LY2880070

Cholinekinase-α proteinandphosphatidylcholine but not phosphocholine are required for breast cancer cell survival

Noriko Moria*, Flonné Wildesa, Samata Kakkada, Desmond Jacoba, Meiyappan Solaiyappana, Kristine Glundea,b and Zaver M. Bhujwallaa,b*

Abstract

High levels of total choline and phosphocholine (PC) are consistently observed in aggressive cancers. Choline kinase (Chk) catalyzes choline phosphorylation to produce PC in phosphatidylcholine (PtdCho) biosynthesis. PtdCho is the most abundant phospholipid in eukaryotic cell membranes and plays a dual role as the structural component of membranes and as a substrate to produce lipid second messengers such as phosphatidic acid and diacylglycerol. Chk-α, but not Chk-β, is overexpressed in various cancers, and is closely associated with tumor progression and invasiveness. We have previously shown that downregulation of mRNA using small interfering RNA (siRNA) against Chk-α (siRNA-Chk) or Chk short hairpin RNA, and the resultant decrease of Chk-α protein levels, significantly reduced proliferation in breast cancer cells and tumors. A novel potent and selective small-molecule Chk-α inhibitor, V-110711, that inhibits the catalytic activity of Chk has recently been developed. Here, we used triple negative MDAMB-231 and SUM149 breast cancer cells to further investigate the role of Chk-α in cancer, by examining Chk-α protein levels, cell viability/proliferation, choline phospholipid and lipid metabolism, lipid droplet formation, and apoptosis, following treatment with V-11-0711. Under the conditions used in this study, treatment with V-11-0711 significantly decreased PC levels but did not reduce cell viability as long as Chk-α protein and PtdCho levels were not reduced, suggesting that Chk-α protein and PtdCho, but not PC, may be crucial for breast cancer cell survival. These data also support the approach of antitumor strategies that destabilize Chk-α protein or downregulate PtdCho in breast cancer treatment. Copyright © 2015 John Wiley & Sons, Ltd.

Keywords: breast cancer; choline kinase; MRS; phosphocholine; phosphatidylcholine; lipid metabolism; lipid droplets

INTRODUCTION

The-ever-increasing evidence of this “cholinic phenotype” in multiple cancers (8), mechanisms underlying the alterations in choline phospholipid metabolism are still not fully understood. One molecular mechanism that is clearly emerging is the overexpression of choline kinase-alpha (Chk-α), the enzyme that catalyzes the phosphorylation of Cho to yield PC in phosphatidylcholine (PtdCho) biosynthesis (Kennedy pathway) (9,10). PtdCho is the most abundant phospholipid in eukaryotic cell membranes and plays a dual role as the structural component of membranes and as the substrate that produces lipid second messengers such as phosphatidic acid and diacylglycerol (11). PC itself has been shown to be mitogenic in NIH3T3 cells (12), and Chkdependent PC was shown to be essential for growth-factorinduced DNA synthesis and to act as a second messenger in eukaryotic cells (12,13).
The regulation of choline phospholipid metabolism by oncogenic signaling pathways includes the involvement of the RAS and phosphatidylinositol 3-kinase/AKT pathways, and transcription factors (8). In mammalian cells, Chk has at least three isoforms, Chk-α1, Chk-α2, and Chk-β (10). Chk-α1 and Chk-α2 MRS studies frequently detect altered choline phospholipid metabolism, especially elevated phosphocholine (PC) and total choline (tCho: PC + glycerophosphocholine (GPC) + free choline (Cho)) signals in cultured human cancer cells, human cancer xenografts, biopsies, and human tumors (1–7). Despite the isoforms derive from alternative splicings of the chk-α gene, whereas Chk-β is derived from a separate gene, chk-β (10). Chkα is overexpressed at the mRNA and protein levels in ovarian, breast, and bladder cancer cell lines, and Chk-α, but not Chk-β, is closely associated with tumor progression and invasiveness (7,14,15). As a result, Chk-α has been the target for the development of novel pharmacological or gene therapy interventions. We previously observed that downregulation of mRNA using small interfering RNA (siRNA) against Chk (siRNA-Chk) and the resultant decrease of Chk-α protein levels significantly reduced proliferation in breast cancer cells (16,17) and tumors (2). We also observed that the combination of siRNA-Chk with 5-FU treatment resulted in an additional reduction of cell viability and proliferation in breast cancer cells (17). MN58b and TCD-717, which are competitive inhibitors with Cho at the Cho binding pocket, are being developed as anticancer drugs, and combination treatment with 5-FU resulted in synergistic antitumoral effect (18). TCD-717 has entered Phase I clinical trials against solid tumors (Study of Intravenous TCD-717 in Patients with Advanced Solid Tumors, NCT01215864). Recently, V-11-0711 (Vertex Pharmaceuticals (Europe) Limited, Abingdon, Oxfordshire, UK), a novel potent and selective small-molecule Chk-α inhibitor with an IC50 of 20 nM that acts by inhibiting the catalytic activity of Chk-α, has been developed (19). In HeLa cells, V-11-0711 significantly reduced PC, as detected by liquid chromatography–tandem mass spectrometry, but did not cause cell death, whereas downregulation of Chk-α by siRNA-Chk caused apoptosis (19).
To further investigate the role of Chk-α in breast cancer, here we treated triple negative MDA-MB-231 and SUM149 breast cancer cells with V-11-0711, and examined its effects on Chk-α protein levels, cell death, and choline phospholipid and lipid metabolism. Under the conditions used, V-11-0711 significantly reduced PC levels but did not reduce cell proliferation, levels of Chk-α protein, PtdCho, or fatty acids in either MDA-MB-231 cells (at 0.1 μM, 1 μM, and 10 μM) or SUM149 cells (at 0.1 μM). In contrast, cell proliferation was reduced when Chk-α protein level was downregulated by siRNA-Chk in the presence or absence of V-11-0711 in both cell lines. These data are consistent with results obtained with HeLa cells (19).
SUM149 cells, which had a lower basal level of Chk-α protein and PC, were more sensitive to V-11-0711 at higher concentrations than MDA-MB-231 cells. Decreased cell viability and PtdCho levels were observed in SUM149 cells following treatment with 1 μM and 10 μM V-11-0711. Additionally, with 10 μM V-11-0711 treatment, fatty acid levels and lipid droplet (LD) formation increased, as did activation of apoptosis and reduction of Chk-α protein levels.
These data demonstrate that reduction of PC has little effect on the proliferation of these breast cancer cells under the conditions described, as long as Chk-α protein levels and PtdCho levels are not reduced. Chk-α protein and PtdCho, but not PC, may be essential in cancer cell survival and proliferation. The data support the development of strategies that destabilize Chk-α protein and/or downregulate PtdCho level in breast cancer.

EXPERIMENTAL DETAILS

Cell culture and treatment

MDA-MB-231 human breast cancer cells were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA) and were grown in RPMI-1640 medium (Sigma-Aldrich, St Louis, MO, USA) supplemented with 10% fetal bovine serum (FBS, Gemini Bio-Products, West Sacramento, CA, USA). SUM149 cells were purchased from Asterand (Detroit, MI, USA) and were grown in DMEM/F-12 (1:1) medium (Mediatech, Manassas, VA, USA) supplemented with 5% FBS, 5 μg/mL insulin (Life Technologies, Grand Island, NY, USA), and 0.5 μg/mL hydrocortisone (Sigma-Aldrich). Cells were maintained at 37 °C in a humidified atmosphere with 5% CO2 in air. All cell lines were tested routinely for mycoplasma contamination. Cells were treated with 0.1% v/v DMSO (vehicle control, Sigma-Aldrich), DharmaFECT (DFECT, transfection reagent, Thermo Fisher Scientific, Lafayette, CO, USA), 0.1 μM, 1 μM or 10 μM V-11-0711 (Vertex Pharmaceuticals (Europe) Limited), and/or 25 nM siRNA in culture media for 48 h.

Cell count and extraction

4 × 106 MDA-MB-231 or 3 × 106 SUM149 cells were seeded in T175 flasks and two to five flasks were used for each treatment. Cells were cultured for 24 h in culture medium and were treated with 0.1% DMSO, 0.1 μM, 1 μM, or 10 μM V-11-0711 for 48 h. Adherent cells were collected by trypsinization and live cell numbers were counted using a hemacytometer after staining dead cells with trypan blue. To obtain % viable cells, the average of the live cell number in each flask was compared with control (DMSO treated) cell number. At least 107 cells were used for cell counting and extraction. Water-soluble as well as lipid extracts were obtained from DMSO, 0.1 μM, 1 μM, or 10 μM V-11-0711 treated cells using the dual-phase extraction method (16). Briefly, the pelleted cells were mixed with 4 mL of ice-cold methanol and vigorously vortexed. After keeping samples on ice for 15 min, 4 mL of chloroform was added, vortexed vigorously, and kept on ice for a further 10 min. Finally, 4 mL of water was added and shaken well. Samples were stored at 4 °C overnight for phase separation and later centrifuged at 20 000 g at 4 °C for 30 min. The water–methanol phase containing water-soluble cellular metabolites such as free Cho, PC, and GPC was treated with about 100 mg of chelex beads (Sigma-Aldrich) to remove any divalent cations. After removing the beads, methanol was evaporated using a rotary evaporator. The remaining water phase was lyophilized. The chroloform phase (lipid phase) was collected in the tube and chroloform was evaporated using nitrogen gas. Both phases of cell extracts were stored at 20 °C until use.

MRS

Water-soluble extracts were resuspended in 0.6 mL of deuterated water (D2O) containing 2.4 × 107 mol of 3-(trimethylsilyl) propionic 2,2,3,3-d4 acid (TSP; Sigma-Aldrich) as an internal standard for MR spectral analysis. Lipid phase cell extracts were resuspended in 0.4 mL of chroloform-D with 0.05% v/v tetramethylsilane (TMS; Cambridge Isotope Laboratories, Andover, MA, USA) and 0.2 mL of methanol-D4 with 0.05% v/v TMS (Cambridge Isotope Laboratories). Fully relaxed 1H MR spectra of water-soluble extracts and lipid phase extracts were acquired on a Bruker AVANCE 11.7 T spectrometer (Bruker BioSpin, Billerica, MA, USA) with flip angle = 30°, sweep width = 10 000 Hz, repetition time = 11.2 s, block size = 32k, and scans = 128.
MR spectra were analyzed using Bruker XWIN-NMR 3.5 software (Bruker BioSpin) as previously described (20). Signal integrals of –N(CH3)3 of Cho at about 3.208 ppm, PC at about 3.226 ppm, and GPC at about 3.235 ppm in water-soluble extracts were determined, normalized to cell number and cell volume, and compared with the standard. To determine concentrations of water-soluble extracts, peak integrations (Imet) from 1H spectra for PC, GPC, and Cho were compared with that of the internal standard TSP (ITSP) according to the equation

ITSPNcellVcell

In this equation, [metabolite] represents the intracellular concentration of the metabolite of interest expressed as mmol/L (mM). ATSP is the number of moles of TSP in the sample, N is the cell number, and V is cell volume (MDA-MB-231, 2050 μm3, and SUM149, 1981 μm3). I represents the signal integral of the selected peak from 1H spectra. Because the number of protons contributing to the signals of PC, GPC, and Cho and the TSP peak is the same, a correction for differences in the number of protons was not required. To determine the cell volume, cell size was determined by trypsinizing the cells, measuring the diameter (d) of 100 randomly selected cells using an optical microscope, and calculating as [(4π/3)(d/2)3]. The resulting metabolite concentrations of PC, GPC, and tCho were averaged for at least three independent experiments.
In lipid phase samples, signal integrals of total methyl groups (–CH3) in fatty acids and cholesterol at about 0.89 ppm, methylene groups (–(CH2)n–) in acyl chains of fatty acids at about 1.27 ppm, and the choline group (–N+(CH3)3) of PtdCho at about 3.22 ppm were determined and normalized to cell number, and compared with the internal standard, TMS. To determine the arbitrary units (A.U.) of metabolites from lipid phase samples, peak integrations (Imet) from 1H spectra for methyl and methylene groups in fatty acids and for PtdCho were compared with that of TMS (ITMS) and normalized by the number of cells (Ncell) used for cell extraction according to the equation

Immunoblot analysis

Total protein from cells was extracted 48 h post treatment using RIPA buffer (50 mM Tris, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton-100, 1% sodium deoxycholate, 0.1% SDS, 1 mM phenylmethylsulfonyl fluoride, 1 mM sodium orthovanadate, 1 mM sodium fluoride, 1 mM dithiothreitol) and a protease inhibitor cocktail at 1:200 dilution (Sigma-Aldrich). Cell lysates were kept on ice for 30 min, and then centrifuged at 16 000 g for 30 min at 4 °C. Protein concentrations were estimated using the Bio-Rad DC assay (Bio-Rad, Hercules, CA, USA). SDS-PAGE was run on a 4–20% reducing gel loaded with equal amounts of total protein (11–50 μg) from samples and the proteins were transferred to a nitrocellulose membrane (Bio-Rad) at 4 °C. After blocking in 5% milk–TBST (TBS Tween), the membrane was separately probed with a custom-made polyclonal antibody against Chk-α (Proteintech Group, Chicago, IL, USA) as previously described (16) or monoclonal antibody against caspase-3 (Cell Signaling Technology, Danvers, MA, USA). Mouse monoclonal antibodies against GAPDH and betaactin (Sigma-Aldrich) were used as loading controls. Secondary antibodies used were horseradish peroxidase-conjugated antimouse and anti-rabbit IgG (GE Healthcare UK, Little Chalfont, Buckinghamshire, UK). Reactions were recorded on Blue Bio film (Denville Scientific, Metuchen, NJ, USA) following use of Super Signal West Pico Substrate (Pierce Biotech, Rockford, IL, USA). Chk-α was detected at approximately 48 kDa. Caspase-3 and cleaved caspase-3 were detected at approximately 35 kDa and 17 kDa respectively. The films were scanned and densitometry was carried out using the Gel Analysis method in ImageJ (NIH, Bethesda, MD, USA). Relative density changes (Chk-α/GAPDH) in Chk-α were normalized to the Chk-α immunoreactive band in control cells, which was set to unity using data from at least three cell lysates.

Cell viability/proliferation assay by CCK-8

4000 cells were seeded in each well of a 96 well plate and cultured overnight. Twenty-four hours later, cells were treated with DMSO, 0.1 μM (SUM149) or 1 μM (MDA-MB-231) of V-11-0711 for 48 h. 25 nM siRNA was transfected transiently using D-FECT with or without V-11-0711 for 48 h. After 48 h treatment, the reagent was changed to fresh culture medium and cells were cultured for three additional days. Cell Counting Kit-8 assays (CCK-8, Dojindo Molecular Technologies, Rockville, MD, USA) were performed at 48 h treatment and at 3 days growth in culture media after 48 h treatment, and were performed using manufacturer’s instructions. In the CCK-8 colorimetric assay the amount of formazan dye, generated by the activities of dehydrogenases in cells, is directly proportional to the number of living cells. Untreated, 0.1% DMSO treated, and negative control siRNA treated cells were used as negative controls. Values from each group were compared with the average of untreated values and shown as viability (%). At least three independent experiments were performed and the average of untreated cells was considered as 100%. RNA interference experiments siRNA targeting Chk-α1 and Chk-α2 (siRNA-Chk) was designed with the sequence 5′-CAUGCUGUUCCAGUGCUCC-3′ and purchased as a duplex from Thermo Fisher Scientific. 25 nM siRNA-Chk was used for SUM149 cells and MDA-MB-231 cells. The same concentrations of negative control siRNA (Shanghai GenePharma, China) were used as a negative control. D-FECT was utilized for transient transfection as per the manufacturer’s instructions. Briefly, cells were cultured for 24 h in antibiotic free culture media and transiently transfected with 25 nM siRNA using D-FECT for 48 h.

Nile red staining of LDs and observation with fluorescence microscope

Cells were grown on glass chamber slides (Thermo Fisher Scientific) in culture medium at 37 °C overnight. After 48 h treatment with DMSO or 0.1 μM, 1 μM, or 10 μM V-11-0711, cells were washed with PBS and fixed with 4% paraformaldehyde. After this, cells were washed with PBS and incubated at 1:20 000 dilution in PBS with a 25 mg/mL Nile red solution (Sigma-Aldrich) for 10 min in the dark at room temperature. Cells were washed with
PBS and mounted with ProLong Gold antifade reagent (Life Technologies), which included 4′,6-diamidino-2-phenylindole (DAPI) for nuclear staining. Cells were imaged with an Olympus FV1000 MPE multiphoton laser scanning microscope (Olympus, Center Valley, PA, USA), using a red filter (excitation 810 nm, emission 640 nm) for Nile red-stained LDs and a blue filter (excitation 810 nm, emission 455 nm) for DAPI-stained nuclei. A 40× LUMPLFL N lens (numerical aperture = 0.8, water immersion) was used to acquire confocal z-stack images. The number of LDs per cell was quantified on maximum intensity projected images from the z-stacks using customized in-house software. More than 100 cells were randomly selected for analysis from each treatment. At least three independent experiments were performed and analyzed for each treatment.

Statistical analysis

Data were expressed as mean ± standard deviation (SD). The significances of differences between the control and treated groups were evaluated by two-tailed unpaired Student t-tests. P values of less than 0.05 were considered significant unless otherwise stated. Each experiment was carried out with at least three individual repeats.

RESULTS

Levels of choline phospholipid metabolites in water-soluble extracts determined by MRS

To investigate choline phospholipid metabolites in water-soluble extracts, 1H MR spectra were obtained after 48 h treatment of MDA-MB-231 cells and SUM149 cells with V-11-0711. Representative spectra demonstrating changes in choline metabolites following treatment with V-11-0711 are shown in Figure 1A, B for MDA-MB-231 and SUM149 cells respectively. As evident in the spectra, all doses of V-11-0711 reduced PC levels compared with controls in both cell lines. Data from multiple experiments are summarized in Figure 1C for MDA-MB-231 cells and Figure 1D for SUM149 cells. All three doses of V-11-0711 resulted in a significant decrease of PC (P = 0.003 for 0.1 μM, 0.0009 for 1 μM, 0.0006 for 10 μM in MDA-MB-231 and P = 0.002 for 0.1 μM, 0.00001 for 1 μM, 0.00001 for 10 μM in SUM149 cells) and tCho (P = 0.002 for 0.1 μM, 0.002 for 1 μM, 0.0002 for 10 μM in MDA-MB-231 and P = 0.003 for 0.1 μM, 0.00004 for 1 μM, 0.00004 for 10 μM in SUM149 cells). In addition, GPC decreased significantly at 1 μM (P = 0.0002) and 10 μM (P = 0.0001) concentrations of V-11-0711 in SUM149 cells, and at 10 μM (P = 0.019) V11-0711 concentration in MDA-MB-231 cells. PC levels in MDA-MB-231 control cells (5.92 ± 2.80 mM) were about five times higher than in SUM149 control cells (1.13 ± 0.29 mM).

Effect of V-11-0711 treatment on cell viability in MDA-MB-231 and SUM149 cells

To investigate the effect of V-11-0711 on cell viability, cell numbers were counted after cells were treated with 0.1–10 μM V-11-0711 for 48 h. Under these conditions MDA-MB-231 cells showed no significant reduction of viability following treatment with up to 10 μM V-11-0711 compared with 0.1% DMSO treated control cells (Fig. 2). SUM149 cells showed no significant reduction of viability following treatment with 0.1 μM V-11-0711, but a significant reduction of viability was observed following 48 h treatment with 1 μM (P = 0.027) and 10 μM V-11-0711 (P = 0.0003) (Fig. 2). The reduction of SUM149 cell viability was dose dependent at 1 μM and 10 μM V-11-0711. Overall, SUM149 cells were more sensitive to V-11-0711 treatment compared with MDA-MB-231 cells.

Effect of V-11-0711 treatment on Chk-α protein level in MDA-MB-231 and SUM149 cells

Figure 1. (A, B) Representative 1H MR spectra from the 3.20–3.25 ppm region of water-soluble extracts from (A) MDA-MB-231 cells and (B) SUM149 cells after treatment with 0.1% DMSO (control) and different doses of V11-0711. Spectra were acquired with 30° flip angle, 10 000 Hz sweep width, 11.2 s repetition time, 32k block size and 128 scans. (C, D) Millimolar levels of PC, GPC and tCho quantified from 1H MR spectra of (C) MDA-MB-231 cells and (D) SUM149 cells. Control (n = 8 for C, D) ( ), 0.1 μM V-11-0711 (n = 3 for C, n = 4 for D) ( ), 1 μM V-11-0711 (n = 3 for C, n = 5 for D) ( ), and 10 μM V-11-0711 (n = 3 for C, D) ( ). Values represent mean ± SD. * P < 0.05 and ** P < 0.01 compared with control. Figure 3A–C shows representative images of Chk-α antibody immunoblots from MDA-MB-231 and SUM149 cells. MDA-MB-231 control cells contained higher Chk-α protein than SUM149 cells (Fig. 3A). To investigate the effect of V-11-0711 on Chk-α protein levels, cells were treated with V-11-0711 for 48 h and cell lysates were prepared. Although not statistically significant, treatment of MDA-MB-231 cells tended to increase Chk-α protein levels approximately 1.4-fold compared with control cells, in a dose dependent manner (Fig. 3B, D). There was no significant change of Chk-α protein level in SUM149 cells following treatment with 0.1 μM and 1 μM V-11-0711 (Fig. 3C, E). Treatment with 10 μM V-11-0711 resulted in a 50% decrease of Chk-α protein level (Fig. 3C, E), although this was not statistically significant. Changes in cell viability after transient siRNA-Chk transfection with or without V11-0711 treatment in MDA-MB-231 and SUM149 cells Since the significant decrease of PC following V-11-0711 treatment did not affect cell viability, especially in MDA-MB-231 cells, we investigated if downregulating Chk-α reduced cell proliferation with or without V-11-0711 treatment. CCK-8 assays were performed after cells were transfected with siRNA-Chk in the presence or absence of V-11-0711. Concentrations of 1 μM were used for MDA-MB-231 cells and 0.1 μM for SUM149 cells, since these concentrations did not affect cell viability and did not decrease Chk-α protein levels, but significantly lowered PC levels. There were no significant changes in viability of MDA-MB-231 cells following treatment with 25 nM negative control siRNA and 1 μM V-11-0711 alone, compared with DMSO treated control cells (Fig. 4A), consistent with the earlier data. There was about 20% reduction in viability of MDA-MB-231 cells after transfection with siRNA-Chk with or without V-11-0711 compared with negative control siRNA treated MDA-MB-231 cells at 48 h treatment. By 3 days, a significant decrease of cell viability was observed in MDA-MB-231 cells after 48 h siRNA-Chk treatment compared with negative control siRNA treated cells (P = 0.0001) (Fig. 4A). The presence or absence of V-11-0711 did not affect the reduction of cell viability. Knockdown of Chk-α protein was confirmed using immunoblot analysis after 48 h siRNA-Chk transfection with or without V-11-0711 (Fig. 4C) in MDA-MB-231 cells. Similar to MDA-MB-231 cells, treatment of SUM149 cells with 25 nM negative control siRNA and 0.1 μM V11-0711 did not result in a significant reduction of viability compared with DMSO treated control cells (Fig. 4B). Significant decreases of viability following transfection with siRNA-Chk, with or without V11-0711, compared with negative control siRNA treated cells were observed at both 48 h treatment (P = 0.013 or 0.015 respectively) and 3 days after 48 h treatment (P = 0.039 or 0.015 respectively) (Fig. 4B). As with MDA-MB-231 cells, there was no significant difference between treatment with siRNA-Chk only and treatment with siRNA-Chk and V11-0711. Immunoblot assays confirmed the downregulation of Chk-α protein by siRNA-Chk in SUM149 cells (Fig. 4D). Levels of lipid metabolites in lipid extracts by MRS To investigate the levels of lipid metabolites following treatment with V-11-0711, 1H MR spectra of lipid-soluble extracts were obtained at 48 h from treated MDA-MB-231 cells (Fig. 5A, B, E, G) and SUM149 cells (Fig. 5C, D, F, H). PtdCho and fatty acid levels of MDA-MB-231 cells did not change significantly following V11-0711 treatment (Fig. 5E, G) although slight decreases of PtdCho levels were observed with V-11-0711 treatments. The levels of PtdCho in SUM149 cells decreased significantly following treatment with 1 μM (P = 0.005) and 10 μM (P = 0.011) V-11-0711, but not with 0.1 μM V-11-0711 (Fig. 5F). There was no change of fatty acid levels with 0.1 μM and 1 μM V-11-0711 treatment of SUM149 cells, but the level of fatty acids increased with 10 μM V-11-0711 treatment (–(CH2)n–; P = 0.059) (Fig. 5H). Effect of V-11-0711 treatment on LD number in MDA-MB-231 and SUM149 cells To further understand the changes in fatty acids observed with V-11-0711 treatment, cells were stained with Nile red (21,22) to evaluate changes in LDs. In MDA-MB-231 cells, V-11-0711 treatment did not change the number of LDs per cell (Fig. 6A, C). In SUM149 cells, 10 μM V-11-0711 treatment significantly increased the number of LDs per cell by about 1.8-fold (P = 0.029) compared with control cells (Fig. 6B, D). Effect of V-11-0711 treatment on caspase-3 protein level in MDA-MB-231 and SUM149 cells To detect the induction of apoptosis by V-11-0711, levels of caspase-3 and cleaved caspase-3 were determined by immunoblot analysis. Activation of caspase-3 is detected from its cleavage by initiator caspases after apoptotic signaling events have occurred (23). In MDA-MB-231 cells, the levels of full-length caspase-3 did not decrease after V-11-0711 treatment at all concentrations, and no band was observed at 17 kDa (Fig. 7A). In SUM149 cells, treatment with 1 μM V-11-0711 induced the appearance of a faint band from cleaved caspase-3, but caused no detectable change of full-length caspase-3 protein. However, full-length caspase-3 decreased and cleaved caspase-3 clearly appeared after treatment with 10 μM V-11-0711 (Fig. 7B). DISCUSSION Increased PC and tCho levels are observed as frequently as increased glycolysis in cancer (1–8). These alterations in choline metabolites are primarily due to increased levels of Chk-α but not Chk-β in cancer cells (7,14,15). Unraveling the role of Chk-α in cancer can provide new insights into the malignant phenotype and lead to the development of treatment strategies targeting this enzyme in particular, and choline phospholipid metabolism in general. Chk-α is crucial for early embryo development, as shown in mouse studies, and disruption of the chk-α gene is lethal for embryonic development (24). Mice that lack Chk-β can survive to adulthood, but develop hindlimb muscular dystrophy and forelimb bone deformity (25,26). Chk-β therefore does not compensate for the loss of Chk-α, which suggests that Chk-α is necessary to maintain PtdCho biosynthesis. Approximately 75% of estrogen receptor (ER)-negative breast cancers have increased Chk activity, while 29% of ER-positive breast cancers have increased Chk activity (4). These data suggest that Chk inhibition may especially benefit ER-negative breast cancer patients. Here we have investigated the effects of a Chk-α specific inhibitor, V-11-0711, on choline and lipid metabolism, Chk-α protein levels, and the viability of triple (ER/progesterone receptor/Her2-neu) negative MDA-MB-231 and SUM149 breast cancer cells. In previous studies, treatment of cancer cells with a Chk inhibitor such as MN58b have shown a significant decrease of proliferation (27) following treatment with 5 μM or higher doses. Here, despite the profound decrease of PC at 0.1, 1, and 10 μM V-11-0711 in MDA-MB-231 cells and 0.1 μM V-11-0711 in SUM149 cells, neither MDA-MB-231 nor SUM149 cells showed decreases of cell viability or Chk-α protein levels, suggesting that decrease of PC alone did not decrease cell viability. V-11-0711 is a highly specific Chk-α inhibitor with an IC50 of 20 nM, and an 11fold lower activity against Chk-β (IC50 220 nM). The decrease of PC following treatment with V-11-0711 was most likely due to the inhibition of the catalytic activity of Chk-α, since V-11-0711 binds in the Cho binding pocket of the enzyme and competes with ATP as described for V-11-023907 (28), and shows excellent selectivity against a panel of 50 kinases (19). However we cannot rule out the possibility of other factors such as the inhibition of Cho transport or increased PC consumption contributing to these changes, as these were not examined here. Since the agent acts on the catalytic activity of the enzyme, we did not anticipate significant changes in Chk protein levels, as confirmed in the immunoblots. We previously observed a significant decrease of cell viability following downregulation of Chk-α with siRNA (2,16,17). Here we confirmed that Chk-α downregulation with siRNA significantly decreased cell viability. The presence or absence of V11-0711 (1 μM for MDA-MB-231 and 0.1 μM for SUM149 cells) did not modify this decrease. These results are also consistent with previous studies using HeLa cells (19). These data highlight the importance of Chk-α protein levels but not its catalytic product, PC, in modifying cell viability. Chk-α activity increases with overexpression of epidermal growth factor receptor (EGFR) and non-receptor tyrosine kinase (c-Src), and this increase requires c-Src kinase activity and protein complex formation of Chk-α with EGFR and c-Src (29). One role of Chk-α in cancer may be the formation of a protein complex with EGFR and c-Src, independently of its catalytic activity. Accumulating evidence of essential non-catalytic properties of protein kinases, which include scaffolding of protein complexes, competitive protein interactions, allosteric effects on other enzymes, subcellular targeting, and DNA binding, has been identified (30). Both EGFR and c-Src are overexpressed in a variety of human tumors and are closely associated with signal transduction cascades such as MAPK, Akt, and JNK pathways, which lead to cell proliferation and DNA synthesis (31–33). Investigating the association of Chkα with EGFR and c-Src would provide additional insights into the role of Chk-α in cancer. Despite having higher Chk-α levels as well as higher PC and tCho compared with SUM149 cells, in these experiments MDAMB-231 cells did not show any decrease of cell viability even at a concentration of 10 μM V-11-0711. In contrast, SUM149 cells, with lower basal levels of Chk-α and PC, showed a significant reduction of cell viability with 1 and 10 μM V-11-0711 treatments. The Chk-α protein level decreased when SUM149 cells were treated with 10 μM V-11-0711. At the higher doses of 1 μM and 10 μM non-specific effects may come into play. Treatment of SUM149 cells with 1 and 10 μM V-11-0711 was accompanied by a significant decrease of PtdCho and cell viability, but without a reduction of the Chk-α protein at the 1 μM concentration. Since the reduction of PtdCho synthesis has been shown to trigger apoptosis, it is possible that the decrease of PtdCho in SUM149 cells may have induced apoptosis that became clearly evident at the 10 μM dose. The induction of apoptosis was evident from the detection of cleaved caspase-3 in the immunoblot assay. Treatment with V-11-0711 resulted in a significant decrease of GPC at 1 and 10 μM in SUM149 cells, and at 10 μM in MDA-MB231 cells. This decrease may have been caused by the reduction of PtdCho, at least in SUM149 cells, since GPC is produced by lysophospholipase (LPL) from Lyso-PtdCho followed by breakdown of PtdCho with phospholipase A (PLA). Treatment of cells with V-11-0711 at these higher non-specific doses may have affected GPC related enzymes such as LPL, PLA, and glycerophosphodiester phosphodiesterase (34–36). We additionally investigated the effect of V-11-0711 on lipid metabolism in MDA-MB-231 and SUM149 cells. As mentioned earlier, PtdCho levels in SUM149 cells decreased after 1 and 10 μM V-11-0711 treatments. The PC decrease did not play a role in the decrease of PtdCho since both MDA-MB-231 cells and SUM149 cells showed a significant PC reduction at low doses (and high doses in the case of MDA-MB-231 cells) without a reduction of PtdCho. When SUM149 cells were treated with concentrations of V-11-0711 that significantly reduced viability (1 μM and 10 μM), PtdCho levels also significantly decreased. At a concentration of 1 μM, V-11-0711 caused a significant reduction of cell viability without decreasing Chk-α protein in SUM149 cells. This reduction of cell viability was accompanied by a reduction of PtdCho that may have triggered apoptosis (37–40), although only a faint band of cleaved caspase-3 was observed. The effect of PC decrease on cell viability may be mediated through different pathways as compared with the effect of PtdCho decrease on cell viability. We have previously observed that downregulation of Chk-α protein reduced PC and cell viability without affecting PtdCho levels in MDA-MB-231 cells (16). Similar observations have been made in a human osteosarcoma xenograft (41). Fatty acid levels in MDA-MB-231 cells were stable after treatment with all three concentrations of V-11-0711. In SUM149 cells, fatty acids increased after 10 μM V-11-0711 treatment, which coincided with a decrease in cell viability, increased LD formation, and caspase-3 activation. Increases in 1H MRS visible lipids have been observed at low pH and high cell density (42), following cancer therapy (43,44), and following the induction of apoptosis (45). MR visible lipids are formed from de novo synthesis of triglycerides (42). Fatty acids detected in MR spectra include the fatty acids from the plasma membrane and mobile lipids. The significant increase in fatty acids\ observed in SUM149 cells following treatment with 10 μM V-11-0711 was most likely due to the increased numbers of LDs in these cells. While LD formation has been often used as an indicator of breast cancer cell differentiation (16,46), it has also been shown to correlate with increased lipogenesis following induction of apoptosis (45,47), and may be due to the inhibition of mitochondrial fatty acid β-oxidation and re-direction of fatty acids away from oxidation and into lipid synthesis (47). Other studies have shown that, in apoptotic cells, iPLA2 activation induced by caspase-3 proteolysis increased the release of fatty acids from PtdCho (40,48). We confirmed the activation of apoptosis from the increase of cleaved caspase-3 observed in SUM149 cells treated with 10 μM V-11-0711. One interesting fact to emerge from these studies is that the studied triple negative breast cancer cells were able to survive a profound decrease of PC potentially because PtdCho levels were stable. Multiple enzymes other than Chk-α, such as CTP:PC cytidylyltransferase, diacylglycerol cholinephosphotransferase in the Kennedy pathway, and PtdCho-specific phospholipases in the catabolic pathway (8), maintain the level of de novo PtdCho. Additionally, the phosphatidylethanolamine (PtdE) kinase methylation pathway, through which PtdE N-methyltransferase (PEMT) catalyzes the conversion of PtdE to PtdCho, can maintain the PtdCho level (11). Studies have shown that the PtdCho generated from the Kennedy and the PEMT pathways differ functionally, and PtdCho from the Kennedy pathway is more important in somatic cells (38,41,49,50). Further investigation of those possibilities is required to better understand how these cells survived a significant reduction of a membrane precursor. 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