Kaempferol induces apoptosis in glioblastoma cells through oxidative stress

Reactive oxygen species (ROS) are released by the normal oxidative metabolism in eukaryotic cells, and cellular antioxidants such as superoxide dismutase (SOD-1) and thioredoxin (TRX-1) detoxify these species. However, ROS overproduction or a decrease in the antioxidative capacity of cells can cause oxidative stress (1). Most cancer cells are under oxidative stress associated with increased metabolic activity and production of ROS (2). Although increased ROS production plays an important role in maintaining cancer phenotype due to their stimulating effects on cell growth and proliferation (3), high levels of ROS can also cause cellular damage (4). ROS-stressing agents can effectively kill cancer cells by increasing the intracellular ROS to a toxic level (5). Human cancer cells with intrinsic oxidative stress are highly sensitive to ROS stress (6, 7) and promoting ROS generation in mitochondria can effectively kill them (4).

The anticancer activity of flavonoids, a group of polyphenolic compounds, has been well shown (8). Flavonoids exhibit anticancer activity by elevating ROS generation (9) and by inhibiting thioredoxin reductase, which is critical for maintaining the cellular redox environment (10). The flavonoid kaempferol is known to possess cancer chemopreventive properties (11, 12) and to inhibit migration and invasiveness of glioma cells (13). A heightened oxidative stress seems to be characteristic of gliomas, with elevation in ROS production increasing the resistance of cells to chemotherapy (14, 15). Because kaempferol alters ROS generation (16, 17), we investigated whether kaempferol could be used as a redox-modulating agent to induce apoptosis in glioma cells by increasing ROS accumulation while concomitantly disabling the antioxidant defense mechanism. Because the anticancer drug doxorubicin exerts its cytotoxic effect on tumor cells through generation of ROS (18), we also investigated whether kaempferol could potentiate the toxic effects of doxorubicin by elevating oxidative stress.

Glioblastoma cell lines LN229, U87MG, and T98G were obtained from the American Type Culture Collection. Cells were cultured in DMEM (Life Technologies, Inc.) supplemented with 10% fetal bovine serum, 100 units/L penicillin, and 100 μg/mL streptomycin (Life Technologies). On attaining semiconfluence, medium was depleted of fetal bovine serum and all treatments were carried out in serum-free medium. The cells were treated with 50 μmol/L kaempferol (in DMSO) for different intervals of time (24, 48, 72, and 96 h). DMSO-treated cells were used as controls. To study whether kaempferol alters the sensitivity of cells to doxorubicin, the cells were treated with 1 μmol/L doxorubicin in the presence and absence of kaempferol for 24 h. To determine the involvement of ROS in cell death, ROS-specific inhibitor N-acetylcysteine (NAC) was added to cells either alone or in combination with kaempferol. For all treatments involving NAC, 1 mmol/L NAC was administered simultaneously with kaempferol. All reagents were purchased from Sigma unless otherwise stated.

Cell viability was assessed using the [3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium, inner salt] (MTS; Promega) assay as described earlier (19). Following treatment of cells (1 × 10³) with kaempferol for different time intervals in 96-well plates, 20 μL of MTS solution were added. After 4-h incubation, the absorbance reflecting reduction of MTS by viable cells was determined at 490 nm. Values were expressed as a percentage relative to those obtained in controls.

Glioma cells (10⁴) were treated with kaempferol, doxorubicin, or NAC, either alone or in different combinations for different time intervals, in eight-well chamber slides (Nunc). Following treatment, apoptotic cells were identified using In Situ Cell Death Detection Kit, TMR red (Roche). Briefly, cells were fixed with 4% paraformaldehyde in PBS and blocked with 4% bovine serum albumin containing 0.02% Triton X-100, and then fixed cells were incubated in the terminal deoxyribonucleotidyl transferase–mediated dUTP nick end labeling (TUNEL) mix (terminal deoxynucleotidyl transferase in storage buffer and TMR red–labeled nucleotide mixture in reaction buffer) for 1 h at room temperature. The slides were mounted with Vectashield mounting medium containing 4′,6-diamidino-2-phenylindole (Vector Laboratories, Inc.). Cell death was determined by counting the number of TUNEL-positive cells (red) that colocalized with 4′,6-diamidino-2-phenylindole (blue) from multiple fields.

Intracellular ROS generation in cells treated with kaempferol or doxorubicin, or a combination of both, and in kaempferol-treated cells transfected with SOD-1 or TRX-1, or both small interfering RNA (siRNA), was assessed using 2′,7′-dichlorodihydrofluorescein diacetate (CM-H₂DCFDA; Molecular Probes) as previously described (20) with some modifications. Briefly, cells were incubated with H₂DCFDA (5 μmol/L) for 30 min at 37°C and washed twice with PBS, and then fluorescence intensity of 5 × 10⁵ cells was measured with a spectrofluorometer (excitation, 500 nm; emission, 530 nm).

LN229 and U87MG cells were transfected with SOD-1 and TRX-1 siRNA as previously described (21). Briefly, 18 h before transfection, 2 × 10⁴ cells were seeded onto 24-well plates in medium without antibiotics and transfection of siRNAs was carried out with Lipofectamine 2000 (Life Technologies-Invitrogen) as per manufacturer's instruction. All transfections were carried out with 50 nmol/L duplex siRNA. After 48 h, the viability of transfected cells was determined by MTS assay. In addition, cells were harvested 48 h following transfection and Western blot analysis was done to determine the level of SOD-1 and TRX-1 in transfected cells. Transfected cells were then either left untreated or treated with 50 μmol/L kaempferol for additional 48 h in fresh medium and cell viability was determined by MTS assay. The siRNA duplexes were synthesized by Proligo. Transfection with nonspecific siRNA against Luciferase GL2 was done as control. The sequences of siRNA used were as follows: siTRX, 5′-AUGACUGUCAGGAUGUUGCdTdT-3′ (sense) and 5′-GCAACAUCCUGACAGUCAUdCdC-3′ (antisense); siSOD-1, 5′-GGCCUGCAUGGAUUCCAUGdTdT-3′ (sense) and 5′CAUGGAAUCCAUGCAGGCCdTdT-3′ (antisense).

Protein was isolated from control and kaempferol-treated cells as previously described (22). Twenty micrograms of each sample were electrophoresed on a 6% to 10% polyacrylamide gel and Western blot analysis was done, as previously described (22), with SOD-1 and TRX-1 antibodies (Lab Frontier). Antibodies were purchased from Santa Cruz Biotechnology unless otherwise mentioned. Secondary antibodies were purchased from Vector Laboratories. After addition of chemiluminescence reagent (Amersham), the blots were exposed to ChemiGenius Bioimaging System (Syngene) for developing and images were captured using Genesnap software (Syngene). Absorbance measurements were made using Gentools software (Syngene). The blots were stripped and reprobed with anti–β-tubulin to determine equivalent loading as described (23).

Mitochondrial membrane potential (ΔΨ_m) was measured using JC-1 probe (20). To measure ΔΨ_m, control and kaempferol-treated cells were labeled for 40 min with 6.5 μmol/L JC-1 at 37°C, washed, and resuspended in medium; fluorescence was recorded at 590 and 527 nm. The ratio of the reading at 590 to 527 nm was considered as the relative ΔΨ_m value. JC-1 accumulates as J-aggregates (590 nm; red) only in metabolically active mitochondria and depolarization of mitochondrial membranes leads to JC-1 monomer formation (527 nm; green). Control and kaempferol-treated cells were also labeled for 10 min with 10 μmol/L JC-1 at 37°C and cells were examined by fluorescence microscopy (20).

Membrane potential of control and kaempferol-treated cells was assessed by flow cytometry using negatively charged DiBAC₄(3) oxonol dye (Molecular Probes; ref. 24). Briefly, 10⁶ cells/mL were equilibrated for 1 min in PBS followed by the addition of 150 nmol/L oxonol. After equilibration for 2 min, oxonol fluorescence intensity of 5 × 10⁴ cells was measured by FACSCalibur (Becton Dickinson) with acquisition of fluorescence emission at the FL1 spectrum versus SSC, as previously described (25). As a positive control for membrane depolarization, cells were incubated with 150 mmol/L KCl for 2 min at 4°C before addition of oxonol dye (24).

Membrane fluidity of glioma cells treated with kaempferol for 96 h was determined using the fluorescent probe 1,6-diphenyl-1,3,5-hexatriene (DPH) as described (26). Briefly, 2 × 10⁵ control and kaempferol-treated cells were incubated with 1 μmol/L DPH in PBS for 2 h at 37°C, following which cells were washed and resuspended in PBS, and the DPH probe bound to the cell membrane was excited at 365 nm and the emission was recorded at 430 nm by Spectrofluorometer (Varian). The fluorescence anisotropy (r) value was calculated using the equation r = [(I_|| − I_⊥) / (I_|| + 2I_⊥)], where I_|| and I_⊥ are the fluorescence intensities oriented parallel and perpendicular, respectively, to the direction of polarization of the excited light. The microviscosity parameter [(r_o / r) − 1]⁻¹ was calculated in each case knowing the maximal limiting fluorescence anisotropy r_o, which is 0.362 for DPH.

Cytometric bead array kit (CBA kit, BD Biosciences) was used to quantitatively measure cytokine and chemokine levels in the supernatant collected from control and cells treated with kaempferol in the presence or absence of NAC. Human apoptosis kit (BD Biosciences) was used to quantitatively measure the expression levels of active caspase-3, cleaved poly(ADP-ribose) polymerase (PARP), and Bcl-2 in protein lysates collected from control and kaempferol-treated cells. Using 50 μL of human inflammation or apoptosis standard and sample dilutions, the assay was done and analyzed on FACSCalibur (Becton Dickinson) as described (27).

Cells were grown in medium containing fetal bovine serum and, on reaching semiconfluence, fetal bovine serum was removed and cells were grown in serum-free medium. A plastic pipette tip was drawn across the center of the culture to produce a clean wound area 12 h after serum depletion. Cells were then left untreated or treated with kaempferol in the presence and absence of NAC for 48 and 72 h and cell movement in the wound area was examined. The migration distance between the leading edge of the migrating cells and the edge of the wound at the beginning and end of each indicated time point was taken by light microscopy.

In studies where doxorubicin accumulation was examined, control and kaempferol-treated U87MG cells (1 × 10⁶) were incubated with 1 μmol/L doxorubicin for 30 min at 37°C and 5% CO₂. Cells were subsequently harvested and washed, and intracellular doxorubicin accumulation was measured by flow cytometry at the FL2 emission spectrum (on FACSCalibur) immediately at t = 0 min for accumulation and at t = 120 min on further incubation at 37°C and 5% CO₂ to allow for efflux as described (25).

All comparisons between groups were done using two-tailed paired Student's t test. P < 0.05 was considered to be significant.

To determine whether kaempferol affects viability of glioma cells, MTS assay was done on LN229, U87MG, and T98G cells treated with kaempferol for 24, 48, 72, and 96 h. A time-dependent decrease in cell viability was observed in LN229 cells treated with kaempferol, with significant 36% and 45% reductions in viability occurring at 72 and 96 h, respectively, as compared with control (Fig. 1A). A similar trend in loss of viability was observed in U87MG and T98G on kaempferol treatment (Fig. 1A).

Because maximal cell death was observed at 96 h posttreatment with kaempferol, we carried out fluorescence-activated cell sorting (FACS) analysis to detect the levels of active caspase-3, cleaved PARP, and Bcl-2 in these cells. Expression of proapoptotic molecules active caspase-3 and cleaved PARP was elevated by 2.3- and 140-fold in kaempferol-treated U87MG as compared with control (Fig. 1B). This was accompanied by a 3.5-fold decrease in the expression of antiapoptotic protein Bcl-2 (Fig. 1B). A similar trend in the expression of active caspase-3, cleaved PARP, and Bcl-2 was observed in LN229 and T98G on kaempferol treatment (Fig. 1B).

Because ROS have been suggested as a possible mediator of apoptosis induced by kaempferol, we examined ROS production in kaempferol-treated cells using the fluorescent probe DCFDA. A gradual increase in ROS production as compared with control was observed with increasing exposure to kaempferol in U87MG, T98G, and LN229 cells. A significant 3-, 3.5-, and 4.5-fold increase in ROS production was observed in U87MG treated with kaempferol for 48, 72, and 96 h, respectively, as compared with control (Fig. 2A). A similar trend in ROS generation was also observed in T98G and LN229 (Fig. 2A), with ROS production reaching maximum at 96 h.

To determine whether increased production of ROS following kaempferol treatment was critical in inducing apoptosis, TUNEL staining was done on cells treated with ROS inhibitor NAC in the presence of kaempferol for 96 h. No significant difference in the percentage of apoptotic cell was observed between NAC-treated cells and control (Fig. 2B). However, the ∼5-fold increase in apoptotic cells observed in all three cell lines treated with kaempferol was significantly reduced to that of control values when kaempferol treatment was supplemented with NAC (Fig. 2B). The cytotoxicity of kaempferol was abolished in the presence of ROS inhibitor.

We next determined the status of SOD-1 and TRX-1, which are involved in maintaining cellular redox balance, in kaempferol-treated gliomas with elevated ROS levels. Western blot analysis revealed a decrease in SOD-1 and TRX-1 expression in gliomas with increasing exposure to kaempferol (Fig. 2C). While the expression of SOD-1 and TRX-1 in U87MG and T98G treated with kaempferol for 24 and 48 h were comparable to control, a decrease in expression was observed at 72 and 96 h (Fig. 2C). Densitometric analysis revealed an ∼2.7- and 2.0-fold decrease in TRX-1 and SOD-1 expression, respectively, in cells treated with kaempferol for 96 h as compared with control. A similar trend was observed in LN229 cells (data not shown).

Because down-regulation of SOD-1 (28) and TRX-1 (29) expression increases sensitivity of tumor cells to cytotoxic agents, we investigated whether knocking down SOD-1 and TRX-1 expression using siRNA would increase sensitivity of glioma cells to kaempferol-induced apoptosis. The expression of SOD-1 and TRX-1 was down-regulated by >70% in cells transfected with SOD-1 and TRX-1 siRNA at 48 h, as determined by Western blot analysis (Fig. 2D). The nonspecific siRNA had no effect on SOD-1 and TRX-1 levels. After 48 h of culture, cells transfected with nonspecific siRNA, SOD-1, TRX-1, or both SOD-1 and TRX-1 siRNA were treated with or without kaempferol for an additional 48 h. SOD-1 and TRX-1 siRNA–transfected cells had decreased cell viability as compared with nontransfected cells or cells transfected with nonspecific siRNA. Kaempferol treatment decreased the viability of SOD-1 and TRX-1 siRNA–transfected cells, on average, by 60% and 55%, respectively, as compared with untreated SOD-1– and TRX-1–transfected control cells (viability set at 100%; Fig. 2D). Viability was further decreased in kaempferol-treated cells transfected with a combination of both SOD-1 and TRX-1 siRNA. The decrease in viability in kaempferol-treated cells transfected with nonspecific siRNA was not significant.

In addition, SOD-1 and TRX-1 siRNA–transfected cells had higher average (∼2-fold) levels of ROS compared with untransfected control. Importantly, kaempferol treatment increased the levels of ROS further (∼3-fold) in SOD-1 and TRX-1 siRNA–transfected cells (Fig. 2D). Thus, SOD-1 and TRX-1 down-regulation augmented kaempferol-induced cytotoxicity in glioma cells by elevating ROS levels.

Because ROS generation is inversely correlated with ΔΨ_m (30), we investigated mitochondrial changes in terms of alterations in ΔΨ_m, which is indicative of the status of mitochondrial function. Measurement of JC-1 aggregate (indicative of intact mitochondria) and J-monomer (indicative of de-energized mitochondria) formation (expressed as the ratio of 590:527 fluorescence) showed a gradual but significant decrease in ΔΨ_m with progressive treatment of U87MG and T98G cells with kaempferol as compared with control (Fig. 3A). Intact mitochondrial function, as indicated by the accumulation of JC-1 orange fluorescence, was observed in control and kaempferol-treated U87MG cells at 24 h (Supplementary data).¹

Because increased ROS generation causes prolonged membrane hyperpolarization (31) and because kaempferol causes ROS associated alterations of plasma membrane (32), the effect of kaempferol on Δψ_p of glioma cells was investigated. FACS analysis using membrane potential–sensing fluorescent dye oxonol indicated a significant (∼2-fold) decrease in the percentage of depolarized cells (indicated by the decrease in relative fluorescence intensity of oxonol) in cells treated with kaempferol for 96 h, as compared with control (Fig. 3C). Addition of KCl depolarized the cells significantly, consistent with the generalized effect of KCl on Δψ_p (data not shown; ref. 24).

As ROS production affects plasma membrane fluidity (33) and because alterations in plasma membrane fluidity occur in cells exposed to various apoptotic stimuli (34), the membrane fluidity of control and kaempferol-treated glioma cells was determined using the fluorescent probe DPH. A significant 3- and 11-fold decrease in the membrane microviscosity of kaempferol-treated U87MG and LN229 cells, respectively, was observed as compared with control. Because microviscosity and fluidity are inversely correlated, these results indicated an increase in membrane fluidity of cells treated with kaempferol for 96 h (Fig. 3C).

Because proinflammatory cytokines and chemokines can influence feed forward cycle of inflammation, redox balance, and migration associated with tumor progression, we carried out cytometric bead array to investigate the cytokine/chemokine profile of glioma cells treated with kaempferol for 96 h. A general trend toward decrease in the release of proinflammatory mediators was observed in kaempferol-treated gliomas. There was a significant 10- and 2-fold decrease in interleukin (IL)-6, ∼8- and 2-fold decrease in IL-8, ∼1.7- and 2-fold decrease in monocyte chemoattractant protein-1 (MCP-1), and ∼2-fold (P < 0.05) decrease in regulated on activation, normal T-cell expressed and secreted (RANTES) expression in kaempferol-treated U87MG (Fig. 4A) and T98G (Fig. 4B) as compared with control.

To determine whether the ability of kaempferol to regulate cytokine/chemokine levels was ROS dependent, the expression of these proinflammatory mediators was determined in cells treated with kaempferol in the presence of NAC. Treatment with NAC alone had no significant effect on the release of cytokines/chemokines from glioma cells. However, treatment with a combination of NAC and kaempferol reversed the inhibitory effect of kaempferol on proinflammatory cytokine/chemokine release because the level of proinflammatory mediators released from glioma treated with this combination was almost comparable to that of untreated control (Fig. 4A and B).

Because both ROS and chemokines are crucial regulators of cancer cell migration and because ROS, IL-8, MCP-1, and RANTES levels were significantly altered in kaempferol-treated cells, we investigated the migration ability of gliomas in the absence and presence of kaempferol using wound healing assay. There was an obvious decrease in cell migration, as shown by the difference in the wound area covered in kaempferol-treated cells as compared with control. Although untreated cells migrated to cover the wound area as expected, treatment with kaempferol inhibited motility of T98G at different time points investigated (Fig. 5). The effects of kaempferol on wound healing were dependent on ROS because the ability of cells treated with a combination of NAC and kaempferol to migrate across the wound area was almost comparable to that of untreated control. A similar trend was observed in U87MG cells (data not shown).

Glioblastoma treatment is most often based on a combination of several drugs. Because doxorubicin, often used in chemotherapeutic regimen, induces apoptosis in glioblastoma (35), we evaluated whether kaempferol could enhance the cytotoxic effect of doxorubicin. Approximately 4- and 5-fold increase in apoptosis was observed in U87MG treated with kaempferol or doxorubicin, respectively, for 24 h, as compared with control. Interestingly, kaempferol potentiated the toxic effect of doxorubicin because a dramatic 18-fold increase in apoptotic cells was observed when U87MG cells were treated with doxorubicin in conjunction with kaempferol for 24 h (Fig. 6A).

To determine whether enhanced oxidative stress could have accounted for the synergistic interaction between kaempferol and doxorubicin, the level of ROS was determined in cells treated with doxorubicin or kaempferol, or a combination of both, for 24 h. Whereas a significant ∼2.0- and 3.1-fold increase in ROS was observed in U87MG cells treated with kaempferol and doxorubicin, respectively, treatment with a combination of both further elevated ROS level by 4.5-fold as compared with control (Fig. 6B).

To determine whether kaempferol potentiated the cytotoxic effect of doxorubicin by enhancing its accumulation, we carried out FACS analysis to determine the ability of U87MG to efflux doxorubicin in treated cells relative to control. In comparison with control, which exhibited 40% doxorubicin efflux over a time course of 120 min following accumulation of doxorubicin, only 8% of doxorubicin was effluxed from cells treated with kaempferol for 96 h (Fig. 6C).

The degree of oxidative stress in a cell reflects a balance between the rate of ROS production and the activity of scavenging systems that detoxify them (5). Increased basal oxidative stress in transformed cells makes them highly dependent on their antioxidant systems to counteract the damaging effect of ROS (36) and this makes them susceptible to further oxidative insults (5, 36). Diverse chemotherapeutic agents can induce apoptosis in cancer cell lines through increased ROS generation (36, 37). Kaempferol is known to induce apoptosis in numerous cancer cell lines; however, its effect on glioma cells is not well understood. Our results show that kaempferol induces apoptosis in human glioma cells through increased ROS generation.

Increased ROS accumulation in kaempferol-treated gliomas was accompanied by a decrease in the levels of redox active proteins SOD-1 and TRX-1, which are involved in maintaining cellular redox balance. SOD-1 acts as a major defense against ROS by detoxifying the superoxide anion, and knockdown of SOD-1 expression induces cell death in malignant cells (38). Increased levels of TRX-1 in a variety of malignant tumors may contribute to the resistance of cancers to therapy by scavenging ROS, which are generated by various anticancer agents (39). Not only does down-regulation of SOD-1 expression increases sensitivity of tumor cells to the cytotoxic agents (28) but the anticancer activity of some flavonoids has also been attributed to their ability to inhibit thioredoxin reductase (10). Importantly, SOD and TRX have been implicated in the pathogenesis of astrocytomas (40). Our findings that siRNA-mediated down-regulation of SOD-1 and TRX-1 in gliomas increases ROS generation and enhances sensitivity to kaempferol-induced apoptosis are consistent with previous reports that inhibition of SOD (7) and TRX (29) enhances sensitivity of tumor cells to ROS-generating anticancer agents. Weakening of the antioxidant defense mechanism may contribute to the sensitivity of glioma to ROS because reduced levels of SOD-1 and TRX-1 might not be sufficient for handling, scavenging, and detoxifying the excessive ROS generated in kaempferol-treated glioma.

Kaempferol-mediated increase in ROS generation and decrease in ΔΨ_m were concomitant with apoptosis, suggesting that severe accumulation of ROS in oncogenically transformed cells causes oxidative damage to mitochondrial membranes and impairs membrane integrity, leading to cell death (36, 37). Because membrane potential influences resistance of cancer cells to chemotherapeutics (24), it is possible that an altered Δψ_p plays a role in kaempferol-induced apoptosis. The fluidity of plasma membrane plays an important role in chemotherapy-induced cell death (41, 42) because changes in multidrug resistance are associated with alterations in the fluidity (43). An increase in membrane fluidity in kaempferol-treated cells could have increased the sensitivity of cells to doxorubicin, as previously reported (43). The inhibition of glioma cell migration by kaempferol in a ROS-dependent manner is consistent with previous reports of ROS involvement in tumor migration (44). ROS is known to down-regulate the expression of proinflammatory genes (45). Because MCP-1 increases aggressiveness of gliomas (46) and because both IL-6 (47) and IL-8 (48) contribute to proliferation and progression of glioblastoma, it is possible that kaempferol inhibits glioma cell migration by suppressing the release of proinflammatory cytokines/chemokines.

Because ROS toxicity induced by certain chemotherapeutic agents can be an effective means of eradicating malignant cells, it is useful to consider effective ways of achieving significant synergy through combination of agents with similar ability to alter redox conditions. Doxorubicin, often included in chemotherapy regimens, has several mechanisms of action; one of which is the formation of free radicals leading to oxidative stress (49). Kaempferol potentiated the toxic effect of doxorubicin by amplifying ROS generation. The ability of kaempferol to enhance intracellular accumulation of doxorubicin by reducing its efflux agrees with previous studies that flavonoids reverse multidrug resistance involved in regulating the efflux of chemotherapeutic drugs from cells (50). It is possible that kaempferol exacerbated doxorubicin-induced damage in glioma cells by coupling enhanced oxidative stress with increased accumulation of doxorubicin. Increased efficacy of kaempferol in inducing apoptosis of human glioma cells through modulation of cellular redox balance, in conjunction with other ROS-generating anticancer therapeutic modalities, suggests that a strategy combining these agents warrants further investigation in glioblastoma.

We thank Uttam Kumar Saini for excellent technical assistance, Prof. Vijayalakshmi Ravindranath (Director, National Brain Research Centre) for her continuous support, and Dr. Anirban Basu (National Brain Research Centre) for his valuable suggestions.