Proc Amer Assoc Cancer Res, Volume 47, 2006
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To improve prostate cancer (PCa) radiotherapy, we have previously shown that purified genistein, the major component of soy, was a potent radiosensitizer and potentiated tumor cell killing in vitro and in vivo . Pre-treatment with genistein enhanced radiation-induced cell killing of PC-3 human PCa cells, and genistein administered before and after tumor irradiation enhanced primary tumor growth inhibition and controlled metastasis to lymph nodes using an orthotopic metastatic PCa model of PC-3 cells in nude mice. Paradoxically, in this model, single therapy with purified genistein caused increased spontaneous metastasis to para-aortic regional lymph nodes. To clarify whether a formulation of soy (43% genistein + 21% daidzein + 2% glycitein) representative of the soy pills used to treat patients abrogates increased metastasis observed with pure genistein, we investigated the effect of soy alone and combined with radiation on PCa in vitro and in vivo . We found that the formulation of soy was more potent than pure genistein at inhibition of PC-3 cell colony formation and potentiated radiation-induced cell killing in a clonogenic assay. Furthermore, soy was more potent than pure genistein at decreasing activation of NF-κB, a transcription factor critically involved in cancer cell survival. Like pure genistein, soy also abrogated radiation-induced activation of NF-κB. We also investigated the effect of soy or genistein alone and combined with radiation on the expression of APE1/Ref-1, a DNA repair/redox protein implicated in tumor cell survival and radioresistance. Soy and genistein alone were effective at downregulating APE1/Ref-1 expression, with soy being more potent, while each considerably reduced APE1/Ref-1 upregulation in response to radiation. Based on studies demonstrating a link between APE1/Ref-1 redox activity and NF-κB, we propose a mechanism of increased cell death following treatment with soy isoflavones by decreasing APE1/Ref-1 redox-mediated activation of NF-κB. This hypothesis was confirmed by concomitant changes in cleaved PARP protein, a marker for apoptosis. Moreover, soy treatment alone did not promote increased metastasis to lymph nodes in contrast to purified genistein in our in vivo model. The combination of soy with prostate tumor irradiation led to greater control of primary tumor growth and metastasis than soy or radiation alone, similar to our previous findings with pure genistein. Histologically, tumors treated with soy and radiation confirmed greater tumor destruction with extensive apoptosis, fibrosis and abnormalities in residual tumor cells, e.g., atypical giant cells, than tumors treated with soy or radiation alone. Our studies demonstrate the safety and potential for the use of soy formulation to enhance clinical strategies for prostate cancer radiotherapy and suggest potential targets for novel therapeutic agents.
Apurinic/apyrimidinic (AP) endonuclease (APE) is a multifunctional protein possessing both DNA repair and redox regulatory activities. In base excision repair (BER), APE is responsible for processing spontaneous, chemical, or monofunctional DNA glycosylase-initiated AP sites via its 5′-endonuclease activity and 3′-"end-trimming" activity when processing residues produced as a consequence of bifunctional DNA glycosylases. In this study, we have fully characterized a mammalian model of APE haploinsufficiency by using a mouse containing a heterozygous gene-targeted deletion of the APE gene (Apex+/–). Our data indicate that Apex+/– mice are indeed APE-haploinsufficient, as exhibited by a 40–50% reduction (p < 0.05) in APE mRNA, protein, and 5′-endonuclease activity in all tissues studied. Based on gene dosage, we expected to see a concomitant reduction in BER activity; however, by using an in vitro G:U mismatch BER assay, we observed tissue-specific alterations in monofunctional glycosylase-initiated BER activity, e.g. liver (35% decrease, p < 0.05), testes (55% increase, p < 0.05), and brain (no significant difference). The observed changes in BER activity correlated tightly with changes in DNA polymerase β and AP site DNA binding levels. We propose a mechanism of BER that may be influenced by the redox regulatory activity of APE, and we suggest that reduced APE may render a cell/tissue more susceptible to dysregulation of the polymerase β-dependent BER response to cellular stress. Apurinic/apyrimidinic (AP) endonuclease (APE) is a multifunctional protein possessing both DNA repair and redox regulatory activities. In base excision repair (BER), APE is responsible for processing spontaneous, chemical, or monofunctional DNA glycosylase-initiated AP sites via its 5′-endonuclease activity and 3′-"end-trimming" activity when processing residues produced as a consequence of bifunctional DNA glycosylases. In this study, we have fully characterized a mammalian model of APE haploinsufficiency by using a mouse containing a heterozygous gene-targeted deletion of the APE gene (Apex+/–). Our data indicate that Apex+/– mice are indeed APE-haploinsufficient, as exhibited by a 40–50% reduction (p < 0.05) in APE mRNA, protein, and 5′-endonuclease activity in all tissues studied. Based on gene dosage, we expected to see a concomitant reduction in BER activity; however, by using an in vitro G:U mismatch BER assay, we observed tissue-specific alterations in monofunctional glycosylase-initiated BER activity, e.g. liver (35% decrease, p < 0.05), testes (55% increase, p < 0.05), and brain (no significant difference). The observed changes in BER activity correlated tightly with changes in DNA polymerase β and AP site DNA binding levels. We propose a mechanism of BER that may be influenced by the redox regulatory activity of APE, and we suggest that reduced APE may render a cell/tissue more susceptible to dysregulation of the polymerase β-dependent BER response to cellular stress. Apurinic/apyrimidinic (AP) 1The abbreviations used are: AP, apurinic/apyrimidinic; APE, apurinic/apyrimidinic endonuclease; BER, base excision repair; β-pol, polymerase β; MFG, monofunctional glycosylase; dRP, 5′-deoxyribose 5-phosphate; EMSAs, electrophoretic mobility shift assays; DTT, dithiothreitol; PBS, phosphate-buffered saline; BSA, bovine serum albumin; RT, reverse transcriptase; I.D.V., integrated density value; ROPS, random oligonucleotide primed synthesis; ASB, aldehyde-reactive probe slot blot. endonuclease (APE) is a multifunctional protein involved in the maintenance of genomic integrity and in the regulation of gene expression. After the initial discovery in Escherichia coli (1Verly W.G. Paquette Y. Can. J. Biochem. 1972; 50: 217-224Google Scholar), APE was purified from calf thymus DNA and extensively characterized as an endonuclease that cleaves the backbone of double-stranded DNA containing AP sites (2Ljungquist S. Lindahl T. J. Biol. Chem. 1974; 249: 1530-1535Google Scholar, 3Ljungquist S. Andersson A. Lindahl T. J. Biol. Chem. 1974; 249: 1536-1540Google Scholar). APE homologues were subsequently identified and characterized in many organisms, including yeast as APN1 (4Popoff S.C. Spira A.I. Johnson A.W. Demple B. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 4193-4197Google Scholar), mice as Apex (5Seki S. Akiyama K. Watanabe S. Hatsushika M. Ikeda S. Tsutsui K. J. Biol. Chem. 1991; 266: 20797-20802Google Scholar, 6Seki S. Ikeda S. Watanabe S. Hatsushika M. Tsutsui K. Akiyama K. Zhang B. Biochim. Biophys. Acta. 1991; 1079: 57-64Google Scholar), and humans as HAP1 (7Robson C.N. Hickson I.D. Nucleic Acids Res. 1991; 19: 5519-5523Google Scholar). In addition to its major 5′-endonuclease activity, APE also expresses minor 3′-phosphodiesterase, 3′-phosphatase, and 3′ → 5′-exonuclease activities (8Wilson III, D.M. Sofinowski T.M. McNeill D.R. Front. Biosci. 2003; 8: D963-D981Google Scholar), the biological significance of which is controversial (9Lebedeva N.A. Khodyreva S.N. Favre A. Lavrik O.I. Biochem. Biophys. Res. Commun. 2003; 300: 182-187Google Scholar). Independent of its discovery as a DNA repair protein, APE was also characterized as REF-1, for redox factor-1, a redox activator of cellular transcription factors (10Xanthoudakis S. Curran T. EMBO J. 1992; 11: 653-665Google Scholar, 11Xanthoudakis S. Miao G. Wang F. Pan Y.C. Curran T. EMBO J. 1992; 11: 3323-3335Google Scholar, 12Evans A.R. Limp-Foster M. Kelley M.R. Mutat. Res. 2000; 461: 83-108Google Scholar). Although the molecular detail of APE redox activity is still unclear (13Ordway J.M. Eberhart D. Curran T. Mol. Cell. Biol. 2003; 23: 4257-4266Google Scholar), the discovery of APE as a regulator of transcriptional activity may underscore the importance of its involvement in cellular stress-response pathways. APE is the primary enzyme responsible for recognition and incision of non-coding AP sites in DNA arising as a consequence of spontaneous, chemical, or DNA glycosylase-mediated hydrolysis of the N-glycosyl bond initiated by the base excision repair (BER) pathway. These lesions are particularly common, arising at the rate of ∼50,000–200,000 AP sites per cell per day under normal physiological conditions (14Nakamura J. Walker V.E. Upton P.B. Chiang S.-Y. Kow Y.W. Swenberg J.A. Cancer Res. 1998; 58: 222-225Google Scholar, 15Nakamura J. Swenberg J.A. Cancer Res. 1999; 59: 2522-2526Google Scholar). Unrepaired AP sites may threaten genomic stability by serving as blocks to DNA replication (16Schaaper R.M. Kunkel T.A. Loeb L.A. Proc. Natl. Acad. Sci. U. S. A. 1983; 80: 487-491Google Scholar), by stalling RNA polymerase II during transcription (17Yu S.-L. Lee S.-K. Johnson R.E. Prakash L. Prakash S. Mol. Cell. Biol. 2003; 23: 382-388Google Scholar), or by promoting topoisomerase II-mediated double strand breaks (18Wilstermann A.M. Osheroff N. J. Biol. Chem. 2001; 276: 46290-46296Google Scholar). Of the repair pathways available to a cell (19Friedberg E.C. Nature. 2003; 421: 436-440Google Scholar, 20Christmann M. Tomicic M.T. Roos W.P. Kaina B. Toxicology. 2003; 193: 3-34Google Scholar), BER is the main pathway responsible for repairing AP sites in DNA. Initiation of BER is made possible by recognition of a damaged base by either a monofunctional or bifunctional DNA glycosylase, in addition to AP site recognition by APE. In monofunctional glycosylase-initiated BER (MFG-BER), a damaged or improper base is recognized and removed by enzymatic hydrolysis of the N-glycosyl bond resulting in the formation of an AP site. AP sites serve as a substrate for APE, which incises the DNA backbone immediately 5′ to the AP site via its 5′-endonuclease activity, producing a single strand break with a normal 3′-hydroxyl group and an abnormal 5′-deoxyribose 5-phosphate (dRP) residue (21Friedberg E.C. Walker G.C. Siede W. DNA Repair and Mutagenesis.2nd. Ed. American Society for Microbiology, Washington, D. C.1995: 208-270Google Scholar). DNA polymerase β (β-pol) inserts a new base followed by the coupled β-pol-mediated excision of the abnormal 5′-dRP, removal of which has been shown to be rate-limiting (22Srivastava D.K. Vande Berg B.J. Prasad R. Molina J.T. Beard W.A. Tomkinson A.E. Wilson S.H. J. Biol. Chem. 1998; 273: 21203-21209Google Scholar). In bifunctional glycosylaseinitiated BER, the damaged base is recognized and removed by a damage-specific DNA glycosylase followed by incision of the DNA backbone by the associated AP lyase activity, yielding a normal 5′-terminal deoxynucleoside-5′-phosphate residue and an abnormal 3′-terminal α,β-unsaturated aldehyde residue that must be processed prior to repair completion (21Friedberg E.C. Walker G.C. Siede W. DNA Repair and Mutagenesis.2nd. Ed. American Society for Microbiology, Washington, D. C.1995: 208-270Google Scholar). The removal of the abnormal 3′-blocking lesion by APE 3′-phosphodiesterase activity is believed to be rate-limiting (23Izumi T. Hazra T.K. Boldogh I. Tomkinson A.E. Park M.S. Ikeda S. Mitra S. Carcinogenesis. 2000; 21: 1329-1334Google Scholar); however, this rate-limiting role is controversial (24Cappelli E. Degan P. Frosina G. Carcinogenesis. 2000; 21: 1135-1141Google Scholar, 25Bogliolo M. Cappelli E. D'Osualdo A. Rossi O. Barbieri O. Kelley M.R. Frosina G. Anticancer Res. 2002; 22: 2797-2804Google Scholar, 26Cappelli E. D'Osualdo A. Bogliolo M. Kelley M.R. Frosina G. Environ. Mol. Mutagen. 2003; 42: 50-58Google Scholar). After APE recognition of AP sites, BER may proceed by one of two pathways: (i) β-pol-mediated single nucleotide insertion, similar to MFG-BER, or (ii) >1 nucleotide strand-displacement synthesis, required to process modified (i.e. reduced, oxidized) AP sites and involves components of the DNA replication machinery (27Demple B. DeMott M.S. Oncogene. 2002; 21: 8926-8934Google Scholar). Completion of BER requires the nick sealing activity of DNA ligase complexes (28Tomkinson A.E. Chen L. Dong Z. Leppard J.B. Levin D.S. Mackey Z.B. Motycka T.A. Prog. Nucleic Acids Res. Mol. Biol. 2001; 68: 151-164Google Scholar). A review of structural studies has proposed a model of BER requiring highly intimate, yet transient protein-protein interactions among BER enzymes to ensure proper damage repair, with APE being at the center of activity (29Wilson S.H. Kunkel T.A. Nat. Struct. Biol. 2000; 7: 176-178Google Scholar). For example, x-ray cross-complementing protein (XRCC1), a protein with no known enzymatic activity, functions as both a scaffold protein and modulator of BER via functional and physical interaction with APE, bridging the incision and nick-sealing steps of BER (30Vidal A.E. Boiteux S. Hickson I.D. Radicella J.P. EMBO J. 2001; 20: 6530-6539Google Scholar). APE has been shown also to interact with β-pol, recruiting it to the incised AP site and enhancing its rate-limiting dRPase activity (31Bennett R.A.O. Wilson III, D.M. Wong D. Demple B. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 7166-7169Google Scholar); this activity is also believed to be involved in the repair of oxidative base lesions (32Allinson S.L. Dianova I.I. Dianov G.L. EMBO J. 2001; 23: 6919-6926Google Scholar). The functional importance of β-pol in oxidative damage repair may be due to the interaction of APE with bifunctional DNA glycosylases responsible for recognizing and removing these lesions. For example, APE has been shown to stimulate 8-oxoguanine DNA glycosylase (OGG1) turnover and enhance its glycosylase activity while minimizing its associated AP lyase activity (33Hill J.W. Hazra T.K. Izumi T. Mitra S. Nucleic Acids Res. 2001; 29: 430-438Google Scholar, 34Vidal A.E. Hickson I.D. Boiteux S. Radicella J.P. Nucleic Acids Res. 2001; 29: 1285-1292Google Scholar), with XRCC1 accelerating this process (35Marsin S. Vidal A.E. Sossou M. Menissier-de Murcia J. Le Page F. Boiteux S. de Murcia G. Radicella J.P. J. Biol. Chem. 2003; 278: 44068-44074Google Scholar), thus eliminating a potentially rate-limiting step of APE and potentiating MFG-BER. Similar results have also been obtained with other bifunctional DNA glycosylases such as endonuclease III (hNTH1), responsible for recognition and removal of ring-saturated pyrimidines (36Marenstein D.R. Chan M.K. Altamirano A. Basu A.K. Boorstein R.J. Cunningham R.P. Teebor G.W. J. Biol. Chem. 2003; 278: 9005-9012Google Scholar). These studies, in addition to a recent mathematical model of BER throughput (37Sokhansanj B. Rodrigue G.R. Fitch J.P. Wilson III, D.M. Nucleic Acids Res. 2002; 30: 1817-1825Google Scholar), suggest a preference for β-pol-mediated MFG-BER in vivo. The objective of the research described in this study is to characterize in more detail the phenotype of an APE heterozygous knockout (Apex+/–) mouse reported previously (38Meira L.B. Devaraj S. Kisby G.E. Burns D.K. Daniel R.L. Hammer R.E. Grundy S. Jialal I. Friedberg E.C. Cancer Res. 2001; 61: 5552-5557Google Scholar), and to address the effect of APE haploinsufficiency on BER capacity. It is important to note that homozygous deletion of the APE gene (Apex–/–) is embryonic lethal, but heterozygous mice survive and are fertile (38Meira L.B. Devaraj S. Kisby G.E. Burns D.K. Daniel R.L. Hammer R.E. Grundy S. Jialal I. Friedberg E.C. Cancer Res. 2001; 61: 5552-5557Google Scholar, 39Ludwig D.L. MacInnes M.A. Takiguchi Y. Purtymun P.E. Henrie M. Flannery M. Meneses J. Pedersen R.A. Chen D.J. Mutat. Res. 1998; 409: 17-29Google Scholar, 40Xanthoudakis S. Smeyne R.J. Wallace J.D. Curran T. Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 8919-8923Google Scholar). Because the sequences encoding both the DNA repair and redox regulatory activities of APE are disrupted in Apex–/– mice, it is unclear whether one or both of these activities are necessary for embryogenesis. Although the role of APE in the redox activation of p53 and other cellular transcription factors suggests its importance in signal transduction pathways, the embryonic lethality observed for APE and three other BER genes (β-pol, DNA ligase I, and XRCC1) suggests a critical role for BER during embryogenesis (41Friedberg E.C. Meira L.B. DNA Repair. 2003; 2: 501-530Google Scholar). Recent studies have implicated a role for p53 in the regulation of the BER pathway (42Smith M.L. Seo Y.R. Mutagenesis. 2002; 17: 149-156Google Scholar); therefore, it is inviting to suggest that APE repair activity in general, and perhaps APE redox regulatory activity in particular, is the reason for the embryonic lethality observed when APE is deficient. Here we present evidence that half the gene dosage of APE results in tissue-specific alterations in MFG-BER and suggest that APE redox activity, as opposed to repair per se, potentiates this phenotypic effect. Data obtained from Apex+/– mice may have relevant transnational implications because APE variants have been identified in the human population (43Hadi M.Z. Coleman M.A. Fidelis K. Mohrenweiser H.W. Wilson III, D.M. Nucleic Acids Res. 2000; 28: 3871-3879Google Scholar), and variants in DNA repair have been associated with increased risk for disease such as cancer (44de Boer J.G. Mutat. Res. 2002; 509: 201-210Google Scholar, 45Mohrenweiser H.W. Wilson III, D.M. Jones I.M. Mutat. Res. 2003; 526: 93-125Google Scholar). The experiments were performed on young (3–6 months) male C57BL/6-specific pathogen-free mice in accordance with the National Institutes of Health guidelines for the use and care of laboratory animals. The Wayne State University Animal Investigation Committee approved the animal protocol. Mice were maintained on a 12-h light/dark cycle and fed a standard lab diet and water ad libitum. The mice were sacrificed by cervical dislocation, and the organs to be studied were flash-frozen in liquid nitrogen and stored at –70 °C for later enzyme studies and Western blot analyses. Tissues for total RNA isolation and RT-PCR analysis were immediately homogenized in TRIzol® Reagent (Invitrogen) according to the manufacturer's protocol. The APE heterozygous knockout (Apex+/–) mice were developed in Friedberg's laboratory as described previously (38Meira L.B. Devaraj S. Kisby G.E. Burns D.K. Daniel R.L. Hammer R.E. Grundy S. Jialal I. Friedberg E.C. Cancer Res. 2001; 61: 5552-5557Google Scholar). In order to obtain the requisite number of animals for the study, the Apex+/– mice were inbred and maintained on a 12-h light/dark cycle and fed a standard lab diet and water ad libitum. The mice appeared normal, were fertile, and there was no retardation in food intake, weight gain, or growth rate; however, it was observed that pups were not produced in expected Mendelian ratios, with heterozygote births predominating, similar to previous observations (39Ludwig D.L. MacInnes M.A. Takiguchi Y. Purtymun P.E. Henrie M. Flannery M. Meneses J. Pedersen R.A. Chen D.J. Mutat. Res. 1998; 409: 17-29Google Scholar). A three-primer PCR strategy was employed to genotype the animals generated by Apex+/– intercrosses as described previously (38Meira L.B. Devaraj S. Kisby G.E. Burns D.K. Daniel R.L. Hammer R.E. Grundy S. Jialal I. Friedberg E.C. Cancer Res. 2001; 61: 5552-5557Google Scholar). The level of APE mRNA was measured using RT-PCR analysis using AccessQuick™ RT-PCR System (Promega, Madison, WI) according to the manufacturer's protocol. Total cellular RNA was isolated from selected tissues using TRIzol® Reagent (Invitrogen), and RNA concentration was determined by measuring UV absorption at 260/280 nm. Oligonucleotide primers specific for the mouse Apex gene (forward, 5′-CTCAAGATATGCTCCTGGAA-3′; reverse, 5′-GGTATTCCAGTCTTACCAGA-3′) were designed using GeneFisher Interactive Primer Design Tool (Bielefeld, Germany) and synthesized by Sigma-Genosys (The Woodlands, TX). RT-PCR thermal cycling conditions were as follows: 48 °C for 45 min, 1 cycle; 95 °C for 2 min, 1 cycle; 95 °C for 1 min, 52 °C for 1 min, 70 °C for 2 min, 22 cycles; and 70 °C for 5 min, 1 cycle. The 350-bp RT-PCR product was stained with ethidium bromide and analyzed on a 2% agarose gel. Intensity of the bands was detected and quantified using a ChemiImager™ system (AlphaInnotech, San Leandro, CA) and expressed as the integrated density value (I.D.V) per μg of RNA used per reaction. Data were normalized based on the amount of β-actin present in each sample. Isolation of crude nuclear extract was accomplished using Sigma CelLytic™ NuCLEAR™ extraction kit (Sigma). All samples and tubes were handled and chilled on ice, and all solutions were made fresh according to the manufacturer's protocol. Resultant nuclear extracts were dialyzed against 1 liter of dialysis buffer (20 mm Tris-HCl, pH 8.0; 100 mm KCl; 10 mm NaS2O5; 0.1 mm DTT; 0.1 mm phenylmethylsulfonyl fluoride; 1 μg/ml pepstatin A) for 4–6 h at 4 °C using Slide-A-Lyzer® minidialysis units (Pierce). Protein concentrations were determined using Protein Assay Kit I (Bio-Rad). Western analysis was performed using 200 μg of crude nuclear extract isolated from selected tissues according to standard protocol. Protein levels were determined using manufacturer recommended dilutions of monoclonal antisera developed against mouse APE/REF-1 (Novus Biologicals, Littleton, CO), and monoclonal antisera developed against rat β-pol (Ab-1 Clone 18S, NeoMarkers, Fremont, CA). The bands were detected and quantified using a ChemiImager™ system (AlphaInnotech, San Leandro, CA) after incubation in SuperSignal® West Pico Chemiluminescent Substrate (Pierce). Data are expressed as the I.D.V. of the band per μg of protein loaded. The 5′-endonuclease activity of APE was analyzed using a quantitative in vitro assay that measures the incision of a 26-mer duplex oligonucleotide substrate containing a synthetic tetrahydrofuran (F) AP site (upper strand, 5′-AATTCACCGGTACCFTCTAGAATTCG-3′; lower strand, 5′-CGAATTCTAGAGGGTACCGGTGAATT-3′) as described previously (46Wilson III, D.M. Takeshita M. Grollman A.P. Demple B. J. Biol. Chem. 1995; 270: 16002-16007Google Scholar). Briefly, 2.5 pmol of radio-end-labeled and purified duplex AP DNA substrate was incubated with 100 ng of crude nuclear extract-selected tissues in a 10-μl reaction mixture containing 50 mm Hepes, pH 7.5; 50 mm KCl; 10 mm MgCl2; 2mm DTT; 1 μg/ml BSA; and 0.05% Triton X-100. The reaction mixtures were incubated for 15 min at 37 °C and stopped by the addition of 50 mm EDTA. Assay products (5 μl) were added to 15 μl of loading dye (95% formamide, 5% glycerol, 10 mg of xylene cyanol, 10 mg of bromphenol blue) and heated at 95 °C for 5 min. Aliquots (3 μl) were run on a 15% denaturing 19:1 acrylamide/bisacrylamide gel (SequaGel® Sequencing System, National Diagnostics, Atlanta, GA) at 55 °C, soaked in fixing solution (10% glacial acetic acid; 10% methanol), wrapped in Saran wrap, and exposed to a Molecular Imaging Screen (Bio-Rad). Endonuclease activity (presence of a 14-mer band) was visualized and quantified using a Molecular Imager® System (Bio-Rad) by calculating the relative amount of the 14-mer oligo product with the unreacted 26-mer substrate (product/(product + substrate)). Data are expressed as machine counts per ng of protein. The principle of this assay is to measure MFG-BER activity. Radio end-labeled and purified 30-bp oligonucleotides (upper strand, 5′-ATATACCGCGGUCGGCCGATCAAGCTTATTdd-3′; lower strand, 3′-ddTATATGGCGCCGGCCGGCTAGTTCGAATAA-5′) containing a G:U mismatch and an HpaII restriction site (CCGG) were incubated in a reaction mixture (100 mm Tris-HCl, pH 7.5, 5 mm MgCl2; 1 mm DTT, 0.1 mm EDTA, 2 mm ATP, 0.5 mm NAD, 20 μm dNTPs, 5 mm diTrisphosphocreatine, 10 units of creatine phosphokinase) with 50 μg of crude nuclear extract isolated from selected tissues. The reaction mixtures were incubated for 30 min at 37 °C, followed by 5 min at 95 °C to stop the reaction. The duplex oligonucleotides were allowed to reanneal for 1 h at room temperature before being briefly centrifuged to pellet the denatured proteins. Repair of the G:U mismatch to a correct G:C base pair was determined via treatment of the duplex oligonucleotide with 20 units of HpaII (Promega, Madison, WI) for 1 h at 37 °C and analysis by electrophoresis on a 20% denaturing 19:1 acrylamide/bisacrylamide gel (SequaGel® Sequencing System, National Diagnostics, Atlanta, GA). Repair activity (presence of a 16-mer band) was visualized and quantified using a Molecular Imager® System (Bio-Rad) by calculating the ratio of the 16-mer product with the 30-mer substrate (product/substrate). Data are expressed as machine counts per μg of protein. The DNA binding ability of nuclear extracts isolated from selected tissues was determined using electrophoretic mobility shift assays (EMSAs). Nuclear extracts (20 μg) were incubated with 4× reaction buffer (final concentrations: 50 mm Hepes, pH 8.0; 100 mm NaCl; 10 mm EDTA; 1 mm DTT; 9.5% glycerol v/v) plus 1 μg of BSA and 2 μg of poly(dI-dC) for 5 min at room temperature. A radio-end-labeled and purified oligonucleotide probe containing an AP site (0.0125 pmol) was added to the reaction mix and incubated for 30 min at room temperature. Negative controls (all components except nuclear extract) were included in all experiments. In competitive assays, 100× molar excess of unlabeled DNA was added to the reaction mixture. The protein-DNA complex was resolved on a 5% non-denaturing polyacrylamide gel in 0.5× TBE buffer. Reaction products were visualized and quantified using a Molecular Imager® System (Bio-Rad). TEMPO Extraction of DNA—DNA for the aldehyde-reactive probe slot blot (ASB) assay was extracted according to the method described by Hofer and Moller (47Hofer T. Moller L. Chem. Res. Toxicol. 1998; 11: 882-887Google Scholar) with some modifications. This method minimizes artifactual DNA damage by using 20 mm TEMPO in all solutions and reagents and by minimizing heat treatment of DNA. Briefly, tissue was homogenized in ice-cold PBS and centrifuged (2000 × g at 4 °C for 5 min). Resultant supernatant was decanted and pellet resuspended in lysis buffer (Applied Biosystems, Foster City, CA). Proteinase K (30 units, Ambion, Austin, TX) was added, and samples were incubated overnight at 4 °C, followed by phenol/chloroform and Sevag (chloroform/isoamyl alcohol, 24:1) extraction. Extracted DNA was precipitated using 7.5% 4 m NaCl and 2 volumes of 100% cold ethanol and centrifuged (2000 × g at 4 °C for 5 min). Resultant pellet was washed in 70% ethanol and resuspended in ice-cold PBS and rehydrated at 4 °C. The samples were incubated with RNase A (2 μg) and RNase T1 (1,000 units, Ambion, Austin, TX) for 30 min at 37 °C, and resultant DNA was cold ethanol-precipitated, resuspended in deionized water at 4 °C, and stored at –70 °C. Gravity Tip Extraction of DNA—DNA for the random oligonucleotide primed synthesis (ROPS) assay was isolated using Qiagen (Valencia, CA) gravity tip columns as described in the manufacturer's protocol. This method generates large fragments of DNA (up to 150-kb) while minimizing shearing. ASB—The ASB assay was carried out as described previously (14Nakamura J. Walker V.E. Upton P.B. Chiang S.-Y. Kow Y.W. Swenberg J.A. Cancer Res. 1998; 58: 222-225Google Scholar) with slight modifications. TEMPO-extracted DNA (8 μg) was incubated in 30 μl of PBS with 2 mm aldehyde-reactive probe (ARP) (Dojindo Laboratories, Kumamoto, Japan) at 37 °C for 10 min. DNA was cold ethanol-precipitated (as described above) and resuspended in 1× TE buffer overnight at 4 °C. DNA was heat-denatured at 100 °C for 5 min, quickly chilled on ice, and mixed with equal amount of 2 m ammonium acetate. DNA was immobilized on a nitrocellulose membrane (Schleicher & Schuell) by using a Invitrogen Filtration Manifold system. The membrane was washed in 5× SSC for 15 min at 37 °C and then baked under vacuum at 80 °C for 30 min. The dried membrane was incubated in a hybridization buffer (final concentrations: 20 mm Tris, pH 7.5; 0.1 m NaCl; 1 mm EDTA; 0.5% casein w/v; 0.25% BSA w/v; 0.1% Tween 20 v/v) for 30 min at room temperature. The membrane was then incubated in the same hybridization buffer containing 100 μl of streptavidin-conjugated horseradish peroxidase (BioGenex, San Ramon, CA) at room temperature for 45 min. Following incubation in horseradish peroxidase, the membrane was washed three times for 5 min each at 37 °C in TBS, pH 7.5 (final concentrations: 0.26 m NaCl; 1 mm EDTA; 20 mm Tris, pH 7.5; 0.1% Tween 20). The membrane was incubated in ECL (Pierce) for 5 min at room temperature and visualized using a ChemiImager™ system (AlphaInnotech, San Leandro, CA). Standards containing known amounts of AP sites (14Nakamura J. Walker V.E. Upton P.B. Chiang S.-Y. Kow Y.W. Swenberg J.A. Cancer Res. 1998; 58: 222-225Google Scholar) were used to determine the relative level of AP sites in liver DNA. Data are expressed as number of AP sites per 106 nucleotides. ROPS—The relative number of 3′-OH group-containing DNA strand breaks was quantified using a Klenow(exo–) incorporation assay based on the ability of Klenow to initiate DNA synthesis from 3′-OH ends of single strand DNA (48Basnakian A.G. James S.J. DNA Cell Biol. 1996; 15: 255-262Google Scholar). Gravity tip extracted DNA (0.25 μg) was heat-denatured at 100 °C for 5 min and added to the Klenow reaction buffer (0.5 mm dTTP, 0.5 mm dGTP, and 0.5 mm dATP; 0.33 μm dCTP, 1 μl of Klenow(exo–)) with 10× Klenow buffer per manufacturer's protocol (New England Biolabs, Beverly, MA) and 5 μCi of [α-32P]ATP (3000 Ci/mmol, PerkinElmer Life Sciences). Reaction mixtures were incubated at 16 °C for 30 min, and the reaction was stopped with the addition of 25 μl of 12.5 mm EDTA, pH 8.0. Samples (5 μl) were spotted onto scored and numbered Whatman DE81 paper and allowed to air dry. The spotted chromatography paper was washed five times for 5 min in 0.5 m Na2HPO4 (dibasic) followed by a brief rinse in deionized water two times and then allowed to air-dry. Paper was cut and placed into scintillation vials with 2.5 ml of ScintiVerse mixture (Fisher). Incorporation of [α-32P]dCTP was quantified using a Packard scintillation counter. Statistical significance between means was determined using analysis of variance followed by the Fisher's least significant difference test where appropriate (49Sokal R.R. Rohlf F.J. Biometry. W. H. Freeman & Co., New York1981: 169-176Google Scholar). A p value less than 0.05 was considered statistically significant. In order to elucidate the molecular effects of gene-targeted disruption of the mouse APE gene (Apex), our laboratory has characterized in detail a transgenic "knockout" mouse containing a heterozygous deletion of the APE gene (Apex+/–). As reported previously, the Apex–/– mice are embryonic lethal, whereas the Apex+/– animals are fertile, appear normal, and exhibit reduced APE mRNA and APE protein levels in mouse embryonic fibroblasts and brain cells as compared with their Apex+/+ counterparts (38Meira L.B. Devaraj S. Kisby G.E. Burns D.K. Daniel R.L. Hammer R.E. Grundy S. Jialal I. Friedberg E.C. Cancer Res. 2001; 61: 5552-5557Google Scholar). In a series of experiments, we were able to confirm whether APE heterozygosity would cause these mice to exhibit haploinsufficiency with respect to APE in brain, liver, and testes. In order to determine whether loss of a functional allele of APE would result in reduced expression of this gene, we have quantified the expression of APE via RT-PCR analysis. By using total RNA isolated from liver, we observed a 40–50% decrease in APE mRNA (Fig. 1A) in Apex+/– mice as compared with their normal, wild-type (Apex+/+) counterparts. Because differences in APE mRNA may not necessarily reflect differences in APE protein levels, we also measured APE protein levels using Western blot analysis. We show a corresponding 40–50% decrease in APE protein in liver (Fig. 1B). To confirm that changes in APE expression (i.e. changes in mRNA and protein levels) result in changes in APE enzymatic activity, we measured APE 5′-endonuclease activity using a 26-bp oligonucl
Wang, Y., Raffoul, J. J., Che, M., Doerge, D. R., Joiner, M. C., Kucuk, O., Sarkar, F. H. and Hillman, G. G. Prostate Cancer Treatment is Enhanced by Genistein In Vitro and In Vivo in a Syngeneic Orthotopic Tumor Model. Radiat. Res. 166, 73–80 (2006).Pretreatment with genistein, a bioactive component of soy isoflavones, potentiated cell killing induced by radiation in human PC-3 prostate cancer cells in vitro. Using an orthotopic xenograft in nude mice, we demonstrated that genistein combined with prostate tumor irradiation caused greater inhibition of primary tumor growth and increased control of spontaneous metastasis to para-aortic lymph nodes, increasing mouse survival. Paradoxically, treatment with genistein alone increased metastasis to lymph nodes. This observation is of concern in relation to soy-based clinical trials for cancer patients. To address whether this observation is because nude mice have an impaired immune system, these studies were repeated in orthotopic RM-9 prostate tumors in syngeneic C57BL/6 mice. The combination of genistein with radiation in this model also caused a greater inhibition of primary tumor growth and spontaneous metastasis to regional para-aortic lymph nodes, whereas treatment with genistein alone showed a trend to increased lymph node metastasis. Data from the syngeneic and xenograft models are comparable and indicate that the combination of genistein with radiotherapy is more effective and safer for prostate cancer treatment than genistein alone, which promotes metastatic spread to regional lymph nodes.
Dietary intake of foods rich in antioxidant properties is suggested to be cancer protective. Foods rich in antioxidant properties include grape ( Vitis vinifera ), one of the world’s largest fruit crops and most commonly consumed fruits in the world. The composition and cancer-protective effects of major phenolic antioxidants in grape skin and seed extracts are discussed in this review. Grape skin and seed extracts exert strong free radical scavenging and chelating activities and inhibit lipid oxidation in various food and cell models in vitro . The use of grape antioxidants are promising against a broad range of cancer cells by targeting epidermal growth factor receptor (EGFR) and its downstream pathways, inhibiting over-expression of COX-2 and prostaglandin E2 receptors, or modifying estrogen receptor pathways, resulting in cell cycle arrest and apoptosis. Interestingly, some of these activities were also demonstrated in animal models. However, in vivo studies have demonstrated inconsistent antioxidant efficacy. Nonetheless, a growing body of evidence from human clinical trials has demonstrated that consumption of grape, wine and grape juice exerts many health-promoting and possible anti-cancer effects. Thus, grape skin and seed extracts have great potential in cancer prevention and further investigation into this exciting field is warranted.
Cancer chemoprevention using natural or synthetic compounds to prevent or suppress the development of cancer, is an area of active investigation. Many compounds belonging to diverse chemical classes have been identified as potential chemopreventive agents, including vitamins and minerals, naturally occurring phytochemicals, and synthetic compounds. Understanding the molecular mechanisms of cancer chemoprevention is not only important for the safe application of these compounds in populations of patients at high risk for cancer, but also allows for further development of novel treatment regimens for cancer patients.
This special issue contains original research as well as review articles that are intended to stimulate the continuing efforts to understand the use of dietary agents in cancer chemoprevention and treatment. The lead article by S. N. Saldanha and T. O. Tollefsbol provides a comprehensive review of dietary agents that have shown strong chemopreventive and therapeutic properties in vitro. They also discuss the design and modification of these bioactive compounds for pre-clinical and clinical applications.
Dietary intake of foods rich in antioxidant compounds has been suggested to be cancer protective. However, randomized clinical trials and epidemiologic studies on the association between intake of foods rich in antioxidants and cancer incidence have yielded mixed results. M. Y. Wei and E. L. Giovannuci discuss the epidemiologic considerations of lycopene as a chemopreventive agent, including measurement of lycopene, its major source in the diet, and the assessment of prostate cancer incidence and progression, with particular emphasis on the effect of PSA screening on this association. K. Zhou and J. J. Raffoul discuss the composition and cancer-protective effects of major phenolic antioxidants in grape skin and grape seed extracts. M. A. Parasramka and S. V. Gupta provide original research demonstrating the anticancer properties of garcinol alone, or combined with curcumin, on pancreatic cancer cells. Garcinol, a polyisoprentylated benzophenone extracted from the rind of the fruit Garcinia indica, a plant found in tropical regions, has antioxidant and anti-inflammatory properties and its role as anticancer agent is thoroughly discussed in the review from N. Saadat and S. V. Gupta.
Two manuscripts discussing the effect of dietary agents on DNA repair capacity are also part of this special issue. In a manuscript by J. J. Raffoul et al., the potential for targeting the DNA base excision repair enzyme APE1/Ref-1 using dietary agents such as soy isoflavones, resveratrol, curcumin, ascorbate, and alpha-tocopherol is discussed. The potential for these natural compounds to be combined with chemotherapy or radiotherapy for the more effective treatment of cancer are also reviewed. A proposed mechanism of action is discussed and an attempt is made to delineate which of the two activities of APE1/Ref-1 (DNA repair versus redox activation of cellular transcription factors) is responsible for the observed effects. The second manuscript by R. Rosati et al. reviews the role for dietary folate in the prevention of colorectal cancer. Data are presented which demonstrate that inhibition of DNA repair is protective in the development of preneoplastic colon lesions, both when folate is depleted and when it is not. This manuscript is a comprehensive review of the literature and provides a critical analysis of the experimental designs used in folate and colorectal cancer research.
Two additional manuscripts detailing the ability of dietary agents to sensitize cancer cells to chemotherapy and radiotherapy are included in this special issue. An original research article by S. Duangmano et al. demonstrates that curcurbitacin B, a plant phytochemical, inhibited breast cancer cell proliferation in a dose-dependent manner and caused radiosensitization of human breast cancer cells via G2/M cell cycle arrest. Furthermore, an original research article by K. Sahin et al. demonstrate that genistein, a soy isoflavone, sensitizes cervical cancer cells to cisplatin via inhibition of NF-kappa B and Akt/mTOR cell signaling pathways.
This special issue concludes with a report of a clinical study demonstrating the prevention of anthracycline-induced cardiac toxicity through supplementation with selenium in a group of pediatric cancer patients.
Research efforts aimed at understanding the role of dietary agents and phytochemicals in cancer prevention and treatment are likely to yield high-impact results that have the potential for immediate clinical applications. Furthermore, combination of phytochemicals and nutritional agents with therapies for advanced cancers, including radiotherapy and chemotherapy, would benefit from a complementary and safe approach using dietary agents to mitigate the adverse effects of these therapies on normal tissues while enhancing the therapeutic efficacy. Elucidation of the mechanisms of interaction between dietary agents and conventional cancer treatments will have a major impact on understanding the molecular mechanisms of cancer chemoprevention and will ultimately result in clinical use of dietary agents as an adjunct to standard cancer treatment.
Julian J. Raffoul
Omer Kucuk
Fazlul H. Sarkar
Gilda G. Hillman
The mechanism by which folate deficiency influences carcinogenesis is not well established, but a phenotype of DNA strand breaks, mutations, and chromosomal instability suggests an inability to repair DNA damage. To elucidate the mechanism by which folate deficiency influences carcinogenicity, we have analyzed the effect of folate deficiency on base excision repair (BER), the pathway responsible for repairing uracil in DNA. We observe an up-regulation in initiation of BER in liver of the folate-deficient mice, as evidenced by an increase in uracil DNA glycosylase protein (30%, p < 0.01) and activity (31%, p < 0.05). However, no up-regulation in either BER or its rate-determining enzyme, DNA polymerase β (β-pol) is observed in response to folate deficiency. Accordingly, an accumulation of repair intermediates in the form of DNA single strand breaks (37% increase, p < 0.03) is observed. These data indicate that folate deficiency alters the balance and coordination of BER by stimulating initiation without subsequently stimulating the completion of repair, resulting in a functional BER deficiency. In directly establishing that the inability to induce β-pol and mount a BER response when folate is deficient is causative in the accumulation of toxic repair intermediates, β-pol-haploin-sufficient mice subjected to folate deficiency displayed additional increases in DNA single strand breaks (52% increase, p < 0.05) as well as accumulation in aldehydic DNA lesions (38% increase, p < 0.01). Since young β-polhaploinsufficient mice do not spontaneously exhibit increased levels of these repair intermediates, these data demonstrate that folate deficiency and β-pol haploinsufficiency interact to increase the accumulation of DNA damage. In addition to establishing a direct role for β-pol in the phenotype expressed by folate deficiency, these data are also consistent with the concept that repair of uracil and abasic sites is more efficient than repair of oxidized bases. The mechanism by which folate deficiency influences carcinogenesis is not well established, but a phenotype of DNA strand breaks, mutations, and chromosomal instability suggests an inability to repair DNA damage. To elucidate the mechanism by which folate deficiency influences carcinogenicity, we have analyzed the effect of folate deficiency on base excision repair (BER), the pathway responsible for repairing uracil in DNA. We observe an up-regulation in initiation of BER in liver of the folate-deficient mice, as evidenced by an increase in uracil DNA glycosylase protein (30%, p < 0.01) and activity (31%, p < 0.05). However, no up-regulation in either BER or its rate-determining enzyme, DNA polymerase β (β-pol) is observed in response to folate deficiency. Accordingly, an accumulation of repair intermediates in the form of DNA single strand breaks (37% increase, p < 0.03) is observed. These data indicate that folate deficiency alters the balance and coordination of BER by stimulating initiation without subsequently stimulating the completion of repair, resulting in a functional BER deficiency. In directly establishing that the inability to induce β-pol and mount a BER response when folate is deficient is causative in the accumulation of toxic repair intermediates, β-pol-haploin-sufficient mice subjected to folate deficiency displayed additional increases in DNA single strand breaks (52% increase, p < 0.05) as well as accumulation in aldehydic DNA lesions (38% increase, p < 0.01). Since young β-polhaploinsufficient mice do not spontaneously exhibit increased levels of these repair intermediates, these data demonstrate that folate deficiency and β-pol haploinsufficiency interact to increase the accumulation of DNA damage. In addition to establishing a direct role for β-pol in the phenotype expressed by folate deficiency, these data are also consistent with the concept that repair of uracil and abasic sites is more efficient than repair of oxidized bases. In human studies, folate deficiency is associated with cancers of the lung, cervix, brain, esophagus, pancreas, breast, colon, and liver (reviewed in Refs. 1Glynn S.A. Albanes D. Nutr. Cancer. 1994; 22: 101-119Crossref PubMed Scopus (128) Google Scholar and 2Choi S.W. Mason J.B. J. Nutr. 2000; 130: 129-132Crossref PubMed Scopus (803) Google Scholar). In support of the epidemiology, the role of folate in the development of both colon and liver cancer has been experimentally demonstrated in animal studies. Folate deficiency enhances the carcinogenic effect of dimethylhydrazine (3Cravo M.L. Mason J.B. Dayal Y. Hutchinson M. Smith D. Selhub J. Rosenberg I.H. Cancer Res. 1992; 52: 5002-5006PubMed Google Scholar), whereas folate supplementation is protective (4Kim Y.I. Salomon R.N. Graeme-Cook F. Choi S.W. Smith D.E. Dallal G.E. Mason J.B. Gut. 1996; 39: 732-740Crossref PubMed Scopus (154) Google Scholar). Folate/methyl deficiency results in hepatocarcinogenesis (5Hoover K.L. Lynch P.H. Poirier L.A. J. Natl. Cancer Inst. 1984; 73: 1327-1336PubMed Google Scholar). Additionally, livers of folate/methyl-deficient rats accumulate preneoplastic changes in response to folate deficiency (6James S.J. Miller B.J. Basnakian A.G. Pogribny I.P. Pogribna M. Muskhelishvili L. Carcinogenesis. 1997; 18: 287-293Crossref PubMed Scopus (97) Google Scholar). It is suggested that the carcinogenic properties of folate deficiency are related to a reduction in S-adenosylmethionine levels altering DNA methylation status and/or to depletion of thymidylate resulting in increased uracil content of DNA. Additionally, folate is required for de novo purine biosynthesis. Which of these factors may be responsible for inducing carcinogenesis is presently unknown. Whereas the underlying mechanism connecting folate deficiency to cancer remains unknown, it is clear that folate deficiency induces a phenotype suggestive of an inability to repair DNA damage. The accumulation of strand breaks, mutations, and chromosomal instability observed in response to folate deficiency all suggest that DNA repair capacity is inhibited. In support of this, folate-deficient cells and animals inefficiently repair alkylation damage (7Branda R.F. Hacker M. Lafayette A. Nigels E. Sullivan L. Nicklas J.A. O'Neill J.P. Environ. Mol. Mutagen. 1998; 32: 33-38Crossref PubMed Scopus (16) Google Scholar). Folate deficiency acts synergistically with ethane methyl sulfonate in Chinese hamster ovary cells (8Branda R.F. Lafayette A.R. O'Neill J.P. Nicklas J.A. Mutation Res. 1999; 427: 79-87Crossref PubMed Scopus (13) Google Scholar), suggesting an inability to repair ethane methyl sulfonate-induced damage. Folate depletion in human lymphocytes sensitizes to oxidative damage induced by hydrogen peroxide (9Duthie S.J. Hawdon A. FASEB J. 1998; 12: 1491-1497Crossref PubMed Scopus (262) Google Scholar). Human colon epithelial cells grown in folate-deficient medium are unable to repair damages induced by methylmethane sulfonate and hydrogen peroxide (10Duthie S.J. Narayanan S. Blum S. Pirie L. Brand G.M. Nutr. Cancer. 2000; 37: 245-251Crossref PubMed Scopus (183) Google Scholar). Folate deficiency impairs the ability of neurons and colonocytes to repair DNA damage (11Kruman I.I. Kumaravel T.S. Lohani A. Pedersen W.A. Cutler R.G. Druman Y. Haughey N. Lee J. Evans M. Mattson M.P. J. Neurosci. 2002; 22: 1752-1762Crossref PubMed Google Scholar, 12Choi S.W. Kim Y.I. Weitzel J.N. Mason J.B. Gut. 1998; 4: 93-99Crossref Scopus (115) Google Scholar). These data suggest that the pathway responsible for repairing these damages may be ineffective when folate is limiting by demonstrating a persistence of DNA damage but stop short of directly measuring DNA repair capacity. The objectives of this study are to directly measure the effects of folate deficiency on DNA repair capacity and to begin identifying the molecular mechanisms responsible for precipitating a phenotype of cancer susceptibility when folate is deficient. Uracil has been shown to accumulate in response to folate deficiency (9Duthie S.J. Hawdon A. FASEB J. 1998; 12: 1491-1497Crossref PubMed Scopus (262) Google Scholar, 10Duthie S.J. Narayanan S. Blum S. Pirie L. Brand G.M. Nutr. Cancer. 2000; 37: 245-251Crossref PubMed Scopus (183) Google Scholar, 13Wickramasinghe S.N. Fida S. Blood. 1994; 83: 1656-1661Crossref PubMed Google Scholar, 14Blount B.C. Mack M.M. Wehr C.M. MacGregor J.T. Hiatt R.A. Wang G. Wickramasinghe S.N. Everson R.B. Ames B.N. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 3290-3295Crossref PubMed Scopus (1215) Google Scholar, 15Duthie S.J. Grant G. Narayanan S. Br. J. Cancer. 2000; 83: 1532-1537Crossref PubMed Scopus (62) Google Scholar). The DNA repair pathway for removal of uracil is the base excision repair (BER) 1The abbreviations used are: BER, base excision repair; UDG, uracil-DNA glycosylase; Ape, AP endonuclease; β-pol, polymerase β; ASB, aldehyde-reactive probe slot blot; ADL, aldehydic DNA lesion; DTT, dithiothreitol; I.D.V., integrated density value; ROPS, random oligonucleotideprimed synthesis; TEMPO, 2,2,6,6-tetramethylpiperidin-oxyl. pathway. The BER pathway repairs small, non-helix-distorting lesions in the DNA. In the process of repairing uracil, the following sequence of events occurs. Uracil-DNA glycosylase (UDG), a monofunctional glycosylase, excises uracil from the DNA backbone, creating a transient abasic site. AP endonuclease 1 (Ape1) cleaves the DNA backbone to allow for incorporation of a correct nucleotide by DNA polymerase β (β-pol). This step results in the transient formation of both a 3′-OH-containing DNA single strand break and a deoxyribose phosphate flap containing an aldehydic group. β-pol also performs the rate-limiting step of deoxyribose flap excision (16Srivastava D.K. Vande Berg B.J. Prasad R. Molina J.T. Beard W.A. Tomkinson A.E. Wilson S.H. J. Biol. Chem. 1998; 273: 21203-21209Abstract Full Text Full Text PDF PubMed Scopus (354) Google Scholar), at which point only the 3′-OH group remains until ligation occurs (ligase 1 or ligase 3-XRCC1 complex), at which point the strand break is fully resolved. Typically, BER functions as a tightly coordinated sequence of enzymatic events such that damage is removed, repair is completed, and intermediate products generated during repair do not accumulate. Mice with gene-targeted disruptions in the rate-limiting step of BER exhibit a dysregulation in this coordination and accumulate DNA single strand breaks both spontaneously and in response to carcinogen exposure, have a reduced DNA damage threshold, and exhibit genomic instability in the forms of both mutation induction and chromosomal damage (17Cabelof D.C. Guo Z. Raffoul J.J. Sobol R.W. Wilson S.H. Richardson A. Heydari A.R. Cancer Res. 2003; 63: 5799-5807PubMed Google Scholar). The phenotype of folate deficiency includes accumulation of strand breaks (9Duthie S.J. Hawdon A. FASEB J. 1998; 12: 1491-1497Crossref PubMed Scopus (262) Google Scholar, 10Duthie S.J. Narayanan S. Blum S. Pirie L. Brand G.M. Nutr. Cancer. 2000; 37: 245-251Crossref PubMed Scopus (183) Google Scholar, 15Duthie S.J. Grant G. Narayanan S. Br. J. Cancer. 2000; 83: 1532-1537Crossref PubMed Scopus (62) Google Scholar, 18Melnyk S. Pogribna M. Miller B.J. Basnakian A.G. Pogribny I.P. James S.J. Cancer Lett. 1999; 146: 35-44Crossref PubMed Scopus (96) Google Scholar, 19Kim Y.I. Shirwadkar S. Choi S.W. Puchyr M. Wang Y. Mason J.B. Gastroenterology. 2000; 119: 151-161Abstract Full Text Full Text PDF PubMed Scopus (85) Google Scholar), reduced tolerance to DNA-damaging agents, increased mutation induction (7Branda R.F. Hacker M. Lafayette A. Nigels E. Sullivan L. Nicklas J.A. O'Neill J.P. Environ. Mol. Mutagen. 1998; 32: 33-38Crossref PubMed Scopus (16) Google Scholar, 8Branda R.F. Lafayette A.R. O'Neill J.P. Nicklas J.A. Mutation Res. 1999; 427: 79-87Crossref PubMed Scopus (13) Google Scholar, 20Branda R.F. Lafayette A.R. O'Neill J.P. Nicklas J.A. Cancer Res. 1997; 57: 2586-2588PubMed Google Scholar), and chromosomal instability (21MacGregor J.T. Schlegel R. Wehr C.M. Alperin P. Ames B.N. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 9962-9965Crossref PubMed Scopus (69) Google Scholar, 22Everson R.B. Wehr C.M. Erexson G.L. MacGregor J.T. J. Natl. Cancer Inst. 1988; 80: 525-529Crossref PubMed Scopus (109) Google Scholar) (i.e. folate deficiency induces a phenotype identical to BER deficiency). The objective of this study was to directly determine the effect of folate deficiency on base excision repair capacity. Because BER is DNA damage-inducible (23Fornace Jr., A.J. Zmudzka B. Hollander M.C. Wilson S.H. Mol. Cell. Biol. 1989; 9: 851-853Crossref PubMed Scopus (155) Google Scholar, 24Cabelof D.C. Raffoul J.J. Yanamadala S. Guo Z. Heydari A.R. Carcinogenesis. 2002; 23: 1419-1425Crossref PubMed Google Scholar), we expected to observe an induction in the activity of this repair pathway in response to the DNA damage induced by folate deficiency. However, in response to folate deficiency we fail to observe an up-regulation in either BER or the rate-determining enzyme in the pathway, β-pol. This lack of response by the BER pathway should result in an accumulation of the DNA damage induced by folate deficiency. We provide evidence that the initial step of BER, glycosylase-initiated removal of uracil, is up-regulated in response to folate deficiency, without a coordinating increase in activity of the subsequent steps of repair. This dysregulation of BER induces a state of BER deficiency mimicking that observed in mice heterozygous for β-pol. Our data demonstrate that in response to folate deficiency repair is initiated but not completed, resulting in an accumulation of DNA repair intermediate products that are genotoxic (25Horton J.K. Joyce-Gray D.F. Pachkowski B.F. Swenberg J.A. Wilson S.H. DNA Repair. 2003; 2: 27-48Crossref PubMed Scopus (83) Google Scholar). Since BER deficiency increases cancer susceptibility, this functional BER deficiency in response to folate deficiency may provide an important mechanistic explanation for the increased cancer risk associated with folate deficiency. As such, human polymorphisms and functional mutations within the β-pol gene may interact with folate deficiency to increase cancer risk. This hypothesis is supported by our data demonstrating that folate deficiency results in a greater accumulation of DNA damage in mice haploinsufficient for β-pol. Experiments were performed in young (3–4 months) male C57BL/6-specific pathogen-free mice in accordance with the National Institutes of Health guidelines for the use and care of laboratory animals. The Wayne State University Animal Investigation Committee approved the animal protocol. Mice were maintained on a 12-h light/dark cycle and fed standard mouse chow and water ad libitum. Mice heterozygous for the DNA polymerase β gene (β-pol+/–) were created in Rajewsky's laboratory by deletion of the promoter and the first exon of the β-pol gene (26Gu H. Marth J.D. Orban P.C. Mossmann H. Rajewsky K. Science. 1994; 265: 103-106Crossref PubMed Scopus (1182) Google Scholar). The animals appear to be normal and are fertile; there is no retardation in food intake, weight gain, or growth rate. The genotype of the mice was determined as described previously (17Cabelof D.C. Guo Z. Raffoul J.J. Sobol R.W. Wilson S.H. Richardson A. Heydari A.R. Cancer Res. 2003; 63: 5799-5807PubMed Google Scholar). At 3–4 months of age, 20 β-pol+/+ and 20 β-pol+/– mice were randomly assigned to two dietary groups and were fed AIN93G-purified isoenergetic diets. (Dyets, Inc., Lehigh Valley, PA). The control group received a folate adequate diet containing 2 mg/kg folic acid. The experimental group received a folate-deficient diet containing 0 mg/kg folic acid. Diets were stored at –20 °C. 1% succinyl sulfathiazole was added to all diets. The animals' food intake and body weights were monitored twice weekly to monitor for signs of toxicity (i.e. weight loss), and the experimental diets were continued for 8 weeks. Animals were anesthetized under CO2 and sacrificed by cervical dislocation. Whole blood was collected, and tissues were flash frozen and stored in liquid nitrogen. Serum folate levels were measured using the SimulTRAC-SNB radioassay kit for vitamin B12 (57Co) and folate (125I) per the manufacturer's protocol (ICN Diagnostics, Orangeburg, NY). Blood was collected at time of sacrifice and allowed to clot at room temperature for 60 min. Samples were centrifuged, and serum was collected for immediate analysis of serum folate levels. Standards provided in the kit were used to generate a standard curve for determination of sample folate values. Radioactivity was measured by a γ-counter, and values were calculated as described by the manufacturer for both serum folate and serum B12. TEMPO Extraction of DNA—Liver DNA for the aldehyde-reactive probe slot blot (ASB) assay was extracted according to the method described by Hofer and Moller (27Hofer T. Moller L. Chem. Res. Toxicol. 1998; 11: 882-887Crossref PubMed Scopus (75) Google Scholar) with some modifications. This method minimizes artifactual DNA damage by using 20 mm TEMPO in all solutions and reagents and by minimizing heat treatment of DNA. Briefly, 100 mg of liver tissue was homogenized in 5 ml of ice-cold phosphate-buffered saline with 20 mm TEMPO and centrifuged at 2000 × g at 4 °C for 5 min. Supernatant was decanted, and pellet was resuspended in 2.5 ml of lysis buffer (pH 8.0; Applied Biosystems, Foster City, CA) with 20 mm TEMPO. Proteinase K (30 units; Ambion, Austin, TX) was added, and samples were incubated overnight at 4 °C. DNA was extracted with 2.5 ml 70% phenol/water/chloroform (Applied Biosystems, Foster City, CA) with 20 mm TEMPO. Further extraction with 2.5 ml of sevag (chloroform/isoamyl alcohol, 24:1) with 20 mm TEMPO was completed. DNA was precipitated using 7.5% 4 m NaCl and 2 volumes of 100% cold ethanol, and pellet was washed in 70% ethanol. The pellet was resuspended in 700 μl of phosphate-buffered saline with 20 mm TEMPO and rehydrated at 4 °C. The samples were then treated with RNase A (2 μg) and RNase T1 (1,000 units; Ambion, Austin, TX) at 37 °C for 30 min to digest the RNA contamination. DNA was cold ethanol-precipitated and resuspended in 400 μl of deionized water at 4 °C. DNA was stored at –70 °C. Gravity Tip Column Extraction of DNA—DNA for the random oligonucleotide-primed synthesis (ROPS) assay was isolated using Qiagen (Valencia, CA) gravity tip columns as described in the manufacturer's protocol. This method generates large fragments of DNA (up to 150 kb) while minimizing shearing. Detection of aldehydic DNA lesions (ADLs) was carried out by ASB as described previously (28Nakamura J. Walker V.E. Upton P.B. Chiang S.Y. Kow Y.W. Swenberg J.A. Cancer Res. 1998; 58: 222-225PubMed Google Scholar) with slight modifications. DNA (8 μg) was incubated in 30 μl of phosphate-buffered saline with 2 mm aldehyde reactive probe (Dojindo Laboratories, Kumamoto, Japan) at 37 °C for 10 min. DNA was precipitated by the cold ethanol method (described above) and resuspended in 1× TE buffer overnight at 4 °C. DNA was heat-denatured at 100 °C for 10 min, quickly chilled on ice, and mixed with an equal volume of 2 m ammonium acetate. The nitrocellulose membrane (Schleicher & Schuell) was prewet in deionized water and washed for 10 min in 1 mm ammonium acetate. DNA was immobilized on the pretreated nitrocellulose membrane using an Invitrogen filtration manifold system. The membrane was washed in 5× SSC for 15 min at 37 °C and then baked under vacuum at 80 °C for 30 min. The dried membrane was incubated in a hybridization buffer (20 mm Tris, pH 7.5, 0.1 m NaCl, 1 mm EDTA, 0.5% (w/v) casein, 0.25% (w/v) bovine serum albumin, 0.1% (v/v) Tween 20) for 30 min at room temperature. The membrane was then incubated in fresh hybridization buffer containing 100 μl of streptavidin-conjugated horseradish peroxidase (BioGenex, San Ramon, CA) at room temperature for 45 min. Following incubation in horseradish peroxidase, the membrane was washed three times for 5 min each at 37 °C in TBS, pH 7.5 (0.26 m NaCl, 1 mm EDTA, 20 mm Tris, pH 7.5, 0.1% Tween 20). Membrane was incubated in ECL (Pierce) for 5 min at room temperature and visualized using a ChemiImager™ system (AlphaInnotech, San Leandro, CA). Nuclear proteins for Western analyses and enzymatic activity assays were isolated using the Sigma CelLytic™ NuCLEAR™ extraction kit, a method that disrupts cells with hypotonic buffer, allowing the cytoplasmic fraction to be removed while the nuclear proteins are released from the nuclei by a high salt buffer. All samples and tubes were handled and chilled on ice, and all solutions were made fresh according to the manufacturer's protocol. The extract was snap frozen in liquid nitrogen and stored at –70 °C. In order to remove salt, the crude nuclear extract was dialyzed against 1 liter of dialysis buffer (20 mm Tris-HCl, pH 8.0, 100 mm KCl, 10 mm NaS2O5, 0.1 mm DTT, 0.1 mm phenylmethylsulfonyl fluoride, 1 μg/ml pepstatin A) for 4–6 h at 4 °C using Slide-A-Lyzer® mini-dialysis units suspended in a floatation device (Pierce). The dialyzed nuclear extracts were flash frozen in liquid nitrogen and stored at –70 °C. Protein concentrations of the nuclear extracts were determined according to Bradford using Protein Assay Kit I (Bio-Rad). Western analysis was performed using liver nuclear extracts (50 μg) subjected to 10% SDS-PAGE and transferred to a Hybond™ ECL™ nitrocellulose membrane (Amersham Biosciences) using a Bio-Rad semidry transfer apparatus. Prior to hybridization, the membranes were stained with MemCode (Pierce) to ensure equal transfer of protein to the membrane. Western blot analysis was accomplished using manufacturer-recommended dilutions of antisera developed against UDG (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), p53 (polyclonal antibody 240; Santa Cruz Biotechnology), β-pol (Ab-1 Clone 18S; NeoMarkers, Fremont, CA), and Ape/Ref1 (Novus Biologicals, Littleton, CO). The bands were detected and quantified using a ChemiImager™ system (AlphaInnotech) after incubation in SuperSignal® West Pico chemiluminescent substrate (Pierce). The data are expressed as the integrated density value (I.D.V.) of the band per μg of protein loaded. UDG activity was determined as described by Stuart et al. (29Stuart J.A. Karahalil B. Hogue B.A. Souza-Pinto N.C. Bohr V.A. FASEB J. 2004; 18: 595-597Crossref PubMed Scopus (97) Google Scholar). Briefly, the 20-μl reaction contained 70 mm Hepes (pH 7.5), 1 mm EDTA, 1 mm DTT, 75 mm NaCl, 0.5% bovine serum albumin, 90 fmol of 5′-end-labeled single-stranded uracil-containing oligonucleotide that was 3′-protected by an amino-spacer (5′-ATATACCGCGGUCGGCCGATCAAGCTTATT-3′, MIDLAND, Midland, TX), and 5 μg of liver nuclear extract. Reactions were incubated at 37 °C for 1 h and then terminated by the addition of 5 μg of proteinase K and 1 μl of 10% SDS and incubation at 55 °C for 30 min. DNA was precipitated in glycogen, ammonium acetate, and ethanol at –20 °C overnight, resuspended in a loading buffer containing 80% formamide, 10 mm EDTA, and 1 μg/ml each of bromphenol blue and xylene cyanol FF. Substrate and reaction products were separated on a 20% denaturing sequencing gel. Glycosylase activity (presence of an 11-mer band) was visualized and quantified using a Molecular Imager® System (Bio-Rad) by calculating the relative amount of the 11-mer oligonucleotide product with the unreacted 30-mer substrate (product/product + substrate). The data are expressed as machine counts/μg of protein. Negative controls consisted of the reaction mixture and oligonucleotide in the absence of nuclear extract. 1 unit of uracil DNA glycosylase inhibitor was added to one sample in each reaction to demonstrate that incision activity was the result of UDG specifically and not another uracil-specific glycosylase (i.e. SMUG). The 5′-endonuclease activity of Ape was analyzed using a quantitative in vitro assay that measures the incision of a 26-mer duplex oligonucleotide substrate containing a tetrahydrofuran (F) AP site as previously described (30Wilson III, D.M. Takeshita M. Grollman A.P. Demple B. J. Biol. Chem. 1995; 270: 16002-16007Abstract Full Text Full Text PDF PubMed Scopus (250) Google Scholar). A 26-mer oligonucleotide (5′-AATTCACCGGTACCFTCTAGAATTCG-3′) was 5′-end-labeled and annealed to an equimolar amount of the complementary strand (5′-CGAATTCTAGAGGGTACCGGTGAATT-3′). 2.5 pmol of double strand oligonucleotide was incubated with 100 ng of crude nuclear extract from liver of control and folate-deficient mice for 15 min at 37 °C and stopped by the addition of 50 mm EDTA (reaction mixture; final concentrations: 50 mm Hepes, pH 7.5, 50 mm KCl, 10 mm MgCl2, 2 mm DTT, 1 μg/ml bovine serum albumin, and 0.05% Triton X-100). Reaction products were run on a 15% denaturing sequencing gel. Endonuclease activity (presence of a 14-mer band) was visualized and quantified using a Molecular Imager® System (Bio-Rad) by calculating the relative amount of the 14-mer oligonucleotide product with the unreacted 26-mer substrate (product/product + substrate). The data are expressed as machine counts/ng of protein. BER capacity was determined as described previously (17Cabelof D.C. Guo Z. Raffoul J.J. Sobol R.W. Wilson S.H. Richardson A. Heydari A.R. Cancer Res. 2003; 63: 5799-5807PubMed Google Scholar). Briefly, end-labeled and purified 30-bp oligonucleotides (upper strand, 5′-ATATACCGCGGUCGGCCGATCAAGCTTATT-3′; lower strand, 3′-TATATGGCGCCGGCCGGCTAGTTCGAATAA-5′) containing a G:U mismatch and an HpaII restriction site (GGCC) and protected by a 3′ amino spacer were incubated in a reaction mixture (100 mm Tris-HCl, pH 7.5, 5mm MgCl2,1mm DTT, 0.1 mm EDTA, 2 mm ATP, 0.5 mm NAD, 20 μm dNTPs, 5 mm di-Tris-phosphocreatine, 10 units of creatine phosphokinase) with 50 μg of nuclear extract isolated from liver of control and folate-deficient mice. The reaction mixtures were incubated for 30 min at 37 °C, followed by 5 min at 95 °C to stop the reaction. The duplex oligonucleotides were allowed to reanneal for 1 h at room temperature and spun down to pellet the denatured proteins. The duplex oligonucleotides present in the supernatant were treated with 20 units of HpaII (Promega, Madison, WI) for 1 h at 37 °C and separated by electrophoresis on a 20% denaturing sequencing gel. Repair activity (presence of a 16-mer band) is visualized and quantified using a Molecular Imager® System (Bio-Rad) by calculating the ratio of the 16-mer oligonucleotide product with the 30-mer substrate (product/substrate). The data are expressed as machine counts/μg of protein. The relative number of 3′-OH group-containing DNA strand breaks was quantified using a Klenow(exo–) incorporation assay based on the ability of Klenow to initiate DNA synthesis from a 3′-OH (31Basnakian A.G. James S.J. DNA Cell Biol. 1996; 15: 255-262Crossref PubMed Scopus (50) Google Scholar). DNA was heat-denatured at 100 °C for 5 min, and 0.25 μg of DNA was added to 15 μl of a Klenow reaction buffer (0.5 mm dTTP, 0.5 mm dGTP, 0.5 mm dATP, 0.33 μm dCTP, 5 units of Klenow(exo–) (New England Biolabs, Beverly, MA)) with 10× Klenow buffer per the manufacturer's protocol (New England Biolabs) and 5 μCi of [α-32P]dCTP (3000 Ci/mmol; PerkinElmer Life Sciences). Reaction mixtures were incubated at 16 °C for 30 min, and the reaction was stopped with the addition of 25 μl of 12.5 mm EDTA, pH 8.0. Samples were spotted (5 μl) onto scored and numbered Whatman DE81 chromatography paper and allowed to airdry. The chromatography paper was then washed five times for 5 min each time in 0.5 m Na2HPO4 (dibasic) to remove unincorporated [α-32P]dCTP and then rinsed twice briefly in water and allowed to air-dry. Paper was cut and placed into scintillation vials with 2.5 ml of Scinti Verse mixture (Fisher). Incorporation of [α-32P]dCTP was quantified using a Packard scintillation counter. Statistical significance between means was determined using analysis of variance followed by Fisher's least significant difference test where appropriate (32Sokal R.R. Rohlf F.J. Biometry. W.H. Freeman and Co., New York1981: 169-176Google Scholar). A p value less than 0.05 was considered statistically significant. In an extensive study, we have carefully characterized the effect of folate deficiency on weight gain/loss and plasma folate levels in C57BL/6 mice. Throughout the 8-week feeding study, the animals' food intake and body weights were monitored twice weekly. Long term folate deficiency can result in toxicity, evidenced primarily by weight loss. Importantly, in our studies, no difference in food intake or weight gain/loss was observed (Fig. 1A). It is essential to demonstrate that the experimental diet (0 mg/kg folic acid) resulted in decreased serum folate levels. As expected, a significant decrease in the level of serum folate in the folate-deficient mouse was observed. The 0 mg/kg folic acid group had serum folate levels 93% lower than the control animals (p < 0.001), such that the folate levels of the deficient animals approached zero (Fig. 1B). The addition of 1% succinyl sulfathiazole allowed attainment of this severity of folate deficiency by preventing intestinal production of folates. Since vitamin B12 can alter one-carbon metabolism through its participation in the methionine synthase reaction, it was important to determine whether our results might be confounded by changes in serum B12 levels. Importantly, no differences were observed in B12 levels between control and deficient groups (data not shown). There was no effect of β-pol heterozygosity on weight or serum folate levels in response to folate deficiency (data not shown). This is the first investigation to directly measure the effect of folate deficiency on BER capacity. It is the BER pathway that holds primary responsibility for removing folate-induced damage from DNA. Previously we (24Cabelof D.
We previously showed that genistein, the major bioactive component of soy isoflavones, acts as a radiosensitizer and potentiates prostate tumor cell killing by radiation in vitro and in animal tumor models in vivo. However, when given alone in vivo, pure genistein promoted increased lymph node metastasis, which was not observed with a soy isoflavone mixture consisting of genistein, daidzein, and glycitein. In this study, we show that soy inhibit tumor cell growth and potentiates radiation-induced cell killing in vitro like pure genistein. In an orthotopic model, combining soy isoflavones with tumor irradiation inhibited prostate tumor growth. To determine the molecular mechanisms by which soy isoflavones potentiate radiotherapy, we investigated apurinic/apyrimidinic endonuclease 1/redox factor-1 (APE1/Ref-1) and nuclear factor kappaB (NF-kappaB), two signaling molecules involved in survival pathways. Soy isoflavones decreased APE1/Ref-1 expression in vitro, whereas radiation up-regulated it. Pretreatment with soy isoflavones followed by radiation inhibited APE1/Ref-1 expression. APE1/Ref-1 decrease correlated with decreased DNA-binding activity of NF-kappaB mediated by soy isoflavones and radiation, thus promoting cell killing. In vivo treatment of prostate tumors with soy isoflavones and radiation down-regulated APE1/Ref-1 protein expression and NF-kappaB activity, confirming the molecular alterations observed in vitro. The down-regulation of APE1/Ref-1 and NF-kappaB by isoflavones, in vitro and in vivo, supports our hypothesis that these markers represent biological targets of isoflavones. Indeed, a 2-fold increase in APE1/Ref-1 expression, obtained by cDNA transfection, resulted in a 2-fold increase in NF-kappaB DNA-binding activity, and both of which were down-regulated by soy isoflavones, confirming the cross-talk between these molecules and, in turn, causing radiosensitization.
