Molecular Cancer Therapeutics CTRC-AACR San Antonio Breast Cancer Symposium Bridging the Lab and the Clinic in Cancer Medicine
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Cancer Research Clinical Cancer Research
Cancer Epidemiology Biomarkers & Prevention Molecular Cancer Therapeutics
Molecular Cancer Research Cancer Prevention Research
Cancer Prevention Journals Portal Cancer Reviews Online
Annual Meeting Education Book Meeting Abstracts Online
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Sankaranarayanapillai, M.
Right arrow Articles by Ronen, S. M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sankaranarayanapillai, M.
Right arrow Articles by Ronen, S. M.
Mol Cancer Ther. 2006;5:1325-1334
© 2006 American Association for Cancer Research

Research Articles: Therapeutics

Detection of histone deacetylase inhibition by noninvasive magnetic resonance spectroscopy

Madhuri Sankaranarayanapillai1, William P. Tong1, David S. Maxwell1, Ashutosh Pal1, Jihai Pang2, William G. Bornmann1, Juri G. Gelovani1 and Sabrina M. Ronen1

1 Experimental Diagnostic Imaging and 2 Pharmaceutical Development Center, The University of Texas M.D. Anderson Cancer Center, Houston, Texas

Requests for reprints: Sabrina M. Ronen, Experimental Diagnostic Imaging, The University of Texas M.D. Anderson Cancer Center, 57-3D, 1515 Holcombe Blvd., Houston, TX 77030-4009. Phone: 713-563-4617; Fax: 713-563-4894. E-mail: sronen{at}di.mdacc.tmc.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Histone deacetylase (HDAC) inhibitors are new and promising antineoplastic agents. Current methods for monitoring early response rely on invasive biopsies or indirect blood-derived markers. Our goal was to develop a magnetic resonance spectroscopy (MRS)–based method to detect HDAC inhibition. The fluorinated lysine derivative Boc-Lys-(Tfa)-OH (BLT) was investigated as a 19F MRS molecular marker of HDAC activity together with 31P MRS of endogenous metabolites. In silico modeling of the BLT-HDAC interaction and in vitro MRS studies of BLT cleavage by HDAC confirmed BLT as a HDAC substrate. BLT did not affect cell viability or HDAC activity in PC3 prostate cancer cells. PC3 cells were treated, in the presence of BLT, with the HDAC inhibitor p-fluoro-suberoylanilide hydroxamic acid (FSAHA) over the range of 0 to 10 µmol/L, and HDAC activity and MRS spectra were monitored. Following FSAHA treatment, HDAC activity dropped, reaching 53% of control at 10 µmol/L FSAHA. In parallel, a steady increase in intracellular BLT from 14 to 32 fmol/cell was observed. BLT levels negatively correlated with HDAC activity consistent with higher levels of uncleaved BLT in cells with inhibited HDAC. Phosphocholine, detected by 31P MRS, increased from 7 to 16 fmol/cell following treatment with FSAHA and also negatively correlated with HDAC activity. Increased phosphocholine is probably due to heat shock protein 90 inhibition as indicated by depletion of client proteins. In summary, 19F MRS of BLT, combined with 31P MRS, can be used to monitor HDAC activity in cells. In principle, this could be applied in vivo to noninvasively monitor HDAC activity. [Mol Cancer Ther 2006;5(5):1325–34]


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Acetylation and deacetylation of nucleosomal core histones play an important role in the modulation of chromatin structure and the regulation of gene expression. The acetylation status of histones is controlled by the activities of histone acetyltransferases and histone deacetylases (HDAC), which, respectively, catalyze removal or addition of acetyl groups onto the {varepsilon}-amino of lysine residues in the histone tail. Disruptions in HDACs and histone acetyltransferases have been associated with cancer development (1, 2). Conversely, it has been shown that HDAC inhibitors (HDACI) lead to differentiation, growth arrest, and/or apoptosis in treated cells and tumors (1, 3–6). Consequently, HDACIs are currently in clinical trials and show promising results in several different tumor types (1, 2, 7–12). The exact mechanism of action of HDACIs is not entirely clear. Reactivation of silenced tumor suppressor genes occurs following HDAC inhibition in some cases (13, 14). However, HDACIs can lead not only to gene stimulation but also to gene repression (15, 16). Nonetheless, many of the modulated genes mediate proliferation, cell cycle progression, or apoptosis and include p21/WAF1, caspases, p53, vascular endothelial growth factor, HER-2/neu, and bcr/abl (17–21). In addition, acetylation of nonhistone proteins is likely involved in the activity of HDACIs, and HDAC substrates include pRb, E2F, and heat shock protein (Hsp) 90 (22).

As mentioned, several HDACIs are currently in clinical trials. However, at present, there is no direct noninvasive means to measure drug delivery to the tumor tissue, drug-target interaction, or molecular response. Response to HDACIs in clinical trials is correlated with acetylation of peripheral blood mononuclear cells or acetylation of histones in tumor biopsy specimens (7, 9, 10, 12). Blood tests are well tolerated by patients but provide only an indirect indicator of drug delivery and activity at the tumor site. Biopsies reliably assess drug action but are surgically invasive. A further difficulty is that response in many cases is associated with tumor stasis, rather than shrinkage (7), limiting the use of traditional imaging methods. Determining the appropriate, biologically relevant, drug dose and assessing drug action at the tumor site, in vivo, present a challenge (23, 24). A noninvasive method of assessing drug delivery to, and the effect on, the intended molecular target is therefore needed.

Magnetic resonance spectroscopy (MRS) presents a noninvasive nondestructive method, which can provide longitudinal pharmacokinetic and pharmacodynamic biomarkers of drug delivery and action at defined anatomic locations in individual cancer patients. 19F MRS has been used in studies of fluorinated chemotherapeutic agents in cells, animal models, and patients (25–27) and also provides a tool to assess different physiologic variables, including oxygenation, pH, and gene expression (28–30). In addition, MRS can monitor changes in cellular metabolites that are associated with clinical response to traditional chemotherapy or radiotherapy. An increase in choline-containing metabolites, as detected using either 31P or 1H MRS, is associated with cell transformation, and a drop in those metabolites is typically associated with response to treatment (31–33). More recently, 31P MRS has been used to identify biomarkers of response to novel targeted therapies. We have shown that MRS can detect several metabolic changes associated with inhibition of the mitogen-activated protein kinase pathway (34, 35) and the phosphatidylinositol 3-kinase pathway (36) as well as response to the Hsp90 inhibitor 17-allylamino-17-demethoxygeldanamycin (17AAG; refs. 37, 38).

Several of the genes and proteins modulated by HDAC inhibition are expected to lead to MRS detectable changes. These include down-regulation of receptor tyrosine kinases and their downstream effector molecules (16, 17, 20, 21, 39), which have been associated with a drop in phosphocholine (34, 36, 40); modulation of p53 (19), which could affect phosphocholine levels (41); and Hsp90 acetylation and inhibition (22, 42, 43), which could lead to increased phosphocholine and glycerophosphocholine (37). Thus, 31P MRS could be used to monitor metabolic changes associated with inhibition of HDAC. However, any metabolic changes observed in the 31P spectrum represent indirect and often nonspecific downstream events. Specific direct indicators of drug activity on the intended molecular target are therefore needed to complement the downstream metabolic changes.

We show here for the first time the use of a fluorinated HDAC substrate, the lysine derivative Boc-Lys-(Tfa)-OH (BLT), as a specific spectroscopic indicator to directly monitor HDAC inhibition. We show that intracellular BLT levels, as determined by 19F MRS, are negatively correlated with HDAC activity. In addition, using 31P MRS, we show that phosphocholine levels also negatively correlate with HDAC activity. Our results indicate that MRS can be used as a method for assessing HDAC inhibition and its downstream signaling and metabolic effects. This methodology could eventually be translated to the clinic where drug delivery and molecular activity would be monitored noninvasively over time leading to optimized drug scheduling and dosing.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
In silico Modeling of BLT Docking into HDAC
Docking was done using the FlexX 1.13.5 software as provided in Sybyl7.1 (Tripos, Inc., St. Louis, MO) running on a 4-Processor R16000 SGI Tezro. Unless otherwise noted, all defaults were used in the docking experiment. The structure of BLT was drawn into Sybyl using the Sketch module, and types were modified to correspond to the protonated state at pH 7. Charges were not assigned to the molecule, because FlexX uses formal charges that are assigned during the actual run. The crystal structure of HDAC8 complexed with suberoylanilide hydroxamic acid (SAHA) was retrieved from the RCSB (ID code 1T69; ref. 44). The binding site was then defined as 6.5 Å around the SAHA ligand. In the customized setting, the zinc atom was added as a template. For H142 and H143, the histidines were selected to be in the {delta} protonated state. This particular protonation state was found to be necessary for proper docking of the hydroxamic acid–based inhibitors. The CSCORE method (45) was used in the ranking of the 30 requested configurations.

MRS Studies of BLT Cleavage
To confirm that MRS can be used to monitor cleavage of the HDAC substrate, 0.6 mmol/L BLT (Advance Chem-Tech, Louisville, KY) was incubated alone or with 28 units recombinant HDAC-8 (Biomol, Plymouth Meeting, PA) in a total volume of 500 µL HDAC assay buffer (Biomol; composed of 25 mmol/L Tris-HCl, 137 mmol/L NaCl, 2.7 mmol/L KCl, 1 mmol/L MgCl2, formulated to maintain HDAC activity). 19F MRS was used to monitor the decrease in BLT and buildup of the cleavage product TFA by acquiring 1H-decoupled spectra at 10-minute intervals on an Avance DPX300 Bruker spectrometer (Bruker Biospin, Rheinstetten, Germany) using a 30° flip angle, 3-second relaxation delay, and 128 scans. A sealed insert containing C6F6 served as a quantification and chemical shift reference (–164.9 ppm relative to CFCl3).

Cell Culture, Cell Proliferation, and HDAC Activity
PC3 human prostate cancer cells were routinely cultured in DMEM/F-12 (Life Technologies, Grand Island, NY) supplemented with 10% FCS (Hyclone, Logan, UT) and 10,000 units/mL penicillin, 10,000 µg/mL streptomycin, and 25 µg/mol amphotericin B (Life Technologies) at 37°C in 5% CO2.

To assess the effect on cell proliferation of BLT, the colorimetric WST-1 cell proliferation assay (Roche, Indianapolis, IN) was used following the manufacturer's instructions. Briefly, 20 x 103 cells per well were seeded in 96-well plates. Cells were then treated for 24 hours with BLT at 5, 10, 20, 50, 100, 200, 500 µmol/L and 1, 2, 5, and 10 mmol/L or with matched DMSO (1:20,000 to 1:10). Subsequently, cells were incubated for 4 hours with the WST-1 cell proliferation reagent and absorbance was read at 440 nm using a Tecan Freedom Evo liquid handler equipped with the SAFIRE monochromator-based microplate reader (Tecan U.S., Research Triangle Park, NC).

The effect on cell proliferation of p-fluoro-SAHA (FSAHA; synthesized in-house based on a previously described method; ref. 46), the fluorinated derivative of the clinically relevant HDACI SAHA, was also determined using the WST-1 assay as described above. Cells were treated with 2, 5, and 10 µmol/L FSAHA either in the presence of 1 mmol/L BLT or in the presence of DMSO (in which BLT had to be dissolved first due to its low solubility in growth medium). Note that controls were treated with DMSO to clearly identify effects that are due to the compound being investigated rather than its vehicle (DMSO). FSAHA dose was based on previous findings (4).

The effect on HDAC activity of BLT and the HDACIs FSAHA and SAHA (also synthesized in-house based on the previously described method; ref. 46) was determined using the Fluor-de-Lys fluorometric assay (Biomol) following the manufacturer's instructions. Briefly, 20 x 103 cells per well were seeded in 96-well plates and incubated for 24 hours with (a) 1 mmol/L BLT, (b) 2, 5, and 10 µmol/L FSAHA, (c) 2, 5, and 10 µmol/L SAHA, (d) 2, 5, 7, 8, 9, and 10 µmol/L FSAHA in the presence of 1 mmol/L BLT, and (e) 2, 5, 7, 8, 9, and 10 µmol/L FSAHA in the presence of vehicle (DMSO). The Fluor-de-Lys substrate was then added for 1 hour, medium was removed, cells were rinsed with PBS and incubated for 10 minutes with the Fluor-de-Lys developer, and fluorescence was read at 460 nm using the Tecan microplate reader as above. Results were normalized to cell density as determined using the WST-1 assay in the same 96 well plate.

MRS Studies of HDAC Activity
For MRS studies, PC3 cells were treated for 24 hours with FSAHA at 2, 5, 7, 8, 9, and 10 µmol/L in the presence of 1 mmol/L BLT or with 1 mmol/L BLT alone. Cells (~1 x 107–1.5 x 107) were then extracted using the dual-phase extraction method as described previously (37, 47). Briefly, cells were extensively rinsed with ice-cold saline to remove any residual extracellular BLT and medium. Cells were then fixed in 10 mL ice-cold methanol, scraped off the surface of the culture flask, collected into glass tubes, and vortexed. Ice-cold chloroform (10 mL) was then added followed by 10 mL ice-cold deionized water. Following phase separation and solvent removal, the water-soluble fraction was reconstituted in 250 µL deuterium oxide and 250 µL DMSO for 19F magnetic resonance measurements. To perform the 31P magnetic resonance measurement, 100 µL EDTA and 50 µL methylene diphosphonic acid in deuterium oxide were added to a final concentration of 10 and 0.35 mmol/L, respectively. The number of cells extracted was determined by counting a separate flask of cells. 19F MRS of the water-soluble metabolites were recorded as above. Metabolite concentrations were determined by integration and comparison with the area of the external C6F6 reference, normalizing to cell number and correcting for saturation effects. Correction for saturation effects was achieved by also acquiring a quantitative inverse gated fully relaxed spectrum on two different samples and calculating the correction factors that need to be applied to the partially relaxed spectra. We determined that the T1 relaxation of BLT is 1 second and that of C6F6 is 3 seconds. The fully relaxed spectrum was therefore acquired using a 90° flip angle and a 15-second relaxation delay (5 times the longest T1). 31P MR spectra were recorded on an Avance DRX500 Bruker spectrometer (Bruker Biospin) using a 30° flip angle and 3-second relaxation delay. Metabolite concentrations were determined by integration and comparison with the area of the internal methylene diphosphonic acid reference, normalizing to cell number and correcting for saturation effects (correction factors were determined as above by acquiring a fully relaxed quantitative spectrum using a 90° flip angle and a 30-second relaxation delay).

To monitor the fluorinated metabolites, the lipid phase was reconstituted in 500 µL CDCl3. To monitor the fluorinated metabolites in cellular protein, the protein pellet obtained during cell extraction was dissolved in 1 mL of 0.5 mol/L NaOH and heated to 60°C for 1 hour. Samples were then analyzed by 19F MRS as above.

Analysis of TFA in Extracellular Medium
To assess buildup of TFA in medium, samples of extracellular medium were collected and analyzed using gas chromatography-mass spectroscopy. To be able to use standard capillary gas chromatography-mass spectroscopy, TFA had to be derivatized. This was done using a modification of the procedure of Scott et al. (48) as follows. Sample (1 mL) or TFA standard in medium was treated with 20 mg NaCl, 60 µL HCl, and 30 µL each of 100 mmol/L 1,3-dicyclohexylcarbohiimide (Sigma-Aldrich Co., St. Louis, MO) and 2,4-difluoroaniline (Sigma-Aldrich) and adjusted to a final volume of 1.320 mL in ethyl acetate. Mixture was vortexed for 60 minutes at room temperature, an additional 50 mg NaCl was added, and sample was briefly vortexed. Following phase separation, the organic layer was removed and stored. The aqueous phase was further extracted twice with 200 µL ethyl acetate. All three ethyl acetate extracts were combined, treated with 50 µL of 3 mol/L HCl and 50 µL saturated anhydrous sodium sulfate, and vortexed and another 20 mg anhydrous sodium sulfate was added and mixed. The organic phase was then evaporated to dryness under a dry nitrogen gas stream. The residue was dissolved in 250 µL toluene and transferred into sample vials for analysis. Sample extract (3 µL) was analyzed using an Agilent (Wilmington, DE) 6890N GC coupled to 5973N MSD in splitless injection mode. TFA was resolved using a Supleco (St. Louis, MO) Omegawax 250 capillary column (30 m x 0.25 mm). Column temperature was 100°C for 1 minute, ramping to 230°C at 25°C/min, and then held at 230°C for 2 minutes. Detection was done in EI-positive mode monitoring the m/z 225 ion.

Western Blot Analysis of Protein Levels
PC3 cells were lysed using cell lysis buffer [0.1% NP40, 50 mmol/L HEPES (pH 7.4), 250 mmol/L NaCl, 1 mmol/L DTT, 1 mmol/L EDTA, 1 mmol/L NaF, 10 mmol/L ß-glycerophosphate, 0.1 mmol/L sodium orthovanadate, 1 µL/mL protease inhibitor cocktail set III (Calbiochem, La Jolla, CA), 1 mmol/L phenylmethylsulfonyl fluoride]. Lysates were centrifuged at 12,000 rpm for 10 minutes (4°C), the protein supernatant was collected, and total protein concentrations were determined using Bio-Rad DC protein assay reagents (Bio-Rad, Hercules, CA). Proteins were separated by SDS-PAGE using 10% gels and transferred electrophoretically to 0.45 µm nitrocellulose membranes. Membranes were blocked in blocking buffer containing 5% nonfat dry milk in TBS (pH 7.6) and 0.1% Tween 20 and incubated overnight at 4°C with primary antibodies as follows: c-Raf (1:1,000; Cell Signaling Technology, Danvers, MA), cyclin-dependent kinase 4 (cdk4; 1:2,000; Cell Signaling Technology), Hsp70 (1:2,000; Stressgen, Victoria, British Columbia, Canada), and glyceraldehyde-3-phosphate dehydrogenase (1:5,000; Stressgen). This was followed by 1-hour incubation with horseradish peroxidase–conjugated secondary anti-rabbit (Cell Signaling Technology) and anti-mouse (Cell Signaling Technology) antibodies at dilutions of 1:1,000 and 1:2,000, respectively. Membranes were washed with enhanced chemiluminescence reagents (LumiGLO & Peroxide, Cell Signaling Technology) for 1 minute and exposed to Hyperfilm (Amersham Biosciences, Piscataway, NJ), which was developed on a Konica SRX-101 automatic developer (Konica, Tokyo, Japan).

Statistical Analysis
All results represent the average of at least three experiments and are expressed as mean ±SD. Data were analyzed by Wilcoxon-Mann-Whitney rank test and P < 0.05 was considered significant. A Spearman's rank correlation was used to analyze correlations. KaleidaGraph (Synergy Software, Essex Junction, VT) and Statistica (Statsoft, Tulsa, OK) software were used.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
BLT Is a HDAC Substrate
First, it was necessary to test the hypothesis that a fluorinated compound composed of a modified lysine could serve as a MRS-detectable substrate of HDAC and could therefore be used to assess HDAC inhibition. We investigated the commercially available BLT and first performed computer modeling to estimate the BLT-HDAC interaction. The known structure of HDAC8 was used (44). The top-ranking consensus scored configuration of BLT docked with HDAC8 is shown in Fig. 1 . The interaction of the carbonyl group with the Y306 is consistent with the proposed model of how acetylated lysine would interact in the catalytic site (49). In the docked structure, there is a hydrogen bond to the backbone carbonyl of G151 and a hydrogen bond between side chain NH and D101. The aliphatic side chain interacts with two phenylalanine groups (F152 and F208) and the interaction is seen in the complex of the SAHA as well (44). The Boc group consistently selected to position nearby F207 and F208 rather than the area occupied in the reported complex with SAHA. This may result from the asymmetry that is present within the BLT and not within SAHA, which forces the choice between optimizing the orientation of the carboxyl group and the Boc group.


Figure 1
View larger version (45K):
[in this window]
[in a new window]
  		Figure 1. HDAC8 crystal structure with docked BLT into catalytic site. Hydrogen bonds are designated with a line. Red, oxygen; blue, nitrogen; pink, fluorine. The element zinc is gold. Inset, BLT.

 
Next, 19F MRS was used to confirm that recombinant HDAC8 does indeed cleave BLT in vitro. As illustrated in Fig. 2 , a drop in BLT levels was detected over time accompanied by an increase in TFA levels, consistent with cleavage of BLT by HDAC8 to form TFA and the 19F magnetic resonance invisible Boc-Lys. No changes in BLT levels or any buildup of TFA could be detected in the control sample, which contained no HDAC8. This confirmed that BLT is indeed a substrate of HDAC8 and that its cleavage by HDAC can be monitored by MRS.


Figure 2
View larger version (14K):
[in this window]
[in a new window]
  		Figure 2. Sequential 19F MRS spectra of BLT in the presence of recombinant HDAC8 in vitro. Spectra are the result of 128 1H-decoupled scans acquired using a 30° flip angle and a 3-second relaxation delay. Spectra show a drop in BLT as it is cleaved by HDAC8 to form TFA. Inset, full time course of the experiment.

 
BLT Does Not Affect Cell Viability or HDAC Activity
Before using BLT as a marker of HDAC activity in cells, it was necessary to rule out its toxicity. The WST-1 assay was used to investigate the effect on PC3 cells of a range of BLT concentrations from 5 µmol/L to 10 mmol/L compared with matched DMSO controls. BLT did not significantly affect cell proliferation compared with controls up to a concentration of 10 mmol/L (P < 0.03 for 10 mmol/L and P > 0.4 for all other concentrations from 5 µmol/L to 5 mmol/L; data not shown). We therefore chose to perform MRS experiments with a BLT concentration of 1 mmol/L, which was expected to lead to a MRS detectable signal. Cell numbers following 24-hour treatment with 1 mmol/L BLT represented 95 ± 4% of controls (P > 0.1).

Next, it was necessary to confirm that 1 mmol/L BLT did not affect HDAC activity in cells. Using the Fluor-de-Lys assay, we determined that incubation of PC3 cells for 24 hours with 1 mmol/L BLT resulted in HDAC activity levels of 102 ± 9% (P > 0.3) relative to DMSO-treated controls, indicating no statistically significant effect of 1 mmol/L BLT on HDAC activity.

Inhibition of HDAC Activity and Cell Proliferation by HDACI Treatment Is Not Affected by the Addition of BLT
Before using MRS of BLT to assess the effect of HDACIs, it was necessary to confirm that the addition of BLT will not modify the biological effects of the HDACI. To this end, we investigated both cell proliferation and HDAC activity. Figure 3A illustrates our findings. Treatment with FSAHA, the fluorinated derivative of the HDACI SAHA, resulted in a significant drop in cell proliferation relative to control at all three doses investigated down to 87 ± 5% at 2 µmol/L, 79 ± 3% at 5 µmol/L, and 66 ± 6% at 10 µmol/L (P < 0.03 for all three doses). Importantly, the presence of BLT did not further affect cell proliferation. In the presence of BLT, cell proliferation dropped to 86 ± 1% at 2 µmol/L, 77 ± 7% at 5 µmol/L, and 63 ± 2% at 10 µmol/L (P < 0.03 relative to controls and P > 0.5 relative to FSAHA). Figure 3B illustrates the effect on HDAC activity of FSAHA and FSAHA in the presence of 1 mmol/L BLT. The lower concentration of FSAHA (2 µmol/L) did not lead to a statistically significant drop in HDAC activity (93 ± 16%), but the higher concentrations of FSAHA resulted in significant inhibition of HDAC activity (58 ± 14% at 5 µmol/L and 41 ± 8% at 10 µmol/L; P < 0.03). Again, the presence of BLT did not significantly alter the effect of FSAHA treatment (down to 88 ± 12%, 69 ± 9%, and 53 ± 5% for 2, 5, and 10 µmol/L, respectively; P > 0.1 relative to FSAHA).


Figure 3
View larger version (34K):
[in this window]
[in a new window]
  		Figure 3. Effect on cell proliferation (A) and HDAC activity (B) of HDACI treatment. PC3 cells were treated with 2, 5, and 10 µmol/L FSAHA or with FSAHA in the presence of 1 mmol/L BLT (FSAHA + BLT). Cell proliferation was determined using the WST-1 assay and HDAC activity was determined using the Fluor-de-Lys assay. Bars, SD. *, P < 0.05, compared with DMSO-treated controls. n.s., not significant (P > 0.05).

 
Because SAHA is being investigated in clinical trials, it was necessary to confirm that the biological effects of its fluorinated derivative FSAHA were comparable with those of SAHA. The effects of SAHA and FSAHA on HDAC activity were compared at 2, 5, and 10 µmol/L. The effect of SAHA on HDAC activity was comparable with that observed with FSAHA at 2 and 5 µmol/L (84 ± 10% at 2 µmol/L and 51 ± 3% at 5 µmol/L; P > 0.7 relative to FSAHA). However, treatment with 10 µmol/L SAHA resulted in a greater inhibition of HDAC activity compared with 10 µmol/L FSAHA (29 ± 3%; P < 0.03 relative to FSAHA). Nonetheless, we reasoned that if MRS can detect the effect of FSAHA on HDAC activity, it would also be able to detect the potentially larger effect of SAHA.

19F MRS of BLT Can Be Used to Assess HDAC Inhibition in Cells
Figure 4A illustrates the 19F MRS spectra recorded from extracts of PC3 cells. Control cells cultured in the presence of 1 mmol/L BLT contained 14 ± 4 fmol/cell BLT. This value increased significantly to 32 ± 4 fmol/cell in cells treated with 10 µmol/L FSAHA in the presence of BLT (P < 0.0002), consistent with intracellular uncleaved BLT levels being higher when HDAC is inhibited. No signal could be detected from FSAHA, which was expected to resonate at –121 ppm. In addition, no 19F signal was observed from the lipid phase or the protein pellet of the cells.


Figure 4
View larger version (10K):
[in this window]
[in a new window]
  		Figure 4. A, representative 1H-decoupled 19F MR spectra of PC3 cell extracts. Cells were treated for 24 h with 10 µmol/L FSAHA in the presence of 1 mmol/L BLT (top) or with 1 mmol/L BLT alone (bottom). Average BLT content in the FSAHA-treated cells was 32 fmol/cell compared with 14 fmol/cell in controls. Spectra are the result of 128 scans acquired using a 30° flip angle and a 3-second relaxation delay. Reference (Ref) was C6F6. B, intracellular BLT levels as a function of HDAC inhibition. BLT levels were determined by 19F MRS as in A. HDAC inhibition was determined using the Fluor-de-Lys assay. Bars, SD. Line, best linear fit. Spearman's rank correlation indicates that BLT levels negatively correlated with HDAC activity ({rho} = –0.75; P < 0.05).

 
Interestingly, TFA levels observed in both treated and control cells remained, within experimental error, unchanged. The average TFA concentration observed in control cells was 1.1 ± 0.6 versus 1 ± 1 fmol/cell in cells treated with 10 µmol/L FSAHA. We therefore speculated that TFA produced inside the cell was removed into the extracellular compartment. To test this hypothesis, gas chromatography-mass spectroscopy was used to determine the levels of TFA present in cellular growth medium. In control cells, the level of TFA was 5 ± 0.4 µg/mL but was only 3.4 ± 0.7 µg/mL in the medium obtained from 10 µmol/L FSAHA-treated cells (P < 0.03).

To confirm that the intracellular BLT levels detected by MRS are indicative of inhibition of cellular HDAC, we next monitored HDAC activity and cellular BLT levels in cells treated with a range of FSAHA concentrations from 2 to 10 µmol/L. HDAC activity dropped significantly for all concentrations greater than 2 µmol/L. An increase in BLT levels was observed for all FSAHA concentrations investigated and reached statistical significance when HDAC activity dropped to 74% relative to controls. Furthermore, a negative correlation was observed between HDAC activity and the level of intracellular BLT ({rho} = –0.75; P < 0.05; Fig. 4B).

31P MRS Can Be Used to Assess HDAC Inhibition in Cells
Because HDAC inhibition leads to modulation of several genes that are associated with MRS detectable metabolic changes, we questioned whether response to treatment with HDACIs is also detectable in the 31P MRS of treated cells. 31P MRS was used to investigate the same PC3 samples investigated by 19F MRS. As illustrated in Fig. 5A , treatment with 10 µmol/L FSAHA resulted in a significant increase in phosphocholine levels from 7 ± 1 to 16 ± 2 fmol/cell (P < 0.01). None of the other metabolites observed in the 31P MRS spectrum were altered. As in BLT, phosphocholine levels for cells treated with lower concentrations of FSAHA also showed an increase relative to controls, reaching statistical significance when HDAC activity dropped to 74% relative to controls. Phosphocholine levels also negatively correlated with HDAC activity ({rho} = –0.86; P < 0.02; Fig. 5B).


Figure 5
View larger version (11K):
[in this window]
[in a new window]
  		Figure 5. A, representative 1H-decoupled 31P MR spectra of PC3 cell extracts. Cells were treated for 24 h with 10 µmol/L FSAHA in the presence of 1 mmol/L BLT (top) or with 1 mmol/L BLT alone (bottom). Average phosphocholine (PC) content in the FSAHA-treated cells was 15 fmol/cell compared with 7 fmol/cell in controls. Spectra are the result of 3,000 scans acquired using a 30° flip angle and a 3-second relaxation delay. Reference was methylene diphosphonic acid. B, phosphocholine levels as a function of HDAC inhibition. Phosphocholine levels were determined by 31P MRS as in A. HDAC inhibition was determined using the Fluor-de-Lys assay. Bars, SD. Line, best linear fit. Spearman's rank correlation indicates that phosphocholine levels negatively correlated with HDAC activity ({rho} = –0.86; P = 0.02).

 
31P MRS Changes Are Consistent with Depletion of Hsp90 Client Proteins cdk4 and c-Raf-1
The increase in phosphocholine observed in our spectra following response to HDACI treatment is an unusual observation and only previously reported following treatment with the Hsp90 inhibitor 17AAG (37). Hsp90 inhibition has also been reported following HDACI treatment. To test the hypothesis that the 31P MRS changes observed here were associated with inhibition of Hsp90, we monitored the levels of the Hsp90 client proteins c-Raf-1 and cdk4. Figure 6 indicates depletion of these two Hsp90 client proteins following treatment with FSAHA. However, induction of Hsp70, which has also been reported following inhibition of 17AAG, was not observed in our cells (Fig. 6).


Figure 6
View larger version (32K):
[in this window]
[in a new window]
  		Figure 6. Representative Western blot analysis of c-Raf-1, cdk4, Hsp70, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH). PC3 cells were treated with vehicle (DMSO), 1 mmol/L BLT, or 10 µmol/L FSAHA in the presence of 1 mmol/L BLT. Glyceraldehyde-3-phosphate dehydrogenase served as a loading and transfer control.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The recent shift to development of antineoplastic therapies that target specific alterations in cancerous cells has been fueled by the clinical successes of tyrosine kinase inhibitors, such as imatinib, gefinitib, and trastuzumab (50, 51). HDACIs are a novel class of anticancer drugs that target HDACs. Modulations in gene expression resulting from the activities of HDACs and their counterpart histone acetyltransferases have been associated with cell transformation (52, 53). Conversely, response to HDACIs has been associated with alterations in expression of several oncogenes and results in induction of differentiation, growth arrest, or apoptosis (1–3, 16, 54). Response to HDACIs in clinical trials has been promising (1, 2, 7, 11, 12). Unfortunately, current methods to monitor response in vivo rely on either surgically invasive biopsies or blood-derived markers (7, 12). There is therefore a need for a direct, reliable, and noninvasive method to monitor drug delivery and response to HDACIs at the tumor site.

Here, we have investigated the fluorinated lysine derivative BLT as a MRS marker of HDAC activity. BLT is detectable by 19F MRS and its cleavage by HDAC was expected to produce TFA and Boc-Lys. In silico and in vitro studies confirmed that BLT is a substrate of HDAC8 and therefore most likely a substrate of other class I and II HDACs. BLT did not affect cell viability or HDAC activity. Most importantly, we show here that the intracellular levels of BLT, as measured by 19F MRS, are correlated with cellular HDAC activity.

Fluorine in the body is in the form of solid fluorides with very short T2 relaxation times, producing wide and virtually nondetectable MRS peaks. In vivo 19F MRS therefore presents the advantage that there are no naturally observable fluorinated molecules. Consequently, exogenously administered fluorine-containing compounds are observed without interference (28). By introducing a fluorinated HDAC substrate, it is therefore straightforward to monitor its fate and thus assess HDAC activity directly in the target tissue.

We have shown previously that 31P MRS provides a noninvasive method for the detection of metabolic biomarkers associated with response to targeted therapies (34, 37). Here, we have applied this methodology to monitor the downstream metabolic effects correlated with HDAC inhibition, complementing the use of 19F MRS to monitor drug activity. We show that phosphocholine levels increase following HDAC inhibition and are correlated with the level of this inhibition. This provides a downstream metabolic biomarker of tumor response to HDACI treatment, further confirming activity of the drug on its target.

MRS is a noninvasive method that can be readily translated to the clinic. Our investigations therefore concentrated on a derivative of the clinically relevant HDACI SAHA (10). We chose to concentrate our studies on FSAHA, rather than SAHA, because we reasoned that if a significant level of FSAHA accumulates intracellularly it would be possible to simultaneously monitor both the delivery of FSAHA and its effect on HDAC activity by 19F MRS. FSAHA could not be detected in any of our spectra. Therefore, we conclude that the intracellular level of FSAHA is below MRS detection level (~0.1 fmol/cell). Current phase I trials (10) found that the mean plasma concentration of SAHA in vivo was <600 mg/mL, which is also expected to be below detection level.

The inhibitory effect of FSAHA was slightly lower at 10 µmol/L than that of SAHA. We have not investigated the reasons for this difference. Nonetheless, this observation does not affect the value of our MRS findings. Because we are able to detect the effects of the less potent fluorinated inhibitor, we believe we would also be able to detect the effects of the parent inhibitor SAHA or other HDACIs of equal or greater efficacy.

Surprisingly, TFA, the cleavage product of BLT, was constant in the intracellular compartment of control and HDACI-treated cells. However, an analysis of the extracellular medium showed that TFA was lower by an average 1.6 µg/mL in the medium of 10 µmol/L FSAHA-treated cells compared with controls. On average, intracellular BLT increased by 18 fmol/cell in those cells. If any TFA produced through cleavage of BLT by HDACs is removed into the extracellular medium, this would lead to TFA levels lower by 1.3 µg/mL in the medium of HDAC-inhibited cells. This number is consistent with the TFA levels determined experimentally in our extracellular medium. We conclude that TFA produced by cleavage of BLT is removed from the intracellular compartment into the medium, in line with previous findings (55). In vivo, depending on the rate of TFA clearance, it is possible that both BLT and TFA will be present in the tumor region. However, due to the small chemical shift difference between BLT and TFA, monitoring TFA is expected to be difficult.

In addition to directly assessing HDAC activity by 19F MRS of BLT, a unique metabolic biomarker of response was afforded by using 31P MRS to monitor the intrinsic cellular metabolites. Inhibition of cell growth following chemotherapeutic treatment as well as signaling inhibition are typically associated with a magnetic resonance visible drop in phosphocholine levels (31, 34–36, 56). The increase in phosphocholine observed here is therefore unusual and has been observed previously only following response to treatment with 17AAG (37, 38). 17AAG causes inhibition of Hsp90, which results in depletion of its client proteins, including cdk4 and c-Raf-1, as well as up-regulation of Hsp70 (57, 58). Interestingly, HDACI treatment results in increased Hsp90 acetylation also leading to inhibition of its activity and depletion of client proteins (39, 43, 59). In the specific case of SAHA, a drop in both cdk4 and c-Raf-1 has been observed in some cases (16) but not in others (60). Our Western blot analysis indicates depletion of both cdk4 and c-Raf-1 in treated cells. However, we did not observe any up-regulation of Hsp70. Thus, it is not entirely clear if the depletion of cdk4 and c-Raf-1 is a direct result of HDAC inhibition or occurs subsequent to Hsp90 acetylation following HDAC inhibition. Nonetheless, we believe that the increase in phosphocholine observed by MRS is associated with the depletion of cdk4 and c-Raf-1. The mechanism linking cdk4 and c-Raf-1 with modulation of phosphocholine remains to be elucidated, but the results described here following HDACI treatment are entirely consistent with our earlier observations (37). It should be noted that we have also previously observed an increase in glycerophosphocholine following response to 17AAG. In this study, this metabolite remained below detection level; thus, it is not clear if its levels are altered by HDACI treatment.

In future, these methods could be applied in vivo. Full toxicity studies of BLT still need to be done. However, preliminary experiments in our laboratory indicate that an i.p. injection of 100 mg/kg BLT once weekly on three consecutive weeks (a schedule consistent with monitoring response to HDACIs in vivo) results in no detectable toxicity to the animal. Importantly, this dose was sufficient to produce a MRS visible BLT signal in s.c. PC3 tumors with a temporal resolution of 5 minutes at 4.7 T. The BLT signal remained detectable within the tumor region for over 2 hours. As expected, the TFA peak could not be easily resolved in our preliminary in vivo studies. However, based on the data presented here, it is expected that intratumoral BLT levels will be higher in HDACI-treated tumors compared with controls; therefore, tumoral BLT levels could be used to assess HDAC activity in vivo. Future work in our laboratory is aimed at confirming this point. Downstream metabolic biomarkers could also be assessed in our preliminary studies. We were able to acquire a 31P spectrum from PC3 s.c. tumors in 30 minutes, providing a means for monitoring phosphocholine levels. This is consistent with previous studies (37) in which an increase in phosphocholine could be monitored as an indicator of response to 17AAG treatment. 1H MRS, with its greater sensitivity compared with 31P, could also be used to monitor the total choline signal as a downstream metabolic marker of response to HDAC inhibition.

In summary, we present here a dual method for noninvasively monitoring response to HDACIs. 19F MRS of the targeted molecular imaging agent BLT can be used to monitor delivery and activity of HDACIs at the tumor site, whereas 31P MRS can be used to monitor the downstream metabolic consequences of HDAC inhibition. Together, these two MRS methods provide both a direct marker of HDAC inhibition and a downstream biomarker of cellular response to the inhibition. We believe that combining both indicators provides a more powerful tool than a single marker alone, particularly at lower levels of HDAC inhibition when the changes observed in either marker alone are relatively small. The combination of 19F and 31P (or 1H) MRS could thus serve as a reliable noninvasive modality to assess HDAC inhibition.


   Acknowledgments
 
We thank Dr. Kumaralal K. Kaluarachchi for help in recording the nuclear magnetic resonance spectra, Dr. Edward Felix with the analytic methods, and Dr. Andrei Volgin and Galina Mikheeva for help with the biological assays and Western blots.


   Footnotes
 
Grant support: National Cancer Institute core institutional grant P30 CA016672 30 for the support of Core NMR Facility and Core Analytical Facility (The University of Texas M.D. Anderson Cancer Center).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 11/28/05; revised 1/20/06; accepted 3/ 8/06.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Marks P, Rifkind RA, Richon VM, Breslow R, Miller T, Kelly WK. Histone deacetylases and cancer: causes and therapies. Nat Rev Cancer 2001;1:194–202.[CrossRef][Medline]
  2. Drummond DC, Noble CO, Kirpotin DB, Guo Z, Scott GK, Benz CC. Clinical development of histone deacetylase inhibitors as anticancer agents. Annu Rev Pharmacol Toxicol 2005;45:495–528.[CrossRef][Medline]
  3. Marks PA, Richon VM, Miller T, Kelly WK. Histone deacetylase inhibitors. Adv Cancer Res 2004;91:137–68.[CrossRef][Medline]
  4. Butler LM, Agus DB, Scher HI, et al. Suberoylanilide hydroxamic acid, an inhibitor of histone deacetylase, suppresses the growth of prostate cancer cells in vitro and in vivo. Cancer Res 2000;60:5165–70.[Abstract/Free Full Text]
  5. Kuefer R, Hofer MD, Altug V, et al. Sodium butyrate and tributyrin induce in vivo growth inhibition and apoptosis in human prostate cancer. Br J Cancer 2004;90:535–41.[CrossRef][Medline]
  6. Fronsdal K, Saatcioglu F. Histone deacetylase inhibitors differentially mediate apoptosis in prostate cancer cells. Prostate 2005;62:299–306.[CrossRef][Medline]
  7. Kelly WK, Richon VM, O'Connor O, et al. Phase I clinical trial of histone deacetylase inhibitor: suberoylanilide hydroxamic acid administered intravenously. Clin Cancer Res 2003;9:3578–88.[Abstract/Free Full Text]
  8. Sandor V, Bakke S, Robey RW, et al. Phase I trial of the histone deacetylase inhibitor, depsipeptide (FR901228, NSC 630176), in patients with refractory neoplasms. Clin Cancer Res 2002;8:718–28.[Abstract/Free Full Text]
  9. Chavez-Blanco A, Segura-Pacheco B, Perez-Cardenas E, et al. Histone acetylation and histone deacetylase activity of magnesium valproate in tumor and peripheral blood of patients with cervical cancer. A phase I study. Mol Cancer 2005;4:22.[CrossRef][Medline]
  10. Kelly WK, O'Connor OA, Krug LM, et al. Phase I study of an oral histone deacetylase inhibitor, suberoylanilide hydroxamic acid, in patients with advanced cancer. J Clin Oncol 2005;23:3923–31.[Abstract/Free Full Text]
  11. Vigushin DM, Coombes RC. Histone deacetylase inhibitors in cancer treatment. Anticancer Drugs 2002;13:1–13.[CrossRef][Medline]
  12. Villar-Garea A, Esteller M. Histone deacetylase inhibitors: understanding a new wave of anticancer agents. Int J Cancer 2004;112:171–8.[CrossRef][Medline]
  13. Archer SY, Meng S, Shei A, Hodin RA. p21(WAF1) is required for butyrate-mediated growth inhibition of human colon cancer cells. Proc Natl Acad Sci U S A 1998;95:6791–6.[Abstract/Free Full Text]
  14. Blagosklonny MV, Robey R, Sackett DL, et al. Histone deacetylase inhibitors all induce p21 but differentially cause tubulin acetylation, mitotic arrest, and cytotoxicity. Mol Cancer Ther 2002;1:937–41.[Abstract/Free Full Text]
  15. Glaser KB, Staver MJ, Waring JF, Stender J, Ulrich RG, Davidsen SK. Gene expression profiling of multiple histone deacetylase (HDAC) inhibitors: defining a common gene set produced by HDAC inhibition in T24 and MDA carcinoma cell lines. Mol Cancer Ther 2003;2:151–63.[Abstract/Free Full Text]
  16. Mitsiades CS, Mitsiades NS, McMullan CJ, et al. Transcriptional signature of histone deacetylase inhibition in multiple myeloma: biological and clinical implications. Proc Natl Acad Sci U S A 2004;101:540–5.[Abstract/Free Full Text]
  17. Richon VM, Sandhoff TW, Rifkind RA, Marks PA. Histone deacetylase inhibitor selectively induces p21WAF1 expression and gene-associated histone acetylation. Proc Natl Acad Sci U S A 2000;97:10014–9.[Abstract/Free Full Text]
  18. Almenara J, Rosato R, Grant S. Synergistic induction of mitochondrial damage and apoptosis in human leukemia cells by flavopiridol and the histone deacetylase inhibitor suberoylanilide hydroxamic acid (SAHA). Leukemia 2002;16:1331–43.[CrossRef][Medline]
  19. Kim MS, Kwon HJ, Lee YM, et al. Histone deacetylases induce angiogenesis by negative regulation of tumor suppressor genes. Nat Med 2001;7:437–43.[CrossRef][Medline]
  20. Park SH, Lee SR, Kim BC, et al. Transcriptional regulation of the transforming growth factor ß type II receptor gene by histone acetyltransferase and deacetylase is mediated by NF-Y in human breast cancer cells. J Biol Chem 2002;277:5168–74.[Abstract/Free Full Text]
  21. Scott GK, Marden C, Xu F, Kirk L, Benz CC. Transcriptional repression of ErbB2 by histone deacetylase inhibitors detected by a genomically integrated ErbB2 promoter-reporting cell screen. Mol Cancer Ther 2002;1:385–92.[Abstract/Free Full Text]
  22. McLaughlin F, La Thangue NB. Histone deacetylase inhibitors open new doors in cancer therapy. Biochem Pharmacol 2004;68:1139–44.[CrossRef][Medline]
  23. Gelmon KA, Eisenhauer EA, Harris AL, Ratain MJ, Workman P. Anticancer agents targeting signaling molecules and cancer cell environment: challenges for drug development? J Natl Cancer Inst 1999;91:1281–7.[Free Full Text]
  24. Parulekar WR, Eisenhauer EA. Phase I trial design for solid tumor studies of targeted, non-cytotoxic agents: theory and practice. J Natl Cancer Inst 2004;96:990–7.[Abstract/Free Full Text]
  25. Griffiths JR, Glickson JD. Monitoring pharmacokinetics of anticancer drugs: non-invasive investigation using magnetic resonance spectroscopy. Adv Drug Deliv Rev 2000;41:75–89.[CrossRef][Medline]
  26. Klomp DW, Van Laarhoven HW, Kentgens AP, Heerschap A. Optimization of localized 19F magnetic resonance spectroscopy for the detection of fluorinated drugs in the human liver. Magn Reson Med 2003;50:303–8.[Medline]
  27. Wolf W, Presant CA, Waluch V. 19F-MRS studies of fluorinated drugs in humans. Adv Drug Deliv Rev 2000;41:55–74.[CrossRef][Medline]
  28. Yu JX, Kodibagkar VD, Cui W, Mason RP. 19F: a versatile reporter for non-invasive physiology and pharmacology using magnetic resonance. Curr Med Chem 2005;12:819–48.[CrossRef][Medline]
  29. Seddon BM, Maxwell RJ, Honess DJ, et al. Validation of the fluorinated 2-nitroimidazole SR-4554 as a noninvasive hypoxia marker detected by magnetic resonance spectroscopy. Clin Cancer Res 2002;8:2323–35.[Abstract/Free Full Text]
  30. Stegman LD, Rehemtulla A, Beattie B, et al. Noninvasive quantitation of cytosine deaminase transgene expression in human tumor xenografts with in vivo magnetic resonance spectroscopy. Proc Natl Acad Sci U S A 1999;96:9821–6.[Abstract/Free Full Text]
  31. Negendank W. Studies of human tumors by MRS: a review. NMR Biomed 1992;5:303–24.[Medline]
  32. de Certaines JD, Larsen VA, Podo F, Carpinelli G, Briot O, Henriksen O. In vivo 31P MRS of experimental tumours. NMR Biomed 1993;6:345–65.[Medline]
  33. Evelhoch JL, Gillies RJ, Karczmar GS, et al. Applications of magnetic resonance in model systems: cancer therapeutics. Neoplasia 2000;2:152–65.[CrossRef][Medline]
  34. Beloueche-Babari M, Jackson LE, Al-Saffar NMS, Workman P, Leach MO, Ronen SM. Magnetic resonance spectroscopy monitoring of MAPK signaling inhibition. Cancer Res 2005;65:3356–63.[Abstract/Free Full Text]
  35. Ronen SM, Jackson LE, Beloueche M, Leach MO. Magnetic resonance detects changes in phosphocholine associated with Ras activation and inhibition in NIH 3T3 cells. Br J Cancer 2001;84:691–6.[CrossRef][Medline]
  36. Beloueche-Babari M, Jackson L, Salman-Al Saffar N, et al. Identification of magnetic resonance detectable metabolic changes associated with inhibition of phosphoinositide 3-kinase signalling in human breast cancer cells. Mol Cancer Ther 2006;5:187–96.[Abstract/Free Full Text]
  37. Chung YL, Troy H, Banerji U, et al. Magnetic resonance spectroscopic pharmacodynamic markers of the heat shock protein 90 inhibitor 17-allylamino,17-demethoxygeldanamycin (17AAG) in human colon cancer models. J Natl Cancer Inst 2003;95:1624–33.[Abstract/Free Full Text]
  38. Beloueche M, Payne GS, Banerji U, et al. MRS monitoring of the response to 17AAG treatment: from the laboratory to the clinic. Proc 20th ESMRMB; 2003; Rotterdam.
  39. Yu X, Guo ZS, Marcu MG, et al. Modulation of p53, ErbB1, ErbB2, and Raf-1 expression in lung cancer cells by depsipeptide FR901228. J Natl Cancer Inst 2002;94:504–13.[Abstract/Free Full Text]
  40. Podo F. Tumour phospholipid metabolism. NMR Biomed 1999;12:413–39.[CrossRef][Medline]
  41. Mori ND, Natarajan R, Singh K, Bhujwalla ZM. Loss of p53 function in colon cancer cells results in increased phosphocholine and total choline. Proc 10th ISMRM; 2002; Hawaii. p. 28.
  42. Fuino L, Bali P, Wittmann S, et al. Histone deacetylase inhibitor LAQ824 down-regulates Her-2 and sensitizes human breast cancer cells to trastuzumab, taxotere, gemcitabine, and epothilone B. Mol Cancer Ther 2003;2:971–84.[Abstract/Free Full Text]
  43. Bali P, Pranpat M, Bradner J, et al. Inhibition of histone deacetylase 6 acetylates and disrupts the chaperone function of heat shock protein 90: a novel basis of antileukemia activity of histone deacetylase inhibitors. J Biol Chem 2005;280:26729–34.[Abstract/Free Full Text]
  44. Berman HM, Westbrook J, Feng Z, et al. The Protein Data Bank. Nucleic Acids Res 2000;28:235–42.[Abstract/Free Full Text]
  45. Clark RD, Strizhev A, Leonard JM, Blake JF, Matthew JB. Consensus scoring for ligand/protein interactions. J Mol Graph Model 2002;20:281–95.[CrossRef][Medline]
  46. Stowell JC, Huot RI, Van Voast L. The synthesis of N-hydroxy-N'-phenyloctanediamide and its inhibitory effect on proliferation of AXC rat prostate cancer cells. J Med Chem 1995;38:1411–3.[CrossRef][Medline]
  47. Tyagi RK, Azrad A, Degani H, Salomon Y. Simultaneous extraction of cellular lipids and water-soluble metabolites: evaluation by NMR spectroscopy. Magn Reson Med 1996;35:194–200.[Medline]
  48. Scott BF, Mactavish D, Spencer C, Strachan WMJ, Muir DCG. Haloacetic acids in Canadian lake waters and precipitation. Environ Sci Technol 2000;34:4266–72.[CrossRef]
  49. Somoza JR, Skene RJ, Katz BA, et al. Structural snapshots of human HDAC8 provide insights into the class I histone deacetylases. Structure (Camb) 2004;12:1325–34.
  50. Hanahan D, Weinberg RA. The hallmarks of cancer. Cell 2000;100:57–70.[CrossRef][Medline]
  51. Sawyers C. Targeted cancer therapy. Nature 2004;432:294–7.[CrossRef][Medline]
  52. Wolffe AP. Chromatin remodeling: why it is important in cancer. Oncogene 2001;20:2988–90.[CrossRef][Medline]
  53. Cress WD, Seto E. Histone deacetylases, transcriptional control, and cancer. J Cell Physiol 2000;184:1–16.[CrossRef][Medline]
  54. Di Gennaro E, Bruzzese F, Caraglia M, Abruzzese A, Budillon A. Acetylation of proteins as novel target for antitumor therapy: review article. Amino Acids 2004;26:435–41.[Medline]
  55. Xu AS, Kuchel PW. Characterisation of erythrocyte transmembrane exchange of trifluoroacetate using 19F-NMR: evidence for transport via the monocarboxylate transporter. Biochim Biophys Acta 1993;1150:35–44.[Medline]
  56. Leach MO, Verrill M, Glaholm J, et al. Measurements of human breast cancer using magnetic resonance spectroscopy: a review of clinical measurements and a report of localized 31P measurements of response to treatment. NMR Biomed 1998;11:314–40.[CrossRef][Medline]
  57. Workman P. Combinatorial attack on multistep oncogenesis by inhibiting the Hsp90 molecular chaperone. Cancer Lett 2004;206:149–57.[CrossRef][Medline]
  58. Neckers L. Development of small molecule Hsp90 inhibitors: utilizing both forward and reverse chemical genomics for drug identification. Curr Med Chem 2003;10:733–9.[CrossRef][Medline]
  59. Kovacs JJ, Murphy PJ, Gaillard S, et al. HDAC6 regulates Hsp90 acetylation and chaperone-dependent activation of glucocorticoid receptor. Mol Cell 2005;18:601–7.[CrossRef][Medline]
  60. Peart MJ, Smyth GK, van Laar RK, et al. Identification and functional significance of genes regulated by structurally different histone deacetylase inhibitors. Proc Natl Acad Sci U S A 2005;102:3697–702.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 Molecular Cancer TherapeuticsHome page
J. Ross, A. M. Najjar, M. Sankaranarayanapillai, W. P. Tong, K. Kaluarachchi, and S. M. Ronen
Fatty acid synthase inhibition results in a magnetic resonance-detectable drop in phosphocholine
Mol. Cancer Ther., August 1, 2008; 7(8): 2556 - 2565.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Sankaranarayanapillai, M.
Right arrow Articles by Ronen, S. M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sankaranarayanapillai, M.
Right arrow Articles by Ronen, S. M.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Cancer Research Clinical Cancer Research
Cancer Epidemiology Biomarkers & Prevention Molecular Cancer Therapeutics
Molecular Cancer Research Cancer Prevention Research
Cancer Prevention Journals Portal Cancer Reviews Online
Annual Meeting Education Book Meeting Abstracts Online