoposide and fludarabine on the apoptotic cell death in human B-cell lymphoma cell line Namalwa (Philchenkov et al., 2001) which is relatively resistant to apoptosis induction partially due to the presence of antiapoptotic genes conferred by integrated EBV genome (Tarodi et al., 1994). Recently, it was shown that several bioflavonoids, in particular quercetin, possess the activity of natural DNA topoisomerase II inhibitors (Strick et al., 2000) and may induce apoptosis in malignant lymphoid cells (Liesveld et al., 2003). In present studies we have attempted to assess and compare the alterations in MLD by NMR technique in cultured human leukemia and lymphoma cell lines (MT4 and Namalwa) treated with chemotherapeutic agents – etoposide and fludarabine or bioflavonoid – quercetin.

Based on the ratio of the integrated areas of CH2 and CH3 resonance peaks the fraction of apoptotic cells after different types of treatment was estimated. The onset of apoptosis was also assessed by DNA fragmentation assay, cell morphologic analysis and flow cytometry.

2 Materials and methods

2.1 Materials

Deuterium oxide (D2O; 99.97%), trimethylsialopropionic acid (TSP), RPMI-1640 medium, fetal calf serum, Hoecht 33342, quercetin, RNAse and propidium iodide were supplied by Sigma (ALSI, Ukraine). Etoposide (Brystol-Myers Squibb SpA, Italy) and fludarabine phosphate (Schering AG, USA) were also used.

2.2 Cell culture technique

Human B-cell lymphoma line Namalwa and T-ALL line MT4 were obtained from the National Collection of Cell Lines of Institute of Experimental Pathology, Oncology and Radiobiology (Kyiv, Ukraine). The cells were maintained at 37°C in an atmosphere of 95% air and 5% CO2 as suspension cultures in medium supplemented with 10% fetal calf serum. The cultures were passaged every 3–4 days immediately upon reaching maximum cell density. The percentage of living and dead cells was determined by Trypan blue exclusion test.

MT4 cells were treated with etoposide (10μM, 18h) in log-phase of the culture growth. Namalwa cells were incubated with fludarabine phosphate (3μM, 48 or 72h; 15μM, 48h) or quercetin which was supplemented to the medium for each passage (30μM, 1.5 month). The necrotic cell death in Namalwa cells was attained by exposure to 30% DMSO or 20% ethanol for 1h.

2.3 Cell morphologic analysis

For visualization of apoptotic cells with nuclear fragmentation Hoecht staining technique was used. The cells were washed in PBS, concentrated and incubated with 1μg/ml Hoecht 33342 solution for 15min, washed several times with PBS, fixed with 4% paraformaldehyde and analysed by fluorescent microscopy.

2.4 DNA fragmentation assay

Cell DNA was isolated by modified technique (Herrmann et al., 1994). Cells (5×106) were washed in phosphate-buffered saline (PBS) and cell pellet was lysed in buffer containing 50mM Tris–HCl, pH 7.5, 20mM EDTA and 1% NP-40. Lysate was centrifuged at 1500g. Pellet was extracted with the same buffer. Upon centrifugation, combined supernatants were treated with 1% SDS, incubated for 2h with RNAse (5μg/ml) at 56°C and then for 24h with proteinase K (2.5μg/ml) at 42°C. DNA was precipitated with ethanol in the presence of ammonium acetate and dissolved in TE buffer. DNA samples were analysed by electrophoresis in 1% agarose gel.

2.5 Cell flow cytometry

Apoptotic cells were measured by flow cytometric analysis using a Becton Dickinson FACScan and CellQuest software (BD Biosciences, USA) according to the modified technique (Gougeon et al., 1996). Briefly, cell pellets were gently resuspended in hypotonic fluorochrome solution (5μg/ml propidium iodide in 0.1% sodium citrate and 0.1% Triton X-100) and incubated for 1h at 4°C and analysed to determine propidium iodide fluorescence of individual nuclei.

2.6 NMR-spectroscopy

Cells were harvested and washed with PBS then twice with PBS made with D2O to reduce protons signal from H2O. Cells (6–10×107) were suspended in a final volume of 0.6ml of PBS–D2O, transferred to a 5mm NMR tube and placed on ice until analysis. The percentage of viable cells, determined by Trypan blue exclusion test, ranged between 80% and 95%, both before and after NMR analyses. 1H NMR spectra were acquired using a 300MHz Varian Mercury 300BB NMR spectrometer (Varian, USA) at 20°C, 90° flip angle, repetition time 10s, 128 excitations, 16K data points and 5kHz spectral width. A glass capillary with 0.1% solution of TSP in D2O was used as a reference at 0.00ppm for each sample. NMR spectra were obtained using presaturation of the residual water protons in the solvent, and samples were spun at 20Hz to prevent settling of cells during the experiment. The standardized areas of the methylene and methyl protons resonances (at 1.3 and 0.9ppm, respectively) were integrated using VNMR software (Varian, USA) and expressed in relative units.

3 Results

The human leukemia MT4 cells were treated with 10μM of etoposide for 18h and apoptotic cells were simultaneously quantified by fluorescence microscopic enumeration of apoptotic cells using Hoecht 33342 staining, by flow cytometric detection of sub-G1 peak cells, and assessment of MLD by proton NMR. Identical samples were also analysed by qualitative method of DNA gel electrophoresis. The morphologic analysis of cells shows distinctive nuclear fragmentation (Fig. 1). Flow cytometry of cells stained with propidium iodide (Fig. 2) demonstrated that about 60% of cells were apoptotic at 18h. As evident in Fig. 3, presence of sub-G1 peak correlated with DNA ladder pattern on the gel. 1H NMR analysis registered 3.5 fold increase in the intensity of the methylene resonance at 1.3ppm in etoposide-treated cells in comparison with untreated MT4 cells (Fig. 4A). The ratio of CH2/CH3 signal intensity of etoposide-treated MT4 cells has increased from 1.8 to 5.9 in 18h of incubation. The choline resonance signal (at 3.2ppm) in etoposide treated cells has decreased 6.4 times as compared with untreated MT4 cells.

Fig. 1

Fluorescence microphotograph of apoptotic MT4 cells. Cells were treated with 10μM of etoposide for 18h and stained with Hoecht 33342. Nuclear fragments are intensively stained, original magnification ×900.

Fig. 2

Demonstration of apoptosis and necrosis in MT4 cells by flow cytometry. MT4 cells were treated with 10μM of etoposide for 18h (A) or with 20% ethanol for 1h (B). Sub-G1 peak is present in etoposide-treated MT4 cells, but absent in control, untreated cells (C) or ethanol-treated necrotic MT4 cells.

Fig. 3

Demonstration of apoptosis and necrosis in MT4 cells by gel electrophoresis of cellular DNA. The same samples as described in Fig. 2 were analysed by the gel electrophoresis. 1 – DNA size markers; 2 – untreated cells; 3 – etoposide-treated cells; 4 – ethanol-treated cells. DNA fragmentation is evident in line 3, but completely absent in other lines.

Fig. 4

1H NMR spectra (at 300MHz) of malignant lymphoid cells treated with different chemotherapeutic agents. (A) 1 – MT4 cells treated with etoposide (10μM, 18h); 2 – control. (B) Namalwa cells treated with Fludarabine 1 – 15μM, 48h; 2 – 3μM, 72h; 3 – 3μM, 48h; 4 – control. (C) 1 – Quercetin treated Namalwa cells (30μM, 1.5 month); 2 – control. (D) Namalwa cells treated with: 1 – ethanol (20%, 1h); 2 – DMSO (30%, 1h); 3 – control.

Cultured Namalwa cells were incubated with fludarabine phosphate either in concentration of 3μg/ml for 2 and 3 days; or in concentration of 15μg/ml for 2 days. At the time of NMR analysis most of cells were not stained with Trypan blue. The results of a 1H NMR spectroscopy are presented in Fig. 4B. The increase in methylene protons signal intensity was observed only on the third day of incubation with a fludarabine phosphate in concentration of 3μg/ml. The ratio of CH2/CH3 signal intensity for untreated Namalwa cells was 1.6 and 1.8 for the cells grown 2 days in the presence of fludarabine phosphate in concentration of 3μg/ml. On the third day of incubation the CH2/CH3 signal intensity ratio increased to 4.4. Incubation of Namalwa cells for 2 days in the presence of higher concentration of fludarabine phosphate did not cause considerable changes in the intensity of methylene protons resonance. However, CH2/CH3 signal intensity ratio was slightly elevated to the level of 2.5. The level of the choline signal intensity did not change significantly after all the tested types of fludarabine phosphate treatment.

The incubation of Namalwa cells in continuous presence of quercetin resulted in 2 fold increase of the methylene protons signal intensity and the CH2/CH3 signal intensity ratio was 3.7 (Fig. 4C). In parallel, the choline resonance signal intensity was decreased 4.1 fold as compared with untreated cells.

In order to demonstrate that changes in the mobile membrane domains registered by 1H NMR are specific for apoptosis we have studied the CH2/CH3 signal intensity ratio when Namalwa cells were treated with 20% ethanol or 30% DMSO which produced a necrotic cell death as determined by morphological criteria. The ratio of integrated areas of the CH2/CH3 resonances in the 1H NMR spectra did not change after treatment with ethanol or DMSO.

4 Discussion

The apoptosis-associated changes in the MLD of T- and B-lymphoid malignant cell lines have been studied by 1H NMR spectroscopy. Research was conducted in vitro in Namalwa B-cell line originating from Burkitt's lymphoma that is relatively resistant to apoptosis induced by chemotherapeutic agents and in the MT4 T-cell line that is more sensitive to apoptosis-inducing agents.

The cells were treated by agents with direct or indirect DNA damaging properties: the inhibitor of topoisomerase II – etoposide, fludarabine phosphate and natural modulator of apoptosis with wide range of action – flavonoid quercetin. Nonapoptotic cell death was induced by treatment with ethyl alcohol and DMSO that cause necrosis of cells. 1H NMR detection of MLD in treated cells was based on estimation of the CH2/CH3 signal intensity ratio. Cellular origin of CH2 and CH3 resonances is related to lipid turnover and cell membrane structure and originates mainly from mobile fatty acyl chains of tissue triacylglycerides (TG) with lesser contributions from free fatty acids (FFA) and cholesteryl esters (CE). Chemical assays on total cell extract demonstrated that, in spite of their different MLD levels, fibroblasts and their H-ras-transformed variant are characterized by very similar TG, CE and FFA contents (Ferretti et al., 1999). These results indicate that, besides the overall capability of the cells to synthesize and store lipids, other factors including globular plasma membrane microdomains and subcellular lipid compartmentalization may modulate the NMR detection of MLD domains in cells.

1H NMR spectral changes associated with significant rise in the methylene signal intensity have been shown associated with apoptotic cell death in a variety of cell lines. The ratio of CH2/CH3 signal intensity measured by proton NMR correlates with the fraction of apoptotic cells determined by DNA fragmentation and morphologically.

Experiments were conducted on the cultured lines of human malignant lymphoid cells, incubated in the presence of various inducers of apoptosis. Previously it was shown that CH3 signal intensity is well defined in untreated cells and arises, in part, from non-lipid CH3 protons such as from nucleic acids and other compounds. CH3 peak height did not change with the onset of apoptosis but CH2 resonance signal intensity was increased up to 6 times of CH3 signal. The ratio of the CH2/CH3 signal intensity observed by 1H NMR is directly proportional to the percentage of apoptotic cells in vitro (Blankenberg et al., 1997). The analysis of morphology of MT4 cells treated with etoposide shows the irreversible structural changes distinctive for apoptosis development. In parallel, assessment of MLD domains by 1H NMR spectroscopy shows the increased amount of methylene protons, 3.3 fold elevated value of the CH2/CH3 signal intensity ratio and decreased signal from choline protons. Based on dose–response data (not shown) such changes are corresponding to the 60%–80% of apoptotic cells, coinciding with morphological data of the late stage of apoptosis.

The fludarabine phosphate is a nucleoside analog, acting as a metabolic inhibitor of DNA synthesis, and inducing apoptosis under certain conditions. The delayed apoptosis requires up to 72h of treatment at lower dose of fludarabine. Based on 1H NMR assessment of the CH2/CH3 signal intensity ratio, presence of the 30%–40% cells undergoing apoptosis was expected, which coincides with flow cytometry estimates, namely 15.1% in 48h and 49.2% in 72h. Interestingly that treatment of Namalwa cells for 2 days with higher concentration of fludarabine phosphate did not cause considerable changes in intensity of CH2 signal. Also fludarabine treatment did not affect the signal intensity of choline protons.

The effect of bioflavonoid quercetin, as a factor, that in high concentrations is the inducer of apoptosis and natural inhibitor of DNA-topoisomerase II in malignant cells was studied in vitro in the cultured Namalwa cells at doses not resulting in the immediate apoptotic response. It was shown that permanent presence of quercetin in cultured medium slowed the rates of cells growth, morphologically showing separate apoptotic cells, on each passage the percent of the lost cells was about 15%. The 1H NMR analysis reveals the considerably elevated levels of CH2 protons resonance and the CH2/CH3 signal intensity ratio. In contrast, significant reduction of protons signal that belongs to choline was observed. Registered difference in the CH2/CH3 signal intensity ratio corresponds to 20%–30% of apoptotic cells.

The induction of necrosis in Namalwa cells resulted in acute necrotic response, based on morphological criteria, in 95% and 60% of DMSO or ethanol-treated cells, respectively.

The results of the study demonstrate that upon the different apoptotic stimuli provided by the agents directly or indirectly affecting DNA structure or DNA synthesis and replication, the patterns of alterations in NMR visualized LMD are essentially the same irrespective of the specific mechanisms of apoptosis induction and DNA damage. This holds true both for acute exposure to the toxic doses of chemotherapeutics such as etoposide or fludarabine and for chronic exposure to quercetin as apoptosis-inducing agent possessing the activity of the natural DNA topoisomerase II inhibitor. The specified NMR visualized patterns are evident prior to the apoptotic cell death both in rapid and delayed evolution of apoptotic events. The signal intensity ratio of characteristic peaks is proportional to the number of apoptotic cells detected by other techniques. It is of importance that the technique was effective in recording rather small apoptotic percentage in the case of the continuous chronic exposure to non-toxic doses of quercetin. The complete patterns of lipid domain alterations need further analysis.

Acknowledgments

We thank Dr. N. Khranovskaya for her help with the flow cytometry. This work was in part supported by grants from: U.S. Civilian Research & Development Foundation (RESC 20-7; UR2-1028-KV-03) and the Fundamental Research Foundation of Ministry of Education and Science of Ukraine (F7/551-2001).

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Received 14 July 2004/2 November 2004; accepted 11 November 2004

doi:10.1016/j.cellbi.2004.11.008
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