p53 Modulates Hsp90 ATPase Activity and Regulates Aryl Hydrocarbon Receptor Signaling

The aryl hydrocarbon receptor (AhR) is a member of the basic-helix-loop-helix family of transcription factors. It binds with high affinity to polycyclic aromatic hydrocarbons (PAH) which are found in tobacco smoke, diesel engine emissions, and charbroiled food (1). Studies in genetically engineered mice suggest a role for AhR in carcinogenesis (2, 3). Mice engineered to be AhR-deficient were protected against PAH-induced skin tumors (2). In transgenic mice, a constitutively active AhR rendered mice more susceptible to carcinogens (3).

In the absence of ligand, the AhR is present in the cytosol in a complex that includes a dimer of the chaperone Hsp90. Once a PAH such as benzo[a]pyrene (B[a]P) binds to the AhR, it translocates to the nucleus and forms a heterodimer with the AhR nuclear transporter (ARNT). The activated AhR–ARNT complex binds to the upstream regulatory regions of genes containing xenobiotic response elements (XRE), resulting in the induction of a network of genes including CYP1A1 and CYP1B1. Consistent with this mechanism, levels of CYP1A1 and CYP1B1 are increased in the oral and bronchial mucosa of smokers (4–6). CYP1A1 and CYP1B1 can convert B[a]P to a toxic metabolite, anti-7, 8-dihydroxy-9, 10-epoxy-7, 8, 9, 10-tetrahydrobenzo[a]pyrene (BPDE; ref. 7), which covalently binds to DNA, forming bulky DNA adducts that induce mutations (8). In this regard, metabolically activated PAHs, including BPDE, have been reported to form adducts within the p53 tumor suppressor gene that correspond to p53 mutational hot spots in lung cancer (9, 10).

The p53 tumor suppressor gene encodes a homotetrameric transcription factor which is activated in response to a variety of cell stressors (11–14). Under these conditions, the p53 protein is stabilized, initiating a transcriptional program that may result in apoptosis, senescence, DNA repair, or cell-cycle arrest (13). More than 50% of cancers contain mutant p53 (15). p53 mutations have also been found in premalignant lesions (16–18). Recently, we showed that p53 regulated Hsp90 ATPase activity via effects on Aha1 raising the possibility that Hsp90 client proteins such as AhR might be affected (19). Although PAH metabolites such as BPDE have been suggested to cause p53 mutations (9, 10), little is known about the potential of p53 to modulate the Aha1–Hsp90–AhR signaling axis resulting in changes in levels of CYP1A1 and CYP1B1 and PAH metabolism. Hence, this study had several objectives. First, we investigated whether p53 regulated AhR-dependent expression of CYP1A1 and CYP1B1 in cell lines derived from the aerodigestive tract. Second, we used p53 heterozygous mutant epithelial cell lines from patients with Li–Fraumeni syndrome (LFS) to determine whether a similar mechanism was operative. Finally, we investigated whether CP-31398, a p53 rescue agent (20, 21), suppressed B[a]P -mediated induction of CYP1A1 and CYP1B1 and DNA adduct formation. Taken together, this study provides new insights into mechanisms by which p53 regulates AhR signaling and ultimately PAH metabolism, which may be important for both understanding chemical carcinogenesis and developing chemopreventive strategies.

Keratinocyte growth media (KGM) and keratinocyte basal media (KBM) were obtained from Lonza. RPMI, FBS, and Lipofectamine 2000 were from Invitrogen. Antibody to β-actin, Lowry protein assay kits, lactate dehydrogenase (LDH) release assay kit, B[a]P, α-naphthoflavone (αNF), and ATP were from Sigma Chemical. Antibodies to CYP1A1, AhR, p53, and Hsp90 were from Santa Cruz Biotechnology. Antibody to Aha1 was from Abcam. Antiserum to CYP1B1 was a generous gift of Dr. Craig B. Marcus (Oregon State University, Corvallis, OR). CP-31398 and PU-H71 were obtained from Tocris Bioscience. Western Lighting Plus ECL was purchased from Perkin Elmer. Nitrocellulose membranes were from Schleicher and Schuell. RNA was prepared with a kit from QIAGEN. PCR primers were synthesized by Sigma-Genosys. Murine leukemia virus reverse transcriptase and Taq polymerase were purchased from Applied Biosystems. NADH, phosphoenolpyruvate, pyruvate kinase, and LDH were from Boehringer Mannheim. 17-Allylamino-17-demethoxygeldanamycin (17-AAG) was from Cayman Chemicals. Reagents for the luciferase assay, pSVβgal, and plasmid DNA isolation kits were from Promega Corp. The XRE-luciferase construct was a gift from Dr. Michael S. Denison (University of California, Davis, CA). Aha1 expression vector was obtained from GeneCopoeia. p53-luciferase plasmid was from Panomics. A BPDE (anti) standard was obtained from the National Cancer Institute Carcinogen Repository at the Midwest Research Institute (Kansas city, MO).

MSK-Leuk1 cells were established from a premalignant dysplastic leukoplakia lesion adjacent to a squamous cell carcinoma of the tongue (22). Sequencing indicated a GC to AT transition in exon 8 (position 14,525) in one allele of p53, resulting in a glu to lys mutation in codon 286 (data not shown). Cells were routinely maintained in KGM, grown to 70% confluence, and trypsinized with 0.125% trypsin–2 mmol/L EDTA solution. BEAS2B, NCI-H1770, and NCI-H1299 cell lines were obtained from the American Type Culture Collection (ATCC) and maintained according to ATCC instructions. The human colon carcinoma cell line EB-1 was kindly provided by Dr. Arnold J. Levine (Princeton University, Princeton, NJ; refs. 23, 24). This cell line carries wild-type p53 under the control of an inducible metallothionein promoter that exhibits little or no detectable basal activity but sustains high levels of p53 expression following stimulation with zinc chloride. These cells were maintained in RPMI media with 10% FBS and supplemented with 0.5 g/L geneticin (G418). The HME32, HME50, and IUSM/LFS/HME cells have been described previously and were provided by Dr. Brittney-Shea Herbert (Indiana University School of Medicine, Indianapolis, IN; refs. 25, 26). Cell toxicity was assessed by measurements of trypan blue exclusion and LDH release. There was no evidence of cell toxicity under the conditions used. Separate experiments were not done to confirm the authenticity of the cell lines used in our experiments.

Immunoprecipitation was carried out with a kit from Upstate Biotechnology according to the manufacturer's instructions. The immunoprecipitates were then subjected to Western blot analysis.

Cell lysates were prepared by treating cells with lysis buffer as described previously (27). Lysates were sonicated for 8 minutes on ice and centrifuged at 14,000 × g for 10 minutes at 4°C to sediment the particulate material. The protein concentration of the supernatant was measured by the method of Lowry (28). SDS-PAGE was conducted under reducing conditions on 10% PAGE. The resolved proteins were transferred onto nitrocellulose sheets and then incubated with antisera to CYP1A1, CYP1B1, p53, AhR, Hsp90, Aha1, and β-actin. Secondary antibody to immunoglobulin G conjugated to horseradish peroxidase was used. The blots were then incubated with the ECL Western Blot Detection System, according to the manufacturer's instructions.

Total RNA from cell lysates was isolated with the RNeasy Mini Kit (QIAGEN). RNA quantification and quality assessment was done by a NanoDrop 2000c (Thermo Scientific) and 2100 Bioanalyzer (Agilent Technologies). RNA (1 μg) was reverse transcribed with murine leukemia virus reverse transcriptase and oligo d(T)16 primer. The resulting cDNA was then used for amplification. Each PCR reaction volume was 20 μL and contained 5 μL cDNA, 2× SYBR Green PCR master mix, and forward and reverse primers. Primers used were: CYP1A1, forward 5′-CCTGCTAGGGTTAGGAGGTC-3′, reverse 5′-GCTCAGCCTAGTTCAAGCAG-3′ and CYP1B1, forward 5′-ACGTACCGGCCACTATCACT-3′, reverse 5′-CTCGAGTCTGCACATCAGGA-3′. Primers for Aha1 and β-actin, an endogenous normalization control, were purchased from Qiagen. Experiments were carried out with a 7500 real-time PCR system (Applied Biosystems). Relative expression was determined by ΔΔC_T (relative quantification) analysis.

Cells were grown to 60% to 70% confluence in 6-well dishes and then transfected using Lipofectamine 2000 for 24 hours. Subsequently, the medium was replaced with serum-free medium for another 24 hours. Luciferase and β-galactosidase were measured in cellular extracts. MSK-Leuk1 cells stably expressing vector alone or Aha1 were selected by growing the cells in 500 μg/mL hygromycin.

MSK-Leuk1 cells were seeded in KGM for 24 hours before transfection. Two micrograms of siRNA oligonucleotides was transfected using DharmaFECT 4 transfection reagent according to the manufacturer's instructions.

Chromatin immunoprecipitation (ChIP) assays were performed with a kit (Upstate Biotechnology) according to the manufacturer's instructions. The cells (2 × 10⁶) were cross-linked in a 1% formaldehyde solution for 10 minutes at 37°C. The cells were then lysed in 200 μL of SDS buffer and sonicated to generate 200- to 1,000-bp DNA fragments. After centrifugation, the cleared supernatant was diluted 10-fold with ChIP buffer and incubated with 1.5 μg of the indicated antibody at 4°C. Immune complexes were precipitated, washed, and eluted as recommended. DNA–protein cross-links were reversed by heating at 65°C for 4 hours, and the DNA fragments were purified and dissolved in 50 μL of water. Ten microliters of each sample was used as a template for PCR amplification. The forward and reverse primers used for amplifying the CYP1A1 promoter are: 5′-ACCCGCCACCCTTCGACAGTTC-3′ and 5′-TGCCCAGGCGTTGCGTGAGAAG-3′. Forward and reverse primers used to amplify the CYP1B1 promoter are: 5′-GTTCCCTTATAAAGGGAG-3′ and 5′-CTGCGATGGAAGCCGTTG-3′ (27). PCR was performed at 94°C for 30 seconds, 60°C for 30 seconds, and 72°C for 45 seconds for 30 cycles. The PCR products generated from the ChIP template were sequenced, and the identity of the CYP1A1 and CYP1B1 promoters was confirmed.

The ATPase assay was based on a regenerating coupled enzyme assay (29). Hsp90 was immunoprecipitated from cell lysates and the pellet was resuspended and used for assay. Reaction was conducted in a 1-mL assay containing 100 mmol/L Tris-HCl pH 7.4, 20 mmol/L KCl, 6 mmol/L MgCl₂, 0.8 mol/L ATP, 0.1 mmol/L NADH, 2 mmol/L phosphoenolpyruvate, 0.2 mg pyruvate kinase, 0.05 mg L-LDH, and Hsp90 immunoprecipitated from cell lysates. Equal amounts of Hsp90 protein were used in each treatment group. Sufficient NADH was added to give an initial absorbance of 0.3 at 340 nm before addition of Hsp90 and activity was detected as a decrease in absorbance. Hsp90 ATPase activity is expressed as pmol/min/mg protein.

MSK-Leuk1 cells were grown to 70% confluence and placed in 10 mL KBM. Cells were pretreated with vehicle, CP-31398, or αNF for 2 hours. Subsequently, cells received 1 μmol/L B[a]P for 12 hours before cell harvest. Medium (1 mL) was collected from each sample before cell harvest for the measurement of B[a]P-tetrols. DNA was isolated and then hydrolyzed in 0.1 mol/L HCl at 90°C for 2 hours. This treatment releases B[a]P-tetrols from the N2-BPDE adducts (30, 31). After acid hydrolysis, the samples were cooled to room temperature and applied to a Restek Pinnacle II, 3 μm, 150 × 2.1 mm C18 HPLC column. Aliquots of the hydrolysate were eluted in a mobile phase of 33% acetonitrile containing 10 mmol/L ammonium acetate, pH 6.0, at a flow rate of 0.12 mL/min. The eluate was analyzed with the fluorescence detector set at 344 nm excitation and 400 nm emission. A Shimadzu HPLC system consisting of an LC-20AD solvent delivery system, an SIL-10Ai autoinjector, an SPD-20AV UV-VIS detector, and an RF-10AxL fluorescence detector was used. Quantitation of the adducts was achieved by comparison with standards of the B[a]P-tetrol isomers. These were generated by incubating anti-BPDE in water at room temperature for 30 minutes (32). The major adduct designated BPDE tetrol I-1 (31) was produced in the cultured cells (30). Only trace amounts of the minor adduct, BPDE tetrol I-2, were detected. B[a]P-tetrols in the media result from the spontaneous decomposition of BPDE. Tetrols released into the media were quantified by a similar method of comparison with the standard BPDE tetrols.

Comparisons between groups were made by Student t test. A difference between groups of P < 0.05 was considered significant.

Here, we evaluated the potential role of p53 as a determinant of AhR signaling. Initially, we used EB-1 cells, a cell line that expresses Zn⁺⁺ inducible wild-type p53 (23, 24). EB-1 cells were transfected with a p53-luciferase reporter construct. Treatment with ZnCl₂ caused a dose-dependent increase in p53 expression and a corresponding increase in p53-luciferase activity (Fig. 1A). Next, to determine whether p53 regulated AhR-mediated gene expression, we carried out transient transfections using an XRE-luciferase construct. This reporter construct was selected because AhR binds to XREs in the promoters of CYP1A1 and CYP1B1, leading to altered transcription. The same concentration range of ZnCl₂ that activated p53-luciferase caused a dose-dependent decrease in XRE-luciferase activity (Fig. 1B) and a reduction in amounts of CYP1A1 and CYP1B1 mRNA and protein in the EB-1 cells (Fig. 1C and D).

B[a]P, a PAH that binds to the AhR, is a prototypic inducer of CYP1A1 and CYP1B1. As shown in Fig. 1E, levels of CYP1A1 and CYP1B1 were lower in B[a]P-treated EB-1 cells in which ZnCl₂ was used to overexpress p53. Because PAHs are believed to contribute to smoking-related epithelial malignancies, we next evaluated the importance of p53 in regulating CYP1A1 and CYP1B1 expression in cell lines derived from the upper aerodigestive tract. MSK-Leuk1 cells were derived from an oral premalignant lesion and express one mutant allele of p53 (22). Silencing of p53 in MSK-Leuk1 cells led to higher levels of CYP1A1 and CYP1B1 under basal conditions and following treatment with B[a]P (Fig. 1F). Three lung cell lines that vary in p53 status (BEAS2B, wild-type; NCI-H1770, mutant; NCI-H1299, null) were transfected with either p53-luciferase or XRE-luciferase constructs (Fig. 1G and H). Consistent with the findings in EB-1 cells, an inverse relationship was observed between p53 activity and XRE-luciferase activity. Here too, higher levels of CYP1A1 and CYP1B1 were found in cells that were null for p53 or expressed mutant p53 under both basal conditions and following treatment with B[a]P (Fig. 1I and J).

We next investigated the effects of CP-31398, a p53 rescue compound, on AhR-dependent gene expression. Because CP-31398 has the potential to have off-target effects (33), it was important to establish the concentration range that activated p53. Transient transfections were carried out to determine the concentration range of CP-31398 that activated p53 in MSK-Leuk1 cells. As shown in Fig. 2A, CP-31398 caused dose-dependent activation of p53. Notably, B[a]P-mediated induction of CYP1A1 and CYP1B1 was suppressed by the same concentrations of CP-31398 that activated p53 (Fig. 2B). Similarly, CP-31398 activated p53 and blocked B[a]P-mediated induction of CYP1A1 and CYP1B1 in NCI-H1770 cells (Fig. 2C and D).

Previously, we reported that loss of p53 function led to HSF-1–mediated induction of Aha1, activator of Hsp90 ATPase (19, 33, 34). Because AhR is a client protein of Hsp90, we determined whether the p53–Aha1–Hsp90 axis contributed to the regulation of CYP1A1 and CYP1B1 expression. Silencing of p53 in MSK-Leuk1 cells led to increased levels of Aha1 mRNA (data not shown) and protein (Fig. 3A), increased interactions between Aha1 and Hsp90 (Fig. 3B), increased Hsp90 ATPase activity (Fig. 3C), and elevated levels of CYP1A1 and CYP1B1 (Fig. 1F). Silencing of Aha1 led to reduced Hsp90 ATPase activity and decreased levels of CYP1A1 and CYP1B1 (Fig. 3D and E). In contrast, overexpression of Aha1 in MSK-Leuk1 cells led to increased levels of Hsp90 ATPase activity and enhanced expression of CYP1A1 and CYP1B1 under both basal conditions and following treatment with B[a]P (Fig. 3F and G). Next, experiments were performed to confirm that the observed changes in Hsp90 ATPase activity were important for regulating the expression of CYP1A1 and CYP1B1. Thus, treatment with either 17-AAG or PU-H71, prototypic inhibitors of Hsp90 ATPase activity, suppressed levels of CYP1A1 and CYP1B1 (Fig. 3H).

Given the importance of AhR in regulating the transcription of CYP1A1 and CYP1B1, we next determined the effects of p53 on the intracellular localization of AhR. Because silencing of p53 led to increased levels of CYP1A1 and CYP1B1 (Fig. 1F), we initially evaluated whether silencing p53 affected the localization of AhR. As shown in Fig. 4A, silencing p53 caused a shift in the intracellular localization of AhR from cytosol to nucleus. ChIP assays were performed to determine whether the increased nuclear localization of AhR in p53-deficient cells was associated with increased recruitment of AhR to the CYP1A1 and CYP1B1 promoters. In fact, silencing of p53 led to about a doubling in the recruitment of AhR to the promoters of both genes (Fig. 4B). Because CP-31398, the p53 rescue agent, suppressed B[a]P-mediated induction of CYP1A1 and CYP1B1 (Fig. 2B and D), we evaluated its effects on B[a]P-mediated shuttling of AhR from the cytosol to the nucleus. As shown in Fig. 4C, B[a]P stimulated the translocation of AhR from the cytosol to the nucleus, an effect that was inhibited by CP-31398. Consistent with this finding, CP-31398 also inhibited B[a]P-mediated stimulation of AhR recruitment to the CYP1A1 and CYP1B1 promoters (Fig. 4D).

Levels of Aha1 and Hsp90 ATPase activity are increased in LFS cells (19). Here, we investigated whether these effects were also associated with changes in AhR-dependent gene expression. In fact, levels of CYP1A1 and CYP1B1 were increased in LFS cells compared with wild-type p53 HME32 cells under both basal conditions (Fig. 5A) and following treatment with B[a]P (Fig. 5B–E). The increased levels of CYP1A1 and CYP1B1 were associated with a marked increase in amounts of AhR in the nucleus compared with the cytosol in LFS cells (Fig. 5F).

Both CYP1A1 and CYP1B1 can metabolize PAHs such as B[a]P into reactive species that form DNA adducts (35, 36). Because CP-31398 suppressed B[a]P-mediated induction of CYP1A1 and CYP1B1 (Fig. 2B and D), we postulated that it might suppress B[a]P metabolism to BPDE and the formation of DNA adducts. Treatment with B[a]P led to a marked increase in DNA adducts, an effect that was attenuated by treatment with CP-31398 (Fig. 6A). α-Naphthoflavone, a known AhR antagonist that inhibits B[a]P metabolism (37), served as a control and inhibited DNA adduct formation by more than 90%. The major pathway by which B[a]P is metabolized to a genotoxic metabolite involves its oxidation to BPDE. The epoxidation reactions in this pathway are catalyzed by the cytochrome P450 mixed function oxidase system, with CYP1A1 and/or CYP1B1 playing a major role (38). BPDE is extremely reactive and undergoes spontaneous hydrolysis in aqueous medium to B[a]P-tetrols. To determine whether the observed suppression of DNA adducts mediated by CP-31398 reflected changes in B[a]P metabolism, the levels of B[a]P-tetrols were measured in the cell medium. As shown in Fig. 6B, treatment with CP-31398 caused more than a 75% suppression of B[a]P-tetrol formation (P < 0.05). Taken together, reactivating p53 with CP-31398 appears to inhibit the metabolic activation of B[a]P resulting in the suppression of B[a]P-induced DNA adducts.

p53 regulates Aha1 levels and thereby Hsp90 ATPase activity (19). Several studies have demonstrated that AhR is a client protein of Hsp90 (39, 40). Previously, we showed that inhibiting Hsp90 ATPase activity blocked PAH-mediated induction of CYP1A1 and CYP1B1 transcription and DNA adduct formation (41). Here, we extend upon these observations by demonstrating that p53 is a determinant of CYP1A1 and CYP1B1 levels under both basal conditions and following stimulation with B[a]P. Several lines of evidence support these points. Levels of CYP1A1 and CYP1B1 were increased in LFS-derived p53 heterozygous versus normal epithelial cells. Silencing of p53 led to increased Aha1 levels, elevated Hsp90 ATPase activity, and increased levels of CYP1A1 and CYP1B1. In contrast to the effects of silencing p53, expression of wild-type p53 in EB-1 cells, a p53-null cell line, caused a gene dose-dependent suppression of CYP1A1 and CYP1B1 mRNA and protein. Moreover, overexpression of wild-type p53 or treatment with CP-31398, the p53 rescue compound, blocked B[a]P-mediated induction of CYP1A1 and CYP1B1. Because p53 can regulate Hsp90 ATPase activity, the significance of Hsp90 in regulating AhR signaling was tested. We showed that 17-AAG and PU-H71, prototypic inhibitors of Hsp90 ATPase, downregulated the expression of CYP1A1 and CYP1B1. To further evaluate the functional significance of Aha1, activator of Hsp90 ATPase, in mediating p53-dependent changes in AhR-dependent gene expression, Aha1 was silenced. Silencing of Aha1 led to reduced Hsp90 ATPase activity and downregulated CYP1A1 and CYP1B1. Interestingly, stably overexpressing Aha1 in MSK-Leuk1 cells was associated with increased Hsp90 ATPase activity and elevated levels of CYP1A1 and CYP1B1 both under basal conditions and following treatment with B[a]P. The above changes in levels of CYP1A1 and CYP1B1 reflected changes in gene transcription. This conclusion is supported by evidence that the AhR was primarily in the nucleus rather than the cytosol in LFS-derived p53 heterozygous versus normal epithelial cells. Moreover, silencing of p53 led to a shift in the AhR from cytosol to nucleus in association with increased recruitment of the AhR to the promoters of CYP1A1 and CYP1B1. Consistent with these findings, CP-31398 blocked B[a]P-mediated shuttling of AhR from cytosol to nucleus and the related recruitment of AhR to the promoters of CYP1A1 and CYP1B1. Both endogenous and exogenous ligands can activate AhR-dependent gene expression (4, 42). It is possible, therefore, that p53 modulates the levels of an endogenous ligand leading to the observed effects. An alternate possibility is that these effects of p53 are ligand-independent. For example, p53 might have posttranslational effects on Hsp90 that could impact on its chaperoning activity (27).

It is important to consider the potential implications of these mechanistic findings. Several studies have suggested that PAH metabolites may play a clinically relevant role in the pathogenesis of p53 mutations (9, 10). Possibly, PAH-mediated induction of CYP1A1 and CYP1B1 will contribute to p53 mutations resulting in further increases in xenobiotic metabolism, leading to increased DNA adduct formation and mutagenesis. It is also important to acknowledge that the AhR can suppress antitumor immune responses (43). p53 can influence innate immune responses as part of its tumor suppressor activities (44). Perhaps p53 mutations will contribute to a failure of immune surveillance by stimulating AhR. Whether these mechanisms contribute to the observed increased incidence of second primary tumors in patients with head and neck squamous cell carcinoma who continue to smoke is unknown and warrants consideration (45).

It is also relevant to consider our findings in the context of LFS. LFS is associated with an increased risk of several tumor types including breast cancer (46). Some studies have suggested an increased risk of non–small cell lung cancer in LFS families (47). Previously, using expression arrays, AhR signaling was suggested to be increased in phenotypically normal epithelial cells from LFS p53 mutation carriers (26). The current study extended upon these findings. We found a significant increase in the transcriptional activity of AhR in untreated, normal-appearing epithelial cells taken from LFS carriers leading to both elevated basal levels of CYP1A1 and CYP1B1 and increased expression following treatment with B[a]P. It is possible, therefore, that augmented xenobiotic metabolism, reduced immune surveillance, or both may contribute to the increased cancer risk among patients with LFS. Because CYP1B1 can catalyze the conversion of estrogens into DNA reactive mutagens (48), our data may be relevant for understanding the pathogenesis of tumors including breast cancer among patients with LFS.

There was good concordance between the effects of CP-31398 on levels of B[a]P-tetrols in the medium and DNA adducts. This suggests that levels of B[a]P-tetrols in the medium serve as markers of cellular metabolism of B[a]P to BPDE. Our results suggest that CP-31398, the p53 rescue compound, blocks B[a]P-mediated induction of CYP1A1 and CYP1B1, resulting in reduced B[a]P metabolism and DNA adduct formation. An alternative explanation, such as induction of DNA repair activity by CP-31398, would not explain the significant reduction in levels of B[a]P-tetrols found in the medium following treatment with the p53 rescue agent. On the basis of our findings, it seems predictable that suppressing PAH-mediated induction of CYP1A1 and CYP1B1 should suppress the formation of DNA adducts and thereby inhibit carcinogenesis. However, this may not be the case in vivo. Studies with CYP1A1-knockout mice have suggested that this enzyme is important in detoxification and protection against B[a]P toxicity (49). Moreover, some models of PAH-induced carcinogenesis indicate that activation of AhR signaling may protect against chemical carcinogenesis (50). As mentioned above, AhR plays a significant role in immunosurveillance in addition to xenobiotic metabolism. Moreover, p53 rescue compounds should have multiple antitumor activities in addition to effects on AhR-dependent functions. Administering a p53 rescue compound topically as a rinse or lozenge or even in an aerosol might protect against tobacco smoke–mediated induction of AhR-dependent genes and possibly mutagenesis in the upper aerodigestive tract. Whether such agents will reduce the risk of smoking-related malignancies warrants consideration.

No potential conflicts of interest were disclosed.

Conception and design: L. Kopelovich, A.J. Dannenberg, K. Subbaramaiah

Development of methodology: L. Kopelovich, J.B. Guttenplan, K. Subbaramaiah

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): A. Kochhar, E. Sue, J.B. Guttenplan, K. Subbaramaiah

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): A. Kochhar, L. Kopelovich, J.B. Guttenplan, A.J. Dannenberg, K. Subbaramaiah

Writing, review, and or revision of the manuscript: A. Kochhar, L. Kopelovich, J.B. Guttenplan, B.-S. Herbert, A.J. Dannenberg, K. Subbaramaiah

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): A. Kochhar, L. Kopelovich, A.J. Dannenberg, K. Subbaramaiah

Study supervision: K. Subbaramaiah

Other: J. Guttenplan developed the method for DNA adduct analysis; B.-S. Herbert provided HME cells.

This work was supported by NIH grant NIDCD 5T32DC000027–22 (to A. Kochhar) and the Breast Cancer Research Foundation.

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.