|Home | About | Journals | Submit | Contact Us | Français|
Over 45,000 new cases of oral and pharyngeal cancers are diagnosed and account for over 8,000 deaths a year in the United States. An environmental chemical receptor, the aryl hydrocarbon receptor (AHR), has previously been implicated in oral squamous cell carcinoma (OSCC) initiation as well as in normal tissue-specific stem cell self-renewal. These previous studies inspired the hypothesis that the AHR plays a role in both the acquisition and progression of OSCC, as well as in the formation and maintenance of cancer stem-like cells. To test this hypothesis, AHR activity in two oral squamous cell lines was modulated with AHR prototypic, environmental and bacterial AHR ligands, AHR-specific inhibitors, and phenotypic, genomic and functional characteristics were evaluated. The data demonstrate that: 1) primary OSCC tissue expresses elevated levels of nuclear AHR as compared to normal tissue, 2) Ahr mRNA expression is up-regulated in 320 primary OSCC, 3) AHR hyper-activation with several ligands, including environmental and bacterial ligands, significantly increases AHR activity, ALDH1 activity, and accelerates cell migration, 4) AHR inhibition blocks the rapid migration of OSCC cells and reduces cell chemoresistance, 5) AHR knockdown inhibits tumorsphere formation in low adherence conditions, and 6) AHR knockdown inhibits tumor growth and increases overall survival in vivo. These data demonstrate that the AHR plays an important role in development and progression of OSCC, and specifically cancer stem-like cells. Prototypic, environmental and bacterial AHR ligands may exacerbate OSCC by enhancing expression of these properties.
This study, for the first time, demonstrates the ability of diverse AHR ligands to regulate AHR activity in oral squamous cell carcinoma cells, as well as regulate several important characteristics of oral cancer stem cells, in vivo and in vitro.
Historically, the majority of studies on environmental chemical carcinogenesis have focused on the ability of genotoxic chemicals to damage DNA and to induce mutations [1, 2]. More recent data suggest that environmental carcinogens also contribute to carcinogenesis by activating signaling pathways responsible for cancer progression .
At the apex of one such pathway is the aryl hydrocarbon receptor (AHR). The AHR belongs to the basic helix-loop-helix/Per-ARNT-Sim (bHLH/PAS) family of transcription factors that control many important biological functions, including cell cycle maintenance, homeostasis, early cell differentiation, and stress responses . The AHR has been widely studied for its ability to be activated by environmental dioxins, polychlorinated biphenyls (PCBs), and polycyclic aromatic hydrocarbons (PAHs) , all of which are high priority chemicals on the U.S. Agency for Toxic Substances and Disease Registry list of pollutants of greatest concern to human health (http://www.atsdr.cdc.gov/SPL/resources). Important recent studies indicate that AHR ligands may be more ubiquitous than previously thought, although not necessarily from environmental sources. Rather, ligands that activate the AHR under physiologically “normal” conditions likely derive from the products of mammalian tryptophan or arachidonic acid metabolism pathways [6–9], and from microbial sources [10, 11]. The latter findings are particularly remarkable since they suggest that the microbiome may play an important role in normal and pathologic AHR signaling and, as noted below, cancer.
Over 45,000 new cases of oral squamous cell carcinoma (OSCC) are diagnosed and account for over 8,000 deaths a year in the United States . The disease and its treatments are highly debilitating and the 5-year survival rates of regionally invasive tongue and floor-of-the-mouth cancers range from only 38% to 63% [13, 14]. It has been suggested that chemoresistant oral cancer stem-like cells (CSLC) contribute significantly to these relatively poor outcomes [15–18]. CSLCs are defined as a relatively small subset of chemotherapy- and radiotherapy-resistant, invasive, metastatic tumor cells that are highly efficient at tumor initiation [19–22]. Oral CSLCs also tend to exhibit higher levels of ALDH1, an enzyme associated with chemo- and radio-resistance, than non-CSLCs [16, 20]. Indeed, there is a direct correlation between ALDH levels and disease stage, prognosis, and chemotherapeutic resistance [18, 21]. Consequently, elucidation of pathways through which CSLCs are produced and maintained is an important next step in identifying potential candidates for targeted OSCC therapy.
It has been estimated that 70%–90% of OSCC are linked to tobacco use, with a linear relationship between the number of smoking years and OSCC risk [23, 24]. There are over 60 carcinogens found in tobacco products, including several AHR ligands such as benzo(a)pyrene (B(a)P) and 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) [25, 26]. B(a)P and other ambient environmental PAHs have been implicated in OSCC through their activation of the AHR, subsequent induction of AHR-regulated P450 enzymes (CYP1A1, CYP1B1), and P450-dependent production of DNA-reactive metabolic intermediates . PAHs also likely play a role in OSCC by altering key cell fate decisions and promoting cell immortalization over senescence . Similarly, TCDD, a high affinity and poorly metabolized AHR ligand, likely contributes to carcinogenesis by chronically activating the AHR signaling pathway . Furthermore, “constitutive” or chronic AHR activation in other cell types has been implicated in cancer progression even in the absence of environmental ligands [30, 31]. Therefore, analysis of AHR signaling in OSCC would both increase our understanding of the etiology of the disease in the presence or absence of environmental stimuli, and identify a novel therapeutic target, i.e., the AHR, regardless of disease etiology.
Here, we explore the role of the AHR in OSCC with particular emphasis on outcomes associated with advancing cancer, including production of oral CSLCs and increased migration. To address the possible contribution of environmental and microbial AHR ligands to OSCC, several classes of AHR ligands were used including a genotoxic PAH (B(a)P), a non-genotoxic but persistent halogenated hydrocarbon (TCDD), PAH-containing 2.5 μm airborne particles (PM2.5), and products from Pseudomonas aeruginosa and Porphyromonas gingivalis, the latter a primary etiological agent associated with chronic periodontitis. To assess the feasibility of targeting the AHR signaling pathway as a novel OSCC treatment, we used two OSCC cell lines, a unique OSCC orthotopic xenograft model , a novel AHR inhibitor (CB7993113) , and molecular knockdown techniques.
All studies with surgical OSCC specimens were approved by the Institutional Review Board at the Boston University Medical Campus. Fresh tissues were obtained from patients with moderately differentiated OSCC of the lateral tongue and base of the tongue. Regions of OSCC and adjacent epithelia (AE), defined by an on-site pathology analysis, were snap-frozen at 80°C. Tissues were divided for H&E analyses, biochemistry, and immunofluorescence staining. OCT-embedded fresh tumor tissues were used for preparation of frozen sections (5 μM). One frozen section was set aside for H&E staining, and the remaining sections were processed for immunofluorescence analyses as described below.
For indirect immunofluorescence analyses, sections (5 μM) of OSCC OCT-embedded tissues were blocked with 10% goat serum and incubated with antibodies against AHR followed by secondary antibodies conjugated with FITC. Negative controls lacked primary antibodies and no staining was observed. The slides were mounted in Vectashield and optical sections were analyzed with a Zeiss LSM 510 META confocal microscope. To compare fluorescence intensities between samples, settings were fixed to the most highly stained sample with all other images acquired at those settings. Nuclear and cytosolic AHR were quantified using ImageQuant TL software (GE Healthcare Life Sciences, Pittsburgh, PA) and calculated as nuclear or cytosolic density/nuclear + cytosolic density x 100%.
DMSO, β-naphthoflavone (β-NF), 7,12-dimethylbenz[a]anthracene (DMBA), 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD), benzo(a)pyrene (B(a)P), pyocyanin (PYO), indoxyl sulfate (IS), thiazolyl blue tetrazolium bromide (MTT), and Cisplatin were obtained from Sigma-Aldrich (St. Louis, MO). 6-Formylindolo(3,2-b)carbazole (FICZ), CH223191, and CB7993113 were provided by Dr. M. Pollastri (Northeastern University). AHR-antibody was purchased and used from Cell Signaling (Product #: 13790)(Danvers, MA).
OSCC tissue lysates were fractionated on 4–12% SDS-polyacrylamide gels, transferred onto polyvinylidene difluoride membranes, blocked with 10% nonfat dry milk, and incubated with primary antibodies to AHR and β-actin. Protein-specific detection was carried out with horseradish peroxidase-labeled secondary antibodies and Enhanced Chemiluminescence Plus (Amersham Biosciences).
Frozen stocks of P. gingivalis wild-type strain 381 were grown anaerobically at 37° C on blood agar plates (Remel) for 3–5 days. Blood-heart infusion medium (Becton-Dickinson Biosciences) supplemented with yeast extract (0.5%; Becton-Dickinson Biosciences), hemin (10 mg/ml; Sigma-Aldrich), and menadione (1 mg/ml; Sigma- Aldrich) was inoculated with plate-grown bacteria and cultures grown anaerobically for 16–18 hours. P. gingivalis was then harvested and transferred to DMEM (Mediatech, Herndon, VA) containing 10% FBS (Sigma-Aldrich) for 48 hours, the supernatant was collected, double sterile filtered, diluted 1:10 and used for experiments (PGS).
For P. aeruginosa, DMEM (Mediatech, Herndon, VA) containing 10% FBS (Sigma-Aldrich) was inoculated with the PAO1 strain (PerkinElmer Xen41; isolated after overnight growth on blood agar plates) and cultured while shaking in room air at 37°C for 6 hours; a sterile supernatant was collected by removing bacteria using sedimentation (at 1500g) followed by filtration (with 0.22-μm pores). The P. aeruginosa supernatants (PAS) were confirmed sterile and stored at 4°C until used in experiments.
2.5μm particulate matter samples were collected for 1 week from a ventilation exhaust stack in a northeast US urban highway tunnel. The samples were collected on polyurethane foam using a High Volume Cascade Impactor (HVCI). PM2.5 were released from the collection filter by vigorous washing in PBS followed by sonication and used at a final concentration of 5 mg/cm .
CAL27 cells were purchased from and cultured according to ATCC recommendations (ATCC, Manassas, VA). All experiments were performed within six months of vial thawing from ATCC, which validates its cell lines via STR-profiling and COI analysis. HSC-3 cells were a generous gift from Dr. Roberto Weigert (NIDCR, Bethesda, MD) and were validated by Genetica DNA Laboratories (Burlington, NC) within six months of all experimentation via STR Profiling. Cells were maintained in DMEM (Mediatech, Herndon, VA) containing 10% FBS (Sigma-Aldrich), 100 I.U. penicillin/100 μg/ml streptomycin (Mediatech), 10 μg/ml insulin (Sigma-Aldrich), and 5 μg/ml plasmocin (Invivogen, San Diego, CA).
Cells were dosed with 0.5 μM FICZ, 10 μM CH223191, 10 μM CB7993113, 1 nM TCDD, 1 μM DMBA, 10 μM IS, 10 μM B(a)P, 1 μM PYO, a 1:2 dilution of P. aeruginosa supernatant, a 1:10 dilution of P. gingivalis supernatant, 5 mg/cm PM2.5, and/or vehicle (0.1% DMSO or PBS) for 24 hours. ALDEFLUOR™ assays were performed according to the manufacturer’s instructions (StemCell Technologies, Vancouver, Canada). Cells (106 cells/ml) were treated with 5 μl/ml ALDEFLUOR substrate. Negative controls were treated with 50 mmol/L diethylaminobenzaldehyde (DEAB), an ALDH-specific inhibitor. Samples were incubated for 35 minutes at 37°C, washed, and suspended in ALDEFLUOR assay buffer. 1.5 μg/ml propidium iodide (PI) was added before samples were assayed to quantify viability. Cells were phenotyped with an LSRII flow cytometer (Becton Dickinson Biosciences, San Jose, CA) using DEAB controls as baselines. All data was analyzed using FlowJo (FlowJo LLC, Ashland, OR).
HSC-3 or CAL27 cells were treated with 10 μM CH223191, 10 μM CB7993113, 0.5 μM FICZ, 1 nM TCDD, and/or vehicle (0.1% DMSO or PBS) every 24 hours. After 48 hours, cells were harvested, dosed, and 3 x 103 cells plated in MammoCult Medium (STEMCELL Technologies) containing 0.5 μg/ml hydrocortisone, 2 mM L-glutamine, 100 I.U. penicillin/100 μg/ml streptomycin, and 1% methylcellulose (Sigma-Aldrich) in ultra-low adherent 24-well plates (Corning). Colonies were quantified with a Celigo S Imaging Cytometer (Brooks Automation, Chelmsford, MA) after 8 days. For secondary sphere formation, tumorspheres were mechanically and enzymatically dissociated with TrypLE Express (Gibco) for 10 minutes at 37° C until a single cell suspension was formed. The cells were then re-plated and imaged as above.
Cells were dosed as above for 24 hours. mRNA was extracted using the RNeasy® Plus Mini Kit (Qiagen, Valencia, CA) and cDNA prepared using the GoScript™ Reverse Transcription System (Promega, Madison, WI) with a 1:1 mixture of random and Oligo (dT)15 primers. All RT-qPCR reactions were performed using the GoTaq® RT-qPCR Master Mix System (Promega). Validated primers were purchased from Qiagen: human Cyp1b1 - QT00209496, and Gapdh – QT01192646. RT-qPCR reactions were performed using a 7900HT Fast Real-Time PCR instrument (Applied Biosystems, Carlsbad, CA) with hot-start activation at 95° C for 2 min, 40 cycles of denaturation (95° C for 15 sec) and annealing/extension (55° C for 60 sec). Relative gene expression was determined using the Pfaffl method  and the threshold value for Gapdh mRNA was used for normalization.
HSC-3 or CAL27 cells cultured in 24-well plates were co-transfected with 0.5 μg pGudluc reporter plasmid, provided by Dr. M. Denison (UC, Davis), and CMV-green (0.25 μg) using 1μL TransIT-2020 transfection reagent (Mirus, Madison, WI). Transfection medium was replaced after 24 hours. The cells were left untreated or dosed as above and harvested after 24 hours in Glo Lysis Buffer (Promega, San Luis Obispo, CA). Luciferase activity was determined with the Bright-Glo Luciferase System according to the manufacturer’s instructions (Promega). Luminescence and fluorescence were determined using a Synergy2 multifunction plate reader (Bio-Tek, Winooski, VT).
HSC-3 cells were grown to confluence in 6-well plates. At confluence, media was changed to serum-free media and cells were pre-dosed for 24 hours as above with FICZ, TCDD, B(a)P, PYO and/or PM2.5 with or without 10 μM CH223191 or CB7993113. After 24 hours a p200 pipet tip was used to make an “X” scratch in each well and non-adherent cells were removed with three PBS washes. Serum-containing media was added and cells re-dosed. TScratch software (Tobias Gebäck and Martin Schulz, ETH Zürich) quantified the closure of the scratch at 0 and 24 hours.
CAL27 cells (3 x 105) were injected into the tongues of six-week old female NCr nude mice (CrTac:NCr-Foxn1nu, Taconic, Hudson, NY) as we previously described . Mice began a daily lavage schedule with 50 μl sesame oil alone or CB7993113 dissolved in sesame oil (50 mg/kg/day in 50 μl) 24 hours before tongue injections. Animals were weighed and tumor size was measured with Vernier calipers over a 26 day period. Animals were sacrificed when mice lost 20% of their total body mass. Surviving mice were sacrificed by day 26. Animals were housed at the Association for Assessment and Accreditation of Laboratory Animal Care-certified Boston University Medical Laboratory Animal Science Center and used in accordance with the NIH Guide for the Care and Use of Laboratory Animals. A Boston University Medical Campus Institutional Animal Care and Use Committee-approved protocol was followed.
CAL27 cells were plated at 3,500 cells per a well in 96-well plates. After 24 hours, cells were treated with a 1 μM to 500 μM Cisplatin with or without 10 μM CH223191. Twenty-four hours later, 10 μL of a 10 mg/mL MTT solution was added to each well, and plates were incubated for 3 hours at 37°C. MTT and media were aspirated off and DMSO was added to solubilize the MTT crystals. After 2 hours of solubilization, the plate absorbance was read at 590 nm and 620 nm (reference filter) to determine cell viability.
Statistical analyses were performed with StatPlus (Alexandria, VA) unless otherwise noted. Data are presented as means + standard errors (SE) where applicable. One-way analysis of variants (ANOVAs) (simple) were used to determine significance. For experiments measuring relative fold-changes in gene expression, the Pfaffl method  was used for normalization to Gapdh mRNA levels.
Previous work implicated the AHR in cancer progression and tissue stem cell biology [36, 37]. We, and others, have demonstrated that the AHR is highly expressed and constitutively active in breast cancers and that its activity correlates with tumor aggressiveness . However, the role of the AHR in OSCC cells has not been examined thoroughly. First, AHR protein levels in primary tissue were evaluated by immunofluorescence. A very weak AHR signal was observed in normal tissue and the stain that could be discerned was 87.1% cytosolic (Figure 1A). In contrast, basal or lateral tongue OSCC expressed higher levels of AHR, with 92.9% of the cells having nuclear AHR staining (Figure 1A, arrows), indicating constitutively active AHR. Elevated AHR protein levels in these tumor samples were reflected in a significantly higher level of Ahr mRNA (p<0.001), as documented in the TCGA OSCC dataset, in 320 primary OSCCs (Figure 1B).
To determine if the AHR in oral squamous carcinoma cells was functional, HSC-3 and CAL27 cells, which express significant AHR protein levels (Supplemental Figure S1), were transiently transfected with an AHR-dependent luciferase reporter plasmid (pGudLuc) , treated with various AHR ligands in the absence or presence of AHR inhibitors CH223191  or CB7993113  (10 μM), and luciferase activity assayed. The baseline level of AHR (reporter) activity in both HSC-3 (Figure 2A) and CAL27 (Figure 2B) cells significantly decreased following addition of CH223191 or CB7993113 (p<0.05-0.005), supporting the hypothesis that the AHR was “constitutively active”, presumably because of endogenous ligands in these lines. Conversely, treatment with any of two prototypic AHR ligands, 0.5 μM FICZ or 1 μM β-napthaflavone (β-NF), significantly induced AHR activity (p<0.05-0.005). B(a)P (10 μM), a readily metabolized environmental PAH, TCDD (1 nM), a high affinity, persistent environmental AHR ligand, and PM2.5 (5 mg/cm), all significantly induced AHR activity (p<0.05-0.005). In each case, CH223191 significantly inhibited induction of AHR activity (p<0.05-0.005) (Figure 2A and 2B).
It was recently reported that some bacterial species, including P. aeruginosa, produce physiologically relevant levels of AHR ligands or tryptophan metabolites that are the precursors to AHR ligands [10, 39]. Additionally, recent studies have implicated altered microbiomes and elevated levels of microbes in oral squamous cell carcinoma [40, 41]. To confirm and extend these observations, HSC-3 and CAL27 cells were treated with 10 μM indoxyl sulfate (IS), a tryptophan-derived AHR ligand that requires both bacterial and mammalian enzymes to be produced , 1 μM pyocyanin (PYO), a P. aeruginosa-derived pigment and AHR ligand , P. aeruginosa supernatant (PAS)(diluted 1 to 2), and P. gingivalis supernatant (PGS)(diluted 1 to 10). All of these treatments resulted in significant increases in AHR activity (Figure 2A and 2B). To our knowledge, this is the first demonstration that P. gingivalis, a bacterium commonly found in the oral cavity, produces AHR ligand(s). In all cases, the induced AHR activity was suppressed by addition of 10 μM CH223191 (p< 0.05). It is important to note that the bacterial media did not induce AHR activity.
To determine if an endogenous AHR target gene was similarly affected by AHR ligands and inhibitors, HSC-3 cells were treated with CH223191 or CB7993113 and/or the ligands described above and Cyp1b1 mRNA levels quantified 24 hours later. Both AHR inhibitors significantly decreased baseline Cyp1b1 levels (Figure 2C) (p< 0.05-0.005). Similar data were obtained with CAL27 cells (data not shown). An increase in Cyp1b1 after treatment with “prototypic”, environmental, or bacterial-derived ligands and suppression of this increase with CH223191 (Figure 2C) further suggested the potential for an array of AHR ligands from diverse sources to influence OSCC biology. (Note that all ligands and inhibitors were tested in all cell lines and the trend of increased signal with agonists and a decrease with agonist plus inhibitor were consistent with all ligands and cell lines tested. That is, there were no ligands with which an effect was seen in one cell line and not in another. Data from independent experiments replicated fewer than three times were not included here).
CSLCs contribute to tumor progression and recurrence in multiple cancer types, including OSCCs [15, 16, 19, 17, 18, 20–22]. Previous studies with normal tissue-specific stem cells suggest that the AHR contributed to properties of “stem-ness” [36, 37, 42, 43]. One readily assessed marker of stem-ness is elevated expression of ALDH1, an enzyme implicated in chemoresistance  the activity of which can be quantified via flow cytometry [16, 20]. In contrast to other tumor types, expression of surface CSLC markers, such as CD44 and CD24, is highly variable and not reliable in OSCC [16, 20, 22]. Therefore, a fluorescence-based ALDH1 enzyme activity assay  was used to quantify ALDH1 activity in HSC-3 and CAL27 cells in the absence or presence of diverse AHR ligands and/or AHR inhibitors.
As noted for baseline pGudLuc activity (Figure 2A and 2B) and endogenous Cyp1b1 mRNA levels (Figure 2C), AHR inhibitors significantly reduced the baseline percentage of ALDH1high HSC-3 (Figure 3A)(p<0.005) and CAL27 (Figure 3B)(p<0.05) cells. Conversely, all of the AHR ligands significantly increased the percentage of ALDH1high cells in HSC-3 and/or CAL27 and this increase was inhibited by CH223191 (p<0.05-0.005) for the vast majority of ligands in both cell lines. These data suggest that the AHR influences expression of this OSCC stem cell-associated marker.
ALDH expression has been associated with chemoresistance [44, 45]. Since the AHR appears to influence ALDH expression (Figure 3), we postulated that the AHR may confer relative chemoresistance to OSCC cells. To test this hypothesis, CAL27 cells were treated with titrated doses of a front-line chemotherapeutic, Cisplatin, with or without CH223191 and cell viability was assayed 24 hours later. CH223191 treatment alone had no effect on viability (i.e., >95% viability in untreated and CH223191-treated cultures, data not shown). However, CH223191 significantly (p<0.05) increased sensitivity to Cisplatin in CAL27 cells over a dosing range of 1 μM to 500 μM Cisplatin. The EC50 of Cisplatin alone was 232.5 μM as compared with 84.2 μM for Cisplatin + CH223191 (Figure 4). These results demonstrate that the AHR contributes to chemoresistance.
Oral CSLC can form tumorspheres and produce progenitor cells in ultralow adherence conditions over several passages . To determine if the AHR contributes to this functional readout of CSLCs, HSC-3 (Figure 5A and 5B) and CAL27 (Figure 5C and 5D) cells were cultured in Mammocult media under ultra-low adherence conditions and AHR activity was inhibited with CH223191 or CB7993113. Both the size and total number of HSC-3 or CAL27 tumorspheres were significantly reduced (p<0.05-0.005) by AHR antagonist treatment in primary, secondary (Figures 5), tertiary, and quaternary (not shown) cultures. These results suggest that the AHR regulates tumorsphere formation by influencing: 1) the ability of CSLCs to differentiate into progenitor cells, 2) the ability of CSLCs to (asymmetrically) divide, and/or 3) the ability of progenitor cells to divide. These functional data, together with the phenotypic data presented in Figure 3, are consistent with a role for the AHR in CSLC development.
A key characteristic of invasive and malignant cancer cells is the ability to migrate. Here, the effects of AHR modulation on the ability of HSC-3 cells to migrate were determined in a scratch-wound assay. Wound repair, a measure of cell migration, was significantly inhibited by CH223191 or CB7993113 treatment (Figure 6A and 6B) (p<0.05). Conversely, 0.5 μM FICZ, 10 μM B(a)P, 1 nM TCDD or 1 μM pyocyanin significantly accelerated cell migration (Figure 6C and 6D) (p<0.05-.005). In all cases, AHR agonist-induced increases in migration rates were significantly inhibited by CH223191 treatment (p<0.05-0.005). Combinations of AHR ligands similarly increased migration rates (Supplemental Figure S2). These data suggest that environmental or bacterial-derived AHR ligands accelerate and AHR antagonists reduce OSCC cell migration and metastasis.
To further explore the role of the AHR in the OSCC, 3 x 105 CAL27 cells were injected into tongues of 24 female, NCr nude mice. Twelve mice were lavaged daily with vehicle (sesame oil) alone (controls) and twelve mice were lavaged daily with CB7993113 (50 mg/kg/day). Tumor volume, animal weight, and survival were evaluated over the next 26 days. Tumor volumes in CB7993113-treated mice were significantly (p<0.05-0.0005) smaller than in control mice beginning at day 5 and continuing until the time at which control mice had to be sacrificed (Figure 7A). In parallel, the weights of control mice began to drop on day 9 and were significantly lower than those of CB7993113-treated mice by day 12, after which the first control mouse had to be sacrificed (Figure 7B). Interestingly, the weights of CB7993113-treated mice that survived until day 26 were stable or slightly increased between day 15 and the termination of the experiment on day 26 (Figure 7B and data not shown). These differences in tumor volume and mouse weights translated into significantly better survival of CB7993113-treated mice (83% survival at 26 days) as compared with control mice (25% survival at 26 days) (Figure 7C). These results are consistent with the findings from the in vitro experiments and strongly support the hypothesis that the AHR plays an important role in OSCC.
Cancers of the head and neck have been clearly linked to exposure to environmental carcinogens, most commonly those found in cigarette smoke. A significant mechanism of carcinogenesis is the production of mutagenic compounds, including PAH-derived epoxides. Since many of those epoxides are produced through the catabolic activity of AHR-regulated P450 oxygenases, the AHR has been indirectly linked to OSCC initiation. Notably, upregulated Cyp1b1 in the oral mucosa is a biomarker for smoke exposure and OSCC risk . As demonstrated for the first time here, chronically activated AHR, by what likely are endogenous ligands, may also play an ongoing role in OSCC progression by driving the expression of several phenotypic and functional characteristics of advancing cancers. These studies also suggest the possibility that ongoing exposure to environmental AHR ligands accelerates still further the development of aggressive, difficult to treat OSCCs.
High-level AHR expression in primary cancers was demonstrated by immunofluorescence of primary tumor samples, computational analysis of mRNAs from 320 primary human cancers and 32 adjacent epithelial samples and western blotting (data not shown). The findings that AHR was highly expressed and “constitutively” active, at least in representative OSCC cell lines, was supported by a decrease in baseline pGudLuc reporter activity, a decrease in Cyp1b1 expression, a decrease in the percentage of ALDH1high cells, a decrease in tumorsphere formation, a slowing of cell migration and a decrease in tumorigenesis in vivo following treatment with an AHR antagonist. Consistent with the hypothesis that environmental AHR ligands can exacerbate cancer progression, prototypic AHR ligands as well as environmental AHR ligands increased pGudLuc activity, Cyp1b1 transcription, the percentage of ALDH1high cells, and the rate of cell migration.
One of the outcomes mediated by the AHR in our studies was the acquisition or enhancement of cancer stem cell-like properties including increased percentages of ALDHhigh cells, resistance to Cisplatin, and tumorsphere formation in low adherence conditions. First described in 2004 , OSCC stem cells have an enhanced ability to form tumors in vivo and to resist chemotherapeutic- or radiation-induced cell death [19, 18, 21, 22]. Cells isolated from patients on the basis of high ALDH expression exhibit an increased resistance to radiation therapy and increased tumorigenicity . ALDHhigh cells from OSCC lines exhibit increased invasiveness, quiescence and epithelial to mesenchymal transition (EMT) as assayed in part by migration in a scratch wound assay . In this context, the demonstration of a role for the AHR in the generation and maintenance of CSLCs takes on a particularly important connotation in that it suggests a treatment to reduce the number and/or activity of CSLCs, or to use in combination with traditional therapeutics to increase the sensitivity of CSLC to traditional chemotherapeutics regimens. It has been recently reported that chemoresistance is directly linked to metastatic behavior , and our findings support the importance of increasing chemo-sensitivity in oral cancer treatment to prevent deleterious downstream outcomes.
Perhaps of special note is the ability of P. aeruginosa and P. gingivalis supernatants to activate the AHR, increase the percentage of ALDHhigh cells and accelerate cell migration. Our studies with P. aeruginosa confirm previous work  by demonstrating that P. aeruginosa-derived pyocyanin is a potent AHR ligand and we extend previous studies by demonstrating that both P. aeruginosa supernatant and pyocyanin exhibit AHR-dependent biologic activity in malignant cells (e.g., induction of ALDH activity, increased cell migration). With regard to P. gingivalis, this is the first demonstration of an AHR ligand being produced by this common oral pathogen, a finding that has significant implications for the possible role for P. gingivalis in OSCC. In this vein, it has recently been shown that infection of OSCC with P. gingivalis induces EMT and increases the percentage of CSLCs in OSCC . In those studies, it was suggested that these outcomes were due in part to increased IL-8 expression, a cytokine regulated by the AHR in human macrophages [48, 49]. Current studies are underway to determine the nature of the AHR ligand produced by P. gingivalis and to determine if AHR-dependent IL-8 production leads to an increase in cells with stem cell-like properties.
We recently described a novel orthotopic xenograft model for human OSCC . Here, this model was used to further explore the role of the AHR in OSCC and to begin to assess if AHR inhibition represents a viable, targeted approach to OSCC therapy. Indeed, daily oral lavage with CB7993113, a non-toxic competitive AHR inhibitor , significantly reduced tumor load and weight loss and increased overall survival. The lack of weight loss observed in CB7993113-treated mice over the course of the experiment was consistent with the lack of toxicity seen with this potential therapeutic in vitro . Of note, studies from other laboratories demonstrate that AHR down-regulation blocks B(a)P-induced OSCC development by blocking DNA adduct formation, likely through inhibition of AHR-dependent CYP1A1 and CYP1B1 induction [50, 51]. Therefore, CB7993113 could fairly be viewed as a potential inhibitor of tumor initiation as well as a potential therapeutic.
Collectively, the data presented here strongly suggest that OSCC is characterized by relatively high AHR expression and constitutive activity. AHR inhibition in OSCC cell lines altered population dynamics specifically by reducing the percentage of ALHDhigh cells and the ability of CSLC to form tumorspheres. Relevant to environmental chemical exposures, including exposure to several AHR ligands in cigarette smoke, prototypic and environmental PAHs as well as a halogenated hydrocarbon up-regulated phenotypic and functional markers of cancer stem cells and increased cell migration, all of which increases concern over the possible contribution of environmental chemical exposures to OSCC progression. Furthermore, the demonstration that P. gingivalis produces an AHR ligand(s), suggests that this bacterium may play a role in oral cavity carcinogenesis in the absence of or perhaps in combination with smoke derived carcinogens. Finally, the results suggest an opportunity for targeted therapy for otherwise refractory OSCCs.
Financial Support: Boston University Oral Cancer Research Initiative, Art BeCAUSE, Avon Foundation for Women, Superfund (5 P42 ES007381-18 (Sherr))
The authors would like to thank Bonnie Campbell and Elizabeth Lindsay for their assistance as lab technicians in the Sherr Lab, and Connie Slocum and Mary Zabinski for their assistance in P. Gingivalis and P. Aeruginosa supernatant collection.
There are no actual, potential, or perceived conflict of interest with regard to the manuscript submitted for review from any of the authors listed above.