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. Author manuscript; available in PMC: 2012 Nov 1.
Published in final edited form as: NMR Biomed. 2011 Mar 24;24(9):1159–1168. doi: 10.1002/nbm.1674

Preclinical Study of Treatment Response in HCT-116 Cells and Xenografts with 1H-decoupled 31P MRS

Moses M Darpolor 1, Peter T Kennealey 3, H Carl Le 1, Kristen L Zakian 1, Ellen Ackerstaff 1, Asif Rizwan 1, Jin-Hong Chen 3, Elliot B Sambol 3, Gary K Schwartz 2, Samuel Singer 3, Jason A Koutcher 1,2,4
PMCID: PMC3201722  NIHMSID: NIHMS274558  PMID: 21994185

Abstract

The topoisomerase I inhibitor, irinotecan, and its active metabolite SN-38 have been shown to induce G2/M cell cycle arrest without significant cell death in human colon carcinoma cells (HCT-116). Subsequent treatment of these G2/M-arrested cells with the cyclin-dependent kinase inhibitor, flavopiridol, induced these cells to undergo apoptosis. The goal of this study was to develop a noninvasive metabolic biomarker for early tumor response and target inhibition of irinotecan followed by flavopiridol treatment in a longitudinal study. A total of eleven mice bearing HCT-116 xenografts were separated into two cohorts where one cohort was administered saline and the other treated with a sequential course of irinotecan followed by flavopiridol. Each mouse xenograft was longitudinally monitored with proton (1H)-decoupled phosphorus (31P) magnetic resonance spectroscopy (MRS) before and after treatment. A statistically significant decrease in phosphocholine (p = 0.0004) and inorganic phosphate (p = 0.0103) levels were observed in HCT-116 xenografts following treatment, which were evidenced within twenty-four hours of treatment completion. Also, a significant growth delay was found in treated xenografts. To discern the underlying mechanism for the treatment response of the xenografts, in vitro HCT-116 cell cultures were investigated with enzymatic assays, cell cycle analysis, and apoptotic assays. Flavopiridol had a direct effect on choline kinase as measured by a 67% reduction in the phosphorylation of choline to phosphocholine. Cells treated with SN-38 alone underwent 83±5% G2/M cell cycle arrest compared to untreated cells. In cells, flavopiridol alone induced 5±1% apoptosis while the sequential treatment (SN-38 then flavopiridol) resulted in 39±10% apoptosis. In vivo 1H-decoupled 31P MRS indirectly measures choline kinase activity. The decrease in phosphocholine may be a potential indicator of early tumor response to the sequential treatment of irinotecan followed by flavopiridol in noninvasive and/or longitudinal studies.

Keywords: irinotecan, flavopiridol, choline kinase, colon cancer, 1H-decoupled 31P MRS, apoptosis

INTRODUCTION

The introduction of combination treatments with novel mechanism of action has led to considerable changes in the management of colon cancer. The treatment of human colon carcinoma with irinotecan followed by flavopiridol has demonstrated favorable results in humans (1) and a xenograft model (2). The success of such treatment regimens can be critically dependent on the timing or scheduled treatments of the two drugs. As this treatment development progresses and is translated into the clinic, there is a crucial need for robust tools and advanced techniques to measure the efficacy of such treatment regimes in vivo and to continuously monitor the tumor response to treatment. Early detection of failed treatment will constitute an opportunity to readjust the treatment or advance to a therapy that will improve the outcome of patients.

Previous studies have utilized 1H magnetic resonance spectroscopy (MRS) to quantify apoptotic cell death (3) and 31P MRS to monitor the effect of a choline kinase inhibitor on cells and tumor extracts (4). Scalar or spin-spin coupling, between 1H and 31P nuclei, results in a splitting of the observed signal into characteristic multiplet pattern visible in the 31P MRS spectrum, although not resolvable in vivo. The spectral linewidth of such coupling between 1H and 31P nuclei in phosphomonoesters and phosphodiesters is on the order of 5–10 Hz (5), which is greater than the achievable linewidth of in vivo 31P MRS. Effective decoupling schemes can be used to collapse the multiplet into a singlet thereby increasing the signal-to-noise ratio (SNR) and simplifying the phosphomonoester resonance to separate phosphocholine and phosphoethanolamine peaks. Broadband multiple composite pulse decoupling sequences such as a MLEV-16 scheme (6–9) can be used to improve the spectral resolution and SNR of 31P MRS spectra acquired during in vivo spectroscopy study. For ex vivo measurements, high resolution-magic angle spinning 1H MRS (HR-MAS 1H MRS) (10) is superior to static 1H MRS because it averages out the susceptibility inhomogeneity across heterogenous tissue samples (11) and eliminates the residual dipolar interactions of neighboring protons. As a result, the sensitivity and resolution of the spectra are significantly increased over static spectra, enabling a more accurate quantitative analysis of the biochemical properties of the sample (10,11).

Several alterations in cancer phospholipid metabolism have been previously noted (12–16). Disruption of the cytidinediphosphatecholine (CDP-choline) pathway, for example through depletion of its products, has been linked to apoptosis in a variety of cell systems (17–21). Phosphatidylcholine (PTC) is the most abundant structural component of cell membranes (22–24) and de novo synthesis of PTC is required for cell growth and differentiation. In mammalian cells, synthesis of PTC is mostly governed by the CDP-choline pathway (25). Choline is initially converted to phosphocholine (PC) by choline kinase (CK), and then to CDP-choline by cholinecitidyltransferase (CT). The final step in which CDP-choline is converted to PTC by cholinephosphotransferase (CPT) occurs very rapidly and results in minimal accumulation of the intermediary product, CDP-choline (26).

Experimental studies have shown that tumors with mutation in the ras oncogene and cell lines transformed with ras have an increase in CK activity and consequently have elevated levels of PC (27,28). Inhibition of CK, the initial enzyme in CDP-choline pathway, has been shown to inhibit proliferation of transformed cells and the growth of human colon tumor xenografts (27,29). It is known that the human colon cancer cell line, HCT-116, harbors the ras mutation (30). These findings are the impetus to investigate the use of in vivo marker of tumors with ras mutation and elevated choline kinase activity. Since phosphocholine is a product of choline kinase detectable by 1H-decoupled 31P MRS, this technique can be use to investigate the potential inhibition of choline kinase by flavopiridol in vivo. Treating HCT-116 cells with irinotecan or its active metabolite SN-38 induces G2/M cell cycle arrest with minimal cell death (2,31). Apoptosis in these G2/M-arrested cells was enhanced after subsequent treatment with flavopiridol, a cyclin dependent kinase (CDK) inhibitor (2,31). However, the underlying molecular mechanism is not fully understood and has become an area of active research.

In this study, we hypothesized that the enhancement of apoptosis by flavopiridol in G2/M-arrested HCT-116 cells, harboring the ras oncogene, may be due to the disruption of the CDP-choline pathway, specifically through the inhibition of CK. It would be ideal to develop noninvasive techniques to assess alterations in tumor cell biology, including the induction of apoptosis following a specific cancer therapy. Therefore, we elected to develop and validate a MRS technique that can be used to investigate CK and CT activity through the measurement of PC/choline and PTC/PC ratios, respectively, to evaluate the inhibitory effects of the drugs that target these critical steps in the CDP-choline pathway. As such, in vivo 1H-decoupled 31P MRS and in vitro HR-MAS 1H MRS are proposed as effective techniques for monitoring changes in the major components of the CDP-choline pathway.

EXPERIMENTAL

In vivo 31P MRS of HCT-116 tumor xenografts

All procedures with animals were performed according to the Memorial Sloan-Kettering Cancer Center’s institutional guidelines for use and care of laboratory animals. Athymic nu/nu female mice (NCI, Bethesda, MD, 8–10 weeks of age) were implanted subcutaneously in the flank with 0.2 mL of HCT-116 cancer cell suspension (2.5×106 cells/mL) in a 50:50 mixture of phosphate-buffered saline and matrigel (BD Bioscience, Franklin Lakes, NJ). During tumor growth, the tumor volume was determined by measuring the three orthogonal axes (a, b, c) of each tumor with calipers and by implementing the formula: (π/6) × a×b×c (32). A tumor volume of 150 mm3 to 200 mm3 was considered appropriate for in vivo MRS experiment. Data from longitudinal tumor volume measurements were fitted to an exponential growth rate, y=y0×e(kt), where y is the tumor volume (mm3), yo is the measured volume at the start of MRS experiment, k is the rate constant, and t is the time (day). Tumor doubling time was calculated as 0.69/k. A more sophisticated model that takes into account the pharmacokinetics and pharmacodynamics of the drugs of HCT-116 xenograft after the sequential treatment of irinotecan followed by flavopiridol in mice was recently published (33).

Baseline MRS experiments were performed on each mouse prior to treatment. A cohort of mice (N =7) was treated sequentially with a combination of irinotecan (100 mg/kg, i.p.) followed by flavopiridol (3 mg/kg, i.p.) with an interval of seven hours. Longitudinal MRS of the tumor within the same mouse was performed on the 1st day, 2nd day, and 7th day after the treatment. An equal volume of saline was administered to another cohort of mice (N =4) as controls. All MRS experiments were carried out on a 7 Tesla Avance II spectrometer (Bruker BioSpin MRI GmbH, Ettlingen, Germany) for small animals and equipped with RF-channels operating at 1H/31P Larmor frequencies of 300/121 MHz, respectively. The magnet is equipped with triple-axis gradients of up to 4 mT/m with an internal diameter of approximately 12 cm. The probe was a dual uncoupled-coil design with a three-turn parallel solenoidal coil as the phosphorus coil inside of a Helmholtz coil as the proton coil, and each coil was mounted on a separate LCR circuit (Figure 1). Images of the dual 1H/31P probes are shown (Figure 1A–B) co-mounted unto a platform of a mouse stereotaxic-holder (10.5 cm in diameter) that was custom-built and is compatible with the magnet. The holder had a doubled walled glass-bowl with barbed fittings to accommodate a water heating-cooling system. Experiments were performed on mice (24±2 g) anesthetized with 2% isoflurane in oxygen at a flow rate of 1.5 L/min, and body temperature (37.2±.1°C) was regulated with the water heating-cooling system. The mouse flank tumor was positioned in the custom-built 1H/31P MR probe, secured with a suture, and placed in a water bath to reduce B○ inhomogeneity (34). After the experimental setup in the 7T magnet was completed, each probe was properly tuned and matched to the Larmor frequency (fo) of the respective nucleus (1H or 31P). The quality factor (Q) of each probe was calculate using the expression Q=fo/Δf, where Δf is the bandwidth (full width at half maximum) of the resonant LCR circuit (Figure 1C–D). Typically, the Q’s for the 1H (Figure 1C) and 31P (Figure 1D) probes in the water bath were 33±3 and 63±2, respectively.

Figure 1.

Figure 1

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Photographs of the custom-built 1H and 31P coils [A] co-mounted on a mouse stereotaxic-holder holder [B] with a close-up view of the 3-turned parallel solenoidal coil as the 31P coil inside the Helmholtz coil as the 1H coil. A typical tuned and matched frequency of the resonant LCR circuit at experimental setup for the [C] 1H coil and [D] 31P coil in water bath.

Proton-decoupled one dimensional 31P chemical shift imaging (1D 31P CSI) was performed to measure metabolite signals of the tumor tissue without contributions from the mouse body. The 1D CSI phosphorus acquisition involved a FID-type sequence with a rectangular pulse, power of 25 dB, a 45° flip angle, 64 averages, TR of 1.8 s, acquisition size of 1024 points, spectral width of 10080 Hz, phase encode gradient duration of 0.4 ms, and a slice thickness of 4 mm. During each experiment, a calibration of the flip angle was performed with constant power while modulating the pulse length over a wide range of pulse lengths from 10 µs to 310 µs to determine an accurate 45° flip angle. Optimized decoupling was performed by modulating the decoupling transmitter power with a decoupling offset of 4.8 ppm from the resonance frequency of protons in water (see Supplementary Material). A composite pulse decoupling was implemented with MLEV-16 sequence of cyclic permutations in the supercycle (6–9). A 60 µl sphere of 50 mM methylenediphosphonic acid (Sigma Chemical Co., St. Louis, MO) doped with 10 µM MAGNEVIST®, gadopentetate dimeglumine, (Bayer Healthcare Pharmaceuticals, Germany) was mounted in a fixed position for chemical shift and pulse length calibrations as well a relative quantification of in vivo 31P metabolite signals. The post-processing and quantification of the in vivo 31P CSI data was performed with the XsOsNMR software package (kindly provided by Dr. D.C. Shungu and Mrs. X. Mao, Hatch MR Research Center, Columbia University College of Physicians and Surgeons, New York, NY).

Statistical analysis was performed using GraphPad Prism version 4.0a for Mac OS X (GraphPad Software, San Diego, California USA). Paired student’s t test was performed to compare groups within an experiment. A p-value less than 0.05 was considered statistically significant.

In vitro 1D 1H MAS MR analysis of HCT-116 cells

HCT-116 cells were cultured for 48–72 h to 60% confluence (first log phase) in Roswell Park Memorial Institute medium (RPMI-1640) supplemented with 10% heat-inactivated fetal bovine serum (Invitrogen, Carlsbad, CA), 100 units/mL penicillin and 100 mg/mL streptomycin, and maintained at 37°C in a humidified incubator with 5% carbon dioxide mixed in air. All cultures tested negative for Mycoplasma species. A stock solution of 5 mM SN-38 (Pharmacia, Peapack, NJ) was prepared in DMSO. A flavopiridol (National Cancer Institute, Bethesda, MD) stock solution of 4.5 mM was prepared in water and stored at −20°C. Cell cultures were exposed to drug free medium (N=4), 20 nM of SN-38 treatment alone (N=4), 150 nM of flavopiridol treatment alone (N=4), or to 20 nM of SN-38 followed by 150 nM flavopiridol after 24 h interval (N=4). Cell cultures were trypsinized, harvested, counted with a hemacytometer, and 8×106 cells were resuspended in PBS/D2O (pH=7.4). HCT-116 cells (8×106 cells) were washed twice with PBS/D2O and then placed in a 4-mm-O.D. zirconium rotor for MR analysis. Five µL of 95 mM 3-(trimethylsilyl) propionic 2,2,3,3-d4 acid, sodium salt (TSP) (Cambridge Isotope Laboratories, Inc., Andover, MA) added to the rotor served as internal chemical shift and concentration standard. All spectra were obtained on a Bruker Avance 600 MHz at 20°C using a Bruker 4-mm high-resolution 1H/13C/15N MAS probe. A spinning rate of 5 kHz was used for all experiments. Spectra for each sample were acquired using a π/2 pulse preceded by 2 s water suppression pulse, with accumulation of 128 transients, a repetition delay of 10 seconds, and a spectral width of 7200 Hz. Data were processed with 1 Hz line broadening followed by Fourier transformation. Assignments of metabolites resonance peaks were made by comparing chemical shifts to those of standard compounds measured in PBS/D2O (pH=7.4), and were also based on previously published NMR resonances (35,36). Cellular metabolites were quantified by computing the integral of assigned resonances using Lorentzian line-shapes to fit the 1D 1H spectra. In spectral regions of overlapping resonances, deconvolution was applied using the NMR post-processing software 1D WINNMR 6.1 software (Bruker, Ettlingen, Germany). Peak integrals of metabolites were tabulated, and the ratio of PC/choline was used as an indirect measure of CK activity where as the ratio of PTC/PC was used as an indirect measure of CT activity.

Statistical analysis was performed using GraphPad Prism version 4.0a for Mac OS X (GraphPad Software, San Diego, California, USA). A two-tailed unpaired student’s t test was performed to verify the statistical significance of the means in the compared groups. A p-value less than 0.05 was considered statistically significant. All results are displayed as mean±SEM (standard error of mean) of measures in groups.

Measurement of the effect of flavopiridol on purified choline kinase [EC 2.7.1.32]

To determine if the inhibition of choline kinase (CK) by flavopiridol was due to a direct effect, purified yeast CK [EC 2.7.1.32] from Saccharomyces cerevisiae was examined by solution proton MR spectroscopy. The enzymatic reaction from choline to phosphocholine was carried out in a buffer solution containing 10 mM tris acetate, 13 mM β-mercaptoethanol, 1 mM EDTA and 15 mM magnesium chloride at pH 7.2. A total of 0.2 units/mL purified CK [EC 2.7.1.32] was used with the final concentrations of 2.87 mM choline and 0.86 mM ATP for each experiment (N=4). All reagents were obtained from Sigma Aldrich, St. Louis MO, and dissolved in 95% distilled H2O/5% D2O. Flavopiridol was diluted directly from a 4.5 mM stock solution into the solution for the enzymatic reaction to final concentrations of 0.5 µM and 1.0 µM. The final solutions were transferred to a standard 5 mm liquid NMR tube, the mixture was vortexed vigorously, and placed immediately into the MR spectrometer. Five µL of a 95 mM TSP solution was used in each experiment as an internal chemical shift and concentration standard. Data acquisition was performed on a Bruker Avance 600 MHz high-resolution MR spectrometer at 20°C using Bruker 5-mm TXI liquid probe. Water suppression was achieved with a 2 s pre-saturation pulse. Spectra were obtained every 100 s and averaged over 600 seconds (10 minutes). Spectral analysis was performed using the 1D WINNMR 6.1 software (Bruker, Ettlingen, Germany). Phosphocholine levels were determined from the 1H MR spectra, as described in the previous section, and plotted with respect to time. Linear regression analysis was performed on the data sets to compute the slope or production rate, compute the goodness-of-fit (r2), and compare the slopes or production rate of various flavopiridol treatments. Statistical analysis was performed using GraphPad Prism version 4.0a for Mac OS X (GraphPad Software, San Diego, California USA).

Cell Cycle Analysis

HCT-116 cells were cultured for 48–72 h to 60% confluence (first log phase) and exposed to drug free medium (N=3), 20 nM of SN-38 treatment alone (N=4), 150 nM of flavopiridol treatment alone (N=4), or in combinations (N=4) with 24 h interval. At the conclusion of treatment, the medium from the flasks was collected to ensure capture of floating cells, cell cultures were trypsinized, and segregated into replicates for further analysis. The cell cycle distribution was analyzed by flow cytometry as previously described (37). Briefly, cells were fixed in ice cold 70% ethanol. Following fixation, cell pellets were washed twice with PBS containing 0.05% tween-20 and then suspended in 5 µg/mL of propidium iodide (Calbiochem, San Diego, CA) containing 1 µl/mL of RNAse. Cell cycle distribution was determined following propidium iodide staining using a FACS Calibur™ flow cytometer (BD Immunocytometry Systems, San Jose, CA). The level of apoptosis was quantified by measuring the amount of subdiploid (sub-G1−Go) cells stained with propidium iodide.

Statistical analysis was performed using GraphPad Prism version 4.0a for Mac OS X (GraphPad Software, San Diego, California, USA). A two-tailed unpaired student’s t test was performed to verify the statistical significance of the means in the compared groups. A p-value less than 0.05 was considered statistically significant. All results are displayed as mean percent±SEM (standard error of mean) of measures in groups.

RESULTS

In vivo 1H-decoupled 31P spectroscopy indirectly measures choline kinase activity

Representative 31P MR spectra of one xenograft at baseline and one day after combined irinotecan followed by flavopiridol treatment are shown in Figure 2. There is notable reduction in the phosphocholine (PC) and inorganic phosphorus (Pi) resonance peaks in response to treatment. The split of the inorganic phosphorus (Pi) resonance peak may suggest changes in intracellular pH (38–40) and may reflect two distinct extracellular (Pi-left, low field) and intracellular (Pi-right, up field) compartments (38,40,41). However, we did not investigate the underlying mechanism to these pH changes. In general, the intracellular compartments of tumors exhibit a neutral or slightly alkaline pH (42). The pretreated baseline spectrum, which indicates the split Pi resonance peak is resolved into two resonances, coalesces to a single resonance peak after treatment. It is possible that in the present study, modulation of the Pi resonance peak may be due to the drug induced changes culminating into a decrease in intracellular pH levels or binding of Pi.

Figure 2.

Figure 2

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Representation of in vivo 1H-decoupled 31P MR spectra of HCT-116 xenograft obtained longitudinally from [A] prior to treatment and [B] one day after sequential treatment of irinotecan (100 mg/kg i.p.) followed by flavopiridol (3 mg/kg i.p.). Spectral peaks are labeled as: 1, methylenediphophonic acid (MDP, 50 mM standard); 2, phosphoethanolamine (PE); 3, phosphocholine (PC); 4, inorganic phosphate (Pi); 5, phosphocreatine (PCr); 6, gamma-nucleoside triphosphate (γ-NTP); 7, alpha-nucleoside triphosphate (α-NTP); 8, nicotinamide adenine dinucleotide (NAD/NADH); 9, beta-nucleoside triphosphate (β–NTP), respectively. Also shown for each spectrum, are the spectra fits, the signals fitted (components), and residual spectrum after fitting are shown below each spectrum. Sequential treatment demonstrates a significant reduction in PC and Pi by the 1st day after treatment as compared to baseline PC.

No significant change in phosphoethanolamine levels was found in either the combined treatment cohort (0.90±0.09, N=7, p=0.33) or the saline-treated cohort (1.07±0.24, N=4, p=0.80) as compared to unit baseline levels (Figure 3A). A statistically significant drop in phosphocholine (0.49±0.07, N=7, p=0.0004) and inorganic phosphate (0.65±0.10, N=7, p=0.0103) levels were observed in HCT-116 xenografts one day after combined irinotecan followed by flavopiridol treatment as compared to unit baseline levels (Figure 3B–C). No significant difference was found in phosphocholine (0.84±0.07, N=4, p=0.11) and inorganic phosphate (1.52±0.32, N=4, p=0.20) levels for saline treatment as compared to unit baseline levels (Figure 3B–C). The tumor growth rates of the two cohorts are shown in Figure 3D along with the 95% confidence interval band for the exponential fit with nonlinear regression. Tumor doubling time in mice treated with irinotecan followed by flavopiridol was significantly longer (54.81±0.01 days, r2=0.42) than in the saline-treated cohort (6.86±0.02 days, r2=0.91). A more sophisticated model that takes into account the pharmacokinetics and pharmacodynamics of both drugs (33) also revealed a substantial growth delay of HCT-116 xenograft after the sequential treatment of irinotecan followed by flavopiridol in mice.

Figure 3.

Figure 3

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Longitudinal results of in vivo 1H-decoupled 31P MR spectroscopy of HCT-116 xenografts: [A] phosphoethanolamine, [B] phosphocholine, [C] inorganic phosphate, and [D] tumor growth rate. Data points are displayed as mean±SEM, and the p-values were evaluated by paired t test (nsp-value>0.05, *p-value≤0.05, ** p-value≤0.01, *** p-value≤0.001).

Indirect choline kinase (CK) and cholinecitidyltransferase (CT) activities measured with 1D 1H MAS MR analysis

Representative HR-MAS 1H MR spectra of HCT-116 cells are shown in Figure 4, with an insert of a zoomed region from 2.5 ppm to 3.5 ppm (Figure 4E). The ratio of PC/choline was used as an indirect measure of CK activity in HCT-116 cells (Figure 5). The intervals between the combined treatments of SN-38 followed by flavopiridol showed significantly smaller PC/choline ratio at intervals of 18 h (14.00±0.94, N=4, p=0.0171) and 24 h (7.50±1.50, N=4, p=0.0076) as compared to the group without a time interval (43.90±9.10, N=4) in the combined treatment (Figure 5A). In another set of HCT-116 cells experiments (Figure 5B), the combined SN-38 followed by flavopiridol treatment with 24 h interval (9.23±1.54, N=4, p=0.0022) or flavopiridol treatment alone (16.54±2.69, N=4, p=0.0344) significantly reduced PC/choline ratio as compared to drug free medium exposure (28.46±3.45, N=4). The PC/choline ratio of SN-38 treatment alone (41.54±5.77, N=4, p=0.0997) showed no significant difference than in drug free medium exposure (28.46±3.45, N=4). The treatment with flavopiridol alone (16.54±2.69, N=4, p=0.0077) or in combination with SN-38 (9.23±1.54, N=4, p=0.0016) significantly reduced PC/choline ratio as compared to SN-38 treatment alone (41.54±5.77, N=4). There was no significant difference between flavopiridol treatment alone (16.54±2.69, N=4, p=0.0564) as compared to combined SN-38 followed by flavopiridol treatment with 24 h interval (9.23±1.54, N=4). The results herein may suggest that a flavopiridol treatment alone or in combination with SN-38 significantly reduces CK activity, and that a time interval between administration of SN-38 and favopiridol may be necessary to potentiate its effect on CK activity.

Figure 4.

Figure 4

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Representative high-resolution MAS 1H MR spectra of HCT-116 cells exposed to 20 nM SN-38 for 24 hours and/or 150 nM flavopiridol for 24 hours where ND denote serum-free media and the numeric subscripts designate hours of treatment schedules: [A] drug free medium (ND24→ND24); [B] SN-38 treatment alone (SN-3824→ND24); [C] flavopiridol treatment alone (ND24→FL24); [D] combined SN-38 followed by flavopiridol (SN-3824→FL24); [E] a zoomed region from 2.6 to 3.5 ppm.

Figure 5.

Figure 5

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Choline kinase (CK) activity indirectly determined in vitro by the MR measurement of phosphocholine to choline ratio in HCT-116 cells. [A] HCT-116 cells were exposed to 20 nM SN-38 for 24 hours followed by 150 nM flavopiridol at various time intervals as indicated (N=4, mean±SEM, relative units). [B] HCT-116 cells were exposed to drug free medium (ND24→ND24), SN-38 treatment alone (SN-3824→ND24), flavopiridol treatment alone (ND24→FL24) or combined SN-38 followed by flavopiridol (SN-3824→FL24). All bar graphs are displayed as mean±SEM, and the p-values were evaluated by two-tailed unpaired t test (nsp-value>0.05, *p-value≤0.05, ** p-value≤0.01, *** p-value≤0.001).

The ratio of PTC/PC was used as an indirect measure of CT activity in HCT-116 cells (Figure 6). The results indicate that there is no significant difference in CT activity for flavopiridol treatment alone (1.72±0.24, N=6, p=0.6057) or SN-38 treatment alone (1.14±0.18, N=6, p=0.2064) as compared to drug free medium exposure (1.54±0.23, N=6). Also, there is no significant difference in PTC/PC ratio between drug free medium exposure (1.54±0.23, N=6) and the combined treatment of SN-38 followed by flavopiridol (2.56±0.40, N=6, p=0.0524).

Figure 6.

Figure 6

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Cholinecitidyltransferase (CT) activity indirectly determined in vitro by the MR measurement of phosphatidylcholine to phosphocholine ratio (PTC/PC) in HCT-116 cells exposed to drug free medium (ND24→ND24), SN-38 treatment alone (SN-3824→ND24), flavopiridol treatment alone (ND24→FL24), or SN-38 followed by flavopiridol (SN-3824→ FL24). Box-and-Whiskers graph: the box extends from the 25th percentile to the 75th percentile with a line at the 50th percentile (the median), and the whiskers extend above and below the box to show the highest and lowest values, respectively. The PTC/PC ratio did not differ significantly between treatment groups, and the p-values were evaluated by two-tailed unpaired t test (nsp-value>0.05).

Flavopiridol directly inhibits purified choline kinase [EC 2.7.1.32]

Phosphocholine levels were plotted with respect to time for no flavopiridol, 0.5 µM flavopiridol, and 1.0 µM flavopiridol treatments (Figure 7). The production rate or slope of phosphocholine in response to 1.0 µM flavopiridol treatment (1.46±0.15×10−3 rel.u./min; r2=0.90) was significantly lower than in response to 0.5 µM flavopiridol treatment (3.25±0.22×10−3 rel.u./min; r2=0.94, p<0.0001), and in the untreated control (9.33±0.51×10−3 rel.u./min; r2=0.97, p<0.0001). In addition, the runs test was used to determine if each data set fits a straight line (significantly nonlinear). If the overall production rates or slopes were identical, there is less than a 0.01% chance of randomly choosing data points with slopes this different. Henceforth, we can conclude that the differences between the slopes are extremely significant. This finding suggests that flavopiridol directly inhibits choline kinase activity, and it is known (43) that flavopiridol competitively inhibits CDKs by directly binding to their ATP binding site.

Figure 7.

Figure 7

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Indirect MR measurement of purified choline kinase [EC 2.1.7.32] activity from Saccharomyces cerevisiae. Phosphocholine levels were measured from 1H MR spectra and plotted with respect to time. Linear regression analysis was performed on the data sets to compute the slope or production rate, the goodness-of-fit (r2), and compare the slopes or production rate of various flavopiridol treatments.

Subsequent SN-38 and flavopiridol treatment induces apoptosis

The cell cycle distribution and apoptotic fraction for HCT-116 cells culture exposed to SN-38 alone ([T2], N=4), flavopiridol alone ([T3], (N=4), or the combination of SN-38 followed by flavopiridol ([T4], N=4) are presented in Figure 8. The percent of apoptotic cells from combined SN-38 followed by flavopiridol treatment (39±10%, N=4) was significantly higher than for SN-38 treatment alone (1.00±0.01%, N=4, p=0.009) or flavopiridol treatment alone (5±1%, N=4, p=0.015). The fraction of G2/M arrested cells for SN-38 treatment alone (83±5%, N=4) was significantly higher than for flavopiridol treatment alone (25±1%, N=4, p<0.0001), but not significantly different when compared to combined SN-38 followed by flavopiridol treatment (76±5%, N=4, p=0.360). The apoptotic and G2/M fractions of cells exposed to drug free medium were 0.85±0.15 (N=3) and 19±2 (N=3), respectively. These results indicate that SN-38 facilitates G2/M cell arrest, and follow-up treatment with flavopiridol induces these G2/M-arrested cells to synergistically undergo apoptosis.

Figure 8.

Figure 8

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Cell cycle distribution and percent apoptosis of HCT-116 treated cells with different treatment schedules as drug free medium (ND24→ND24 [T1]), 20 nM SN-38 alone (SN-3824→ND24 [T2]), 150 nM flavopiridol alone (ND24→FL24 [T3]), or SN-38 followed by flavopiridol (SN-3824→FL24 [T4]) as determined by flow cytometry. ND represents serum-free media and subscripts designate hours of treatment. All bar graphs are displayed as mean percent±SEM (N=3–4), and the p-values were evaluated by two-tailed unpaired t test.

DISCUSSION

The results of this study suggest the techniques of 1H-decoupled 31P and HR-MAS 1H MRS can be used to investigate the mechanisms of action of agents that target the key enzymes in the CDP-choline pathway and for assessing apoptosis. A statistically significant decrease in phosphocholine and inorganic phosphate levels were observed in HCT-116 xenografts following combined treatment, which were evidenced within twenty-four hours of treatment completion. A previous report has shown statistically significant growth delays when p21-intact HCT-116 xenografts were treated sequentially with irinotecan and flavopiridol (2). Consistent with this result, we found that a single treatment with irinotecan followed by flavopiridol reduced the doubling time of tumors within a week after treatment. We performed a pilot study (see Supplementary Material) on mice bearing HCT-116 xenograft in cohorts treated with irinotecan alone (N=3), flavopiridol alone (N=3), and in combination as irinotecan followed by flavopiridol (N=3). Our assessment assured no considerable difference was depicted in the baseline 31P MRS spectra compared with that obtained 24 hours after independent treatment with either irinotecan alone or flavopiridol alone. Our pilot data is congruent with previous published work, which showed that flavopiridol treatment alone had no significant effect on HCT-116 xenografts growth (2). Also, a phase II trial of flavopiridol alone in patients with advanced colorectal cancer did not show significant effect (44). Thus, we focused on the combined treatment for the in vivo studies, and we performed the independent and combined treatment regimens in vitro to elucidate the mechanism of such treatment on choline metabolism. Ultimately, our objective was to investigate a detectable in vivo marker for early treatment response of irinotecan followed by flavopiridol, which is the preferred treatment in the clinic rather than either drug alone (1). The 7 h interval in our preclinical study for this sequential treatment of irinotecan followed by flavopiridol was empirically selected based on experiments from our previous study (2). In vitro HCT-116 cell experiments have shown significant treatment effect at longer intervals (18–24 h) between the sequential administration of the two drugs than in the in vivo studies (2,31). Taking into account the pharmacokinetics and pharmacodynamics of the drugs in vivo, a recent study has modeled an optimal period of 8–10 h between doses, which compares favorably with the 7 h interval used in our study (33).

To discern the underlying mechanism for this effect in the xenografts, in vitro HCT-116 cell cultures were investigated with enzymatic assays, cell cycle analysis, and apoptotic assays. The ratio of PC/choline was used as an indirect measure of choline kinase (CK) activity whereas the ratio of PTC/PC was used as an indirect measure of cholinecitidyltransferase (CT) activity. Our study suggests that flavopiridol alone or preceded by treatment with SN-38 or irinotecan significantly reduces CK activity, and that the time interval between the administration of the two drugs plays a pivotal role in potentiating the treatment response of CK activity. On the other hand, there was no significant difference in CT activity for flavopiridol treatment alone or SN-38 treatment alone when compared to untreated controls. The significantly reduced production rate of phosphocholine by purified CK [EC 2.7.1.32] in response to flavopiridol treatment suggests that flavopiridol directly inhibits CK activity, and it is known that flavopiridol competitively inhibits CDKs by directly binding to their ATP binding site (43,45). In addition, the fraction of G2/M arrested cells was significantly higher in the cohort treated with SN-38 alone than in other cohorts. The percent of apoptotic cells was significantly higher in the cohort treated with SN-38 followed by flavopiridol than in other cohorts. These results indicate that SN-38 facilitates G2/M cell arrest in HCT-116 cells, and that follow-up treatment with flavopiridol induces these G2/M-arrested cells to undergo apoptosis, which is in congruence with precedent reports (2,31).

The topoisomerase I inhibitor, irinotecan, and its active metabolite SN-38 have been shown to induce G2/M cell cycle arrest without significant cell death in human colon carcinoma cells (HCT-116) (2,31). Subsequent treatment of these G2/M-arrested cells with flavopiridol induced these cells to undergo apoptosis. Flavopiridol inhibits a wide range of phosphokinases including the cyclin-dependent kinases as well as tyrosine and serine kinases (45). The potent inhibitory effects of flavopiridol are mediated largely through the inhibition of CDKs in cycling cells (45) although flavopiridol is also capable of inducing cell death in non-cycling cells (46). Furthermore, flavopiridol is known to transcriptionally down regulate many anti-apoptotic proteins (47,48). Flavopiridol is able to induce apoptosis in a wide variety of tumor cells in vitro and in vivo, however the percentage of cells undergoing apoptosis varies considerably, and in some systems, may be independent of cyclin-dependent kinase activity (45). These findings may suggest that there are additional mechanisms of action. Our experiments identified CK to be an additional targeted kinase among the many phosphokinases inhibited by flavopiridol. Flavopiridol has been shown to inhibit CDKs by directly binding to the ATP-binding site (43) which lead us to hypothesize that flavopiridol may inhibit CK in a similar manner. In support of this observation, high-resolution proton MR methods were used to quantify PC synthesis rates in a purified solution of yeast CK, which contained constant concentrations of CK and ATP. The addition of 0.5 µM flavopiridol results in a 65% decrease in the PC synthesis rate. Increasing the flavopiridol concentration to 1.0 µM resulted in an 84% decrease in the PC conversion rate. This suggests that flavopiridol is directly inhibiting CK through a mechanism similar to the inhibition of CDKs. The IC50 of less than 0.5 µM for flavopiridol in isolated yeast CK is well within the 500 nM ranges that are readily achievable in humans and is similar to the IC50 found for flavopiridol for CDKs and in fact much lower than the IC50 for EGF receptor kinases, pp60 Src, PKC and Erk-1 kinases (45).

Collectively, this study demonstrates the usefulness of in vivo 1H-decoupled 31P MRS and HR-MAS 1H MRS to evaluate drug-induced metabolic changes in the CDP-choline pathway, which may serve as surrogate marker of tumor cell apoptosis, and tumor regression. This study suggests that flavopiridol potentiates apoptosis in HCT-116 cells by potentially inhibiting CK as one possible mechanism. In vivo 1H-decoupled 31P MRS indirectly measures changes in choline kinase activity and the observed decrease in phosphocholine may be a potential indicator of early tumor response to the sequential treatment of irinotecan followed by flavopiridol in a noninvasive and/or longitudinal study. In light of these results, plans are underway to test whether these MRS changes can be observed clinically in patients treated with sequential irinotecan and flavopiridol therapy.

Supplementary Material

Supplementary Material

ACKNOWLEDGEMENTS

The authors express thanks and appreciation to Ms. Cornelia Matei and the late Dr. Mihai Coman for their assistance with the use and care of animals during experimental procedures. Also, the authors appreciate the time and scientific contributions from Dr. Monica Motwani, Dr. Murray F. Brennan, Dr. Douglas A. Levine, Dr. Rachael O’Connor, and Dr. Penelope DeCarolis. Human colon carcinoma cells, HCT-116, were a gift from Dr. Vogelstein (John Hopkins University, Baltimore, MD).

Financial Support: This work was supported by the NIH/NCI grants R01-CA067819, P50-CA086438, P30-CA008748, R24-CA83084 and the Kirsten Ann Carr Fund.

List of Abbreviations

1H MRS

proton magnetic resonance spectroscopy

31P MRS

phosphorus magnetic resonance spectroscopy

SNR

signal-to-noise ratio

MLEV

Malcolm LEVitt sequences

CDP-choline

cytidinediphosphatecholine

PTC

phosphatidylcholine

PC

phosphocholine

CK

choline kinase

CT

cholinecitidyltransferase

CPT

cholinephosphotransferase

HCT-116

human colon cancer cells

SN-38

active metabolite of irinotecan

NMR

nuclear magnetic resonance

RF

radiofrequency

LCR

inductance capacitance resistance

1D CSI

one dimensional chemical shift imaging

FID

free induction decay

RPMI

Roswell Park Memorial Institute

DMSO

dimethyl sulfoxide

TSP

3-(trimethylsilyl) propionic 2,2,3,3-d4 acid sodium salt

EDTA

ethylenediaminetetraacetic acid

EC

enzyme commission

PBS

phosphate buffered saline

FACS

fluorescence activated cell sorting

DAG

diacylglycerol

EGF

epidermal growth factor

PKC

protein kinase C

SEM

standard error of mean

REFERENCE

  • 1.Shah MA, Kortmansky J, Motwani M, Drobnjak M, Gonen M, Yi S, Weyerbacher A, Cordon-Cardo C, Lefkowitz R, Brenner B, O'Reilly E, Saltz L, Tong W, Kelsen DP, Schwartz GK. A Phase I Clinical Trial of the Sequential Combination of Irinotecan Followed by Flavopirodol. Clinical Cancer Research. 2005;11(10):3836–3845. doi: 10.1158/1078-0432.CCR-04-2651. [DOI] [PubMed] [Google Scholar]
  • 2.Motwani M, Jung C, Sirotnak FM, She Y, Shah MA, Gonen M, Schwartz GK. Augmentation of Apoptosis and tumor regression by flavopiridol in the presence of CPT-11 in Hct116 colon cancer monolayers and xenografts. Clin Cancer Res. 2001;7(12):4209–4219. [PubMed] [Google Scholar]
  • 3.Blankenberg FG, Katsikis PD, Storrs RW, Beaulieu C, Spielman D, Chen JY, Naumovski L, Tait JF. Quantitative analysis of apoptotic cell death using proton nuclear magnetic resonance spectroscopy. Blood. 1997;89(10):3778–3786. [PubMed] [Google Scholar]
  • 4.Al-Saffar NMS, Troy H, Ramirez de Molina A, Jackson LE, Basetti Madhu, Griffiths JR, Leach MO, Workman P, Lacal JC, Judson IR, Chung Y-L. Noninvasive magnetic resonance spectroscopic pharmacodynamic markers of the choline kinase inhibitor MN58b in human carcinoma models. Cancer Research. 2006;66(1):427–434. doi: 10.1158/0008-5472.CAN-05-1338. [DOI] [PubMed] [Google Scholar]
  • 5.Luyten PR, Bruntink G, Sloff FM, Vermeulen JWAH, Van Der Heijden JI, Den Hollander JA, Heerschap A. Broadband proton decoupling in human 31P NMR spectroscopy. NMR in Biomedicine. 1989;1(4):117–183. doi: 10.1002/nbm.1940010405. [DOI] [PubMed] [Google Scholar]
  • 6.Levitt MH, Freeman R. Composite Pulse Decoupling. J Magn Reson. 1981;43:65–80. [Google Scholar]
  • 7.Levitt MH, Freeman R, Frenkiel T. Broadband Heteronuclear Decoupling. J Magn Reson. 1982;47:328–330. [Google Scholar]
  • 8.Levitt MH, Freeman R, Frenkiel T. Supercycles For Broadband Heteronuclear Decoupling. J Magn Reson. 1982;50:157–160. [Google Scholar]
  • 9.Shaka AJ, Keeler J, Freeman R. Evaluation of a new broadband decoupling sequence: WALTZ-16. Journal of Magnetic Resonance. 1983;53(2):313–340. [Google Scholar]
  • 10.Andrew ER, Bradbury A, Eades RG. Removal of dipolar broadening of nuclear magnetic resonance spectra of solids by specimen rotation. Nature. 1959;183:1802–1803. [Google Scholar]
  • 11.Cheng LL, Ma MJ, Becerra L, Ptak T, Tracey I, Lackner A, Gonzalez RG. Quantitative neuropathology by high resolution magic angle spinning proton magnetic resonance spectroscopy. Proc Natl Acad Sci USA. 1997;94(12):6408–6413. doi: 10.1073/pnas.94.12.6408. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Pelech SL, Vance DE. Regulation of phosphatidylcholine biosynthesis. Biochim BiophysActa. 1984;779(2):217–251. doi: 10.1016/0304-4157(84)90010-8. [DOI] [PubMed] [Google Scholar]
  • 13.Podo F. Tumour phospholipid metabolism. NMR Biomed. 1999;12(7):413–439. doi: 10.1002/(sici)1099-1492(199911)12:7<413::aid-nbm587>3.0.co;2-u. [DOI] [PubMed] [Google Scholar]
  • 14.Ackerstaff E, Glunde K, Bhujwalla ZM. Choline phospholipid metabolism: a target in cancer cells? J Cell Biochem. 2003;19(3):525–533. doi: 10.1002/jcb.10659. [DOI] [PubMed] [Google Scholar]
  • 15.Glunde K, Serkova NJ. Therapeutic targets and biomarkers identified in cancer choline phospholipid metabolism. Pharmacogenomics. 2006;7(7):1109–1123. doi: 10.2217/14622416.7.7.1109. [DOI] [PubMed] [Google Scholar]
  • 16.Plathow C, Weber WA. Tumor Cell Metabolism Imaging. Journal of Nuclear Medicine. 2008;49(6):43S–63S. doi: 10.2967/jnumed.107.045930. [DOI] [PubMed] [Google Scholar]
  • 17.Albright CD, Liu R, Bethea TC, Da Costa K-A, Salganik RI, Zeisel SH. Choline deficiency induces apoptosis in SV40-immortalized CWSV-1 rat hepatocytes in culture. FASEB J. 1996;10(4):510–516. doi: 10.1096/fasebj.10.4.8647350. [DOI] [PubMed] [Google Scholar]
  • 18.Cui Z, Houweling M. Phosphatidylcholine and cell death. Biochim Biophys Acta. 2002;1585(2–3):87–96. doi: 10.1016/s1388-1981(02)00328-1. [DOI] [PubMed] [Google Scholar]
  • 19.Cui Z, Houweling M, Chen MH, Record M, Chap H, Vance DE, Terce F. A genetic defect in phosphatidylcholine biosynthesis triggers apoptosis in Chinese hamster ovary cells. J Biol Chem. 1996;271(25):14668–14671. doi: 10.1074/jbc.271.25.14668. [DOI] [PubMed] [Google Scholar]
  • 20.Yen C-LE, Mar M-H, Meeker RB, Fernandes A, Zeisel SH. Choline defiency induces apoptosis in primary cultures of fetal neurons. FASEB J. 2001;15(10):1704–1710. doi: 10.1096/fj.00-0800com. [DOI] [PubMed] [Google Scholar]
  • 21.Yen C-LE, Mar M-H, Zeisel SH. Choline deficiency-induced apoptosis in PC12 cells is associated with diminished membrane phosphatidylcholine and sphingomyelin, accumulation of ceramide and diacylglycerol, and activation of caspase. FASEB J. 1999;13(1):135–142. [PubMed] [Google Scholar]
  • 22.Kent C, Carman GM. Interactions among pathways for phosphatidylcholine metabolism, CTP synthesis and secretion through the Golgi apparatus. Trends Biochem Sci. 1999;24(4):146–150. doi: 10.1016/s0968-0004(99)01365-1. [DOI] [PubMed] [Google Scholar]
  • 23.Carman GM, Henry SA. Phospholipid biosynthesis in the yeast Saccharomyces cerevisiae and interrelationship with other metabolic processes. Prog Lipid Res. 1999;38(5–6):361–399. doi: 10.1016/s0163-7827(99)00010-7. [DOI] [PubMed] [Google Scholar]
  • 24.Yu Y, Sreenivas A, Ostrander DB, Carman GM. Phosphorylation of Saccharomyces cerevisiae choline kinase on Ser30 and Ser85 by protein kinase A regulates phosphatidylcholine synthesis by the CDP-choline pathway. J Biol Chem. 2002;277(38):34978–34986. doi: 10.1074/jbc.M205316200. [DOI] [PubMed] [Google Scholar]
  • 25.Kent C. Eukaryotic phospholipid biosynthesis. Annu Rev Biochem. 1995;64:315–343. doi: 10.1146/annurev.bi.64.070195.001531. [DOI] [PubMed] [Google Scholar]
  • 26.McMaster CR, Bell RM. Phosphatidylcholine biosynthesis via the CDP-choline pathway in Saccharomyces cerevisiae. Multiple mechanisms of regulaion. J Biol Chem. 1994;269(20):14776–14783. [PubMed] [Google Scholar]
  • 27.Hernandez-Alcoceba R, Saniger L, Campos J, Nunez MC, Khaless F, Gallo MA, Espinosa A, Lacal JC. Choline kinase inhibitors as a novel approach for antiproliferative drug design. Oncogene. 1997;15(19):2289–2301. doi: 10.1038/sj.onc.1201414. [DOI] [PubMed] [Google Scholar]
  • 28.Ramirez de Molina A, Penalva V, Lucas L, Lacal JC. Regulation of choline kinase activity by Ras proteins involves Ral-GDS and PI3K. Oncogene. 2002;21(6):937–946. doi: 10.1038/sj.onc.1205144. [DOI] [PubMed] [Google Scholar]
  • 29.Hernandez-Alcoceba R, Fernandez F, Lacal JC. In vivo antitumor activity of choline kinase inhibitors: a novel target for anticancer drug discovery. Cancer Research. 1999;59(13):3112–3118. [PubMed] [Google Scholar]
  • 30.Okada F, Rak JW, St. Croix B, Lieubeau B, Kaya M, Roncari L, Shirasawa S, Sasazuki T, Kerbel RS. Impact of oncogenes in tumor angiogenesis: mutant K-ras up-regulation of vascular endothelial growth factor/vascular permeability factor is necessary, but not sufficient for tumorigenicity of human colorectal carcinoma. Proc Natl Acad Sci USA. 1998;95(7):3609–3614. doi: 10.1073/pnas.95.7.3609. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Ambrosini G, Seelman SL, Qin L-X, Schwartz GK. The cyclin-dependent kinase inhibitor flavopiridol potentiates the effects of topoisomerase I poisons by suppressing Rad51 expression in a p53-dependent manner. Cancer Res. 2008;68(7):2312–2320. doi: 10.1158/0008-5472.CAN-07-2395. [DOI] [PubMed] [Google Scholar]
  • 32.Euhus DM, Hudd C, LaRegina MC, Johnson FE. Tumor Measurement in the Nude Mouse. Journal of Surgical Oncology. 1986;31(4):229–234. doi: 10.1002/jso.2930310402. [DOI] [PubMed] [Google Scholar]
  • 33.Goteti K, Garner CE, Utley L, Dai J, Ashwell S, Moustakas DT, Gonen M, Schwartz GK, Kern SE, Zabludoff S, Brassil PJ. Preclinical pharmacokinetic/pharmacodynamic models to predict synergistic effects of co-administered anti-cancer agents. Cancer Chemother Pharmacol. 2010;66(2):245–254. doi: 10.1007/s00280-009-1153-z. [DOI] [PubMed] [Google Scholar]
  • 34.Ballon D, Mahmood U, Jakubowski A, Koutcher JA. Resolution Enhanced NMR Spectroscopy in Biolgical Systems via Magnetic Susceptibility Matched Sample Immersion Chambers. Magn Reson Med. 1993;30(6):754–758. doi: 10.1002/mrm.1910300615. [DOI] [PubMed] [Google Scholar]
  • 35.Chen J-H, Enloe BM, Weybright P, Campbell N, Dorfman D, Fletcher CD, Cory DG, Singer S. Biochemical Corrfelates of Thiazolidinedione-Induced Adipocyte Differentiation by High-Resolution Magic Angle Spinning NMR Spectroscopy. Magn Reson Med. 2002;48(4):602–610. doi: 10.1002/mrm.10256. [DOI] [PubMed] [Google Scholar]
  • 36.Adosraku RK, Choi GTY, Constantinou-Kokotos V, Anderson MM, Gibbons WA. NMR lipid profiles of cells, tissues, and body fluids: proton NMR analysis of human erythrocyte lipids. J Lipid Res. 1994;35(11):1925–1931. [PubMed] [Google Scholar]
  • 37.McGahon AJ, Martin SJ, Bissonnette RP, Mahboubi A, Shi Y, Mogil RJ, Nishioka WK, Green DR. The end of the (cell) line: Methods for the study of apoptosis in vitro Methods. Cell Biology. 1995;46:153–181. doi: 10.1016/s0091-679x(08)61929-9. [DOI] [PubMed] [Google Scholar]
  • 38.Foyer C, Walker D, Spencer C, Mann B. Observations on the phosphate status and intracellular pH of intact cells, protoplasts and chloroplasts from photosynthetic tissue using phosphorus-31 nuclear magnetic resonance. Biochem J. 1982;202(2):429–434. doi: 10.1042/bj2020429. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Yoshida T, Watari H. Exercise-induced splitting of the inorganic phosphate peak: investigation by time-resolved 31P-nuclear resonance spectroscopy. Eur J Appl Physiol. 1994;69:465–473. doi: 10.1007/BF00239861. [DOI] [PubMed] [Google Scholar]
  • 40.Muruganandham M, Alfieri AA, Matei C, Chen Y, Sukenick G, Schemainda I, Hasmann M, Saltz LB, Koutcher JA. Metabolic signatures associated with a NAD synthesis inhibitor-induced tumor apoptosis identified by 1H-decoupled 31P magnetic resonance spectroscopy. Clin Cancer Res. 2005;11(9):3503–3513. doi: 10.1158/1078-0432.CCR-04-1399. [DOI] [PubMed] [Google Scholar]
  • 41.Griffiths JR, Stevens AN, Iles RA, Gordon RE, Shaw D. 31P-NMR investigation of solid tumours in the living rat. Bioscience Reports. 1981;1(4):319–325. doi: 10.1007/BF01114871. [DOI] [PubMed] [Google Scholar]
  • 42.Stubbs M, Bashford CL, Griffiths JR. Understanding the tumor metabolic phenotype in the genomic era. Curr Mol Med. 2003;3(1):49–59. doi: 10.2174/1566524033361645. [DOI] [PubMed] [Google Scholar]
  • 43.De Azevedo WF, Jr., Mueller-Dieckmann H-J, Schulze-Gahmen U, Worland PJ, Sausville E, Kim S-H. Structural basis for specificity and potency of a flavonoid inhibitor of human CDK2, a cell cycle kinase. Proc Natl Acad Sci USA. 1996;93(7):2735–2740. doi: 10.1073/pnas.93.7.2735. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Aklilu M, Kindler HL, Donehower RC, Mani S, Vokes EE. Phase II study of flavopiridol in patients with advanced colorectal cancer. Ann Oncol. 2003;14:1270–1273. doi: 10.1093/annonc/mdg343. [DOI] [PubMed] [Google Scholar]
  • 45.Sedlacek HH. Mechanisms of action of flavopiridol. Crit Rev Oncol Hematol. 2001;38(2):139–170. doi: 10.1016/s1040-8428(00)00124-4. [DOI] [PubMed] [Google Scholar]
  • 46.Bible KC, Kaufmann SH. Flavopiridol: a cytotoxic flavone that induces cell death in noncycling A549 human lung carcinoma cells. Cancer Research. 1996;56(21):4856–4861. [PubMed] [Google Scholar]
  • 47.Kitada S, Zapata JM, Andreeff M, Reed JC. Protein kinase inhibitors flavopiridol and 7-hydroxy-staurosporine down-regulate antiapoptosis proteins in B-cell chronic lymphocytic leukemia. Blood. 2000;96(2):393–397. [PubMed] [Google Scholar]
  • 48.Wittmann S, Bali P, Donapaty S, Nimmanapalli R, Guo F, Yamaguchi H, Huang M, Jove R, Wang HG, Bhalla K. Flavopiridol down-regulates antiapoptotic proteins and sensitizes human breast cancer cells to epothilone B-induced apoptosis. Cancer Research. 2003;63(1):93–99. [PubMed] [Google Scholar]

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Supplementary Materials

Supplementary Material
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