Affiliations of authors: Cancer Research UK Biomedical Magnetic Resonance Research Group, Department of Basic Medical Sciences, St. George's Hospital Medical School, London, U.K. (Y-LC, HT, MS, JRG); Cancer Research UK Centre for Cancer Therapeutics, Institute of Cancer Research, Sutton, Surrey, U.K. (UB, MIW, IRJ, PW); Cancer Research UK Clinical Magnetic Resonance Research Group, Institute of Cancer Research and Royal Marsden Hospital, Sutton (LEJ, MOL, SMR).
Correspondence to: Yuen-Li Chung, PhD, Cancer Research UK Biomedical Magnetic Resonance Research Group, Department of Basic Medical Sciences, St. George's Hospital Medical School, Cranmer Terrace, London SW17 ORE, U.K. (e-mail: ychung{at}sghms.ac.uk).
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ABSTRACT |
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INTRODUCTION |
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17-Allylamino,17-demethoxygeldanamycin (17AAG), a novel anticancer drug that inhibits Hsp90 (4,5,7), is a member of the benzoquinone ansamycin antibiotic family, which includes the geldanamycins and herbimycins. 17AAG has lower hepatic toxicity than its parent compound, geldanamycin, but the same potent anticancer activity (8-10). 17AAG is now in phase I clinical trials as a potential anticancer drug; the absence of antiproliferative-related toxicity observed to date suggests that 17AAG and related drugs may be combined with cytotoxic agents (4,5).
In previous studies (11,12), human colon cancer cells treated with 17AAG showed depletion of Hsp90 client proteins c-Raf-1 and Akt and inhibition of signal transduction, leading to cell cycle arrest and apoptosis. Depletion of these client proteins and elevation of Hsp70 expression (both mRNA and protein) represent a molecular signature that is diagnostic of Hsp90 inhibition (4,11). However, molecular analysis requires biopsies before and after treatment. Hence, a noninvasive surrogate marker for Hsp90 inhibition and, potentially, for analysis of treatment response would be a more desirable endpoint to use both in clinical trials and eventually in patient management (13).
In vivo phosphorus magnetic resonance spectroscopy (31P-MRS) provides noninvasive biochemical information on both healthy and diseased tissues (14-16). This technique has also been used to study the biochemistry and physiology of tumor cells and solid cancers and to assess tumor response following therapies in both human and animal models (17-24). Markers for tissue bioenergetics, such as nucleoside triphosphate (NTP), inorganic phosphate (Pi), and intracellular pH, as well as various phosphorus-containing components of phospholipid membrane turnover, such as phosphomonoesters (PMEs) and phosphodiesters (PDEs), are readily observed using 31P-MRS.
To identify a noninvasive and robust surrogate marker for Hsp90 inhibition and tumor response to 17AAG treatment, we performed a comprehensive 31P-MRS study in human colon carcinoma models. First, we used 31P-MRS on extracts of three human colon cancer cell lines (SW620, HCT116, and HT29) to determine whether any MR spectral changes are associated with Hsp90 inhibition after 17AAG treatment at concentrations that inhibit cell proliferation. Second, we examined the effect of 17AAG on an MF-1 nude mouse HT29 xenograft model to determine efficacy, to determine whether 31P-MRS changes observed in the cell studies were reproducible in vivo, and to determine whether 31P-MRS could provide a noninvasive pharmacodynamic marker for tumor response in clinical trials. The effect of 17AAG treatment on levels of Hsp90 client proteins and the Hsp70 co-chaperone was also determined to correlate these changes in protein expression with changes in the MR spectrum.
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MATERIALS AND METHODS |
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17AAG (powder or dissolved [25 mg/mL] in dimethyl sulfoxide [DMSO]), 17-amino,17-demethoxygeldanamycin (17AG), NSC683666, and egg phospholipids (vehicle) were provided by Dr. P. Ivy at the National Cancer Institute (Bethesda, MD). The powder form of 17AAG was used in the cell experiments, and the dissolved form of 17AAG was used in the xenograft experiments. Dulbecco's modified Eagle medium, McCoy's medium, fetal calf serum, penicillin, and streptomycin were purchased from Gibco (Paisley, U.K.). Perchloric acid (PCA) and potassium hydroxide were purchased from Merck (Poole, U.K.). Hypnorm was purchased from Jansen Pharmaceuticals (Buckinghamshire, U.K.), and Hypnovel was purchased from Roche (Welwyn Garden City, U.K.). Bicinchonic acid and enhanced chemiluminescence reagents were purchased from Pierce (Rockford, IL). Tris-glycine gels and Tropifluor polyvinylidene fluoride membranes were purchased from Invitrogen (Groningen, The Netherlands) and Immobilon (Bedford, MA), respectively. Primary antibodies SC-133 and SC-260 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA), whereas primary antibodies SPA-810 and MAB-371 were purchased from Stressgen Biotechnologies (Victoria, British Columbia, Canada) and Chemicon International (Temecula, CA), respectively. Secondary anti-rabbit (cat. No. NA9340) and anti-mouse (cat. No. NA931V) antibodies were purchased from Amersham Pharmacia Biotechnology (Buckinghamshire, U.K.). All other chemicals were purchased from Sigma (Poole, U.K.).
Cell Culture and Treatment
All cell lines (HCT116, HT29, and SW620) (American Type Culture Collection, Manassas, VA) were cultured in Dulbecco's modified Eagle medium supplemented with 10% fetal calf serum, 80 U/mL penicillin, and 80 µg/mL streptomycin at 37 °C in 5% CO2. Treatment with 17AAG, aimed at achieving comparable inhibition of cell growth for all three cell lines, was as follows: HCT116 and HT29 cells were treated with 1.36 µM 17AAG for 24 hours at 37 °C, and SW620 cells were treated with 1.36 µM 17AAG for 48 hours at 37 °C. The cells then underwent trypsinization (11,12) and trypan blue exclusion assay (11,12). The effect of 17AAG treatment on cell proliferation was monitored by counting the cells in a vehicle (DMSO, 1 : 5000)-treated control flask and comparing that number with the number of cells in a 17AAG-treated flask. The effect of 17AAG treatment on the cell lines was further monitored by assessing Hsp70 and Hsp90 client protein levels using western blots as described below.
HT29 cells were also treated with either 1.36 µM 17AG (the active metabolite of 17AAG) or 1.36 µM NSC683666 (an inactive benzoquinone ansamycin analog) for 24 hours at 37 °C. The effects of these compounds on cell proliferation (using the method described above) and Hsp70 and Hsp90 client protein expression (using western blots) were determined. HT29 cells were then treated with 10 µM doxorubicin for 24 hours at 37 °C to confirm that the effect of 17AAG and 17AG treatment on cell density and phosphorus metabolites was not due to cell death.
Cell Cycle Analysis
Cell cycle analysis of control and treated cells by flow cytometry was performed on cells (1 x 106) fixed in 70% ethanol, which were then treated with 100 µg/mL RNase A in phosphate-buffered saline and stained with 4 µg/mL propidium iodide (12), using an Elite Enhanced System Performance cell sorter (Beckman Coulter, High Wycombe, U.K.) at 488 nm. The cytometry data were analyzed using WinMdi and Cylchred software (University of Wales College of Medicine, Cardiff, U.K.).
HT29 Xenograft Model
MF-1 nude mice were injected subcutaneously in the flank with 0.2 mL of a suspension of HT29 human colon carcinoma cells (2.5 x 107 cells/mL) that had been grown as a monolayer in cell culture (24). Tumors reached the minimum size required for MRS measurement (i.e., approximately equivalent to a weight of 500 mg) about 4 weeks after cell inoculation. Tumor size was calculated by measuring the length, width, and depth of each tumor using calipers and by using the following formula: l x w x d x (/6). Once a tumor size of
500 mg (mean ± standard error = 484 ± 36 mg [n = 19]) was established, mice were randomly divided into two groups; 14 mice were treated with 17AAG in egg phospholipids (EPL) vehicle at 80 mg/kg intraperitoneally once a day for 4 days and five mice (controls) were treated with EPL vehicle (10% DMSO added) alone following the same regimen. Animals were treated in accordance with local and national ethical requirements and with the United Kingdom Coordinating Committee on Cancer Research Guidelines for the Welfare of Animals in Experimental Neoplasia (25).
In Vitro 31P-MRS of Cell Extracts
To obtain an MR spectrum, 2 x 107 to 4 x 107 cells in logarithmic phase were extracted as previously described (20,26). Briefly, cells were rinsed with ice-cold saline and fixed in 6 mL of ice-cold methanol. Cells were then scraped off the surface of the culture flask, collected into tubes, and vortexed for 30 seconds at room temperature to optimize phospholipid metabolite extraction from the ruptured cells. Chloroform (6 mL) was then added to each tube, followed by an equal volume of de-ionized water. Following phase separation and solvent removal (20,26), samples were stored at -80 °C until analysis. Prior to acquisition of the MRS spectra, the water-soluble metabolites were resuspended in deuterium oxide with 10 mM EDTA (pH 8.2). Proton (1H)-decoupled 31P-MRS spectra were acquired at room temperature on a 500-MHz Bruker spectrometer (Bruker Biospin, Coventry, U.K.). Metabolite contents were determined by peak integration, normalized relative to the peak integral of an internal reference (methylene diphosphonic acid), and corrected for signal intensity saturation (27) and the number of cells extracted per sample.
In Vivo 31P-MRS of HT29 Xenografts
Animals were anesthetized with a single intraperitoneal injection of a Hypnovel/Hypnorm/water (1 : 1 : 2) mixture as previously described (24). Animals were placed in the bore of a Varian 4.7 tesla (T) nuclear magnetic resonance (NMR) spectrometer, and tumors were positioned in the center of a 12-mm two-turn 1H/31P surface coil. Image-selected in vivo spectroscopy-localized 31P-MR spectra of the tumors were obtained at 37 °C as previously described (28). Briefly, a gradient strength of up to 7.5 x 10-4 T/cm was applied with adiabatic pulses of 800 ms, a 90° sincos excitation pulse, and a sech 180° inversion pulse, with a total repetition time of 3 seconds and 600 averages. 31P-MRS of the tumors was performed before treatment (i.e., day 1) and 4 days after treatment (i.e., day 5). 31P-MR spectra were quantified using the VARiable PROjection program (VARPRO) to determine precise chemical shifts and peak integrals (29). After the final 31P-MRS study, animals were killed by cervical dislocation, and the tumors were removed, freeze-clamped, and stored at -80 °C until analysis.
The surface coils used to obtain the 31P-MRS signal from subcutaneous tumors in vivo were of nonuniform spatial sensitivity, so it was not possible to use an internal standard. Lack of spatial sensitivity and problems with baseline definition and overlapping peaks meant that it was difficult to quantitate the concentration of metabolites from the resulting spectra. As a result, the signal intensities observed in the in vivo 31P-MR spectra are expressed as ratios of metabolites (30).
In Vitro 1H- and 31P-MRS of Tumor Extracts
The freeze-clamped tumors were divided into two groups. Half of the tumors were extracted in 6% PCA as previously described (31), and the rest were used for western blot analysis (described below). Neutralized extracts were freeze-dried and reconstituted in 1 mL of deuterium oxide, and the extracts (0.5 mL) were placed in 5-mm NMR tubes. For 1H-MRS, the water resonance was suppressed by using gated irradiation centered on the water frequency. Sodium 3-trimethylsilyl-2,2,3,3-tetradeuteropropionate (50 µL, 5 mM) was added to the samples for chemical shift calibration and quantitation. Immediately before the MRS analysis, the pH of the samples was readjusted to 7 with PCA or potassium hydroxide. Metabolite concentrations of betaine, lactate, alanine, -hydroxybutyrate, creatine, phosphocholine (PC; with a contribution from phosphoethanolamine [PE]), glycerophosphocholine (GPC), valine, glutamate, taurine, and glycine were quantified from the 1H-MR spectra (32). For 31P-MRS, which was carried out after the 1H-MRS study, EDTA (50 µL, 60 mM) was added to each sample for chelation of metals ions, and methylene diphosphonic acid (50 µL, 5 mM) was added to each sample for chemical shift calibration and quantitation. The extract spectra for both the control and the treated animals were acquired under identical conditions.
Protein Extraction and Western Blot Analysis
Cells and xenograft tissue were lysed in 50 µL and 1 mL of lysis buffer (containing 0.1% Nonidet P-40 , 50 mM HEPES [pH 7.4], 250 mM NaCl, 1 mM phenylmethylsulfonyl fluoride, 10 µg/mL aprotonin, 20 µM leupeptin, 1 mM dithiothreitol, 1 mM EDTA, 1 mM NaF, 10 mM -glycerophosphate, and 0.1 mM sodium orthovanadate), respectively. The protein supernatant was collected after centrifugation at 18 000g for 10 minutes at 4 °C. Protein concentrations were determined by bicinchonic acid reagents (Pierce). Proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis using 4%-20% precast Tris-glycine gels and transferred electrophoretically to 0.45 µM polyvinylidene fluoride membranes. The membranes were then blocked in 0.5% casein blocking buffer (10 mM Tris-HCl [pH 7.6], 155 mM NaCl, 0.1% Tween-20, and 0.02% thimerosal) and incubated overnight at 4 °C with primary antibodies, followed by a 1-hour incubation with horseradish peroxidase-conjugated secondary antibodies at room temperature. The membranes were then washed with enhanced chemiluminescence reagent for 1 minute and exposed to hyperfilm (Amersham Pharmacia Biotechnology, Buckinghamshire, U.K.), which was then developed on a Konica SRX-101A automatic developer (Konica, Tokyo, Japan). Band intensities were assessed by visual examination. The primary antibodies used and their dilutions were as follows: c-Raf-1, 1 : 250 (SC-133); Hsp70, 1 : 2000 (SPA-810); Cdk4, 1 : 1000 (SC-260); and glyceraldehyde-3-phosphate dehydrogenase, 1 : 5000 (MAB-371). Secondary anti-rabbit (NA9340) and anti-mouse (NA931V) antibodies were used at a 1 : 1000 dilution (11,12).
Statistical Analysis
Data are presented as the mean and 95% confidence intervals (CIs). For comparison of metabolite concentrations and ratios, t tests were used, with a P value of <.05 considered to be statistically significant. The Mann-Whitney U test was performed for comparison of proportions, with a P value of <.05 considered to be statistically significant. Spearman's rank correlation was used to correlate changes in tumor size with PME/PDE ratios. All statistical tests were two-sided.
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RESULTS |
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SW620, HCT116, and HT29 cells were treated with doses of 17AAG designed to achieve comparable inhibition of cell proliferation for all three cell lines. After 24-hour (HCT116 and HT29 cells) or 48-hour (SW620 cells) incubation, the number of cells per flask was reduced substantially in each cell line to approximately 60% of the number of vehicle-treated control cells (66%, 95% CI = 62% to 70% in SW620; 58%, 95% CI = 39% to 77% in HCT116; and 57%, 95% CI = 56% to 58% in HT29) (data not shown), consistent with decreased proliferation.
17AAG treatment was also associated with depleted levels of the Hsp90 client proteins c-Raf-1 and Cdk4 and with increased Hsp70 levels (Fig. 1, A). These results provide molecular evidence that Hsp90 was inhibited in the 17AAG-treated cells.
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In Vitro 31P-MRS of Cell Extracts
To identify potential noninvasive markers of Hsp90 inhibition, we determined the 31P-MR spectrum of colon cancer cells treated in vitro with 17AAG. Fig. 2 illustrates 31P-MR spectra of control and 17AAG-treated HT29 cells, demonstrating an increase in PC and GPC levels following treatment. A statistically significant doubling of PC levels (relative to those in vehicle-treated cells) was detected in HT29 and HCT116 cells, and a statistically significant 20-fold increase (again, relative to those in vehicle-treated cells) in these levels was detected in SW620 cells. Increased GPC levels were also detected in all three cell lines; the increase was statistically significant for both HCT116 (in which GPC was below detection in controls) and HT29 cells. Changes in NTP levels were not statistically significant. Glycerophosphoethanolamine (GPE) was not detectable in HCT116 cells and did not change (at 0.3 fmol/cell, 95% CI = 0.0 to 0.3 fmol/cell) following 17AAG treatment in SW620 cells but showed a statistically significant increase from 2.0 to 5.0 fmol/cell (difference = 3.0 fmol/cell, 95% CI = 1.1 to 4.9 fmol/cell; P = .01) in HT29 cells. PE levels in cell extracts remained below detection level in all three cell lines (data not shown).
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Previous studies (33-35) have shown that the levels of PMEs can be modulated by the availability of exogenous precursors. The increase observed may have been due to a decrease in cell counts consistent with decreased proliferation, leading to a reduction in choline consumption following treatment. Therefore, it was necessary to rule out the possibility that the alterations observed in PC and GPC levels following 17AAG treatment were not a result of increased choline availability per cell. To eliminate this possibility, HT29 cells were grown at different cell densities, covering the range observed in control and treated flasks. No statistically significant difference in MR spectra was observed for cells grown at densities ranging from 1 x 107 to 8 x 107 cells per flask (data not shown). Thus, the changes in phosphorus metabolites following 17AAG and 17AG treatment are unlikely to be due to the effects of cell density.
In Vivo 31P-MRS of HT29 Xenografts
Previous studies (10) have shown statistically significant growth delays when HT29 human colon tumor xenografts are treated with 17AAG. Consistent with these results, 17AAG treatment for 4 days reduced tumor size to 97% of pretreatment volume, whereas control tumors increased in size to 120% of pretreatment volume (difference = 23%, 95% CI = 7% to 41%; P = .006). Western blots of the excised tumors showed increased levels of Hsp70 in the 17AAG-treated group (Fig. 1, B). Those same studies also demonstrated client protein depletion in association with elevated Hsp70 levels in this tumor model.
In vivo 31P-MR spectra from a representative HT29 tumor before and after 17AAG treatment are shown in Fig. 4. In vivo 31P-MRS of the 17AAG-treated tumor group showed a small, albeit statistically significant, reduction in the -NTP/total phosphorus signal (TotP) ratio, relative to pre-treatment values (P = .03), which was not always immediately apparent on visual inspection of the spectra because of differences in line width. Statistically significant increases in the PME/PDE (P = .02), PME/TotP (P = .05), and PME/
-NTP (P = .03) ratios, relative to pretreatment values, were also observed after 17AAG treatment (Table 2). No statistically significant changes in these ratios were observed after vehicle treatment of tumors. A statistically significant inverse correlation was found between the percentage change in the PME/PDE ratio and the percentage change in tumor size following 17AAG treatment (r = -0.63, 95% CI = -0.88 to -0.13; P = .02).
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Expanded in vitro 31P-MR spectra (i.e., -3 to 6 ppm) of the tumor extracts from a 17AAG-treated and a control vehicle-treated mouse are presented in Fig. 5, A; peaks were assigned as previously described (36). In vitro 31P-MRS of the extracts, in which the PME signal is better resolved than in in vivo experiments, showed statistically significantly elevated levels of PE (P = .02), PC (P = .07), and the (PE + PC)/(GPE + GPC) ratio (P = .04) (equivalent to the PME/PDE ratio in vivo) in the 17AAG-treated tumors relative to the vehicle-treated tumors (Table 3).
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DISCUSSION |
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Inhibition of Hsp90 by 17AAG was demonstrated by induction of Hsp70 expression and reduction of client protein (c-Raf-1 and Cdk4) levels, which is consistent with our previous findings (10-12). The pattern of changes observed following Hsp90 inhibition was also seen in the HT29 xenografts. An antiproliferative effect on cell and tumor growth was demonstrated by cell counts in vitro and by tumor volume in vivo. Tumors from HT29-xenografted mice treated with 17AAG showed a cytostatic response over the treatment period coincident with the observed molecular changes. Our previous studies (10) with HT29 xenografts have shown that this response translates into a prolonged growth delay. It should be noted that the changes observed in the phosphorus-containing metabolites were observed in the same in vitro cell cultures and in vivo HT29 xenografts in which the molecular signature of Hsp90 was also demonstrated. However, it should also be noted that our investigations were performed using a single 17AAG dose level and at a single time point both in vitro and in vivo. Hence, further studies will need to examine more closely the dose-response and time-response relationships of the changes in phospholipid levels in tumor cells and xenografts.
It is well established that 17AAG affects the cell cycle in a cell line-dependent manner (12). Indeed, similar differences in cell cycle effects between HT29 and HCT116 cells have been previously observed (12). These differences in cell cycle effects may relate to a differential response to depletion of Hsp90 client proteins involved in cell cycle control (4). Importantly, the same spectral alterations were observed in HT29 and HT116 cells regardless of the difference in cell cycle changes, indicating that the cell cycle effect is not responsible for the modulation in phospholipid metabolites following 17AAG treatment.
Because the TotP levels in the tumor were unlikely to change substantially during these experiments, the reduced -NTP/TotP ratio following 17AAG treatment suggests that NTP levels in the HT29 tumor xenografts decreased following 17AAG treatment and that tumor bioenergetics were compromised. This finding was not observed in vitro, where absolute quantitation was possible, probably because cultured tumor cells are bathed in well-oxygenated medium, whereas in in vivo experiments, the structurally and functionally disturbed microcirculation impairs delivery of nutrients, including oxygen (38,39).
In vivo, the PME level, whether compared with TotP, PDE, or -NTP levels, increased statistically significantly following 17AAG treatment from pretreatment (Table 2). The principal PME signals in tumors are PC and PE, precursors of phosphatidylcholine and phosphatidylethanolamine, which are major components of biologic membranes (40). In 31P-MRS of the tumor extracts, the PE level increased statistically significantly after 17AAG treatment (P = .02) and, although the PC level also increased with 17AAG treatment, the difference did not reach statistical significance (P = .07; Table 3). In the 1H-MR spectra of the tumor extracts, the PC plus PE level was statistically significantly higher in the 17AAG-treated group than in the vehicle-treated group (P = .04). The singlet (i.e., single peak) at 3.22 ppm in the 1H-MR spectrum arises mainly from nine equivalent protons from the PC molecule with a small (approximately 20%) contribution from the PE molecule, which has only two protons resonating at 3.22 ppm (41). Hence, the statistically significant increase in the peak at 3.22 ppm is likely due to an elevated level of PC.
Our in vivo findings confirm the statistically significant increase in PC levels following 17AAG treatment that we observed for all three colon cancer cell lines in vitro. In tumor cells, the association between increased PC and Hsp90 inhibition by 17AAG treatment was further confirmed by studies using the active 17AAG metabolite 17AG and the inactive analog NSC683666. Changes in the MR spectra similar to those observed with 17AAG treatment were also seen with 17AG treatment but not with NSC683666 treatment. In addition, we ruled out the possibility that the increase in PC levels was due to inhibition of cell growth by monitoring the effect of doxorubicin, a standard chemotherapeutic agent, on cell proliferation of HT29 cells. This treatment caused a comparable decrease in cell number but had no statistically significant effect on PC levels.
Interestingly, although both PC and PE peaks were observed in untreated tumor extracts, only the PC peak was observed in the cultured cell extracts. Similar findings (32) in HT29 tumor xenografts and cultured cells have been attributed to the greater availability of ethanolamine in vivo when compared with cells in culture, where ethanolamine is only present in added serum. The relative amounts of PC and PE, both of which contribute to the PME signal, appear to vary among both experimental models and human tumors in patients (42). Precursor availability may also have influenced the increased GPC levels that we detected in the cell extracts but not in the tumor extracts following 17AAG treatment.
Our finding that PC and PME levels increase during Hsp90 inhibition and arrest of cell growth following 17AAG treatment was unexpected. Indeed, in many previous preclinical and clinical studies (23,35,42-46), PME levels are high in rapidly proliferating tumors and decrease after tumor response to standard chemotherapy or radiotherapy treatment. The time scale in which these changes in PME levels were previously observed (45,46) is similar to the time scale of the changes seen in this study; hence, the increased PME levels that we found are unlikely to be a transient effect. Therefore, our findings with 17AAG treatment cannot be a general consequence of tumor growth inhibition. One possible explanation for the rise in PME levels during tumor growth is that they may indicate membrane synthesis in rapidly dividing cells (42) and that growth inhibition by conventional cytotoxic agents lowers the PC level by inhibiting proliferation. Another possible explanation involves the effects of oncogenic mutations, particularly activation of the Ras signaling pathways (47,48), on phospholipid metabolism. Our previous results (10-12) have shown that the Ras/Raf/mitogen-activated protein kinase kinase/extracellular signal-regulated kinase (Ras/Raf/MEK/ERK) and phosphatidylinositol-3-kinase/serine/threonine protein kinase (PI3K/AKT) pathways are blocked following inhibition of Hsp90 by 17AAG. Whatever the explanation, our finding that 17AAG treatment caused an increase in PC and PME levels is unusual and could have important implications for the understanding of the mechanism of action for 17AAG.
The elevated PC level we observed in vitro and the elevated PME level we observed in vivo could, in principle, arise in three possible ways: 1) increased uptake and phosphorylation of extracellular choline to form PC, 2) de novo synthesis of PC, or 3) mobilization of MR-invisible cholines to PC. Hypothesis 1 is unlikely in view of the study of Liu et al. (49), who showed that geldanamycin inhibits choline kinase and net cellular accumulation of (methyl-14C)choline and (methyl-14C)phosphocholine. Hypothesis 2 is also unlikely, because synthesis of PC in mammals is thought to be confined to the liver. Therefore, hypothesis 3 is the most probable explanation for the elevated levels of PC in vitro and PME in vivo. Hence, the most likely occurrence is that 17AAG promotes the hydrolysis of an MR-invisible pool of phosphatidylcholine, causing an increase in PC both in cell extracts and in vivo. In addition, the increase in intracellular PC might also inhibit choline kinase and thus impede the uptake of extracellular choline, which could explain the results of Liu et al.
Activation of pathways downstream of tyrosine receptor kinases and Ras has been associated with several alterations in enzymes associated with choline metabolism and phosphatidylcholine synthesis (50-52). Choline kinase activation, inhibition of cytidine triphosphate-phosphocholine cytidyltransferase, and activation of phospholipases A2, C, and D have all been reported at different steps in the Ras/Raf/MEK/ERK, PI3K/AKT, and Ral-guanine nucleotide dissociation stimulator signaling pathways (50-55). By depleting proteins involved in those signaling pathways, 17AAG treatment probably affects the enzymes involved in choline metabolism and alters the balance between synthesis and hydrolysis of phosphatidylcholine.
In conclusion, the present study has identified changes in phospholipid metabolism associated with inhibition of the Hsp90 molecular chaperone by 17AAG in human colon cancer cells. The effects of 17AAG treatment on phospholipid metabolites were found to occur both in cell lines treated in culture and in tumor xenografts in mice. The increased PME/PDE ratio was found to correlate with tumor response, indicating that it could act as a potential predictive marker. The treatment-induced changes were observed at 17AAG doses that caused tumor growth inhibition via inhibition of Hsp90, as demonstrated by changes in client protein and Hsp70 levels. Hence, monitoring the pharmacodynamic effects of 17AAG treatment and possibly of other Hsp90 inhibitors on phospholipid metabolism by MRSusing either 31P or the more sensitive 1Hmay provide a noninvasive surrogate marker for Hsp90 inhibition and potentially for tumor response in solid tumors in clinical trials with this drug and other Hsp90 inhibitors.
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NOTES |
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P. Workman is a Cancer Research UK Life Fellow.
Supported by Cancer Research UK grants C12/A1209, C12/A1212, and C1060/A808/G7643.
We thank the University of London Intercollegiate Research Service for the use of their NMR system at Birkbeck College and Dr. P. Ivy at the National Cancer Institute (Bethesda, MD) for providing 17AAG, 17AG, NSC683666, and EPL vehicle.
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Manuscript received February 6, 2003; revised August 27, 2003; accepted September 4, 2003.
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