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Cancer Research 67, 10198, November 1, 2007. doi: 10.1158/0008-5472.CAN-07-2505
© 2007 American Association for Cancer Research
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Molecular Biology, Pathobiology, and Genetics

Cell Growth Inhibition by Okadaic Acid Involves Gut-Enriched Kruppel-like Factor–Mediated Enhanced Expression of c-Myc

Liyue Zhang1,2, Anil Wali1,3,4, Chilakamarti V. Ramana5 and Arun K. Rishi1,2,4

1 John D. Dingell V.A. Medical Center and Departments of 2 Internal Medicine and 3 Surgery and 4 Karmanos Cancer Institute, Wayne State University, Detroit, Michigan and 5 Department of Medicine, Yale University School of Medicine, New Haven, Connecticut

Requests for reprints: Arun K. Rishi, John D. Dingell VA Medical Center, Research 151, Room B 4270, 4646 John R., Detroit, MI 48201. Phone: 313-576-1000, ext. 4492; Fax: 313-576-1112; E-mail: Rishia{at}Karmanos.org.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Human breast cancer (HBC) cell growth suppression by okadaic acid (OA) was previously found to involve elevated expression of oncogenes c-myc and c-fos and apoptosis. Since, c-Myc influences diverse pathways of cell growth, we hypothesized that elevated levels of c-Myc are involved in HBC growth suppression. Here, we investigated whether induction of c-Myc by OA or protein synthesis inhibitor cycloheximide contributed to HBC growth inhibition and the mechanisms involved. OA, cycloheximide, or the chemotherapeutic drug Taxol suppressed HBC cell growth. However, OA or cycloheximide treatments over 6 or 10 h, respectively, induced c-Myc expression. Depletion of c-Myc, on the other hand, resulted in enhanced HBC cell viabilities when exposed to OA or cycloheximide, but not by Taxol. OA induced c-myc transcription by targeting an 80-bp region from positions –11 to +70, relative to the P1 transcription start of mouse c-myc promoter. Gel mobility shift assays revealed binding of HBC cell nuclear proteins to the OA-responsive c-myc promoter fragment, whereas binding of one complex was elevated in the case of the OA-treated or cycloheximide-treated HBC cell nuclear extracts. Database search revealed presence of a consensus sequence for zinc finger protein gut-enriched Kruppel-like factor (GKLF) in OA-responsive region of the c-myc promoter. Mutation of GKLF consensus sequences abrogated OA responsiveness of the c-myc promoter, and OA treatments caused enhanced expression of GKLF in HBC cells. Thus, OA-dependent attenuation of HBC growth is accomplished, in part, by zinc finger transcription factor GKLF-mediated enhanced transcription of c-myc. [Cancer Res 2007;67(21):10198–206]


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The myc oncogene plays a central role in proliferation and malignant transformation of human and animal cells by influencing diverse pathways of growth, differentiation, genomic stability, cell cycle control, and apoptosis (1, 2). The three myc genes N-myc, L-myc, and c-myc are differentially expressed in mammalian development. The c-Myc, however, is the most potent in effecting cellular transformation, as well as causing lymphoid tumors in the transgenic mice (3). The c-myc gene is transcribed as three mRNAs that give rise to c-Myc1, c-Myc2, and c-MycS proteins. The 62-kDa c-Myc2 protein is the major form with the N-terminal transactivation domain that contains two regions called Myc homology boxes I and II, which are highly conserved among members of myc family. The carboxyl terminus of the c-Myc proteins contains a basic region and a helix-loop-helix/leucine zipper (HLH/LZ) domain. The HLH/LZ domain functions in heterodimerization of c-Myc with another transcription factor Max that serves as an obligate heterodimeric partner for both the Myc and Mad families of proteins. The Myc/Max heterodimer binds to the c-Myc consensus sequence called E-box present in the regulatory regions of the c-Myc target genes (1, 4).

The c-Myc regulates cell cycle, in principle, by activating genes that are positive regulators of the cell cycle, as well as concomitantly suppressing expression of growth inhibitory genes. For example, c-Myc stimulates transcription of genes, such as cyclins D1, D2, E, and A; cdk4, e2f1, e2f2, cdc25A, and b; etc. (ref. 1 and references therein). However, only cdk4, e2f2, and cyclins D1 and D2 contain canonical E-box sequences in their regulatory regions. Thus, E-box–independent mechanisms are likely used by c-Myc to activate expression of its target genes. On the other hand, c-Myc suppresses transcription of growth inhibitory genes, such as gadd45, cyclin-dependent kinase inhibitors p21WAF1/CIP1 and p27Kip1 (ref. 1 and references therein). The cellular transformation by c-Myc is thought to be the consequence of its aberrant or genetically altered expression. The role of c-Myc in transformation is further highlighted by the fact that it regulates expression of the human telomerase transcriptase (hTERT) gene through the E-box sequences located in the promoter of the hTERT gene (5, 6). Moreover, estrogen activates hTERT, in part, via its activation of expression of c-Myc (6), whereas Mad causes transcriptional suppression of hTERT (7). Thus, c-Myc–dependent transformation may be attributed to its aberrant expression coupled with its induction of the positive regulators of cell cycle and concomitant suppression of the growth inhibitory genes.

Expression of c-Myc is regulated at transcriptional, posttranscriptional, as well as posttranslational levels. Multiple pathways, including Ras signaling, are known to regulate c-Myc expression at posttranslational level by modulating its phosphorylation at specific residues during cell cycle progression (8). At the late G1 phase of the cell cycle, the phosphorylated c-Myc is targeted by protein phosphatase 2A (PP2A) that results in enhanced dephosphorylation of c-Myc and its subsequent degradation by ubiquitin-mediated pathway (8). Inhibition of PP2A by agents such as okadaic acid (OA2; a specific inhibitor of PP1 and PP2A), on the other hand, interferes with dephosphorylation of c-Myc, resulting in its elevated expression (9). Although global inhibition of PP2A often results in enhanced transformation of cells (10), exposure to OA has also been shown to inhibit cell growth in part by stimulating apoptosis (11, 12). However, the precise nature of the pathways targeted by OA and potential role of c-Myc in OA-dependent cell growth suppression signaling remain to be elucidated.

A number of studies have indicated involvement of c-Myc in cell growth inhibition pathways, including apoptosis. However, the mechanisms underlying growth suppression by c-Myc are yet to be fully defined. The c-Myc–dependent apoptosis concerns mainly the systems in which its expression has been induced by transfection, viral infection, or promoter-driven transgene methodologies (13). Both p53-dependent, as well as independent, pathways have been suggested in c-Myc–induced apoptosis in different experimental systems. To further define the role of c-Myc in cell growth suppression, we used human breast cancer (HBC) cells in conjunction with OA. Additional agents, such as cycloheximide, as well as chemotherapeutic Taxol, that target cellular pathways different from that of OA were also used to investigate involvement of c-Myc in suppression of cell growth induced by these agents. Cycloheximide-dependent growth inhibitory effects involve targeting of peptidyl transferase activity of the 60S ribosome, blocking of translational elongation, and protein synthesis, whereas Taxol inhibits cell growth in part by targeting microtubule assembly, resulting in inhibition of mitosis, cell cycle arrest, and apoptosis (14, 15). All of these agents attenuated HBC cell growth, whereas only OA or cycloheximide caused elevated expression of c-Myc. We speculated that elevated expression of c-Myc serves as one of the triggers leading to cell growth suppression by agents, such as OA or cycloheximide. Our investigation revealed that depletion of c-Myc abrogated the growth inhibitory effects of OA or cycloheximide, but not Taxol. OA targets zinc finger transcription factor gut-enriched Kruppel-like factor (GKLF) to stimulate c-myc, which, in turn, contributes to HBC growth inhibition. Because cell growth signaling pathways are target of many current anticancer therapies, identification of regulators of c-Myc–dependent growth suppression signaling, such as GKLF, has potential to facilitate development of strategies for effective targeting of breast cancer.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials. DMEM, Ham's F-12 medium, and fetal bovine serum (FBS) were purchased from Life Technologies, Inc. The reagent kit for probe labeling, [{alpha}-32P]dCTP (3,000 Ci/mmol), was purchased from NEN. Oligonucleotides for PCR amplification (see below) were purchased from Integrated DNA Technologies, Inc. OA was purchased from Alexis Biochemicals, and cycloheximide and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) were purchased from Sigma Chemical Company. The chemotherapeutic drug Taxol was obtained from Bristol-Myers-Squibb Company. Neomycin and gentamycin were purchased from Life Technologies-Bethesda Research Laboratories.

Cell lines and cell culture. The ER-negative, p53-negative MDA-MB-231, MDA-MB-468, and MDA-MB-435 HBC, HCT-116 colon cancer, and PC-3 prostate cancer cells were cultured and maintained essentially as described before (16, 17). The HBC and PC-3 cells were cultured routinely in DMEM/Ham's F-12 medium (1:1) supplemented with 5% FBS, whereas HCT-116 cells were cultured in RPMI media supplemented with 10% FBS. Cells were plated either in a 100-mm dish at a density of 2 x 106 cells per dish or in six-well tissue culture plates at a density of 2 x 105 cells per well. The cells were transfected with phosphorylated Rous sarcoma virus-ß (pRSV-ß), which encodes for ß-galactosidase, in combination with different luciferase constructs (see below), and ß-galactosidase and luciferase activities were measured essentially as described (17). In addition, MDA-MB-468 HBC cells were transfected with a plasmid encoding c-Myc antisense, and stable sublines were isolated and characterized as detailed before (16).

RNA isolation and Northern and Western blot analyses. Logarithmically growing cells were either untreated or treated with OA, Taxol, or cycloheximide. The cells were then harvested, cellular RNAs were isolated, and 20 µg of each of the RNAs were analyzed by Northern blot analyses, in conjunction with radio-labeled cDNA probes, essentially as described (17). In addition, both the floating and adherent cells were harvested and lysed, and the protein extracts were analyzed by Western immunoblotting with anti–c-myc antibody (PharMingen) and subsequently with anti-actin antibody (Sigma Biochemicals), essentially as per the manufacturer's guidelines and our previously described protocols (18).

Cell viability and apoptosis assays. Cell viabilities were measured by MTT assay as described (16). Cells (2 x 104) were seeded in six-well plates and treated with or without various drugs (see below). Cells were then incubated with MTT stock solution followed by treatment with acidic isopropanol, essentially as described before (16). The assessment of the live cells was derived by measuring the absorbance of the converted dye at a wavelength of 570 nm. In addition, the apoptosis levels were assessed by ELISA-based DNA fragmentation assay (Roche Diagnostics) by measuring the "enrichment factor" indicating the level of apoptosis, essentially by the manufacturer suggested formula and our previously described methods (16).

Flow cytometric analysis. Flow cytometric analysis of DNA content was done to assess the cell cycle phase distribution as described previously (19). Normally growing parental wild-type, the vector plasmid pcDNA3 transfected, or c-Myc–antisense clone 3 and clone 6 sublines were independently stained for DNA content using propidium iodide. After staining, cells were analyzed on a Becton Dickinson FACScan cytometer. The data were analyzed using the multicycle program from Phoenix Flow Systems.

Plasmids and cDNA probes. The plasmids having mouse c-myc promoter (–1,100 to +580, –424 to +580, –140 to +340, and +70 to +160 relative to the P1 transcription site) driving luciferase reporter gene (20) were kindly provided by Dr. George Stark of the Cleveland Clinic. The plasmid pcDNA3-Myc containing the entire c-Myc-2 open reading frame has been described before (16). The c-Myc promoter subfragments were PCR amplified using the plasmid –1,138 to +580 Luc as template, and a combination of oligos Myc 5 (5' CTCCCCGGGCTCCCGAGTTCCCAAAGCAG 3'; sense oligo from positions –140 to –120) and Myc 7 (5' CTCCTCGAGTACTACAGCGAGTCAGAAAA 3'; antisense oligo from positions +156 to +175) or Myc 6 (5' CTCCCCGGGGCGGCCGAGGACCCCTGGCT 3'; sense oligo from positions –11 to +10) and Myc 7. The XmaI plus XhoI cut PCR products were ligated into pGL2-Basic (Promega) vector plasmid to obtain –140 to +175 Luc and –11 to +175 Luc constructs. In addition, c-myc promoter fragment from positions –11 to + 70 was PCR amplified using oligo Myc 6 in combination with oligo Myc 8 (5' CTCAAGCTTCCCTTCAGGAGGCAGGAG 3'; antisense oligo from positions +55 to +72), digested with XmaI plus HindIII, and ligated into vector plasmid pBSK to obtain pBSK–c-Myc clone 1. Next, mutations in the GKLF consensus sequence were introduced by a PCR-based strategy. A 65-nucleotide primer, Myc 9 (5' CTCAAGCTTCCCTTCAGGAGGCAGGAGGGGAGCTGAGATTTTTTTATCGGACCCGGCAGCTGAGA 3'; antisense oligo from positions +17 to +72), was synthesized to introduce mutations at the GKLF consensus sequence site (21, 22). PCR-amplified DNA fragment using oligos Myc 6 and Myc 9 was then subcloned into vector plasmids pBSK to obtain pBSK–c-Myc (mut) clone 1. The inserts of pBSK–c-Myc clone 1 and pBSK–c-Myc (mut) clone 1 plasmids were then subcloned into pGL2-promoter plasmid (Promega) to obtain –11 to +70 (wt) SV40-Luc and –11 to +70 (mut) SV40-Luc constructs, respectively.

In addition, a 310-bp GKLF cDNA fragment was PCR amplified using reverse transcribed RNA from MDA-MB-468 HBC cells and a combination of oligos GKLF-1 (5' AGAGAATTCAGTATTTTTTACTTTTCACAC 3'; sense oligo from positions 1,777 to 1,806; ref. 21) and GKLF-2 (5' AGGGAATTCTGGTCTTCCCTCCCCCAAC 3'; antisense oligo from positions 2,061 to 2,088; ref. 21). The PCR DNA was digested with EcoRI and cloned into vector plasmid pcDNA3 (Invitrogen) to obtain pcDNA3-GKLF clone 1. Further, a 48-bp subfragment of p21WAF1/CIP1 3' untranslated region (UTR; positions +892 to +939; ref. 23) was generated by synthesizing 52-mer each of the sense and antisense oligos. These oligos were phosphorylated by using T4 polynucleotide kinase and annealed, and the double-stranded subfragment was subcloned into vector plasmid pBSK to obtain pBSK-WAF clone 42.C.12. All the plasmids used in this report were sequenced to confirm the validity of the inserts and are summarized in Table 1 . The 310-bp GKLF cDNA fragment of pcDNA3-GKLF clone 1 plasmid was excised by digestion with EcoRI, gel purified, and subsequently used as radio-labeled probe for Northern blot hybridization.


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  		Table 1.

 
Gel electrophoretic mobility shift assays. The c-myc promoter fragment from positions –11 to +72 was radiolabeled by end-filling using Klenow fragment of DNA polymerase and [{alpha}-32P]dCTP, followed by its purification on G-50 sephadex spin column. Nuclear protein extracts from untreated, as well as OA-treated or cycloheximide-treated, HBC cells were prepared, DNA-protein binding was carried out in vitro, and DNA-bound complexes were analyzed by gel electrophoretic mobility shift assays as described before (17, 24, 25). The specific competitor DNA consist gel-purified c-myc promoter fragment from positions –11 to +72, whereas the nonspecific competitor DNA consist 48-bp gel-purified p21WAF1/CIP1 3' UTR fragment of plasmid pBSK-WAF clone 42.C.12. The binding reactions were electrophoresed onto 5% nondenaturing polyacrylamide gels, the gels were dried at 60°C, and DNA-protein binding was visualized by autoradiography.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
OA and cycloheximide enhance c-myc expression in HBC cells. Agents such as OA, cycloheximide, or Taxol attenuate growth of diverse cell types in part by causing apoptosis (11, 12, 14, 15). To investigate mechanisms of HBC growth suppression by these agents, HBC cells were exposed to 120 nmol/L OA, 0.7 mmol/L cycloheximide, or 30 µmol/L Taxol for varying periods of time, followed by determination of cell viability using MTT assay. Treatments with OA for 6 h, cycloheximide for 10 h, or Taxol for 36 h resulted in ~80% loss in viability of HBC cells (Fig. 1A–C ). The loss of cell viability by these agents was due, in part, to apoptotic cell death as evidenced by fragmented cell nuclei of the treated cells (not shown), suggesting that apoptosis contributes to reduced cell viabilities by each of the agent. Next, HBC cells were treated with either 120 nmol/L OA for 6 h or 0.7 mmol/L cycloheximide for 10 h. Untreated, as well as treated, cells were then harvested, and total RNAs were isolated and analyzed by Northern blot hybridization to determine c-myc expression. Treatments with OA or cycloheximide caused increased expression of c-myc mRNA (Fig. 1D, lanes 3, 4, 7, and 8), consistent with our observations of elevated c-myc levels by OA in the ER-negative MDA-MB-435 (17) and ER-positive MCF-7 HBC cells (not shown). Treatments with Taxol, however, failed to increase c-myc mRNA levels in HBC cells (not shown). Data in Fig. 1 suggest that, although all the agents used in this study suppressed growth of HBC cells, c-Myc expression was likely associated with HBC cell growth inhibitory effects of OA or cycloheximide.


Figure 1
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  		Figure 1. Attenuation of HBC cell viabilities and stimulation of c-myc expression by OA and cycloheximide (CHX). HBC cells were treated with noted doses of OA (A), cycloheximide (B), or Taxol (C) for the indicated times. MTT assays were carried out as in Materials and Methods. C, the MTT values are plotted in log scale. Columns, means of four independent experiments; bars, SE; WT, wild-type. D, the indicated wild-type HBC cells were either untreated (control), treated with 120 nmol/L OA for 6 h, or 0.7 mmol/L cycloheximide for 10 h. Expression of c-myc was analyzed by Northern blot hybridization as in Materials and Methods. Ethidium bromide stain of the RNA gel is also shown to assess RNA loading, as indicated by the signals for 28S and 18S rRNAs.

 
c-Myc expression plays a role in HBC growth suppression by OA or cycloheximide. Whether elevated levels of c-Myc play a role in inhibition of HBC cell growth by OA or cycloheximide was investigated by using c-Myc antisense sublines that express reduced levels of c-Myc. Two sublines, c-Myc antisense clone 3 and clone 6 that express 50% reduced levels of c-Myc when compared with their vector-transfected or wild-type counterparts were isolated and characterized by us before (16). Given that c-Myc targets diverse pathways of cell cycle control, we first determined whether loss of c-Myc causes alterations in cell cycle progression of c-Myc antisense sublines. These sublines, along with their wild-type or vector-transfected counterparts, were subjected to flow cytometric analysis after labeling with propidium iodide. Reduced expression of c-Myc did not alter cell cycle phase distribution, as well as DNA content of c-Myc antisense HBC cells (not shown). Next, we determined whether loss of c-Myc interferes with the growth inhibitory effects of OA, cycloheximide, or Taxol. Wild-type MDA-MB-468 HBC cells, the vector pcDNA3 clone 1, or the c-Myc antisense clones 3 and 6 HBC sublines were independently treated with either 120 nmol/L OA for 6 h, 0.7 mmol/L cycloheximide for 10 h, or 30 µmol/L Taxol for 36 h, and their viabilities were determined by MTT assays. As expected, each of the agents caused up to ~80% loss of viability of wild-type or vector-transfected HBC cells. In contrast, treatments with cycloheximide or OA, but not with Taxol, failed to significantly reduce viabilities of c-Myc antisense cells (Fig. 2A–C ). The failure of OA to attenuate viability of c-Myc antisense HBC cells was due, in part, to the reduced apoptosis when compared with OA-dependent apoptosis in the vector-transfected controls (Fig. 2D). Data in Fig. 2 indicate that loss of c-Myc interferes with HBC growth inhibitory effects of OA.


Figure 2
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  		Figure 2. Loss of c-Myc interferes with HBC growth suppression by OA or cycloheximide. Indicated cell types were either untreated (Control) or treated independently with noted doses of OA for 6 h (A), cycloheximide for 10 h (B), or Taxol for 24 h (C). MTT assays were carried out as in Materials and Methods. D, Sublines were treated with 120 nmol/L dose of OA for 6 h, and apoptosis levels were determined by ELISA-based apoptosis assay as in Materials and Methods. Columns, means of four independent experiments; bars, SE.

 
The extent levels of c-Myc contribution to growth inhibitory effects of OA was investigated next by using c-Myc antisense clone 3 and clone 6 HBC cells. These cells were either untreated or independently treated with 120 nmol/L of OA for various periods of time, followed by determination of their viabilities. As shown in Fig. 3A and B , OA treatments for periods of 6 and 12 h resulted in minimal, statistically insignificant loss in their viabilities when compared with the viabilities of their untreated counterparts. Exposure to OA for periods of >18 h, however, resulted in significant loss of their viability, to the extent that only ~20% or less viable cells were present when OA treatments were extended to 36 and 48 h of time periods (Fig. 3A and B). Whether growth suppression of c-Myc antisense cells by OA over 36 to 48 h of time periods involves c-Myc expression was investigated by Western immunoblot analyses. Wild-type HBC cells or their vector-transfected counterparts were either untreated or treated with OA for 6 h, whereas c-Myc antisense sublines were either untreated or treated with OA for 12, 36, and 48 h. The Western blot analysis revealed that although 6-h treatments with OA caused elevated expression of c-Myc in the wild-type or the vector-transfected HBC cells, exposure of c-Myc antisense cells to OA for 12 h failed to increase c-Myc levels. The c-Myc antisense cells that were exposed to OA for 36 or 48 h, on the other hand, displayed elevated levels of c-Myc (Fig. 3C and D). The data in Figs. 2 and 3 suggest that increased expression of c-Myc is involved in OA-dependent HBC growth inhibition, whereas loss of c-Myc interferes with growth inhibitory effects of OA.


Figure 3
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  		Figure 3. Prolonged exposure to OA attenuates viabilities and stimulates c-Myc expression in HBC sublines expressing reduced c-Myc. c-Myc antisense clone 3 (A) or clone 6 (B) HBC sublines were either untreated (control; filled columns) or treated with OA (open columns) for indicated times. MTT assays were carried out as in Materials and Methods. Columns, means of four independent experiments; bars, SE. C and D, HBC cells were either untreated (noted as 0 above respective lanes) or exposed to noted dose and times of OA. Thirty micrograms of each of the protein lysates were analyzed on SDS-PAGE gels, and c-myc expression was determined by immunoblotting as in Materials and Methods. The protein loading was assessed by reprobing the membranes with antiactin antibody.

 
OA targets c-Myc expression, in part, by transcriptional mechanisms. Because OA induced c-Myc mRNA (Fig. 1D), as well as protein (Fig. 3C and D), we speculated that this increase in c-myc expression was due, in part, to transcriptional mechanisms. To test this possibility, we used various c-myc promoter constructs listed in Table 1. HBC cells were transfected with various c-myc promoter constructs in combination with ß-galactosidase expression plasmid. First, the plasmids –1,138 to +580 Luc, –424 to +580 Luc, –140 to +340 Luc, –140 to +175 Luc, –11 to +175 Luc, and +70 to +164 Luc were separately transfected into MDA-MB-468 HBC cells. In certain cases, ~40 h posttransfection, the cells were treated with 120 nmol/L OA for 6 h, and both the controls and treated cultures were harvested and assayed for luciferase activities. All the constructs, except +70 to +164 Luc, elicited ~2.5-fold to 3.5-fold increase in the luciferase activities in the presence of OA (Fig. 4A and B ), suggesting that –11 to +70 region of mouse c-myc gene harbors OA-responsive elements. The OA responsiveness of the –11 to +70 c-myc promoter subfragment was further evaluated by using additional HBC, as well as colon carcinoma HCT-116 and prostate carcinoma PC-3, cells. First, various HBC cells were transfected with the –11 to +175 Luc construct, and luciferase activities in the absence or presence of OA were determined. As expected, OA caused ~3-fold to 5-fold increase in luciferase activities of –11 to +175 Luc construct (Fig. 4C). Whether OA also stimulates c-myc promoter in HCT-116 and PC-3 cells was investigated next. The plasmids –11 to +175 Luc or +70 to +164 Luc were transfected in these cells, and luciferase activities in the absence or presence of OA were determined. Consistent with our data in Fig. 4B, OA stimulated activity of –11 to +175 Luc plasmid in these cells (Fig. 4D). Data in Fig. 4 strongly suggest that OA targets c-myc promoter sequences within positions –11 to +70 in a cell type–independent fashion.


Figure 4
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  		Figure 4. OA regulates c-myc promoter activities. HBC cells (A and B) were transfected with indicated c-myc promoter construct in combination with pRSV-ß plasmid. C, wild-type HBC cells were transfected with noted c-Myc promoter construct in combination with pRSV-ß plasmid. In addition, colon carcinoma HCT-116 cells or prostate carcinoma PC-3 cells (D) were transfected with indicated c-Myc promoter constructs in combination with pRSV-ß plasmid. Approximately 40 h after transfections, the cells were then either untreated (Control) or treated with indicated dose of OA for 6 h, and luciferase and ß-galactosidase activities were measured as in Materials and Methods. The luciferase activities were expressed as light units and normalized to ß-galactosidase activities expressed as absorbance. Columns, means of three independent experiments expressed relative to luciferase activities obtained from untreated controls, which were arbitrarily defined as 1; bars, SE.

 
OA and cycloheximide target HBC nuclear protein(s) that specifically interact with OA-responsive c-myc promoter sequences. HBC cells were either untreated, treated with 120 nmol/L OA for 6 h, or treated with 0.7 mmol/L cycloheximide for 10 h, and the nuclear protein extracts were prepared as in Materials and Methods. Gel electrophoretic mobility shift assays were used to investigate binding of HBC nuclear proteins in vitro with the OA-responsive region of the c-myc promoter. As shown in Fig. 5 , the –11 to +72 subfragment binds multiple complexes present in the nuclear extracts of all the HBC cells (lanes 2, 4, 5, 7, 8, 10, 11, 13, 14, 16, 17, and 19). The DNA-protein(s) binding was competed away by 200-fold molar excess of the unlabeled double-stranded –11 to +72 DNA fragment (lanes 3, 6, 9, 12, 15, and 18), whereas the binding could not be competed away by a 200-fold molar excess of double-stranded 48 bp of cDNA p21WAF1/CIP1 3' UTR (positions 892–939; ref. 23) subfragment (lanes 4, 7, 10, 13, 16, and 19). Increased binding of a high molecular weight complex (location denoted by * in Fig. 5) was noted in the case nuclear protein extracts obtained from HBC cells treated with either 120 nmol/L OA (A, lanes 5, 7, 11, 13, 17, and 19; B, lanes 5 and 7) or with 0.7 mmol/L cycloheximide (B, lanes 11 and 13). The data in Fig. 5 suggest that OA or cycloheximide cause enhanced binding of specific HBC nuclear proteins to the OA-responsive c-myc promoter sequences.


Figure 5
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  		Figure 5. Binding of HBC cell nuclear proteins to OA-responsive c-myc promoter sequences. Preparation of the nuclear proteins, their binding with radio-labeled probe, and analysis of the DNA-protein complexes by electrophoretic mobility shift assays was as detailed in Materials and Methods. Nuclear extracts in (A), lanes 5 to 7, 11 to 13, and 17to 19, and (B), lanes 5 to 7, were derived from the HBC cells treated with 120 nmol/L OA for 6 h, whereas nuclear extracts in lanes 11 to 13 of (B) were derived from HBC cells treated with 0.7 mmol/L cycloheximide for 10 h. Lanes 1 and 20, probe only; lanes 2, 5, 8, 11, 14, and 17, probe and nuclear protein extracts; lanes 3, 6, 9, 12, 15, and 18, probe, nuclear extracts, and 200-fold excess of cold unlabeled probe DNA; lanes 4, 7, 10, 13, 16, and 19, probe, nuclear extracts, and 200-fold excess of cold unlabeled nonspecific fragment DNA. *, location of putative OA-responsive DNA-protein(s) complex.

 
OA increases c-myc expression, in part, by targeting GKLF. Database search revealed the presence of a consensus sequence for zinc finger transcription factor GKLF in the OA-responsive –11 to +70 c-myc promoter subfragment (Fig. 6A ). Whether GKLF consensus sequences are involved in OA-dependent increase in c-myc promoter activities in HBC cells was investigated next. The plasmids SV40-Luc (pGL2-Promoter, Promega), –11 to +70 (wt) SV40-Luc, or –11 to +70 (mut) SV40-Luc in combination with plasmid pRSV-ß were independently transfected into MDA-MB-468 HBC cells. The cells were either untreated or treated with 120 nmol/L OA for 6 h, followed by assays for luciferase and ßGal activities. OA caused ~2.0-fold increase in luciferase activities of –11 to +70 (wt) SV40-Luc, whereas no significant increase in luciferase activities was noted for either SV40-Luc or –11 to +70 (mut) SV40-Luc (Fig. 6B), suggesting that OA stimulates c-myc promoter, in part, by targeting GKLF consensus sequences. The zinc finger transcription factor GKLF was revealed as a target of OA by Northern blot analysis, wherein MDA-MB-468 HBC cells treated with 120 nmol/L OA for 6 h were found to have increased levels of GKLF mRNA (Fig. 6C). Together, the data in Fig. 6 strongly suggest that elevated expression of c-myc by OA involves the zinc finger transcription factor GKLF.


Figure 6
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  		Figure 6. OA regulation of c-myc transcription involves GKLF cis sequences. A, the GKLF consensus sequence, its location on the antisense strand in the c-myc promoter (underlined, bold block letters), as well as c-myc promoter with mutant GKLF cis sequences. B, HBC cells were transfected with the indicated luciferase constructs in combination with pRSV-ß. Approximately 40 h after transfections, the cells were either untreated (–OA) or treated with 120 nmol/L OA (+OA) for 6 h, and luciferase and ß-galactosidase activities were measured as in Materials and Methods. The luciferase activities were expressed as light units and normalized to ß-galactosidase activities expressed as absorbance. Columns, means of three independent experiments expressed relative to luciferase activities obtained from untreated controls, which were arbitrarily defined as 1; bars, SE. C, HBC cells were either untreated (lane 1), treated with 120 nmol/L OA for 0.5, 1, 2, 4, and 6 h (lanes 2, 3, 4, 5, and 6, respectively), followed by RNA extraction and Northern blot hybridization with radiolabeled GKLF cDNA probe as in Materials and Methods. Ethidium bromide stain of the RNA gel is also shown to assess RNA loading, as indicated by the signals for 28S and 18S rRNAs.

 

   Discussion
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We have previously observed that submicromolar doses of OA stimulated c-myc expression in HBC cells (17). Here, we tested a hypothesis that cellular level of c-Myc is a critical regulator of signaling pathways regulating cell growth, and expression of c-Myc beyond a threshold may cause growth suppression of the target cells. By using HBC sublines that express antisense mediated reduced levels of c-Myc (16), we show that loss of c-Myc interferes with their growth attenuation by OA. Although, depletion of c-Myc did not cause alterations in the cell cycle phase distribution or the DNA contents of these cells, prolonged exposure to OA, nevertheless, resulted in elevated c-Myc and their growth suppression (see Fig. 3). These observations support a compelling argument that expression of c-Myc beyond a threshold likely attenuates HBC cell growth.

Given that OA is also a tumor promoter and activates activator protein (AP-1)–mediated signaling in different cell types, including HBC cells (ref. 17 and references therein), it is conceivable that OA stimulation of c-myc transcription entailed aberrant induction of pathways used by mitogenic stimuli. In this context, c-myc transcription was found to be regulated by Src kinase using Rac-dependent pathway (26, 27), by a KH domain protein FKB that binds an element 1.5 kb upstream of the c-myc promoter (28), by growth factor pathways involving mitogen-activated protein kinase (MAPK) signaling, by transcription factors, such as nuclear factor-{kappa}B (NF-{kappa}B), E2Fs (ref. 29, and refs. within), or by cytokine signaling pathways that converge on transcription factors, such as signal transducers and activators of transcription (STAT; ref. 20 and references therein). OA-stimulated c-Myc expression in HBC cells, independent of the transcription factors AP-1, NF-{kappa}B, E2F-1, FKB, or STATs, is supported by the fact that the OA-responsive c-myc promoter sequences lack consensus elements for any of these transcription factors. Furthermore, expression of wild-type extracellular signal-regulated kinase 2 (ERK2) or STAT3 (20) proteins failed to enhance transcriptional activity of –11 to +175 Luc construct in the presence of OA (not shown). In addition, HBC cells transfected with –11 to +175 Luc construct that were either treated with OA, a combination of OA and 30 µmol/L PD98059 (pharmacologic inhibitor of MAP/ERK kinase 1) or OA and 25 µmol/L SB203580 [pharmacologic inhibitor of p38 stress-activated protein kinase (SAPK) {alpha}/ß; ref. 30], also failed to show differences in activation of c-myc promoter (data not shown). Thus, OA stimulation of c-Myc in HBC cells is likely independent of signaling pathways involving STAT3, ERK/MAPK, or p38/SAPK. Since p38 SAPKs are expressed as multiple isoforms, including p38 {alpha}, ß, {gamma}, and {delta}, and SB203580 has been shown to selectively inhibit p38 {alpha}/ß isoforms only (ref. 30 and references therein), it remains to be determined whether p38 isoforms {gamma} and {delta} or c-Jun-NH2-kinase (JNK) family of SAPKs are involved in stimulation of c-myc transcription by OA in HBC cells.

Overexpression of c-Myc in the absence of survival signals (such as growth factors) causes apoptosis (31–34). The induction of apoptosis by c-Myc involves p53-dependent, as well as independent, pathways (13), including ADP ribosylation factor pathway and stimulation of cytochrome c release (35, 36). Expression of c-Myc destabilizes mitochondrial integrity in part by functionally cooperating with Bax to induce apoptosis, as well as potentiates the mitochondrial pathways of apoptosis by acting upstream of apoptosis signal-regulating kinase 1 in the p38 signaling cascade (37, 38). In the cells with defective intrinsic apoptosis signaling, c-Myc expression sensitizes cells to TRAIL-mediated cell death, in part, by blocking NF-{kappa}B activity (39). Since OA or cycloheximide, while targeting distinct intracellular signaling by inhibiting PP1/PP2A or protein synthesis pathway, respectively, causes elevated expression of c-Myc and cell growth inhibition, it is unclear whether and to what extent c-Myc in turn targets mitochondria involving interactions with proteins, such as Bax and cytochrome c release to promote apoptosis. Nevertheless, since the HBC cells used in this report possess a mutant nonfunctional p53 (40), OA-mediated or cycloheximide-mediated effects likely involve p53-independent pathways. Our findings are also consistent with a study describing a role for c-Myc in DNA damage–induced apoptosis of a small cell lung cancer cell line independent of the involvement of p53 (41). A moderate increase in expression of c-Myc, as well as proapoptotic protein Bax, was noted when the small cell lung cancer cell line POGB was exposed to the DNA-damaging agents, such as doxorubicin (200 ng/mL) for 18 to 48 h or {gamma}-radiation (8 Gy) for 18 h (41). Although, doxorubicin causes apoptosis in HBC cells, it, nevertheless, does not induce c-Myc and Bax proteins.6 The extent OA-dependent or cycloheximide-dependent HBC growth inhibition uses DNA damage pathways is unclear. Apoptosis-inducing stimuli, such as UV and Taxol, have been found to activate JNK, which, in turn, regulated c-Myc–mediated apoptosis involving phosphorylation of c-Myc at Ser62 and Ser71 in NIH3T3 cells (42). In addition, oncogene Ras-mediated phosphorylation of c-Myc at Ser62 has been implicated to regulate c-Myc stability in the initial stages of cell proliferation (8, 43). Whether additional posttranslational mechanism(s) are used by OA to regulate c-Myc expression is a subject of our ongoing studies.

Previously, we found that OA-dependent c-myc expression was accomplished, in part, by posttranscriptional mechanisms using adenosine-uridine rich elements (ARE) present in the 3' UTR of the c-myc mRNA (17). OA caused increased binding in vitro of a novel ~75 kDa HBC cell cytosolic protein to c-myc 3' UTR sequences containing AREs. This report shows that OA up-regulates c-Myc by transcriptional mechanisms involving enhanced binding of HBC cell nuclear proteins to the OA-responsive elements of the c-myc promoter (Figs. 4 and 5). Together with our current findings demonstrating involvement of GKLF in regulating c-Myc expression, our studies not only highlight a pathway regulating complex OA-dependent growth suppression mechanisms in cancer cells but also reveal a mediator of signaling with potential to serve as target for design and development of efficacious anticancer therapeutics.


   Acknowledgments
 
Grant support: Detroit Medical Center Institute for Oncology and Allied Diseases Institutional Research grant (A.K. Rishi), Department of Veterans Affairs Medical Research Services (A. Wali and A.K. Rishi), and Susan G. Komen Foundation for Breast Cancer Research (A.K. Rishi).

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.

We thank William Browning of John D. Dingell Veterans Affairs Medical Center Medical Media Department for preparing the illustrations and Drs. Madan Boyanapalli and Adhip Majumdar for the expert technical assistance and critical comments, respectively.


   Footnotes
 
6 Unpublished observation. Back

Received 7/ 5/07. Revised 8/20/07. Accepted 8/31/07.


   References
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 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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