Molecular Cancer Therapeutics CTRC-AACR San Antonio Breast Cancer Symposium
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 Kurz, E. U.
Right arrow Articles by Kroll, D. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Kurz, E. U.
Right arrow Articles by Kroll, D. J.
Vol. 1, 121-131, December 2001     Molecular Cancer Therapeutics
© 2001 American Association for Cancer Research

The Histone Deacetylase Inhibitor Sodium Butyrate Induces DNA Topoisomerase II{alpha} Expression and Confers Hypersensitivity to Etoposide in Human Leukemic Cell Lines1

Ebba U. Kurz2, Sara E. Wilson3, Kelly B. Leader, Brante P. Sampey, William P. Allan, Jack C. Yalowich and David J. Kroll4,5

Department of Pharmaceutical Sciences [E. U. K., S. E. W., K. B. L., B. P. S., D. J. K.], Center for Pharmaceutical Biotechnology [D. J. K.], and Program in Molecular Toxicology and Environmental Health Sciences [B. P. S., D. K. J.], School of Pharmacy, University of Colorado Health Sciences Center, and University of Colorado Cancer Center [E. U. K., D. J. K.], Denver, Colorado 80262, and Department of Pharmacology, University of Pittsburgh School of Medicine and Cancer Institute, Pittsburgh, Pennsylvania 15261 [W. P. A., J. C. Y.]


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The differentiating agent and histone deacetylase inhibitor, sodium butyrate (NaB), was shown previously to cause a transient, 3–17-fold induction of human DNA topoisomerase II{alpha} (topo II{alpha}) gene promoter activity and a 2-fold increase in topo II{alpha} protein early in monocytic differentiation of HL-60 cells. This observation has now been extended to other short chain fatty acids and aromatic butyrate analogues, and evidence is presented that human topo II{alpha} promoter induction correlates closely with histone H4 acetylation status. Because increased topo II{alpha} expression is associated with enhanced efficacy of topo II-poisoning antitumor drugs such as etoposide, the hypothesis tested in this report was whether NaB pretreatment could sensitize HL-60 myeloid leukemia and K562 erythroleukemia cells to etoposide-triggered DNA damage and cell death. A 24–72 h NaB treatment (0.4–0.5 mM) induced topo II{alpha} 2–2.5-fold in both HL-60 and K562 cells and caused a dose-dependent enhancement of etoposide-stimulated, protein-linked DNA complexes in both cell lines. At concentrations with minimal effects on cell cycle kinetics (0.4 mM in HL-60; 0.5 mM in K562), NaB pretreatment also modestly enhanced etoposide-triggered apoptosis in HL-60 cells, as determined morphologically after acridine orange/ethidium bromide staining, and substantially increased K562 growth inhibition and poly(ADP-ribose)polymerase cleavage after etoposide exposure. Therefore, a temporal window may exist whereby a differentiating agent may sensitize experimental leukemias to a cytotoxic antitumor agent. These results indicate that histone deacetylase inhibitors should be investigated for etoposide sensitization of other butyrate-responsive hematopoietic and nonhematopoietic tumor lines in vitro and in vivo.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The transcriptional and posttranscriptional regulation of DNA topo6 II{alpha}, a nuclear enzyme essential for completion of mitosis (1-3), has been a primary focus of our research groups (4-8). Drugs such as the epipodophyllotoxins (etoposide and teniposide), anthracyclines, ellipticines, anthracenediones, and aminoacridine derivatives exert their cytotoxic activity, at least in part, by trapping topo II in a covalent complex with DNA (9). These "cleavable complexes" (10) can act as physical barriers to DNA replication (11-13), cause mitotic catastrophes (14), induce recombination events (15, 16), and/or trigger a cytotoxicity cascade culminating in apoptosis (17-19). Because the class of antitumor drugs converts this essential enzyme into a lethal instrument, it follows that tumor cell populations possessing high levels of topo II{alpha} are more effectively killed by these agents.

Two genes exist for mammalian type II DNA topoisomerases, termed {alpha} and ß, likely attributable to evolutionary gene duplication (20, 21). topo II{alpha} is often the more predominant cellular form (22, 23). Cleavable complexes formed with topo II{alpha} correlate more closely with teniposide cytotoxicity in leukemia cells than do those formed with topo IIß (24), but in other systems topo IIß may also influence drug sensitivity (25, 26). The {alpha} form is expressed in a proliferation-dependent manner, whereas the ß form is expressed independently of growth status (27-29). Therefore, rapidly growing cells that contain high topo II{alpha} levels are effectively killed by the topo II-directed drugs, whereas differentiated or otherwise growth-arrested cells usually possess diminished topo II{alpha} levels and are intrinsically resistant to these agents (9, 30, 31). Similarly, acquired resistance to a topo II-directed agent is often attributable to suppressed topo II{alpha} and/or topo IIß expression (8, 32, 33).

Several transcription factors have been linked to the high, proliferation-dependent expression of topo II{alpha} including c-Myb and B-Myb (5), NF-M (4), NF-Y (34-37), and YB-1 (38). The Sp3 transcription factor is also known to activate the human topo II{alpha} gene (39). Conversely, the p53 tumor suppressor protein represses the human topo II{alpha} gene through the basal transcriptional machinery (40) or via an inverted CCAAT box at position -68 (41). The activated Ras pathway also stimulates both topo II{alpha} activity and expression. topo II{alpha} activity is enhanced via protein interactions with extracellular signal-regulated kinase 2 (42), and topo II{alpha} trans-activation can be driven through a Ets-like enhancer element at position -480 of the topo II{alpha} promoter (43). Finally, hyperthermia also induces topo II{alpha} expression in vitro (44) as a result of increased topo II{alpha} mRNA stability (45).

In previous studies, the serendipitous observation was made that topo II{alpha} transcription and gene expression was transiently induced by the monocytic differentiating agent, NaB (6). NaB has long been recognized as an inhibitor of HDAC activity (46), an effect that is, in turn, associated with selective gene activation or repression (47-49). It was hypothesized that as a result of increased topo II{alpha} expression, NaB-pretreated leukemia cells would exhibit increased sensitivity to etoposide because of increased topo II-mediated DNA damage.

In the present study, we show that several SCFAs and aromatic butyrate analogues also stimulate the activity of a synthetic topo II{alpha} promoter-reporter construct in direct relation to the magnitude of histone deacetylase inhibition. NaB, the most efficacious stimulator of topo II{alpha} promoter activity and histone H4 acetylation, also induces the endogenous expression of pharmacologically competent topo II{alpha} protein in both HL-60 and K562 cells. Consequently, NaB pretreatment sensitizes human leukemia cells to etoposide cytotoxicity measured by increased fraction of morphologically apoptotic cells (HL-60), enhanced growth inhibition (K562), or stimulation of PARP proteolytic cleavage. Our results are discussed in light of other recent reports of a physical relationship between topo II{alpha} and HDACs and indicate that other butyrate-responsive hematopoietic and nonhematopoietic cell lines should be investigated for synergism between HDAC inhibitors and classical cytotoxic agents.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Chemicals and Enzymes
Unless noted otherwise, all cell culture reagents and molecular biology grade enzymes were obtained from Life Technologies, Inc. (Gaithersburg, MD). All SCFAs were obtained from Sigma Chemical Co./Aldrich Chemical (St. Louis, MO) as sodium salts (acetate, propionate, butyrate, and caproate) or free acids (valeric and heptanoic acids), the latter of which were solubilized in equimolar NaOH. Solutions (in 0.9% saline) of sodium phenylacetate and sodium phenylbutyrate manufactured by Elan Pharmaceuticals (San Francisco, CA) were generous gifts of Dr. Andrew Kraft (Division of Medical Oncology, University of Colorado School of Medicine). Buffers and all other chemicals were obtained from United States Biochemical/Amersham Pharmacia Biotech (Arlington Heights, IL). Chemicals were of the highest purity listed by the supplier, and molecular biology-tested reagents were obtained wherever possible.

Mammalian Cell Culture
HL-60 human promyelocytic cells (ATCC CCL 240) or K562 human erythroleukemia cells (ATCC CCL 243) were obtained from the ATCC (Manassas, VA). Cells were maintained as suspension cultures in either RPMI 1640 (HL-60) or Iscove’s modified Minimal Essential Medium (K562; Life Technologies, Inc.) supplemented with 10% fetal bovine serum, 50 units/ml penicillin G, and 50 µg/ml streptomycin sulfate. All cells were maintained at 37°C in a humidified atmosphere containing 5% CO2. Cells were carried as exponentially growing cultures by propagation at 5 x 105 cells/ml every 2–3 days.

Cellular Histone Acetylation Assay
Nuclei were isolated, and histone fractions were prepared from cell nuclei as described (50) with the following modifications. Pelleted nuclei were resuspended in 1.35 ml of sterile, ice-cold water containing 1 mM phenylmethylsulfonyl fluoride and 0.15 ml of 4 N H2SO4 added dropwise with swirling and histones extracted overnight. After centrifugation at 12,000 x g for 10 min, histone supernatants were transferred to 15 ml of Falcon polypropylene tubes and filled with cold acetone/HCl (99:1 acetone:5 N HCl) and incubated at -20°C for 72 h. During the final pelleting of the histone isolates, all samples were initially centrifuged for 10 min at 2,000 x g in a swinging-bucket rotor. The supernatants were aspirated to ~1 ml, mixed by pipetting, then transferred to a 1.5 ml of Eppendorf tube for centrifugation at 12,000 x g for 10 min at 4°C. Supernatants were carefully aspirated, and pellets were lyophilized and then resuspended in loading buffer (8 M urea, 10% glycerol, 0.9 N acetic acid, 5% 2-mercaptoethanol, and 0.25% methyl green) for Triton-acid-urea gel electrophoresis. Triton-acid-urea gels containing 12% polyacrylamide (in 7.5 M urea, 0.37% Triton X-100, 0.9 N acetic acid, 0.125% ammonium persulfate, and 0.125% TEMED) were prerun for 6 h at 115 V until constant amperage was achieved. The wells were loaded with 1 M cysteamine, 7.5 M urea, 0.9 N acetic acid and run for another hour at 115 V. Solubilized histones were boiled for 5 min and then loaded and subjected to electrophoresis at 115 V for 15–18 h. After Coomassie Blue staining, gels were scanned, and acetylated bands were quantified on a Bio-Rad Fluor-S MultiImager (f/11, white light scan, clear filter, 3 s). The intensity of uni-, bi-, and triacetylated histone H4 bands (denoted 1, 2, and 3, respectively) were combined, and data were expressed as a ratio relative to the unacetylated form (denoted 0).

topo II Assays
Western immunoblotting for steady-state topo II{alpha} protein levels from HL-60 nuclear extracts was performed exactly as described by Fraser et al. (6) using a polyclonal antiserum generated in the Yalowich laboratory to a COOH-terminal recombinant topo II{alpha} peptide (7).

Transcriptional activation of a topo II{alpha} promoter-luciferase reporter construct was assessed following cellular electroporation as described previously (6) using the construct, -562TOP2LUC. The promoter region of this plasmid corresponds to positions -562 to +90 of the human topo II{alpha} 5'-flanking region (51), which had been cloned upstream of the firefly luciferase cDNA in pA3LUC. Various concentrations of NaB or other fatty acids (dissolved in sterile water, or 0.9% saline for the aromatic analogues) were incubated in triplicate with transfected cells continuously for 24 h prior to harvest and quantitation of reporter gene activities. topo II{alpha} promoter activity was normalized to the activity derived from an internal control plasmid, pCMV-ßgal, which encodes Escherichia coli ß-gal under control of the CMV immediate/early promoter, as described previously (5, 6).

Etoposide-stabilized topo II-DNA cleavable complexes were quantified using a modification (32) of a K-SDS precipitation assay. Briefly, exponentially growing HL-60 or K562 cells (5 x 105/ml) were incubated for 24 h with [14C]leucine (0.2 µCi/ml; 325 mCi/mmol) and [methyl-3H]]thymidine (0.6 µCi/ml; 6.7 Ci/mmol). In the case of HL-60 cells, the labeling was accompanied by a 24-h exposure to either 0 or 0.4 mM NaB; for K562 cells, the labeling was preceded by a 48-h exposure to 0 or 0.5 mM NaB and continued during the labeling period for a total of a 72 h butyrate exposure. During the final 30 min, cells received 0–100 µM etoposide (Sigma Chemical Co.; dissolved in DMSO) or an equivalent amount of vehicle, in triplicate. Cell suspensions were then pelleted by centrifugation at 1000 x g and lysed with 500 µl of a solution containing 2.5% (w/v) SDS, 10 mM EDTA, and 0.8 mg/ml salmon sperm DNA. The lysate was sheared through a 23-gauge needle 15 times and then incubated at 65°C for 15 min. Protein and protein-associated DNA were precipitated by the addition of 110 µl of 1 M KCl. Precipitates were recovered by centrifugation at 5000 x g and washed three times with a solution containing 10 mM Tris-HCl (pH 8.0), 100 mM KCl, 1 mM EDTA, and 0.1 mg/ml salmon sperm DNA. Precipitable 14C and 3H was assessed by dissolving the pellet in 500 µl of water followed by liquid scintillation counting. Cleavable complex data were expressed as the fold-increase over control in the ratio of 3H cpm/14C cpm and plotted relative to etoposide concentration as the mean fold-increase ± SE.

Apoptosis and Cell Growth Inhibition Measurements
The effect of a 24 h of 0.4 mM NaB pretreatment on etoposide-triggered apoptosis was quantified using the ethidium bromide/acridine orange fluorescence microscopy method of Duke and Cohen as extensively described and documented by Dwyer-Nield et al. (52). HL-60 cells were exposed to various concentrations of etoposide during the final 2–4 h of NaB. Cells were centrifuged at 350 x g for 5 min and then resuspended at ~5 x 105 to 5 x 106 cells/ml of RPMI 1640 plus 10% fetal bovine serum. One µl of a dye mixture containing of 100 µg/ml acridine orange (Sigma Chemical Co.) and 100 µg/ml ethidium bromide (Sigma Chemical Co.) prepared in PBS were placed in the bottom of a 12 x 75-mm glass tube, followed by 25 µl of cell suspension. After gentle agitation, 10 µl of the dye-cell mixture was placed on a clean microscope slide and then covered with a 22-mm2 coverslip. Cells were examined under epi-illumination at x400-x800. After counting 200 cells, the number of each of four cellular states were scored: (a) viable cells with normal nuclei (VN; bright green chromatin with organized structure); (b) viable cells with apoptotic nuclei (VA; bright green chromatin highly condensed or fragmented); (c) nonviable cells with normal nuclei (NVN; bright orange chromatin with organized structure); (d) nonviable cells with apoptotic nuclei (NVA; bright orange chromatin highly condensed or fragmented). For the purposes of this report, data were expressed only for the percentage of apoptotic cells (VA + NVA/total).

For K562 cells, exponentially growing cultures were exposed to 0 or 0.5 mM NaB for 72 h and then pelleted and exposed to increasing concentrations of etoposide in NaB-free medium for 1 h at 37°C. Cells were pelleted and washed twice with drug-free medium and then resuspended at a cell density of 1.2 x 105 cells/ml. Cells were counted 48 h later on a model ZBF Coulter Counter (Coulter Electronics, Hialeah, FL). Cell growth (beyond the starting concentration of 1.2 x 105 cells/ml) in etoposide-treated versus control cells was ultimately expressed as a percentage of inhibition of control cell growth. The 50% growth inhibitory concentration (IC50) was calculated from replicate concentration-response curves generated from three separate experiments.

Cleavage of PARP as a measure of apoptotic caspase activity (53) was also assessed after etoposide exposure in K562 cells pretreated with NaB (0.5 mM for 72 h) by immunoblotting for the Mr 116,000 parent band and the Mr 85,000–89,000 proteolytic cleavage product. Following SDS-PAGE with 7.5% acrylamide and electrotransfer to Immobilon-P membranes, blots were probed with a polyclonal antiserum obtained from Santa Cruz Biotechnology Biochemical (Santa Cruz Biotechnology, CA) at a 1:1000 dilution in TBS-T [25 mM Tris-HCl (pH 7.4), 150 mM NaCl, and 0.05% Tween 20] with 5% nonfat dry milk. Secondary detection of immune complexes was accomplished by probing with a 1:5000 dilution of goat antirabbit-horseradish peroxidase conjugate (Pierce, Rockford, IL), and bands were detected by fluorescence radiography using an enhanced chemiluminescence substrate (NEN).

Statistical Analysis
Where indicated, differences from control values were assessed by one-way ANOVA using Dunnett’s multiple comparison post-hoc test with GraphPad Prism software. Significant differences were indicated where P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Several SCFA Induce Both the Activity of the DNA topo II{alpha} Promoter and Acetylation of Histone H4 in HL-60 Cells
It has been recognized for >20 years that SCFAs other than NaB are capable of enhancing the acetylation state of histones, especially histone H4, by inhibiting cellular histone deacetylase activity (46, 54). Having initially demonstrated that NaB was an efficacious inducer of the human topo II{alpha} promoter (6), we sought to investigate whether this was a property shared by other SCFAs and, if so, whether the magnitude of induction correlated with HDAC inhibitory activity. HL-60 cells were transfected with -562TOP2LUC and treated for 24 h with 1 mM concentrations of each of the SCFAs with carbon chain lengths from two to seven and the aromatic butyrate analogues, phenylacetate and phenylbutyrate (as sodium salts), followed by quantitation of luciferase reporter enzyme activity. Parallel cultures, treated identically, were subjected to analysis of histone acetylation status by electrophoresis on Triton-acid-urea gels.

SCFAs as short as acetate (C2) produced a significant increase in topo II{alpha} promoter activity, which rose substantially as chain length increased, peaking at 9.6-fold with butyrate (C4), then decreasing to 4.3-fold with caproate (C6), and returning to control levels with the seven-carbon fatty acid, heptanoate (Fig. 1). Of the aromatic butyrate analogues, only phenylbutyrate substantially activated this promoter construct in HL-60 cells, although phenylacetate could induce the promoter >2-fold at concentrations >=4 mM (data not shown). The biphasic response seen with the SCFAs is highly reminiscent of the relationship between carbon chain length and histone acetylation observed previously in HeLa cells (54). We therefore investigated whether there was a direct correlation between the magnitude of topo II{alpha} promoter activation produced by the SCFAs and the increase in histone H4 acetylation status.



View larger version (38K):
[in this window]
[in a new window]
  		Fig. 1. Effect of SCFAs and butyrate analogues on topo II{alpha} promoter activity in HL-60 cells. A, the topo II{alpha} promoter-luciferase reporter plasmid, -562TOP2LUC (20 µg), and an internal control plasmid, pCMV-ßgal (1 µg), were cotransfected into HL-60 cells by electroporation, and cells were exposed to the sodium salts of the indicated fatty acids [acetate (C2), propionate (C3), butyrate (C4), valerate (C5), caproate (C6), heptanoate (C7), phenylacetate, or phenylbutyrate] at a concentration of 1 mM for 24 h. Cells were then processed for measurement of luciferase activity, and expression was normalized to that of ß-galactosidase activity produced from the internal control construct. Data are expressed as relative light units divided by milliunits of ß-gal activity; bars, SE.

 
When similarly treated HL-60 cells were subjected to histone acetylation analysis (Fig. 2A), it was clear that the most efficacious topo II{alpha} promoter inducers were also the most effective inhibitors of histone H4 deacetylation. This relationship is demonstrated graphically in Fig. 2B. However, this concordance with topo II{alpha} promoter activation appeared to plateau at the very high levels of histone acetylation caused by butyrate. This observation suggests an upper limit to this relationship, such that above an acetylated:unacetylated histone H4 ratio of 3, little further induction of the topo II{alpha} promoter can be achieved.



View larger version (34K):
[in this window]
[in a new window]
  		Fig. 2. Correlation of increased histone H4 acetylation with topo II{alpha} promoter activation. A, each group of HL-60 cells was treated with each of the fatty acids (1 mM) for 24 h as in Fig. 1. Cells were then processed for isolation of nuclear histones as described in "Materials and Methods," subjected to electrophoresis on a Triton-acid-urea polyacrylamide gel, and stained with Coomassie Blue. Acetylation of histone H4 was quantified by scanning the intensities of the unacetylated band and the total acetylated bands. The intensity of uni-, bi-, and triacetylated histone H4 bands (denoted 1, 2, and 3, respectively) were combined, and data were expressed as a ratio relative to the unacetylated form (0). Differences in loading and/or histone recovery was therefore normalized by expressing acetylation as the ratio of total acetylated H4:unacetylated H4. B, graphic representation of the relationship between histone H4 acetylation and topo II{alpha} promoter activation. The data from Fig. 1 (Y axis) and A in this figure (X axis) are plotted with each point denoted by the associated fatty acid treatment. PA, sodium phenylacetate; PB, sodium phenylbutyrate.

 
Our previous work demonstrated that 0.4 mM NaB treatment of HL-60 cells for 24 h induced topo II{alpha} promoter-driven reporter enzyme activity 3–17-fold but caused only a ~2-fold increase in steady-state topo II{alpha} protein levels (6). A similar magnitude of NaB induction of topo II{alpha} protein levels (2.1 ± 0.3-fold; mean ± SE from three separate experiments) is shown in Fig. 3 by the band at Mr 170,000. Using 24 h pretreatment as the optimal time for observing topo II{alpha} induction, we investigated whether greater NaB concentrations (1–8 mM) would further induce enzyme expression. Although these higher NaB concentrations caused a 15–40-fold induction of reporter enzyme activity derived from the promoter construct, endogenous topo II{alpha} protein levels still increased by only 2.5–2.7-fold at 1 mM NaB and fell to under 2-fold control levels at higher NaB concentrations (data not shown). These results are not entirely surprising in that topo II{alpha} levels rarely vary by more than 2–3-fold in proliferating cells, and forced overexpression of the enzyme is known to cause feedback inhibition of its own transcription (55).



View larger version (36K):
[in this window]
[in a new window]
  		Fig. 3. NaB treatment of HL-60 and K562 cells increases topo II{alpha} protein levels. Logarithmically growing leukemia cells were exposed to NaB (0.4 mM, 24 h for HL-60; 0.5 mM, 72 h for K562) and then lysed directly in SDS loading buffer. Proteins were resolved by electrophoresis on SDS-7.5% acrylamide gels, transferred to Immobilon-P membranes, and then immunoblotted for human topo II{alpha} using an antiserum raised to the COOH terminus of the enzyme. Immunoreactivity was visualized using a goat antirabbit IgG-horseradish peroxidase secondary antibody and chemiluminescent substrate. topo II{alpha} bands were quantified by scanning on a Bio-Rad Fluor-S imaging system.

 
We next sought to learn whether NaB could induce topo II{alpha} protein levels in another commonly used experimental leukemia cell line, K562. Similar to HL-60 cells, K562 cells will differentiate in response to NaB but exhibit more delayed kinetics in both differentiation as well as in response to cytotoxic agents (18). In K562 cells treated with NaB for 72 h, topo II{alpha} expression was induced 2.5 ± 0.2-fold (mean ± SE from five separate experiments; Fig. 3).

Increased topo II{alpha} Expression in HL-60 and K562 Cells Correlates with Increased Etoposide-stabilized, Protein-linked DNA
To assess whether the topo II{alpha} induced by NaB in HL-60 and K562 cells was pharmacologically functional, a classic protein-linked DNA assay was used in the presence of etoposide as an indicator of the extent of cleavable complex formation in intact cells (Fig. 4). Consistent with the NaB-mediated increase in topo II{alpha} expression in HL-60 and K562 cells (Fig. 3), NaB-pretreated HL-60 and K562 cells both displayed a greater magnitude of protein-linked DNA as a function of etoposide concentration, as compared with control cells not exposed previously to NaB (Fig. 4). Therefore, the increased levels of topo II{alpha} detected in Fig. 3 represented a pharmacologically functional pool of enzyme in both leukemic cell lines. However, it should be noted that one limitation of this assay is the inability to determine the topo II isoform increasingly trapped in complexes from NaB-pretreated cells. Although these data are consistent with an increase in topo II{alpha}, conclusive demonstration of the isoform(s) involved will require immunological confirmation.



View larger version (20K):
[in this window]
[in a new window]
  		Fig. 4. NaB effects on etoposide-mediated, protein-linked DNA in HL-60 and K562 cells. A standard K-SDS precipitation assay was used to measure etoposide-stabilized, cleavable complex formation. HL-60 cells (A; n = 3/group) and K562 cells (B; n = 4 with separate experiments performed on separate days) were either left untreated (no NaB) or exposed to 0.4 or 0.5 mM NaB, respectively, as in Fig. 3, and labeled for the final 24 h with [14C]leucine (0.2 µCi/ml) and [3H]thymidine (0.6 µCi/ml). Cells were then treated with the indicated concentrations of etoposide (or vehicle) for 30 min and lysed, and protein-DNA complexes were precipitated as described in "Materials and Methods." Protein-linked DNA was quantitated by liquid scintillation counting to generate protein-linked DNA as a ratio of 3H cpm/14C cpm. The data are expressed as the mean fold-increase in this ratio relative to each DMSO vehicle control; bars, SE. {blacksquare}, no NaB; {blacktriangleup}, 0.4 mM NaB (A) and 0.5 mM NaB (B).

 
NaB Confers Hypersensitivity to Etoposide-triggered Apoptosis (HL-60) and Enhanced Cell Growth Inhibition or PARP Cleavage (K562)
NaB-pretreated HL-60 cells were investigated for their apoptotic response to etoposide relative to untreated controls using the ethidium bromide/acridine orange assay of Cohen and Duke as modified (52). This assay facilitates the microscopic identification of viable and nonviable apoptotic cells and distinguishes apoptotic cells from those that have undergone frank necrosis. HL-60 cells were pretreated for 24 h with 0.4 mM NaB, or sterile water, and then subjected to various concentrations of etoposide for 2 h. Both agents were then washed out, and cells were incubated in drug-free medium for 2 h before processing for staining. As shown in Fig. 5A, NaB pretreatment sensitized cells to the apoptotic effects of 30 and 100 µM etoposide. This phenomenon was characterized further in a time course experiment with 30 µM etoposide (Fig. 5B). At times ranging from 4 to 24 h after drug washout, NaB pretreatment caused a modest sensitization to etoposide. Taken together, these data indicate that NaB induction of topo II{alpha} led to an enhancement of the apoptotic efficacy of etoposide in HL-60 cells.



View larger version (20K):
[in this window]
[in a new window]
  		Fig. 5. Effect of NaB pretreatment on etoposide-mediated apoptosis in HL-60 cells. A, etoposide dose-response experiment. Logarithmically growing HL-60 cells were exposed to 0 or 0.4 mM NaB for 24 h and then incubated with the indicated etoposide concentrations for 2 h. Cells were pelleted by centrifugation, washed twice with drug-free medium, and allowed to incubate for 2 h further in drug-free medium. Apoptotic cells were scored by fluorescence microscopy after staining with ethidium bromide/acridine orange. The mean percentage of apoptotic cells is plotted against etoposide concentration; bars, SE. B, etoposide time course experiment. HL-60 cells were pretreated as described in A and then exposed to 30 µM etoposide for 2 h. Cells were pelleted by centrifugation, washed twice with drug-free medium, and allowed to incubate for 4–24 h further in drug-free medium. At the indicated time points, apoptotic cells were quantified as described above. *, statistical difference from the respective control (P < 0.05) using Dunnett’s multiple comparison test. {square}, control; {triangleup}, 0.4 mM NaB.

 
In contrast to HL-60 cells, the erythroleukemia line K562 does not readily undergo apoptosis that can be distinguished morphologically, and its cell death kinetics are delayed in relation to HL-60 cells (18, 53, 56). Therefore, the effect of NaB pretreatment on K562 responsiveness to etoposide was tested in a cell growth inhibition assay. After a 72-h pretreatment with 0.5 mM NaB, the IC50 for a 1-h etoposide exposure was reduced by 3-fold when compared with control cells (Fig. 6), consistent with the increase in etoposide-mediated DNA damage.



View larger version (15K):
[in this window]
[in a new window]
  		Fig. 6. Effect of NaB pretreatment on K562 cell growth after short exposure to etoposide. Logarithmically growing K562 cells were exposed to 0 or 0.5 mM NaB for 72 h and then incubated with the indicated etoposide concentrations for 1 h. Cells were pelleted by centrifugation, washed twice with drug-free medium, and incubated 48 h further in drug-free medium, at which time total cell number was counted. The percentage of cell growth inhibition in the presence of etoposide was calculated relative to total cell number in groups not treated with etoposide. Data points are expressed as the mean percentage of growth inhibition; bars, SE. Averaging results from seven separate experiments, the 50% inhibitory concentrations (IC50) were 8.7 ± 1.8 µM and 3.1 ± 0.5 µM for cells untreated or exposed previously to 0.5 mM NaB, respectively.

 
Given that NaB pretreatment conferred etoposide hypersensitivity to K562 cells, the activation of caspase activities was also examined by immunoblotting for cleavage of PARP. Very little PARP cleavage was observed after either the NaB pretreatment alone or with 16 or 24 h of exposure to 30 µM etoposide (Fig. 7). In contrast, NaB pretreatment of K562 cells strongly enhanced the extent of PARP cleavage after these etoposide treatments, as evidenced by the increased intensity of the cleaved immunoreactive band at Mr 85,000–89,000.



View larger version (20K):
[in this window]
[in a new window]
  		Fig. 7. Effect of NaB pretreatment on etoposide-stimulated PARP cleavage in K562 cells. Logarithmically growing K562 cells were exposed to 0 or 0.5 mM NaB for 72 h, followed by 30 µM etoposide for 16 or 24 h. Cells (1 x 106 per lane) were harvested, resolved by SDS-PAGE, and immunoblotted for PARP using a polyclonal antiserum (Santa Cruz Biotechnology). Immunoreactivity was visualized using a goat antirabbit IgG-horseradish peroxidase secondary antibody and chemiluminescent substrate.

 
Effect of NaB Pretreatment on HL-60 and K562 Cell Cycle Distribution
Although the NaB enhancement of etoposide cytotoxicity in both leukemia cell lines correlated with topo II{alpha} induction and increased drug-induced protein-linked DNA, the potential contribution of altered cell cycle kinetics to etoposide sensitization was also investigated. topo II-directed drugs are known to be most effective during S-phase but can kill cells to some extent at all phases of the cell cycle (9, 12, 57). Our earlier work demonstrated that a 16-h treatment of HL-60 cells with 0.4 mM NaB could trigger a very transient increase in DNA synthesis (as measured by [3H]thymidine pulse incorporation), which returned to control levels by 24 h (6). Consistent with this earlier finding, the data in Table 1 indicate that 24 h of NaB pretreatment had no substantial effect on HL-60 cell cycle distribution. In fact, the very small but statistically significant increase in G2-M distribution may actually reflect the increased need for topo II{alpha} enzyme activity in chromosomal segregation. Therefore, NaB-mediated etoposide sensitization of HL-60 cells did not appear to result from altered cell cycle kinetics but was rather more likely a result of increased cellular topo II{alpha} content.


View this table:
[in this window]
[in a new window]
  		Table 1 Effect of each NaB treatment on cell cycle distribution of HL-60 and K562 cells

Logarithmically growing cultures of HL-60 or K562 cells were exposed to 0.4 mM NaB for 24 h or 0.5 mM NaB for 72 h, respectively, or an equivalent amount of sterile water for each control. Cells were then processed for flow cytometry, and DNA content was assessed using a Coulter EPICS flow cytometer. Percentages represent the mean value ± SEM (n = 3).

 
The 72-h pretreatment of K562 cells with 0.5 mM NaB caused a modest but significant increase in S-phase distribution, accompanied by a decrease of similar magnitude in the G0-G1 fraction (Table 1). Although the increased S-phase fraction may contribute in small part to NaB sensitization of K562 cells to etoposide, it is unlikely to account entirely for either the 3-fold decrease in etoposide IC50 shown in Fig. 6 or the substantial enhancement of PARP cleavage in Fig. 7. Taken together, these data suggest that the prototypical HDAC inhibitor, NaB, can produce a transient hypersensitivity to the antitumor action of etoposide in experimental leukemia cell lines by inducing functional topo II{alpha} expression with a concordant increase in etoposide-stimulated, enzyme-mediated DNA damage.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Given that NaB caused a transient increase in HL-60 cell topo II{alpha} expression (6), it was reasoned that a window may exist to maximize rationally differentiation/cytotoxic drug efficacy. In the present report, NaB-induced topo II{alpha} expression correlated with enhanced etoposide-stimulated, protein-linked DNA in two human leukemia cell lines. On the basis of extensive literature demonstrating a positive relationship between topo II{alpha} levels and cytotoxic efficacy of drugs that target this enzyme, the logical assumption was made that increased cleavable complex formation would, in turn, increase etoposide cytotoxicity in NaB-pretreated cells. Indeed, NaB conferred to HL-60 cells a modest hypersensitivity to etoposide-mediated apoptosis, although in K562 cells there was a marked inhibition of cell growth and a substantial enhancement of PARP cleavage.

Other limited attempts to enhance cellular sensitivity to topo II poisons by pharmacological modulation of topo II levels have met with mixed success. For example, substantial induction of topo II{alpha} and ß levels was observed in HL-60 cells after 96 h of treatment with all-trans retinoic acid (58). However, induction was not accompanied by increases in etoposide-induced DNA cleavage, and in fact, etoposide-induced apoptosis was reduced significantly relative to controls. In contrast, one clever approach in confluent NIH3T3 cells to block topo II{alpha} transcriptional repression by NF-Y with DNA minor groove binding agents effectively induced the enzyme and caused a dramatic 10–20-fold reduction in the etoposide IC50 (35). These investigators argued that manipulation of topo II levels in plateau phase cell populations is more likely to represent situations encountered in vivo than with exponentially growing cultures.

In the present study, a question remains concerning the precise mechanism by which HDAC inhibitors induce topo II{alpha} expression and why there is discord between the magnitude of promoter activation and steady-state enzyme levels. Although our data support a direct relationship between topo II{alpha} promoter activation (from a transfected plasmid construct) and the degree of histone H4 acetylation, there are likely to be promoter-specific consequences of HDAC inhibition that mediate the magnitude, if any, of endogenous gene activation. For example, the high potency HDAC inhibitor, trapoxin, induces only 2% of genes studied (59). Ectopic expression of the histone acetyltransferase P/CAF (as a mimic of NaB treatment) failed to activate topo II{alpha} gene expression in HL-60 cells,7 suggesting that global activation of histone acetylation is unlikely to account for enhanced topo II{alpha} expression. In other systems, increasing data suggest the cooperation of specific acetylase/deacetylase enzymes with specific transcription factors via corepressor or coactivator proteins (60). For example, in transcriptional activation the yeast GCN5 histone acetyltransferase appears to function in a complex of proteins that bridges activators to the basal transcriptional machinery (61). Conversely, in the case of transcriptional repression, heterodimers of Mad and Max interact with SIN3 corepressor proteins to recruit RPD3 histone deacetylase activity to specific promoters, allowing lysine tails of deacetylated histones to displace activating transcription factors (62). Precisely how histone acetylation status affects the assembly of transcriptional activators and repressors on the endogenous human topo II{alpha} gene promoter remains an ongoing focus of our laboratories.

Other investigators have independently identified additional mechanisms by which HDAC inhibitors may converge on topo II{alpha} regulation. HDAC1 was shown recently to interact directly with topo II{alpha} (63). These investigators also demonstrated that the potent fungal natural product, TSA, could sensitize CCRF-CEM cells and a number of Chinese hamster ovary cell lines to etoposide (63). However, topo II{alpha} levels or activity were not assessed after TSA treatment. Shortly thereafter, Turner’s group independently confirmed the interaction of topo II{alpha} and HDAC1, but these investigators demonstrated paradoxically that TSA treatment confers resistance to etoposide-induced HL-60 cell apoptosis (64). However, this end point was only measured at a single high concentration (100 µM) of etoposide. The present report and these two other recent studies all differ in the type and phasing of HDAC inhibitor treatment and etoposide exposure, making it difficult at present to generalize about the utility of combining these agents. In related work, it should also be noted that TSA was shown recently to induce the activity of the mouse topo II{alpha} gene promoter in mouse 3T3 fibroblasts (65), but this study did not investigate TSA effects on endogenous topo II{alpha} levels or cellular sensitivity to topo II poisons. We have subsequently observed a 3–6-fold induction of the human topo II{alpha} promoter with 24-h exposure to TSA at 100–300 nM in several human tumor cell lines,8 suggesting that this effect is not limited to the murine gene. Whether TSA behaves in our system with regard to increased enzyme expression and etoposide cellular sensitivity is the subject of ongoing study.

The use of differentiating agents, including NaB, in clinical management of neoplastic disease has shown some promise (66-68), especially when combined with cytotoxic chemotherapy (69). However, enthusiasm has been limited because of the short half-life of fatty acids and a lack of mechanistic rationale for their combination with cytotoxic agents. A search for butyrate analogues possessing a more favorable pharmacokinetic profile than NaB has led to clinical trials with phenylacetate, phenylbutyrate, and tributyrin (70) as well as pivalyloxymethyl butyrate (AN9). Newer fungal agents (TSA, as well as trapoxin and apicidin) with nanomolar inhibitory action on HDAC may also represent useful therapeutics. However, most clinical work to date has used the aromatic butyrates. Phenylbutyrate has been used in the treatment of ß-thalassemia because of its ability to induce fetal hemoglobin (71) and in children with urea-cycle disorders (72) because of its glutamine-scavenging activity. In fact, phenylbutyrate was used recently to facilitate the birth of a child harboring a heterozygous mutation for ornithine transcarbamylase (73).

With regard to cancer therapy, others have investigated HDAC inhibitors for efficacy as single-agent antitumor drugs. This approach arose originally from the observation that a number of nonhematopoietic cell lines (including melanoma and carcinoma of the breast, ovary, lung, prostate, and colon) were responsive to the differentiating effects of butyrate or butyrate analogues (74-78). A synthetic benzamide HDAC inhibitor (MS-27-275) also possesses antitumor activity in a variety of human tumor xenografts in nude mice (79). Similarly, sodium phenylbutyrate can trigger apoptosis in prostate carcinoma cells (80). In colon carcinoma, NaB causes cell cycle arrest by inducing p21 (78), but this effect does not require p53 (81, 82). The latter finding is provocative in that the HDAC inhibitors may activate normally p53-dependent apoptotic machinery in cells lacking functional p53 (83, 84). Some of the cytostatic effects of phenylacetate and phenylbutyrate may also be mediated by their action as ligands for the peroxisome proliferator-activated receptor {gamma} (85). Although it remains to be demonstrated whether any of these other cell types also exhibit NaB-inducible topo II{alpha} expression or whether peroxisome proliferator-activated receptor {gamma} affects topo II{alpha} expression, HDAC inhibitors are already being tested in the clinic against several solid tumor types. Finally, HDAC inhibitors may have antitumor effects that may only be apparent when in vivo models are used. For example, TSA was shown recently to act as an angiogenesis inhibitor in human tumor xenografts of nude mice in part by suppressing hypoxia-responsive tumor suppressor genes (86).

Although our work has focused on the ability of HDAC inhibitors to influence topo II-dependent drug action, evidence suggests that HDAC inhibitors may also be useful combination chemotherapy agents via actions independent of and/or downstream from topo II-mediated DNA damage. Phenylbutyrate, but not NaB, enhanced doxorubicin cytotoxicity in culture, presumably by a loss of numerous cellular antioxidant defenses (87), although topo II{alpha} levels were not quantified. In another study (88), pivalyloxymethylbutyrate (AN9) was shown to enhance anthracycline efficacy but via decreased rate of anthracycline aglycone formation and a reduction in NAD(P)H:quinone oxidoreductase and cytochrome P-450 reductase activities. Oxidative stress attributable to reactive oxygen species has also been implicated in butyrate sensitization of colorectal carcinoma cells to tumor necrosis factor-{alpha} and Fas ligand (89). Reactive oxygen species may also contribute to p53-independent induction of p21 during growth arrest (83). In addition, NaB potentiates apoptosis in retinoblastoma cells treated with vincristine or cisplatin, although the mechanism of this effect was not elucidated (90). One possible explanation, particularly with regard to vincristine, is that NaB can cause G2 arrest in human breast carcinoma cells and drive DNA endoreduplication, resulting in polyploidy and delayed cell death (91).

In summary, more detailed understanding of the biological consequences of altering histone acetylation may lead to opportunities for synergistic combination chemotherapy between HDAC inhibitors and classical cytotoxic agents. It should also be appreciated that HDAC inhibitors possess other nonhistone activities that may contribute to their single-agent and combination antitumor effects, and moreover, some of these effects may be unique to the HDAC inhibitor in question. Our current work details a direct link between HDAC inhibitors, the induction of topo II{alpha}, and the consequent enhancement of etoposide-triggered DNA damage. However, further studies on HDAC inhibitors in post-DNA damage processing events may provide insights for maximizing the anticancer efficacy of these agents.


   Acknowledgments
 
We dedicate this work to the memory of Dr. Alan P. Wolffe (1959–2001), whose untimely passing (reported in Science, 293: 1065, 2001) represents a great loss to the field of epigenetic transcriptional control by histone acetylation. The foundation work of Drs. Tessa L. Brandt and David J. Fraser are recognized in providing the hypothesis tested in this report. The technical assistance of Pegge M. Halandras and Charles T. Nuttelman are also recognized for the generation of preliminary data leading to the current studies. We thank Dr. J. John Cohen (University of Colorado School of Medicine, Division of Immunology) for access to his fluorescence microscope. D. J. K. also thanks Dr. Kenneth N. Kreuzer (Duke University Medical Center) and the members of his laboratory for their constructive comments and stimulating discussion of these studies.


   Footnotes
 
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.

1 This work was supported by Grant RPG-97-084-01-CDD from the American Cancer Society, a grant from the Cancer League of Colorado (both to D. J. K.), NIH Grant CA74972 (to J. C. Y.), and an American Cancer Society/Brooks Trust Fellowship Award (to S. E. W.). The experiments from the University of Colorado Cancer Center Flow Cytometry Core Facility were supported by NIH Grant P30 CA46934 from the National Cancer Institute. Back

2 Present address: Department of Biological Sciences, University of Calgary, Calgary, Alberta, T2N 1N4 Canada. Back

3 Present address: Department of Biological Sciences, Stanford University, Stanford, CA 94305. Back

4 Present address: Division of General Internal Medicine, Department of Medicine, Duke University Medical Center, Duke Comprehensive Cancer Center, Durham, NC 27710. Back

5 To whom requests for reprints should be addressed, at Duke University Medical Center, Box 3020, Microbiology, Durham, NC 27710. Phone: (919) 668-0281; Fax: (919) 309-4042; Email: kroll001{at}mc.duke.edu Back

6 The abbreviations used are: topo, topoisomerase; NaB, sodium butyrate; HDAC, histone deacetylase; IC50, drug concentration that inhibits growth by 50%; PARP, poly(ADP-ribose)polymerase; ATCC, American Type Culture Collection; CMV, cytomegalovirus; ß-gal, ß-galactosidase; SCFA, short-chain fatty acid; TSA, trichostatin A. Back

7 D. J. Kroll, unpublished observation. Back

8 J. C. Yalowich and D. J. Kroll, manuscript in preparation. Back

Received 8/13/01; revised 9/19/01; accepted 9/25/01.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Holm, C., Goto, T., Wang, J. C., and Botstein, D. DNA topoisomerase II is required at the time of mitosis in yeast.Cell , 41:553 –563,1985 .[CrossRef][Medline]
  2. Holm, C., Stearns, T., and Botstein, D. DNA topoisomerase II must act at mitosis to prevent nondisjunction and chromosome breakage.Mol. Cell. Biol. , 9:159 –168,1989 .[Abstract/Free Full Text]
  3. Uemura, T., Ohkura, H., Adachi, Y., Morino, K., Shiozaki, K., and Yanagida, M. DNA topoisomerase II is required for condensation and separation of mitotic chromosomes in S. pombe.Cell , 50:917 –925,1987 .[CrossRef][Medline]
  4. Brandt, T. L., and Kroll, D. J. NF-M trans-activates the human DNA topoisomerase II{alpha} promoter independently of c-Myb in HL-60 cells.Leuk. Res. , 21:711 –720,1997 .[CrossRef][Medline]
  5. Brandt, T. L., Fraser, D. J., Leal, S., Halandras, P. M., Nelson, A. R., and Kroll, D. J. c-Myb trans-activates the human DNA topoisomerase II{alpha} gene promoter.J. Biol. Chem. , 272:6278 –6284,1997 .[Abstract/Free Full Text]
  6. Fraser, D. J., Brandt, T. L., and Kroll, D. J. Topoisomerase II{alpha} promoter trans-activation early in monocytic differentiation of HL-60 human leukemia cells.Mol. Pharmacol. , 47:696 –706,1995 .[Abstract]
  7. Ritke, M. K., Allan, W. P., Fattman, C., Gunduz, N. N., and Yalowich, J. C. Reduced phosphorylation of topoisomerase II in etoposide-resistant human leukemia K562 cells.Mol. Pharmacol. , 46:58 –66,1994 .[Abstract]
  8. Ritke, M. K., Roberts, D., Allan, W. P., Raymond, J., Bergoltz, V. V., and Yalowich, J. C. Altered stability of etoposide-induced topoisomerase II-DNA complexes in resistant human leukaemia K562 cells.Br. J. Cancer , 69:687 –697,1994 .[Medline]
  9. Corbett, A. H., and Osheroff, N. When good enzymes go bad: conversion of topoisomerase II to a cellular toxin by antineoplastic drugs.Chem. Res. Toxicol. , 6:585 –597,1993 .[CrossRef][Medline]
  10. Nelson, E. M., Tewey, K. M., and Liu, L. F. Mechanism of antitumor drug action: poisoning of mammalian DNA topoisomerase II on DNA by 4'-(9-acridinylamino)-methanesulfon-m-anisidide.Proc. Natl. Acad. Sci. USA , 81:1361 –1365,1984 .[Abstract/Free Full Text]
  11. Catapano, C. V., Carbone, G. M., Pisani, F., Qiu, J., and Fernandes, D. J. Arrest of replication fork progression at sites of topoisomerase II-mediated DNA cleavage in human leukemia CEM cells incubated with VM-26.Biochemistry , 36:5739 –5748,1997 .[CrossRef][Medline]
  12. D’Arpa, P., Beardmore, C., and Liu, L. F. Involvement of nucleic acid synthesis in cell killing mechanisms of topoisomerase poisons.Cancer Res. , 50:6919 –6924,1990 .[Abstract/Free Full Text]
  13. Hong, G., and Kreuzer, K. N. An antitumor drug-induced topoisomerase cleavage complex blocks a bacteriophage T4 replication fork in vivo. Mol.Cell. Biol. , 20:594 –603,2000 .
  14. Lock, R. B., and Stribinskiene, L. Dual modes of death induced by etoposide in human epithelial tumor cells allow Bcl-2 to inhibit apoptosis without affecting clonogenic survival.Cancer Res. , 56:4006 –4012,1996 .[Abstract/Free Full Text]
  15. Berger, N. A., Chatterjee, S., Schmotzer, J. A., and Helms, S. R. Etoposide (VP-16-213)-induced gene alterations: potential contribution to cell death.Proc. Natl. Acad. Sci. USA , 88:8740 –8743,1991 .[Abstract/Free Full Text]
  16. Neece, S. H., Carles-Kinch, K., Tomso, D. J., and Kreuzer, K. N. Role of recombinational repair in sensitivity to an antitumour agent that inhibits bacteriophage T4 type II DNA topoisomerase.Mol. Microbiol. , 20:1145 –1154,1996 .[CrossRef][Medline]
  17. Hashimoto, H., Chatterjee, S., and Berger, N. A. Inhibition of etoposide (VP-16)-induced DNA recombination and mutant frequency by Bcl-2 protein overexpression.Cancer Res. , 55:4029 –4035,1995 .[Abstract/Free Full Text]
  18. Ritke, M. K., Rusnak, J. M., Lazo, J. S., Allan, W. P., Dive, C., Heer, S., and Yalowich, J. C. Differential induction of etoposide-mediated apoptosis in human leukemia HL-60 and K562 cells.Mol. Pharmacol. , 46:605 –611,1994 .[Abstract]
  19. Kamesaki, S., Kamesaki, H., Jorgensen, T. J., Tanizawa, A., Pommier, Y., and Cossman, J. bcl-2 protein inhibits etoposide-induced apoptosis through its effects on events subsequent to topoisomerase II-induced DNA strand breaks and their repair.Cancer Res. , 53:4251 –4256,1993 .[Abstract/Free Full Text]
  20. Lang, A. J., Mirski, S. E., Cummings, H. J., Yu, Q., Gerlach, J. H., and Cole, S. P. Structural organization of the human TOP2A and TOP2B genes.Gene (Amst.) , 221:255 –266,1998 .[CrossRef][Medline]
  21. Sng, J. H., Heaton, V. J., Bell, M., Maini, P., Austin, C. A., and Fisher, L. M. Molecular cloning and characterization of the human topoisomerase II{alpha} and IIß genes: evidence for isoform evolution through gene duplication.Biochim. Biophys. Acta , 1444:395 –406,1999 .[Medline]
  22. Tsai-Pflugfelder, M., Liu, L. F., Liu, A. A., Tewey, K. M., Whang-Peng, J., Knutsen, T., Huebner, K., Croce, C. M., and Wang, J. C. Cloning and sequencing of cDNA encoding human DNA topoisomerase II and localization of the gene to chromosome region 17q21–22.Proc. Natl. Acad. Sci. USA , 85:7177 –7181,1988 .[Abstract/Free Full Text]
  23. Jenkins, J. R., Ayton, P., Jones, T., Davies, S. L., Simmons, D. L., Harris, A. L., Sheer, D., and Hickson, I. D. Isolation of cDNA clones encoding the ß isozyme of human DNA topoisomerase II and localisation of the gene to chromosome 3p24.Nucleic Acids Res. , 20:5587 –5592,1992 .[Abstract/Free Full Text]
  24. Qiu, J., Catapano, C. V., and Fernandes, D. J. Formation of topoisomerase II{alpha} complexes with nascent DNA is related to VM-26-induced cytotoxicity.Biochemistry , 35:16354 –16360,1996 .[CrossRef][Medline]
  25. Brown, G. A., McPherson, J. P., Gu, L., Hedley, D. W., Toso, R., Deuchars, K. L., Freedman, M. H., and Goldenberg, G. J. Relationship of DNA topoisomerase II{alpha} and ß expression to cytotoxicity of antineoplastic agents in human acute lymphoblastic leukemia cell lines.Cancer Res. , 55:78 –82,1995 .[Abstract/Free Full Text]
  26. Dereuddre, S., Delaporte, C., and Jacquemin-Sablon, A. Role of topoisomerase IIß in the resistance of 9-OH-ellipticine-resistant Chinese hamster fibroblasts to topoisomerase II inhibitors.Cancer Res. , 57:4301 –4308,1997 .[Abstract/Free Full Text]
  27. Kimura, K., Saijo, M., Ui, M., and Enomoto, T. Growth state- and cell cycle-dependent fluctuation in the expression of two forms of DNA topoisomerase II and possible specific modifications of the higher molecular weight form in the M phase.J. Biol. Chem. , 269:1173 –1176,1994 .[Abstract/Free Full Text]
  28. Sullivan, D. M., Latham, M. D., and Ross, W. E. Proliferation-dependent topoisomerase II content as a determinant of antineoplastic drug action in human, mouse, and Chinese hamster ovary cells.Cancer Res. , 47:3973 –3979,1987 .[Abstract/Free Full Text]
  29. Woessner, R. D., Mattern, M. R., Mirabelli, C. K., Johnson, R. K., and Drake, F. H. Proliferation- and cell cycle-dependent differences in expression of the 170 kilodalton and 180 kilodalton forms of topoisomerase II in NIH-3T3 cells.Cell Growth Differ. , 2:209 –214,1991 .[Abstract]
  30. Davies, S. M., Robson, C. N., Davies, S. L., and Hickson, I. D. Nuclear topoisomerase II levels correlate with the sensitivity of mammalian cells to intercalating agents and epipodophyllotoxins.J. Biol. Chem. , 263:17724 –17729,1988 .[Abstract/Free Full Text]
  31. Kaufmann, S. H., McLaughlin, S. J., Kastan, M. B., Liu, L. F., Karp, J. E., and Burke, P. J. Topoisomerase II levels during granulocytic maturation in vitro and in vivo.Cancer Res. , 51:3534 –3543,1991 .[Abstract/Free Full Text]
  32. Ritke, M. K., and Yalowich, J. C. Altered gene expression in human leukemia K562 cells selected for resistance to etoposide.Biochem. Pharmacol. , 46:2007 –2020,1993 .[CrossRef][Medline]
  33. Webb, C. D., Latham, M. D., Lock, R. B., and Sullivan, D. M. Attenuated topoisomerase II content directly correlates with a low level of drug resistance in a Chinese hamster ovary cell line.Cancer Res. , 51:6543 –6549,1991 .[Abstract/Free Full Text]
  34. Isaacs, R. J., Harris, A. L., and Hickson, I. D. Regulation of the human topoisomerase II{alpha} gene promoter in confluence-arrested cells.J. Biol. Chem. , 271:16741 –16747,1996 .[Abstract/Free Full Text]
  35. Tolner, B., Hartley, J. A., and Hochhauser, D. Transcriptional regulation of topoisomerase II{alpha} at confluence and pharmacological modulation of expression by bis-benzimidazole drugs.Mol. Pharmacol. , 59:699 –706,2001 .[Abstract/Free Full Text]
  36. Wang, H., Jiang, Z., Wong, Y. W., Dalton, W. S., Futscher, B. W., and Chan, V. T. Decreased CP-1 (NF-Y) activity results in transcriptional down-regulation of topoisomerase II{alpha} in a doxorubicin-resistant variant of human multiple myeloma RPMI 8226.Biochem. Biophys. Res. Commun. , 237:217 –224,1997 .[CrossRef][Medline]
  37. Falck, J., Jensen, P. B., and Sehested, M. Evidence for repressional role of an inverted CCAAT box in cell cycle-dependent transcription of the human DNA topoisomerase II{alpha} gene.J. Biol. Chem. , 274:18753 –18758,1999 .[Abstract/Free Full Text]
  38. Shibao, K., Takano, H., Nakayama, Y., Okazaki, K., Nagata, N., Izumi, H., Uchiumi, T., Kuwano, M., Kohno, K., and Itoh, H. Enhanced coexpression of YB-1 and DNA topoisomerase II{alpha} genes in human colorectal carcinomas.Int. J. Cancer , 83:732 –737,1999 .[CrossRef][Medline]
  39. Mo, Y. Y., Wang, Q., and Beck, W. T. Down-regulation of topoisomerase II{alpha} in CEM cells selected for merbarone resistance is associated with reduced expression of Sp3.Cancer Res. , 57:5004 –5008,1997 .[Abstract/Free Full Text]
  40. Sandri, M. I., Isaacs, R. J., Ongkeko, W. M., Harris, A. L., Hickson, I. D., Broggini, M., and Vikhanskaya, F. p53 regulates the minimal promoter of the human topoisomerase II{alpha} gene.Nucleic Acids Res. , 24:4464 –4470,1996 .[Abstract/Free Full Text]
  41. Wang, Q., Zambetti, G. P., and Suttle, D. P. Inhibition of DNA topoisomerase II{alpha} gene expression by the p53 tumor suppressor.Mol. Cell. Biol. , 17:389 –397,1997 .[Abstract]
  42. Shapiro, P. S., Whalen, A. M., Tolwinski, N. S., Wilsbacher, J., Froelich-Ammon, S. J., Garcia, M., Osheroff, N., and Ahn, N. G. Extracellular signal-regulated kinase activates topoisomerase II{alpha} through a mechanism independent of phosphorylation.Mol. Cell. Biol. , 19:3551 –3560,1999 .[Abstract/Free Full Text]
  43. Chen, G., Templeton, D., Suttle, D. P., and Stacey, D. W. Ras stimulates DNA topoisomerase II{alpha} through MEK: a link between oncogenic signaling and a therapeutic target.Oncogene , 18:7149 –7160,1999 .[CrossRef][Medline]
  44. Matsuo, K., Kohno, K., Sato, S., Uchiumi, T., Tanimura, H., Yamada, Y., and Kuwano, M. Enhanced expression of the DNA topoisomerase II gene in response to heat shock stress in human epidermoid cancer KB cells.Cancer Res. , 53:1085 –1090,1993 .[Abstract/Free Full Text]
  45. Goswami, P. C., Roti Roti, J. L., and Hunt, C. R. The cell cycle-coupled expression of topoisomerase II{alpha} during S phase is regulated by mRNA stability and is disrupted by heat shock or ionizing radiation.Mol. Cell. Biol. , 16:1500 –1508,1996 .[Abstract]
  46. Boffa, L. C., Vidali, G., Mann, R. S., and Allfrey, V. G. Suppression of histone deacetylation in vivo and in vitro by sodium butyrate.J. Biol. Chem. , 253:3364 –3366,1978 .[Abstract/Free Full Text]
  47. Struhl, K. Histone acetylation and transcriptional regulatory mechanisms.Genes Dev. , 12:599 –606,1998 .[Free Full Text]
  48. Grunstein, M. Histone acetylation in chromatin structure and transcription.Nature (Lond.) , 389:349 –352,1997 .[CrossRef][Medline]
  49. McMurry, M. T., and Krangel, M. S. A role for histone acetylation in the developmental regulation of V(D)J recombination.Science (Wash. DC) , 287:495 –498,2000 .[Abstract/Free Full Text]
  50. Mullen, J. R., Kayne, P. S., Moerschell, R. P., Tsunasawa, S., Gribskov, M., Colavito-Shepanski, M., Grunstein, M., Sherman, F., and Sternglanz, R. Identification and characterization of genes and mutants for an N-terminal acetyltransferase from yeast.EMBO J. , 8:2067 –2075,1989 .[Medline]
  51. Hochhauser, D., Stanway, C. A., Harris, A. L., and Hickson, I. D. Cloning and characterization of the 5'-flanking region of the human topoisomerase II{alpha} gene.J. Biol. Chem. , 267:18961 –18965,1992 .[Abstract/Free Full Text]
  52. Dwyer-Nield, L. D., Thompson, J. A., Peljak, G., Squier, M. K., Barker, T. D., Parkinson, A., Cohen, J. J., Dinsdale, D., and Malkinson, A. M. Selective induction of apoptosis in mouse and human lung epithelial cell lines by the tert-butyl hydroxylated metabolite of butylated hydroxytoluene: a proposed role in tumor promotion.Toxicology , 130:115 –127,1998 .[CrossRef][Medline]
  53. Kaufmann, S. H., Desnoyers, S., Ottaviano, Y., Davidson, N. E., and Poirier, G. G. Specific proteolytic cleavage of poly(ADP-ribose) polymerase: an early marker of chemotherapy-induced apoptosis.Cancer Res. , 53:3976 –3985,1993 .[Abstract/Free Full Text]
  54. Leder, A., and Leder, P. Butyric acid, a potent inducer of erythroid differentiation in cultured erythroleukemic cells.Cell , 5:319 –322,1975 .[CrossRef][Medline]
  55. 55. Asano, T., An, T., Mayes, J., Zwelling, L. A., and Kleinerman, E. S. Transfection of human topoisomerase II{alpha} into etoposide-resistant cells: transient increase in sensitivity followed by down-regulation of the endogenous gene.Biochem. J. , 319 (Pt. 1):307 –313,1996 .
  56. Martins, L. M., Mesner, P. W., Kottke, T. J., Basi, G. S., Sinha, S., Tung, J. S., Svingen, P. A., Madden, B. J., Takahashi, A., McCormick, D. J., Earnshaw, W. C., and Kaufmann, S. H. Comparison of caspase activation and subcellular localization in HL-60 and K562 cells undergoing etoposide-induced apoptosis.Blood , 90:4283 –4296,1997 .[Abstract/Free Full Text]
  57. Chow, K. C., and Ross, W. E. Topoisomerase-specific drug sensitivity in relation to cell cycle progression.Mol. Cell. Biol. , 7:3119 –3123,1987 .[Abstract/Free Full Text]
  58. Aoyama, M., Grabowski, D. R., Isaacs, R. J., Krivacic, K. A., Rybicki, L. A., Bukowski, R. M., Ganapathi, M. K., Hickson, I. D., and Ganapathi, R. Altered expression and activity of topoisomerases during all-trans retinoic acid-induced differentiation of HL-60 cells.Blood , 92:2863 –2870,1998 .[Abstract/Free Full Text]
  59. Van Lint, C., Emiliani, S., and Verdin, E. The expression of a small fraction of cellular genes is changed in response to histone hyperacetylation.Gene Exp. , 5:245 –253,1996 .
  60. Burgess, S. M., Ajimura, M., and Kleckner, N. GCN5-dependent histone H3 acetylation and RPD3-dependent histone H4 deacetylation have distinct, opposing effects on IME2 transcription, during meiosis and during vegetative growth, in budding yeast.Proc. Natl. Acad. Sci. USA , 96:6835 –6840,1999 .[Abstract/Free Full Text]
  61. Candau, R., Zhou, J. X., Allis, C. D., and Berger, S. L. Histone acetyltransferase activity and interaction with ADA2 are critical for GCN5 function in vivo.EMBO J. , 16:555 –565,1997 .[CrossRef][Medline]