Since the discovery of CHD1L in 2008, it has emerged as an oncogene implicated in the pathology and poor prognosis of a variety of cancers, including gastrointestinal cancers. However, a mechanistic understanding of CHD1L as a driver of colorectal cancer has been limited. Until now, there have been no reported inhibitors of CHD1L, also limiting its development as a molecular target. We sought to characterize the clinicopathologic link between CHD1L and colorectal cancer, determine the mechanism(s) by which CHD1L drives malignant colorectal cancer, and discover the first inhibitors with potential for novel treatments for colorectal cancer. The clinicopathologic characteristics associated with CHD1L expression were evaluated using microarray data from 585 patients with colorectal cancer. Further analysis of microarray data indicated that CHD1L may function through the Wnt/TCF pathway. Thus, we conducted knockdown and overexpression studies with CHD1L to determine its role in Wnt/TCF-driven epithelial-to-mesenchymal transition (EMT). We performed high-throughput screening (HTS) to identify the first CHD1L inhibitors. The mechanism of action, antitumor efficacy, and drug-like properties of lead CHD1L inhibitors were determined using biochemical assays, cell models, tumor organoids, patient-derived tumor organoids, and in vivo pharmacokinetics and pharmacodynamics. Lead CHD1L inhibitors display potent in vitro antitumor activity by reversing TCF-driven EMT. The best lead CHD1L inhibitor possesses drug-like properties in pharmacokinetic/pharmacodynamic mouse models. This work validates CHD1L as a druggable target and establishes a novel therapeutic strategy for the treatment of colorectal cancer.
The integrity of the genome is maintained by conformational changes to chromatin structure that regulate accessibility to DNA for gene expression and replication. Chromatin structure is maintained by posttranslational modifications of histones and rearrangement of nucleosomes (1–3). ATP-dependent chromatin remodelers are enzymes that alter chromatin by changing histone composition, and evicting or translocating nucleosomes along DNA. Their activity plays a critical role in cellular function by regulating gene expression and the accessibility of DNA for replication, transcription, and DNA repair (4, 5). Dysregulation of chromatin remodeling is associated with human disease, particularly cancer (6, 7). In the last decade, the chromatin remodeler known as chromodomain helicase/ATPase DNA binding protein 1–like (CHD1L), also known as amplified in liver cancer 1 (ALC1), has emerged as an oncogene implicated in the pathology of prominent human cancers (8, 9). CHD1L is also involved in multidrug resistance, ranging from upregulation of drug resistance efflux pumps (e.g., ABCB1; ref. 10) to PARP1-mediated DNA repair (11, 12) and antiapoptotic activity (13, 14). Moreover, amplification or overexpression of CHD1L are correlated with poor prognoses for patients, including low overall survival (OS) and metastatic disease (15–17). Therefore, the multifunctional oncogenic mechanisms of CHD1L make it an attractive therapeutic target in cancer.
While the cancer-driving mechanisms of CHD1L have been studied in liver (13, 14), breast (18), and lung (10) cancer, little is known about the pathologic mechanisms associated with CHD1L in colorectal cancer. A majority of patients with colorectal cancer possess mutations in the Wnt signaling pathway, leading to aberrant T-cell factor/lymphoid enhancer factor transcription, denoted henceforth as TCF transcription or TCF complex (19, 20). The TCF complex is orchestrated by TCF4 (a.k.a. TCFL2), which is activated through interactions with an array of coactivators such as β-catenin, PARP1, and CREB Binding protein (CBP; ref. 21). Recently, TCF4 was shown to be a specific driver of both early metastasis from adenomas (i.e., polyps) and from late-stage metastatic patients (mCRC; refs. 16, 17). Moreover, we and others have shown that TCF transcription functions as a master regulator of epithelial-to-mesenchymal transition (EMT; refs. 22–24), a process that can transform relatively benign epithelial tumor cells into mesenchymal cells with increased cancer stem cell (CSC) stemness and other malignant properties that drive metastatic colorectal cancer (25). Currently, no drug has been clinically approved that target the Wnt/TCF pathway (26). To this end, we have evaluated the clinicopathologic characteristics of CHD1L in colorectal cancer, which led us to hypothesize that CHD1L is a druggable target involved in TCF transcription.
In this study, we propose a mechanism for CHD1L-mediated TCF transcription. Utilizing high-throughput screening (HTS), we have identified the first small-molecule inhibitors of CHD1L. We show that lead inhibitors are able to prevent TCF transcription, reverse EMT, and other malignant properties in a variety of cell models including tumor organoids and patient-derived tumor organoids (PDTO). The top lead CHD1L inhibitor displays drug-like pharmacologic properties, including in vivo pharmacokinetic and pharmacodynamic profiles, important for translational development toward the treatment of colorectal cancer and other cancers.
Materials and Methods
Clinicopathologic characterization of CHD1L
Transcriptome expression data of 585 patients with colorectal cancer from the CIT cohort (GEO: GSE39582) were used for in silico validation (GSE39582; ref. 27). Gene expression analyses were performed by the Affymetrix Human Genome U133 Plus 2.0 Array. RMA was used for data preprocessing and COMBAT for batch correction. Signal intensity was log2 normalized. The CHD1L cutoff for colorectal cancer risk stratification based on disease-specific survival was determined by the receiver operating characteristic (ROC) curve. Cutoff for CHD1L expression was set to 6.45. Differences in OS were estimated by the Kaplan–Meier method and compared using the log-rank test. For the comparison of categorical variables, we used the Fisher exact test. The Mann–Whitney U test was used for two groups of continuous variables and in case of more than two groups data has been analyzed by the Kruskal–Wallis test. For all two-sided P values, the unadjusted significance level of 0.05 was applied.
The CHD1L cutoff and clinicopathologic characteristics have been evaluated by multiple Cox regression analysis. Only variables that were significant in univariate analyses had been integrated in the Cox regression model using the Wald forward algorithm for significance determination. All variables including more than two groups had been categorized and the stepwise entry criterion for covariates was P < 0.05 and the removal criterion was P > 0.1. Statistical analysis had been performed using SPSS (IBM), GraphPad Prism, JMP (SAS), and R Studio.
University of Colorado Cancer Center patient sample RNA-sequencing analysis
RNA-sequencing (RNA-seq) data were from colorectal cancer patient tumor xenograft explants were obtained from the University of Colorado Cancer Center gastrointestinal (UCCC GI) tumor tissue bank, and analyzed as described previously (28). Briefly, gene expression was log2 normalized and measured by FPKM (Fragments Per Kilobase of transcript per Million mapped reads). The Wnt signaling pathway defined by the Kyoto Encyclopedia of Genes and Genomes (KEGG) was used as the gene set in this study. Samples with expression of CHD1L <1 FPKM were considered low expression and were removed from this study. Genes with significant Spearman correlations (P < 0.05) were displayed as heatmap using matrix2png (gene-wise Z-normalized).
Cell lines were purchased directly from ATCC and used as indicated. Engineered cell lines previously reported were short tandem repeat profiled for authenticity. All cell lines were tested for bacterial and Mycoplasma contamination before use. Deidentified patient sample cells were obtained from the UCCC GI tissue bank, which are maintained, cataloged, and annotated by the tissue bank.
CHD1L overexpression and shRNA knockdown
Full-length CHD1L was synthesized in a pGEX-6P-1 plasmid (GenScript). The CHD1L sequence flanked by EcoRI and NotI was digested out and ligated to a lentiviral backbone to create pCDH1-CMV-CHD1L-EF1-puro plasmid for overexpression of CHD1L in human colorectal cancer cells. Mission shRNA (scrambled) and TRCN0000013469 and TRCN0000013470 (sh69 and sh70) specific for CHD1L were purchased from Sigma-Aldrich. Virus was produced in HEK293T cells using TransIT-293 reagent (Mirus), and plasmids pHRdelta8.9 and pVSV-G. Colorectal cancer cells were transduced with overexpression or shRNA knockdown virus and selected with 2 μg/mL puromycin for 7 days.
Western blot analysis
Colorectal cancer cell lines and homogenized tumor tissue samples from mice were resuspended in RIPA lysis buffer [20 mmol/L Tris-HCl (pH 7.5), 150 mmol/L NaCl, 1 mmol/L Na2 EDTA, 1 mmol/L EGTA, 1% NP-40, 1% sodium deoxycholate, 2.5 mmol/L sodium pyrophosphate, 1 mmol/L β-glycerophosphate, 1 mmol/L Na3VO4, 0.1 mmol/L PMSF]. Protein concentration was determined using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). Forty micrograms of sample were run on 10% Bis-Tris gels. Following electrophoresis, the proteins were transferred to a nitrocellulose membrane. The membranes were blocked at room temperature with 5% nonfat milk in TBS/tween 20 (TBST) for 1 hour at room temperature. Membranes were washed three times with TBST. Blots were incubated with the appropriate primary antibody in 5% nonfat milk in TBST overnight at 4°C. Membranes were washed three times with TBST and then incubated with appropriate secondary antibody for 1 hour. Membranes were washed again with TBST three times. Blots were exposed using SuperSignal West Pico PLUS Chemiluminescent Substrate (Thermo Fisher Scientific) and imaged using a Bio-Rad ChemiDoc Imaging System.
TOPflash TCF transcriptional reporter assay
TOPflash (Millipore) was used to evaluate TCF transcriptional activity in colorectal cancer cells. A total of 20,000 cells per well were plated into 96-well white plates and transfected with TransLT1 reagent (Mirus). Cells were incubated with transfection mix for 24 hours. Next, cells were washed with PBS and a 1:1 ratio of PBS: One-Glo luciferase reagent Promega was added and the luminescence was detected within 10 minutes. A duplicate experiment was conducted to measure cell viability using CellTiter-Glo (Promega), which was used to normalize TOPflash luminescence to obtain the fold change in TCF activity. Experiments were replicated two times (n = 3 for each experiment). A duplicate plate was generated to measure the total protein content with BCA assay at the endpoint. One-Glo signal (TCF activity) was normalized to the protein amount first before the data of CHD1L knockdown or overexpression groups were normalized to the scr or EV controls, respectively.
Nuclear cell lysates were generated from untreated SW620 cells. For the input control, 100 μL of 1 mg/mL nuclear extract was saved and used as the input. Immunoprecipitation (IP) was conducted with Dynabeads Protein-A IP Kit (Thermo Fisher Scientific). Briefly, 300 μg of lysate incubated with 2 μg of the anti-TCF4 and anti-CHD1L IP antibody, anti-rabbit IgG, and anti-mouse IgG were used as nonspecific binding controls and were rotated at 4°C for 2 hours. After preincubation, 50 μL of beads were transferred to the preincubated antibody/lysate mixture followed by overnight incubation at 4°C. The flow through was collected and the beads were washed three times with PBST. Proteins were eluted with 20 μL of 50 mmol/L glycine (pH = 2.8) at 70°C for 10 minutes.
Using detailed methods described previously (23), cells were cross-linked with 1.42% formaldehyde for 15 minutes and quenching with 125 mmol/L glycine for 5 minutes. Cells were lysed with Szak's RIPA buffer and sonicated. The IP steps were conducted at 4°C as follows: 50 μL of protein A/G agarose beads were prewashed with cold Szak RIPA buffer and incubated with 1 mg of lysate for 2 hours. Salmon sperm DNA (0.3 mg/mL) was added and incubated for 2 hours. Lysate (100 μL) was set aside as the input control. Anti-CHD1L (2 μg) was added to the remainder and incubated overnight. Beads were washed and the supernatant was aspirated to 100 μL followed by the addition of 200 μL of 1.5×-Talianidis elution buffer. To elute immunocomplexes and reverse crosslink, 12 μL of 5 mol/L NaCl was added and the mixture was incubated at 65°C for 16 hours. The supernatant was mixed with 20 μg of proteinase K and incubated for 30 minutes at 37°C. DNA was extracted with phenol/chloroform and precipitated with ethanol. The IP product was amplified with PowerUp SYBR Green Master Mix (Applied Biosystems) using known published primers (23).
Colony formation was assessed after CHD1L knockdown in SW620 cells or overexpression in DLD1 cells as described previously using the well-established crystal violet or the area methods (23, 24). Cells were plated at 1,000 cells/well in 6-well plates and medium was changed two times per week over a 10-day time course. To assess CHD1L inhibitors for their ability to suppress CSC stemness, HCT116- or CHD1L-overexpressing DLD1 cell lines were pretreated in monolayer cultures for 24 hours with vehicle control (0.5% DMSO) or CHD1L inhibitors at the concentrations indicated. Pretreated viable cells were plated at 1,000 cells/well in 6-well plates or 200 cells/well in a 24-well plates. For the crystal violet method (23), colonies were analyzed as reported previously. For the area method (24), colonies were analyzed using the Incucyte S3 2018A software with the following parameters modified from default: (i) for HCT116 cells segmentation adjustment = 0.6; Min area (μm2) = 3 × 104; Max area (μm2) = 1.6 × 106; Max eccentricity = 0.9; (ii) for DLD1CHD1L-OE cells segmentation adjustment = 1; Min area (μm2) = 1 × 104; Max area was not constrained; Max eccentricity = 0.95. Experiments were replicated two times (n = 2 for each experiment).
Tumor organoid culture
As described previously (23, 24), cell lines were cultured as tumor organoids using phenol red–free RPMI1640 containing 5% FBS and by seeding 5,000 cells/well into uncoated 96-well U-bottom Ultra Low Attachment Microplates (PerkinElmer) followed by centrifugation for 15 minutes at 1,000 rpm to promote cells' aggregation. A final concentration of 2% Matrigel was added and tumor organoids were allowed to self-assemble over 72 hours under incubation (5% CO2, 37°C, humidity) before treatment, and maintained under standard cell culture conditions during treatment time courses.
VimPro-GFP and EcadPro-RFP reporter 3D high-content imaging assays
Stable VimPro-GFP or EcadPro-RFP SW620 reporter cells were generated using pCDH-VimPro-GFP-EF1-puro virus or pCDH-EcadPro-mCherry-EF1-puro virus as reported previously (23, 24). The stable fluorescently labeled reporter cells were used to generate tumor organoids as described herein. Tumor organoids were treated with CHD1L inhibitors at 10 μmol/L for an additional 72 hours. Following treatment, tumor organoids were stained with 16 μmol/L of Hoechst 33342 for 1 hour (nuclei stain). Images were taken with a 5× air objective. Z-stacks were set at 26.5 μm apart for a total of 15 optical slices. Imaging and high-content analysis were performed using an Opera Phenix and Harmony software (PerkinElmer). Nuclei were identified within each layer and cells were found with either GFP or mCherry channel. The fluorescence intensities of each channel were calculated and thresholds were set based on the background intensities. Percentages of GFP or mCherry RFP–positive cells were calculated and normalized to the DMSO-treated group.
Tumor organoid cytotoxicity
SW620 tumor organoids were cultured as described herein. CellTox Green cytotoxicity assay solution was prepared per manufacturer's protocol (Promega). Briefly, tumor organoids were treated for 72 hours with CellTox Green reagent (0.5×) and various doses of CHD1L inhibitors over a range of 0–100 μmol/L. Organoids were imaged using the Opera Phenix with excitation at 488 nm and emission at 500–550 nm. Mean intensity of the whole well was utilized for calculating cytotoxicity with Lysis Buffer (Promega) as 100% cytotoxicity control and 0.5% DMSO as 0% cytotoxicity control. Intensity values were normalized to these controls using GraphPad Prism.
HCT116 cells were plated at 60,000 cells/well into an ImageLock 96-well plate (Sartorius) and allowed to attach overnight. A wound was created in all wells using the WoundMaker then washed two times with PBS. The plate was brought to 4°C using a Corning XT Cool Core to avoid polymerization of Matrigel during the preparation of the invasion conditions (Corning). Wells were coated with 50 μL of 50% Matrigel in RPMI1640 media. Plates were centrifuged at 150 rpm at 4°C for 3 minutes, using a swing bucket rotor to ensure even Matrigel coating with no air bubbles. Afterwards, plates were placed on a CoolSink XT prewarmed inside a cell culture incubator (5% CO2, 37°C, humidity) for 10 minutes to evenly polymerize the Matrigel, followed by the addition of CHD1L inhibitors dissolved in 50 μL of RPMI1640 media containing 5% FBS. Finally, the plate was placed in an Incucyte S3 live cell imager (Sartorius) for 48 hours. The wound was imaged every hour using the phase contrast channel and 10× objective in wide mode.
Cloning and purification of recombinant CHD1L
Cat-CHD1L (residues 16-61) and fl-CHD1L (residues 16-879) constructs were a generous gift from Helena Berglund at the Department of Medical Biochemistry and Biophysics, Karolinska Institute (Stockholm, Sweden). Proteins were expressed in Rosetta 2 (DE3) pLysS cells (Promega) in Terrific Broth (Thermo Fisher Scientific). Cultures were induced with 0.2 mmol/L IPTG at OD600 = 2.0 at 18°C for 16 hours. Cells were harvested and resuspended in buffer-A, containing 20 mmol/L HEPES, pH 7.5, 500 mmol/L NaCl, 50 mmol/L KCl, 20 mmol/L imidazole, 10 mmol/L MgCl2, 1 mmol/L TCEP, 10% glycerol, and 500 μmol/L phenylmethylsulfonylfluoride. Cells were lysed by sonication and cellular debris was removed by centrifugation. The supernatant was loaded onto a NiNTA resin column (Qiagen). Protein bound to the column was washed with 1× with buffer-A, 1× with buffer-A containing 10 mmol/L ATP, and an additional time with buffer-A. Proteins were eluted using buffer-B (buffer-A with 500 mmol/L imidazole) with a gradient from 20 to 500 mmol/L imidazole. Following affinity purification, Cat-CHD1L was dialyzed overnight into 50 mmol/L Tris, pH 7.5, 200 mmol/L NaCl, and 1 mmol/L DTT. Similarly, fl-CHD1L was dialyzed overnight into 20 mmol/L MES, pH 6.0, 300 mmol/L NaCl, 10% glycerol, and 1 mmol/L DTT. Protein was then purified by ion-exchange chromatography, cat-CHD1L was bound to a Q-sepharose column (GE Healthcare) and fl-CHD1L was bound to a S-sepharose column (GE Healthcare), proteins were eluted using a NaCl gradient of 0.2–1 mol/L for cat-CHD1L and 0.3–1 mol/L fl-CHD1L. Pure fractions were pooled, concentrated, and further purified by size-exclusion chromatography using an Superdex 200 column (GE Healthcare) in 20 mmol/L HEPES, pH 7.5, 100 mmol/L NaCl, 1 mmol/L TCEP, 10% Glycerol. Protein purifications were conducted using an ACTA Start FPLC (GE Healthcare).
CHD1L ATPase assay
All reactions were carried out using low volume nonbinding surface 384-well plates (Corning Inc.). 100 nmol/L cat-CHD1L or fl-CHD1L and 200 nmol/L c-Myc DNA or mononucelosome (Active Motif) were added to a buffer containing 50 mmol/L Tris pH 7.5, 50 mmol/L NaCl, 1 mmol/L DTT, 5% glycerol, and the reaction was initiated by the addition of 10 μmol/L ATP (New England Biolabs) to a total volume of 10 μL and incubated at 37°C for 1 hour. ATPase activity was measured by adding 500 nmol/L of Phosphate Sensor (Life Technologies) measuring excitation (430 nm) and emission (450 nm) immediately on an Envision plate reader (PerkinElmer). An inorganic phosphate standard curve was used to convert the fluorescence to [Pi], and enzyme kinetics were determined using GraphPad Prism.
HTS drug discovery for inhibitors of CHD1L
Assay composition was the same as described above using cat-CHD1L, except that the reaction mixture volume was modified to accommodate addition of drug or DMSO. Using a Janus liquid handler (PerkinElmer), compounds dissolved in 100% DMSO were mixed with 50 mmol/L Tris pH 7.5, 50 mmol/L NaCl, 1 mmol/L DTT, 5% glycerol buffer to 200 μmol/L in 10% DMSO. Next, 1 μL of each compound was added to the enzyme mixture to give a final concentration of 20 μmol/L. The negative control used was 1% DMSO and 10 mmol/L EDTA was used as a positive control. Reactions were initiated with the addition of 10 μmol/L ATP and incubated at 37°C for 1 hour. ATPase activity was measured by fluorescence by adding 500 nmol/L Phosphate Sensor. Cat-CHD1L was screened against a 20,000-compound diversity set from Life Chemicals and a Kinase inhibitor library from Selleck Chemicals. Both libraries were prescreened before purchase to remove Pan-assay interference compounds (PAINS).
Patient-derived tumor organoid culture and viability assay
Colorectal cancer patient tumor tissues were obtained from the UCCC GI tissue bank and expanded following established protocols (29). Briefly, cells were seeded at 5,000 cells per well in 96-well plates and cultured by established methods (30) allowing PDTO formation over 72 hours. PDTOs were treated with DMSO (0.5%) or compound 6 with various concentrations for an additional 72 hours to obtain a dose response. PDTO cell viability was measured using CellTiter Blue (Promega). Eighty microliters of media were aspirated from wells and 80 μL of CellTiter Blue was added and incubated for 4 hours and cell viability was measured by fluorescence intensity using 560-nm excitation and 590-nm emission.
Evaluation of apoptosis
SW620 cells were plated at 30,000 cells/well in 96-well plates. Cells were treated with DMSO (negative control), SN-38 (apoptosis positive control), and compound 6 at concentrations indicated for 12 hours. Cells were then rinsed two times with cold PBS, 1× with cold Annexin-V staining buffer (10 mmol/L HEPES, pH 7.4, 140 mmol/L NaCl, 2.5 mmol/L CaCl2) and then incubated with Annexin-V FITC at 1:100 for 30 minutes in the dark. Cells were then rinsed two times with Annexin-V staining buffer and FITC intensity was measured using an Envision plate reader (PerkinElmer).
Evaluation of DNA damage by γH2AX
DLD1CHD1L-OE cells were seeded into a 96-well PerkinElmer Cell Carrier plate and allowed to adhere overnight. Cells were then treated with the appropriate compound at 10 μmol/L (0.5% DMSO) or with CHD1L inhibitor in combination with SN-38 (1 μmol/L), oxaliplatin (10 μmol/L), and etoposide (10 μmol/L). Cells were treated for 6 hours. Media were aspirated and cells were washed with cold PBS. Cells were then fixed with 3% paraformaldehyde for 15 minutes at room temperature and the fixed cells were washed with PBS three times. Cells were blocked for 1 hour at room temperature in 5% BSA and 0.3% Triton X-100 in PBS. Cells were then immunostained with phospho-(S139)-γH2AX rabbit mAb using a 1:800 dilution in 1% BSA, 0.3% Triton X-100 in PBS at 4°C overnight. Primary antibody was aspirated and cells were washed with PBS. Cells were incubated for 2 hours at room temperature with goat anti-rabbit Alexa Fluor Plus 647 fluorescent secondary antibody at a concentration of 5 μg/mL in 1% BSA, 0.3% Triton X-100 in PBS. Cells were then washed with PBS, Hoechst 33342 stain was diluted to a concentration of 1:1,000 in PBS, and added to cells for 15 minutes at room temperature. Cells were then imaged using a 20× water objective on the PerkinElmer Phenix HCS imaging system. Synergy was determined using the coefficient of drug interaction (CDI) equation. CDI = (A+B)/(AB). Synergy was determined with a CDI < 0.8, additivity was 0.8–1.2, and antagonism was defined by a CDI > 1.2.
Aqueous solubility and CLogP
Using a recently reported detailed method (24), aqueous solubility was tested for lead compound 6. The PBS UV absorption spectra were compared with a fully saturated solution in 1-propanol and the solubility of compound 6 in 10% DMSO in PBS (pH 7.4) was determined using linear regression analysis. The solubility in PBS was conducted in duplicate experiments. The consensus LogP (CLogP) values were obtained using the SwissADME web tools (31).
Microsome stability studies
The microsomal stability of compound 6 was determined using female CD-1 mouse microsomes (M1500) purchased from Sekisui XenoTech, following our recently reported method (24). Samples were centrifuged at 20,000 × g for 10 minutes and the supernatant was transferred to an autosampler vial for LC-MS analysis. The following mass transition (m/z, amu) was monitored for compound 6 (molecular weight = 393.5).
In vivo pharmacology
All animal studies were conducted in accordance with the animal protocol procedures approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Colorado Anschutz Medical Campus (Aurora, CO) and Colorado State University (Fort Collins, CO).
Nine-week-old female CD-1 mice, purchased from Charles River, were used for pharmacokinetic studies using methods we recently reported (24). Briefly, the pharmacokinetic studies were designed to cover a range of 0.25 to 24 hours with 3 mice/time point for a total of 21 mice/compound 6. Each mouse was dosed with a single intraperitoneal injection of compound 6 at 50 mg/kg prepared in 100% DMSO. Whole blood was harvested at specific time points and the separated plasma frozen at −80°C for storage or used for LC/MS-MS analysis.
Pharmacodynamics and liver toxicity
Two million VimPro-GFP SW620 cells suspended in 100 μL of a 1:1 mixture of Matrigel and RPMI1640 were injected into the flanks of 9-week-old female athymic nude mice [Nude-Foxn1nu (069); Envigo]. Growth was monitored with caliper measurements three times per week. At four weeks, mice were randomized into 2 groups and treated with 50 mg/kg of compound 6 in 200 μL of vehicle (10% DMSO, 40% PEG 400, 50% PBS pH = 7.4) or with vehicle control. Treatments were administered intraperitoneally once daily over five days. Mice were sacrificed 2 hours after the final dose on day five of the treatment. Tumors and livers were collected for analysis of compound 6 accumulation measured by LC/MS, Western blot analysis measuring effects on EMT and apoptosis, and liver toxicity.
Data were subjected to unpaired two-tailed Student t test with Welch correction statistical analysis or as otherwise stated using Prism (GraphPad). All experiments were replicated three times (n = 3) or as described in the methods.
Clinicopathologic characterization of CHD1L in patients with colorectal cancer
CHD1L expression is correlated with poor prognosis in several cancers but limited information about the pathology of CHD1L in CRC is known. As a result, we sought to expand the pathogenic characterization and mechanisms of pathology for CHD1L in patients with colorectal cancer. The clinicopathologic characteristics of 585 patients with colorectal cancer were analyzed from the Cartes d'Identite des Tumeurs (CIT) program with respect to CHD1L expression (GEO: GSE39582) and are summarized (Supplementary Table S1; ref. 27).
Follow-up information was available for all patients in the CIT cohort over a period of 15 years. For the entire patient cohort, high CHD1L expression is associated with lower OS (P = 0.0167) and median survival (MS) of 8.8 years for high CHD1L patients. Median survival was not reached in the low CHD1L cohort as 72% (115/159) of patients were censored and 26% (42/159) were deceased (Fig. 1A). We evaluated patient data using the TNM staging system. As stage I and IV patients have a high likelihood of survival or death, respectively, we evaluated survival of patients with stage II and III colorectal cancer. High CHD1L expression was associated with a lower OS (P = 0.0191) and MS of 11 years for stage II and III colorectal cancer, again median survival was not reached in the low CHD1L cohort (Fig. 1B). Survival was also analyzed with respect to CHD1L expression for each stage of colorectal cancer. Stage II patients showed a significant difference in survival (P = 00319) with a MS of 11 years, no significant difference was observed for stage I, III, or IV patients (Supplementary Fig. S1). Welch one-way ANOVA analysis of CHD1L expression indicates a significant increase in CHD1L levels from cancer stages I through IV (Fig. 1C). We evaluated patients with stage I and II versus patients with stage III and IV colorectal cancer and observed a significant increase in CHD1L expression in the stage III and IV versus early-stage cohorts (P = 0.0051; Supplementary Fig. S2A). Analysis of CHD1L expression with respect to lymph node metastasis suggests that CHD1L is overexpressed in patients with increased regional lymph node metastasis (N1 P = 0.0128, N2 P = 0.05 compared with N0; Fig. 1D). Although the trend of CHD1L expression was the same for the N3 cohort, no significance was determined due to the limited number of patient samples available. We found no significant difference in CHD1L expression with respect to tumor size, metastasis, or location (Supplementary Fig. S2B–S2D).
Evaluation of CHD1L in colorectal cancer molecular subtypes
The association of CHD1L expression with six molecular subtypes of colorectal cancer (27): C1 (immune system down, n = 116), C2 (deficient mismatch repair, n = 104), C3 (KRAS mutant, n = 75), C4 (CSC, n = 59), C5 (activated WNT pathway, n = 152), and C6 (chromosomal instability normal, n = 60) was investigated (Fig. 1E). There is a significant difference of CHD1L expression among the six molecular subtypes (P < 0.001). CHD1L expression was high in C5, C4, and C3 and low in C2 and C6. The C2 subtype is associated with a decrease in the WNT signaling pathway and deficient for mismatch repair. The C4 and C6 subtypes are associated with poorer relapse-free survival compared with other subtypes. The C4 subtype is associated with increased CSC stemness and the C5 subtype is associated with activated WNT signaling and deregulated EMT pathways. The lower CHD1L expression in the C2 (deficient mismatch repair) subtype is consistent with its known function in DNA damage response (32). In addition, CHD1L expression was lower in patients with deficient mismatch repair than patients without (P < 0.001; Fig. 1F). CHD1L expression was also higher in patients with KRAS mutations (P = 0.049). The expression of CHD1L in the C3, C4, and C5 molecular subtypes prompted us to further investigate the function of CHD1L expression in EMT, CSC stemness, and the WNT/TCF pathway.
CHD1L expression correlates with Wnt/TCF–associated genes
Utilizing a smaller cohort of patients with colorectal cancer (n = 26) from the UCCC GI tumor tissue bank, a similar trend was observed as with the larger CIT cohort where CHD1L expression significantly correlated with late stage and metastatic colorectal cancer compared with early stage and primary colorectal cancer (Supplementary Fig. S3). When analyzing CHD1L expression with genes involved in the KEGG WNT pathway, using Spearman correlation we observed a significant positive correlation with 65 of 125 genes. Among these were well-established genes involved in TCF-transcription such as topoisomerase IIα (TOP2A; r = 0.65, P = 0.004; refs. 23, 24), and TCF4 (r = 0.61, P = 0.0012; Supplementary Fig. S4). We also observed a significant positive correlation between known CSC markers CD44 (r = 0.43, P = 0.038), LGR5 (r = 0.55, P = 0.0075), and CHD1L. When comparing the CIT cohort to the UCCC cohort a significant correlation was observed for TOP2A (r = 0.1275, P = 0.0020) and TCF4 (r = 0.1050, P = 0.011). Consistent with this result, we have shown that TOP2A is a required component of the TCF complex, promoting EMT in colorectal cancer (23, 24). Hence, CHD1L may be involved in TCF transcription and EMT in patients with colorectal cancer.
CHD1L mediates TCF transcription in colorectal cancer
On the basis of the correlation of CHD1L with TCF complex members, we hypothesized that CHD1L has a mechanistic role in TCF transcription. To test this, we utilized SW620 and DLD1 cell lines, which have high and low endogenous CHD1L expression, respectively (Supplementary Fig. S5). We used shRNA to knockdown CHD1L in SW620 cells (SW620CHD1L-KD) and overexpressed CHD1L in DLD1 cells (DLD1CHD1L-OE). Using the TOPflash luciferase reporter (33) transfected into SW620CHD1L-KD or DLD1CHD1L-OE, we determined that overexpression of CHD1L produced a significant increase in TCF transcription (P < 0.0001; Fig. 2A). Conversely, SW620CHD1L-KD cells displayed a significant decrease in TCF transcription (P = 0.0006). These results establish CHD1L as a potential factor directly involved in TCF transcription.
CHD1L directly interacts with the TCF transcription complex
Activation of TCF transcription is a dynamic process that involves the shedding of corepressor proteins, binding of coactivator proteins, and remodeling of the chromatin landscape (1, 21). We performed coimmunoprecipitation (co-IP) studies with TCF4, demonstrating that CHD1L directly binds to the TCF complex (Fig. 2B). CHD1L has been well characterized as a binding partner with PARP1 in DNA damage response (11, 32). PARP1 is also a component of the TCF complex binding to TCF4 and β-catenin (34). Our results demonstrate that CHD1L binds to the TCF complex, which is likely through interactions between TCF4 and PARP1.
To further characterize CHD1L as a component of the TCF complex, we performed chromatin immunoprecipitation (ChIP) of CHD1L to TCF complex WNT response elements (WRE) in SW620 cells (Fig. 2C). CHD1L was enriched at c-Myc, vimentin, slug, LEF1, and N-cadherin WREs, further supporting that CHD1L is functioning directly with the TCF complex. Taken together, the data implicate CHD1L as a critical component of TCF transcription.
CHD1L-mediated TCF transcription promotes EMT and CSC stemness in colorectal cancer
Previously, we have characterized TCF transcription as a master regulator of EMT in colorectal cancer (23). In addition, CHD1L localizes at WREs of EMT effector genes (Fig. 2C). Therefore, we evaluated whether knockdown or overexpression of CHD1L modulates EMT by measuring biomarker expression in SW620CHD1L-KD and DLD1CHD1L-OE cells. Knockdown of CHD1L induced reversion of EMT, decreasing vimentin and slug while increasing E-cadherin expression (Fig. 2D). Conversely, EMT was induced in DLD1CHD1L-OE cells, evidenced by a decrease in E-cadherin and an increase in vimentin and slug expression (Fig. 2E). These results indicate that CHD1L is an EMT effector gene involved in promoting the mesenchymal phenotype in colorectal cancer. A hallmark of EMT is an increase in CSC stemness. To characterize the impact of CHD1L expression on stemness, we performed clonogenic colony formation assays (Fig. 2F; ref. 35). CSC stemness increased with DLD1CHD1L-OE (P = 0.0001) and decreased in SW620CHD1L-KD (P = 0.002) cells measured by colony formation.
HTS identifies the first small-molecule inhibitors of CHD1L
After establishing CHD1L as a driver of TCF-mediated EMT, we sought to identify small-molecule inhibitors of CHD1L and assess their therapeutic potential. Our drug discovery goal was to target CHD1L DNA translocation or interactions with DNA, which are dependent on its catalytic domain ATPase activity (Supplementary Figs. S6 and S7; refs. 36, 37). CHD1L belongs to the SNF2 (sucrose non-fermenter 2) ATPase superfamily of chromatin remodelers that contains a two-lobe ATPase domain. CHD1L also has a macro domain that is unique relative to other chromatin remodelers, which promotes an autoinhibited state through interactions between the macro and the ATPase domains (Supplementary Fig. S7A) (38, 39). However, the macro domain binds to PARP1, the major activator of CHD1L, alleviating autoinhibition (38, 39). Using the methodology by Lehmann and colleagues (38), we purified full-length CHD1L (fl-CHD1L) and the catalytic ATPase domain (cat-CHD1L; Supplementary Fig. S7B). The cat-CHD1L provides for a more robust ATPase assay compared with fl-CHD1L, which is consistent with the report from Lehman and colleagues (38) Therefore, to discover direct inhibitors of CHD1L ATPase we developed a HTS assay in the context of TCF transcription, including cat-CHD1L, c-Myc WRE DNA, ATP, and phosphate-binding protein that fluoresces upon binding inorganic phosphate (Pi). We validated this assay and performed pilot screening against clinically relevant kinase inhibitors (Supplementary Fig. S6). Moreover, the pilot screen found no hits, demonstrating that CHD1L is not a likely target for kinase inhibitors. Thus, we expect hits identified will have limited off target binding of kinases. Once validated, we conducted primary HTS using 20,000 compounds from the Life Chemicals Diversity Set, which were screened at 20 μmol/L in 1% DMSO with 10 mmol/L EDTA as a positive control (Supplementary Fig. S7B). The screen provided robust statistics with an average Z′-factor value of 0.57 ± 0.06 over 64 plates (Supplementary Fig. S7C). The average compound activity was 92.3% ± 17.8. As a result, we set the hit limit to be three SDs from the mean at 39% ATPase activity. This stringent hit limit identified 64 hits, of which 53 hits were confirmed against recombinant CHD1L ATPase activity (Supplementary Fig. S7D).
A subset of seven confirmed hits (compounds 1–7) represent a range of pharmacophores with greater than 50% inhibition against cat-CHD1L ATPase. After hit confirmation, compounds were purchased as new batches in solid form from Life Chemicals for lead validation studies (Supplementary Table S2). Compounds 1–7 were subjected to dose response studies against cat-CHD1L ATPase, which validated these hits as potent CHD1L inhibitors with activity between 900 nmol/L to 5 μmol/L (Fig. 3A). There are striking similarities in structure between the hits, with hit 1–3 and 6–7 being close derivatives, further validating the HTS results. Next, we tested compounds 1–7 in HCT116, SW620, and DLD1CHD1L-OE cells for their ability to inhibit TCF transcription using the TOPflash reporter system (Fig. 3B). Compounds 1–3 were eliminated displaying no significant activity in cells. Compound 4 was also deprioritized due to modest activity with no dose-dependent inhibition of TCF activity. However, compounds 5–7 demonstrate superior dose-dependent activity against TCF transcription in all three colorectal cancer cell lines, advancing as lead CHD1L inhibitors. Notably, we observed decreased inhibition of TCF transcription for 5–7 at the low 2 μmol/L dose in DLD1CHD1L-OE cells, which is evidence of cellular CHD1L target engagement.
CHD1L inhibitors reverse EMT and malignant properties in colorectal cancer
After validating hits 5–7 against CHD1L-mediated TCF transcription, we next evaluated their ability to reverse EMT and other malignant properties in colorectal cancer. E-cadherin and vimentin are putative biomarkers for the epithelial and mesenchymal phenotypes, respectively (40). Loss of E-cadherin and gain of vimentin are also clinical biomarkers of poor prognosis (41–45). Accordingly, we developed and validated lentiviral promoter-driven reporters for E-cadherin (pCDH1-EcadPro-RFP) and vimentin (pCDH1-VimPro-GFP), which faithfully reports E-cadherin and vimentin protein expression (23, 24). SW620 cells transduced with either EcadPro-RFP or VimPro-GFP were cultured as tumor organoids for 72 hours, reaching a diameter of 600 μm. Tumor organoids were treated with compounds 5–7 for an additional 72 hours to determine the effective concentration 50 percent (EC50) for modulating promoter activity. Changes in promoter expression was quantified using a 3D confocal image–based high-content analysis algorithm (Fig. 4A–C; refs. 23, 24). We observed that compounds 5–7 effectively downregulated vimentin promoter activity with EC50 values of 15.6 ± 1.7 μmol/L (5), 4.7 ± 1.5 μmol/L (6), and 12.8 ± 1.3 μmol/L (7). Conversely, E-cadherin promoter activity was upregulated with EC50 values of 11.9 ± 0.3 μmol/L (5), 11.4 ± 0.3 μmol/L (6), and 28 ± 0.003 μmol/L (7). These results are consistent with our hypothesis that small-molecule inhibitors of CHD1L reverse TCF-driven EMT in colorectal cancer. To confirm that CHD1L inhibitors reverse EMT, we looked at the protein expression of two additional putative biomarkers of EMT, slug (mesenchymal), and zona occludens-1 (ZO-1, epithelial); changes in slug and ZO-1 are considered major criteria for EMT (46). SW620 tumor organoids treated with CHD1L inhibitors downregulate slug and upregulate ZO-1, further indicating a reversion of EMT (Fig. 4D).
A hallmark of EMT is an increase in CSC stemness and cell invasion. Therefore, we tested the ability of compounds 5–7 to inhibit migration and invasion in HCT116 and DLD1CHD1L-OE cells. All three compounds demonstrate a significant inhibition of CSC stemness (Fig. 4E). However, compounds 5 and 6 display more potent dose-dependent inhibition. Interestingly, DLD1CHD1L-OE cells form two times more colonies than HCT116 cells, which have moderate CHD1L expression. This observation is consistent with CHD1L's oncogenic and tumorigenic properties. Next, using HCT116 cells with uniform scratch wounds imbedded in 50% Matrigel, we treated cells with CHD1L inhibitors and monitored invasion over 72 hours. Compounds 5–7 exhibited a dose-dependent inhibition of invasion (Fig. 4F; Supplementary Fig. S8), with compound 6 displaying the most potent activity.
Inhibition of CHD1L enhances efficacy of DNA-damaging drugs
CHD1L is known to function in PARP1-mediated DNA damage repair, which is a mechanism of drug resistance to DNA-damaging chemotherapy (10, 32, 39). For example, drug resistance to cisplatin in lung cancer was observed in cells overexpressing CHD1L, and the efficacy of cisplatin was restored after CHD1L knockdown (10). In addition, knockdown of CHD1L alone does not increase in DNA damage (32). To determine whether CHD1L inhibitors could increase the efficacy of DNA-damaging drugs against low CHD1L–expressing DLD1 cells transduced with empty vector (DLD1CHD1L-EV) and overexpressing DLD1CHD1L-OE cells, we evaluated compound 6 alone and in combination with SN-38 (the active pharmacophore of prodrug irinotecan), oxaliplatin, and etoposide. To assess DNA damage we measured the phosphorylation of H2AX (γH2AX) by immunofluorescence (Supplementary Fig. S9), which is used as a biomarker for DNA-damaging chemotherapy (32). Compound 6 alone showed no significant DNA damage when treating cells at 10 μmol/L and measuring γH2AX activity, which is consistent with previously reported CHD1L knockdown studies (32). However, combination treatments in DLD1CHD1L-OE cells with compound 6 synergized with etoposide (10 μmol/L) and SN-38 (1 μmol/L), significantly increasing DNA damage compared with etoposide and SN-38 alone. In DLD1CHD1L-EV cells only the combination of etoposide and compound 6 displayed significant synergy. Under the experimental conditions used, we observed no synergy with oxaliplatin. Nevertheless, SN-38 (i.e., irinotecan) combination therapy, known as FOLFIRI, is a standard-of-care chemotherapy in the treatment of colorectal cancer. Therefore, the enhanced DNA damage that occurs with compound 6 in combination with SN-38 supports our hypothesis that CHD1L inhibitors may increase the efficacy of colorectal cancer standard-of-care DNA-damaging chemotherapies.
CHD1L inhibitors reverse EMT prior to the induction of cell death
CHD1L has been reported to confer antiapoptotic activity by inhibiting activation of caspase-dependent apoptosis (13, 47). In addition, reversal or inhibition of EMT is known to restore apoptotic activity of cancer cells (48). To determine whether CHD1L inhibitors reverse EMT prior to induction of cell death, we monitored E-cadherin expression by EcadPro-RFP reporter activity, and cytotoxicity was measured using the CellTox Green assay. Cells were treated with CHD1L inhibitors for 72 hours and imaged every 2 hours. We observed a significant increase in E-cadherin expression prior to induction of cytotoxicity for compound 6 relative to DMSO (Fig. 5A).
We then evaluated whether CHD1L inhibitors are able to induce apoptosis in colorectal cancer, we performed Western blot from SW620 tumor organoids and observed that E-cadherin becomes cleaved after treatment with compounds 5 and 6 (Fig. 5B), which is a marker of apoptosis (49). Next, using the more potent CHD1L inhibitor 6, we measured increases in cleaved PARP1, cleaved caspase 8, and cleaved caspase 3 relative to DMSO control (Fig. 5C). These results suggest compound 6 induces extrinsic apoptosis that is consistent with E-cadherin–mediated apoptosis through death receptors (48). To further characterize the apoptotic activity of CHD1L inhibitors, we performed annexin-V staining in SW620 cells over 12 hours. Compound 6 induced significant apoptosis compared with DMSO and had similar activity to the positive control SN-38 (Fig. 5D).
CHD1L inhibitors are effective against patient-derived tumor organoids
The use of PDTOs in preclinical drug development has been established as predictive in vitro cell model for clinical efficacy (31). After establishing the ability of compound 6 to reverse EMT and induce apoptosis using cell line–based models, we evaluated the efficacy of compound 6 in PDTOs produced from patient sample CRC102 obtained from the UCCC GI tissue bank (Fig. 5E). Consistent with our results in colorectal cancer cell lines, compound 6 showed potent cytotoxicity in PDTOs with an EC50 of 11.6 ± 2 μmol/L. This result further validates compound 6 as a priority lead CHD1L inhibitor.
In vitro and in vivo pharmacokinetics, pharmacodynamics, and liver toxicity of lead CHD1L inhibitor 6
To assess the drug-like potential and properties of compound 6, we conducted in silico, in vitro, and in vivo pharmacokinetic studies assessing CLogP, aqueous solubility, stability in mouse liver microsomes, and pharmacokinetics in CD-1 mice (Fig. 6; Supplementary Fig. S10). Compound 6 has an excellent balance of lipophilicity (CLogP = 3.2) and aqueous solubility that is relatively stable to liver-metabolizing enzymes, and an excellent pharmacokinetic disposition when administered to CD-1 mice. Compound 6 reaches a high plasma drug concentration CMax (∼30,000 ng/mL) and AUC (∼80,000 ng/mL/h) with a relatively long half-life (t1/2λ) of 3 hours after intraperitoneal (i.p.) administration.
Next, we conducted a second acute in vivo experiment using a maximum tolerated dose of compound 6 (50 mg/kg) administered to athymic nude mice by intraperitoneally once daily over five days. The goals of this experiment were to (i) determine whether compound 6 causes acute toxicity to livers, (ii) accumulates in VimPro-GFP SW620 xenograft tumors, and (iii) to determine pharmacodynamic effects. Compound 6 accumulates in SW620 tumors at a concentration of 10,533 ± 5,579 ng/mL (n = 4). As expected, when comparing the ratio of compound 6 accumulation in tissue/plasma, we observed 2.7 times more accumulation in liver compared with tumor (Fig. 6B). However, there was no apparent liver toxicity resulting from compound 6 at the dose and schedule administered (Fig. 6C; Supplementary Table S2). Overall, there were no significant histologic differences between the livers of vehicle or compound 6–treated mice. The primary histologic change observed were minimal fibrosis and inflammation of the hepatic capsule in both vehicle and compound 6–treated animals. This change suggests a very low grade, subclinical peritonitis, and is consistent with being secondary to intraperitoneal drug administration. We also did not observe any changes in weight loss to animals treated with compound 6 during the course of study.
In accordance with accumulation of compound 6 in tumors, we measured pharmacodynamic effects on tumor tissue by Western blot analysis, indicating a significant downregulation of mesenchymal markers vimentin, vimentin reporter (VimPro-GFP), and slug (Fig. 6D and E). Although not statistically significant, we also observed upregulation of the epithelial marker ZO-1 and induction of cleaved caspase 3, the putative biomarker of apoptosis. Taken together, we observed the pharmacodynamic effects by compound 6 indicating the reversion of EMT and apoptosis in vivo that were consistent with in vitro cell-based antitumor activity of compound 6. In summary, compound 6 displays good pharmacokinetic drug-like properties and the ability to alter EMT and induce apoptosis in vivo with no observed liver toxicity.
CHD1L is amplified (Chr1q21) and overexpressed in many types of cancer (8, 9). CHD1L overexpression has been characterized as a marker for poor prognosis and metastasis in numerous cancers (8, 9, 16, 17). While the collective literature demonstrating CHD1L as an oncogene and driver of malignant cancer is compelling, we set out to test the rigor of the prior research and our hypothesis that CHD1L is an oncogene with potential as a molecular target in colorectal cancer. We conducted in silico analyses of transcriptome data from a large cohort of 585 patients with colorectal cancer obtained over 15 years (27). CHD1L expression correlates with poor survival, with low CHD1L patients living significantly longer than high CHD1L patients (Fig. 1). Using the same cohort, Marisa and colleagues identified six distinct subtypes for improved clinical stratification of colorectal cancer and CHD1L is universally expressed in all subtypes, indicating its potential as a therapeutic target for colorectal cancer. CHD1L also correlated with tumor node metastasis, with increased expression moving from N0 (no regional spread) to N3 (distant regional spread). We then analyzed transcriptome data from a UCCC patient cohort and found that CHD1L expression significantly correlated with stage IV and metastatic colorectal cancer. Literature reports and our own analyses and data demonstrate that CHD1L is an oncogene promoting malignant colorectal cancer and its high expression correlates with poor prognosis and survival of patients with colorectal cancer.
In this study, we have determined a new biological function for CHD1L as a DNA binding factor for the TCF transcription complex required for promoting TCF-driven EMT and other malignant properties (Fig. 2). Using HTS drug discovery, we have identified the first known inhibitors of CHD1L (Fig. 3), and have characterized several lead compounds that display good pharmacologic efficacy in cell-based models of colorectal cancer, including PDTOs. CHD1L inhibitors effectively prevent CHD1L-mediated TCF transcription, leading to the reversion of EMT and other malignant properties, including CSC stemness and invasive potential (Fig. 4). Notably, CHD1L inhibitor 6 displays the ability to induce cell death that is consistent with the reversion of EMT and induction of cleaved E-cadherin mediated extrinsic apoptosis through death receptors (Fig. 5) (48). Furthermore, compound 6 synergizes with SN-38 displaying potent DNA damage induction compared with SN-38 alone, which is consistent with the inhibition of PARP1/CHD1L–mediated DNA repair (Supplementary Fig. S9; refs. 32, 39). Finally, we have characterized compound 6 as a lead CHD1L inhibitor with drug-like physicochemical properties and favorable in vivo pharmacokinetic/pharmacodynamic disposition with no acute liver toxicity (Fig. 6). Therefore, the pharmacophore of compound 6 has excellent potential for drug development.
On the basis of the data presented herein, we propose a mechanism of action for CHD1L-mediated TCF-driven EMT involved in colorectal cancer tumor progression and metastasis (Fig. 6F). We hypothesize that the TCF complex specifically recruits CHD1L to dynamically regulate metastatic gene expression. Central to this hypothesis, we propose that CHD1L binds to nucleosome hindered WREs when directed by the TCF complex via protein interactions with TCF4. Importantly, PARP1 is characterized as the major cellular activator of CHD1L through macro domain binding that releases autoinhibition (38, 39). Moreover, PARP1 is a required component of the TCF complex forming interactions with β-catenin and TCF4 (34). Therefore, it is reasonable to postulate that CHD1L is recruited by the TCF complex and activated by PARP1 and TCF4. Once activated, CHD1L exposes WREs by nucleosome translocation, facilitating TCF complex binding to WREs and transcription of malignant genes promoting EMT. CHD1L inhibitors have a unique mechanism of action by inhibiting CHD1L ATPase activity, which prevents exposure of WREs to the TCF complex, inhibiting transcription of TCF target genes associated with EMT and metastatic colorectal cancer.
In conclusion, since its discovery in 2008, numerous literature reports have characterized the biological role and clinical evaluation of oncogenic CHD1L in various solid tumor types. However, no approaches targeting oncogenic CHD1L are in clinical practice. With this report, we demonstrate that CHD1L is an oncogene and druggable target in colorectal cancer. We have identified the first inhibitors of CHD1L that have contributed fundamental knowledge of the mechanism of action of CHD1L in colorectal cancer and its potential as a therapeutic target. Future work with CHD1L inhibitors will focus on optimization through medicinal chemistry and conducting in vivo antitumor and antimetastatic efficacy studies. However, the work herein lays a foundation for the development of CHD1L-targeted therapeutics as an effective strategy to treat colorectal cancer and other CHD1L-driven cancers.
Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
Conception and design: J.M. Abbott, D.V. LaBarbera
Development of methodology: J.M. Abbott, Q. Zhou, H. Esquer, L. Pike, P.J. Lunghofer, T.M. Pitts, W.A. Messersmith, D.V. LaBarbera
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): J.M. Abbott, Q. Zhou, H. Esquer, L. Pike, S. Rinaldetti, A.D. Abraham, D.A. Ramirez, T.M. Pitts, D.P. Regan, D.L. Gustafson, D.V. LaBarbera
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): J.M. Abbott, Q. Zhou, H. Esquer, L. Pike, S. Rinaldetti, A.D. Abraham, A.-C. Tan, D.L. Gustafson, W.A. Messersmith, D.V. LaBarbera
Writing, review, and/or revision of the manuscript: J.M. Abbott, S. Rinaldetti, A.-C. Tan, D.L. Gustafson, W.A. Messersmith, D.V. LaBarbera
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): J.M. Abbott, D.A. Ramirez, W.A. Messersmith, D.V. LaBarbera
Study supervision: D.V. LaBarbera
This research was supported in part by the Department of Defense Peer Reviewed Cancer Research Program (W81XWH-18-1-0142; to D.V. LaBarbera); The Colorado Cancer Translational Research Accelerator (principal investigator: D.V. LaBarbera; co-principal investigator: D.L. Gustafson); The Skaggs School of Pharmacy and Pharmaceutical Sciences (SSPPS) ADR grant (to D.V. LaBarbera); and The Cancer League of Colorado grant (principal investigator: D.V. LaBarbera; co-principal investigator: W.A. Messersmith). S. Rinaldetti was supported by the Cancer Foundation of Luxembourg. We thank the SSPPS HTS drug discovery and chemical biology core facility, and the UCCC shared resource laboratories, particularly the Pharmacology core facility for microsomal and pharmacokinetic studies. The UCCC is an NIH NCI designated cancer center supported by grant number P30CA046934. We are grateful to Helena Berglund at the Karolinska Institute for generously providing the pNIC-CH2 vector.
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.