(Z)-3,5,4′-Trimethoxystilbene Limits Hepatitis C and Cancer Pathophysiology by Blocking Microtubule Dynamics and Cell-Cycle Progression

Hepatocellular carcinoma (HCC) is the third most common cause of cancer-related death worldwide with a dismal 5-year survival rate of 11% (1, 2). Chronic infection with hepatitis C virus (HCV) is a major risk factor for the development of HCC (1, 3). HCV patients coinfected with other viruses or with metabolic comorbidity (obesity and diabetes) exhibit faster progression of liver disease and are difficult to treat with standard interferon-based treatment regimens (3). The combinations of direct-acting antiviral (DAA) drugs against three key HCV nonstructural proteins (NS3, NS5A, and NS5B) have shown remarkable efficacy (>90%) for curing the infection (4, 5). However, these drugs are inaccessible to millions of patients and a complete recovery of damaged liver by the HCV treatments alone has not been proven.

The majority of HCC cases are normally diagnosed at late stages, and the use of curative surgery or other treatments are less successful (6, 7). Kinase inhibitors have been shown to increase survival only by a few months or are ineffectual in advanced HCC patients (8). The activation of multiple signaling pathways and enrichment of tumor/cancer stem cells (CSC) within the tumor appear to mediate HCC multidrug resistance (9–11). CSCs represent small subpopulations within a tumor that possess self-renewing capabilities and the ability to differentiate into a heterogeneous lineage within the tumor mass (12–14).

We previously demonstrated a positive correlation between the levels of HCV replication and expression of an array of CSC-associated markers, including doublecortin-like kinase 1 (DCLK1; refs. 15, 16). These changes appear to promote cellular dedifferentiation and the gain of CSC properties in HCV-positive cells. We have further demonstrated that knockdown of DCLK1 results in downregulation of HCV replication, cell migration, and epithelial–mesenchymal transition (EMT) in multiple cancer cell lines and in mouse models (15, 17). There is currently no clear choice for targeting CSCs efficiently (18, 19). DCLK1 is considered to be an important target for the treatment of tumors of the liver, pancreas, and colon. HCV also induces EGFR that facilitates HCV entry into the cells and promotes progression of liver diseases (20, 21). The EGFR inhibitor, erlotinib, has been shown to downregulate HCV RNA levels in a mouse model (20). These observations clearly suggest that cellular kinases (DCLK1 and EGFR) are critical components of HCV-induced chronic liver diseases.

The naturally occurring antioxidant, resveratrol (RVT) has been extensively studied for possible health benefits (22, 23), including anticancer effects (24–26). However, its potential benefits could not be demonstrated in patients with malignancy due to poor absorption and extremely low bioavailability (23, 27). The structurally related analogue (Z)-3,5,4′-trimethoxystilbene (Z-TMS), originally isolated from the bark of Virola elongate, has been shown to inhibit microtubule polymerization, induce G₂–M arrest, and inhibit ornithine decarboxylase activity (28, 29). Z-TMS also has long half-life and limited drug clearance when administered intravenously (30, 31). Here, we report mechanism(s) of Z-TMS's antiviral and antitumor activities in several in vitro models and in DEN/CCL4–induced liver injury in C57BL/6 mouse model. Z-TMS exerts its inhibitory effects on tumors by diminishing DCLK1⁺ cell population, interference with DCLK1-microtubule dynamics, cell-cycle arrest at G₂–M phase, promotion of autophagy, and causing nuclear fragmentation. It is considerably effective against erlotinib-resistant T790M-mutant lung carcinoma cells and may evolve as a potential candidate for the treatment of HCV-induced advanced liver diseases, including HCC.

The non–small cell lung adenocarcinoma cells (NSCLC, H1975) and primary human hepatocytes were purchased from ATCC and BD Biosciences, respectively. All other cells (endogenous) were tested before start of the studies using MycoAlert Kit (Lonza) and were found negative for Mycoplasma. The parent cell line Huh7 and human hepatocytes were recently evaluated (January 14, 2016) for Hantaan virus, LCMV, and Mycoplasma using PCR by IDEXX Bioresearch and were negative for these agents. The characteristics of hepatoma Huh7-derived cell lines (Huh7.5, GS5, and FCA4) have been described previously (15, 16, 32, 33). HCV⁺DCLK1⁺ hepatoma cells were generated by transducing G418-resistant FCA4 cells with lentivirus expressing DCLK1 (isoform 1) tagged with RFP at the N-terminus. The cells were cultured as described previously (34). Resveratrol (Sigma) and Z-TMS (Cayman Chemical) were prepared in DMSO and stored at −20°C. EGFR (aa 695–1210) and luciferase assay kits were purchased from Promega. The antibodies were purchased from the following sources: DCLK1 (ab31704), NS5B (35586), actin (ab1801), and Abcam; LC3B (2775), cleaved caspase-3 (Asp175; 9664), and p-Akt (4069), from Cell Signaling Technology; and anti-NS3 antibody (217-A) from Virogen.

The cell culture–generated JFH1 HCV-2a infectious particles (HCVcc) were produced as described previously (35). In brief, JFH1 RNAs were electroporated into Huh7.5 cells and HCVcc particles were determined in cell culture supernatant. Huh7.5 cells were infected with HCVcc at multiplicity of infection of 1 for 48 hours. Subsequently, the HCVcc-infected cells were treated with Z-TMS (1 μmol/L) or equivalent amounts of DMSO in triplicates for 48 hours. The untreated infected cells were considered as positive control, whereas uninfected Huh7.5 cells served as negative control for detection of the HCV RNA by qRT-PCR. The HCV RNA copy numbers per microgram of total RNAs in Z-TMS were calculated and compared with vehicle-treated and -untreated samples. JFH-1/GND RNAs (replication defective JFH1 mutant) were used as negative control. In most of the experiments, HCV-infected cells were serum starved for 4 hours before harvesting. Total RNAs from the HCV replicon–expressing cells were isolated and subjected to RT-PCR as described previously (15).

Cells treated with resveratrol, Z-TMS, or DMSO were washed with PBS, collected, and lysed with mammalian protein extraction reagent (M-PER; Thermo Scientific). Western blots were performed using 30–40 μg of cell lysates (15, 34) and antibodies were diluted as recommended by the manufacturers. Each Western blot analysis was repeated three times. To evaluate target protein to actin ratio, band intensities were calculated using GelQuant software and mean ratios were presented. Cell viability was determined using CellTiter 96 AQueous One Solution (Promega) as recommended by the manufacturer.

H1975 cells were cultured in RPMI1640 media supplemented with 10% FCS. Z-TMS–treated and -untreated H1975 cells in a 96-well culture plate were stained with DRAQ5 and scanned with Operetta (PerkinElmer). Each treatment was done in quadruplets and each well was analyzed for five different fields for the analysis of nuclear fragmentation. The images (average of 5 fields for each well) were analyzed by Columbus software. Wound-healing assay for cell migration was carried out as described previously (34).

Cells treated were with Z-TMS (1 μmol/L) or DMSO for 48 hours, harvested, and washed with PBS. The pelleted cells were resuspended in PBS containing paraformaldehyde (2%) for 1 hour at 4°C, washed twice with ice-cold PBS, resuspended and fixed in 70% ethanol, and incubated overnight at 4°C. The cells were treated with RNase A (200 μg/mL) and propidium iodide (PI, 50 μg/mL; Sigma) for 30 minutes at 37°C in the dark and stored at 4°C until acquisition of the flow cytometry data. The instrument was calibrated with unstained cells and sorted for PI intensity. A total of 10,000 cell events were collected. Flow cytometry data were collected and analyzed using the CELLQuest program (Becton Dickinson).

EGFR kinase assay was performed using the ADP-Glo kinase assay and recombinant EGFR representing amino acids 695–1210 of the wild type using Promega's protocol. The kinase was treated with 1 μmol/L of Z-TMS or erlotinib in triplicates, and luciferase read-out for the EGFR kinase activities were compared with the untreated control. The experiment was repeated twice.

To evaluate the relationship between DCLK1 expression levels and survival of HCC patients, a Kaplan–Meier plot was generated using the liver HCC (LIHC, total 369 cases) data provided by the TCGA Research Network (http://cancergenome.nih.gov/) using the UCSC Genome Browser. Immunohistochemical staining of liver tissues from patients with HCV and HCC were obtained from the LTCDS (Minneapolis, MN; refs. 15, 34).

All mice studies reported here were preapproved (protocols # 12-041 and 15-083) and supervised by the University of Oklahoma Health Sciences Center (OUHSC) Institutional Animal Care and Use Committee, which adheres to the PHS Policy IV.B.3. Athymic nude Balb/c mice were purchased from The Jackson Laboratory and housed in pathogen-free conditions. One million Huh7 cells were washed with PBS three times, resuspended in the same buffer, and injected subcutaneously into the dorsal flanks of 4- to 6-week-old mice as reported previously (34). Tumors were measured with calipers and the volumes were calculated using formula: 0.5 × (length × width²).

Adult immunocompetent C57BL/6 mice (n = 30, 8-week-old, generated by in-house breeding at OUHSC Rodent Barrier Facility) were randomized into 6 groups (5 mice/group). Groups 1 through 3 received two intraperitoneal injections of diethylnitrosamine [DEN, 25 mg/kg body weight (BW), one injection per week] followed by twice-weekly injections of CCl₄ (1 mL/kg BW) for the next 7 weeks. Except group 1, groups 2 and 3 were also treated with 20 and 40 mg/kg BW of Z-TMS dissolved in DMSO, respectively. Group 4 received only Z-TMS (40 mg/kg BW) to determine its possible toxicity. Groups 5 and 6 served as DMSO and untreated controls, respectively, during the same period. At the termination of the experiments, all the animals were euthanized and subjected to necropsy for collection and analysis of blood or tissues of different organs.

All experiments were performed in biological replicates of at least three and repeated to confirm the results. The graphs were presented as mean ± SD. The P values were calculated using the Student t test. Results with P ≤ 0.05 were considered statistically significant.

Published reports suggest potential benefits of resveratrol (3,5,4′-trihydroxy-trans-stilbene, RVT) and its analogues on human health (reviewed in refs. 36, 37). We initially tested the anti-HCV activities of RVT and its natural analogue Z-TMS in hepatoma cell lines (GS5 and FCA4) expressing HCV-1b subgenomic replicons (16, 34). The results suggested that Z-TMS is nearly 100 times more potent in downregulating the levels of HCV NS5B polymerase, which is now considered a core target of direct antiviral therapy (results not shown). Next, FCA4 cells were treated with varying amounts (0, 0.05, 0.1, 0.5, 1.0, and 2.5 μmol/L) of Z-TMS or equivalent amounts of DMSO for 48 hours. The total lysates prepared from drug-treated and -untreated cells were analyzed for NS5B and actin (loading control) levels. The data suggest that Z-TMS inhibits NS5B expression in a dose-dependent manner (Fig. 1A, lanes 2–6). Similar results were obtained with GS5 cells (not shown). The DMSO solvent control had no inhibitory effect on the NS5B levels (lanes 7–11) as compared with the untreated control (lane 1). Under these conditions (0.1–1.0 μmol/L Z-TMS for 48-hour treatment), FCA4 cells as well as normal human hepatocytes remained fully viable (Fig. 1B and C).

To examine whether Z-TMS inhibits HCV replication in an HCV infection model, Huh7.5 cells were infected with tissue culture–generated JFH1 viruses (HCVcc) for 48 hours to allow replication of the viral RNA. The infected cells were then treated with 1 μmol/L of Z-TMS or vehicle for 48 hours. Total RNAs isolated from these cells were subjected to qRT-PCR to determine the levels of HCV replication. As expected, the JFH1 RNA copy number was drastically reduced (∼10 times) by Z-TMS treatment as compared with the DMSO-treated or -untreated cells (Fig. 1D). The results were fully corroborated by the dramatic reduction in NS3 level (Fig. 1E, lane 3) as compared with the controls (lanes 2 and 4).

Microtubules are highly dynamic structures whose regulations are critical for HCV replication, cell division, cell migration, and cell polarity. The CSC marker DCLK1 associates with microtubules and stimulates polymerization of tubulins. We have previously shown that DCLK1 is overexpressed in liver tissues derived from patients with cirrhosis and HCC. Its expression is also correlated with activation of inflammatory and protumorigenic signals as well as hepatoma cell migration (34). These observations led us to investigate the significance of DCLK1 overexpression in patient survival and effects of Z-TMS on DCLK1 interaction with microtubules or microtubule dynamic instability. As shown in Fig. 2A, DCLK1 is overexpressed in liver tissues of HCV-positive patients with cirrhosis and HCC but not in normal liver. The Kaplan–Meier plot suggests that the 5-year survival rate is approximately reduced by three times in HCC patients (n = 369) with hepatic DCLK1 overexpression (Fig. 2B, red and black lines) as compared with the patients with low DCLK1 levels (green line). In a mouse tumor xenograft model described previously (34), large tumors (average size of 4–4.5 cm³) derived from Huh7 cells exhibited extensive staining for human DCLK1 except the one that was smaller in size (0.65 cm³; Fig. 2C). Because of DCLK1′s important roles in microtubule dynamics, HCV replication and tumor growth in addition to the ability of HCV⁺DCLK1⁺ hepatoma cells to form spheroids (15, 16), we examined the effects of Z-TMS on HCV⁺DCLK1⁺ cells. These cells were isolated from FCA4 cells coexpressing recombinant RFP-DCLK1 and HCV subgenomic replicon by FACS method. As shown in Fig. 2D, expression of HCV NS5B polymerase in HCV⁺DCLK1⁺ hepatoma cells was significantly reduced by 1 μmol/L of Z-TMS treatment (lane 3) as compared with controls (lanes 1 and 2). Z-TMS treatment also caused significant inhibition in the proliferative potential of these cells (Fig. 2E). We carried out confocal immunofluorescence microscopy to determine localization of DCLK1 with microtubule in these cells after Z-TMS treatment. The results revealed that Z-TMS induces bundling of microtubules and speckle-like structures of DCLK1–microtubule complexes (Fig. 2F, bottom). However, the untreated and DMSO-treated cells exhibit normal distribution of DCLK1 (red) and tubulin (green) or their complexes (yellow). Collectively, these results indicate a direct relationship of DCLK1 with the survival of HCC patients and possibility of targeting HCV-positive DCLK1-overexpressing hepatoma cells by Z-TMS.

We used a 3D magnetic levitation method (38) to culture GS5 cells as visible cell spheroids/aggregates (Fig. 3A). The levels of NS5A-GFP green fluorescence indicated continued HCV replication in the spheroid-like cultures (Fig. 3A, inset). Immunohistochemical staining of these aggregates revealed different cell phenotypes. Although large cells displayed extensive CK19 expression (circulating tumor cell marker) and cell-to-cell contacts, a low number of unique small cells were DCLK1 positive. This observation clearly suggests heterogeneous cell phenotypes in the spheroids. Z-TMS successfully inhibited growth of this cellular mass (Fig. 3B).

GS5 cells, which express a NS5A-GFP chimeric protein encoded by HCV-1b subgenomic RNAs, and Huh 7.5 cells were treated with 1 μmol/L of Z-TMS for 48 hours and stained with PI for cell-cycle analysis. The results clearly suggest that most GS5 cells and Huh7.5 parent cells were stalled in the G₂–M phase (>90%) with a concomitant reduction in the G₁ and S-phases (Fig. 4A). However, DMSO-treated and untreated cells showed similar cell-cycle distribution. The result shows that the observed change in cell cycle was induced by Z-TMS and not by the DMSO (solvent). Similar Z-TMS effects were observed for other hepatoma cell lines (Huh7 and FCA4, not shown). Next, total lysates of the drug-treated and -untreated cells were subjected to Western blot analysis for detection of proteins involved in cell-cycle checkpoints. Z-TMS treatment caused downregulation of total cyclin-dependent kinase 1 (CDK1) with a concomitant induction of p21^Cip1/Waf1 expression (Fig. 4B, lane 3), which is a potent inhibitor of CDK1 and CDK2. Also, p21^Cip1/Waf1 mediates growth arrest and cellular senescence. Activation of Akt overcomes a G₂–M cell-cycle checkpoint induced by DNA damage (39). Therefore, successful blockage of cancer cells at G₂–M phase should also be accompanied by the downregulation of Akt. Treatment of GS5 cells with Z-TMS (1 μmol/L) resulted in significant downregulation of Akt (Ser⁴⁷³) phosphorylation without affecting total Akt level (Fig. 4C, lane 3). Taken together, Z-TMS appears to induce G₂–M cell-cycle arrest by induction of p21^Cip1/Waf1 and downregulation of Akt signaling.

Because Z-TMS inhibits Akt activation, the status and function of its upstream regulator EGFR) was examined. Our interest in EGFR investigation stems in part from the observation that erlotinib, an EGFR inhibitor, diminishes HCV entry into cells and reduces HCV replication in a mouse model (20). Because of the lack of an EGFR-mutant hepatoma model, we used a well-characterized erlotinib-resistant lung adenocarcinoma (NSCLC) H1975 cells that contains the T790M gatekeeper mutation in EGFR to determine the effect of Z-TMS on these cells. As shown in Fig. 5A, Z-TMS significantly reduced survival of H1975 cells in the 0.1 μmol/L to 1.0 μmol/L range within 48 hours, whereas erlotinib failed to reduce cell survival even though it was fully active against the wild-type EGFR kinase domain (aa 695–1210; Fig. 5A and B). The luciferase-based EGFR activity assay suggests that erlotinib efficiently inhibited (90%) the kinase activities, whereas Z-TMS failed to exhibit strong inhibition in spite of reducing H1975 survival. Confocal immunostaining clearly revealed that only Z-TMS but not erlotinib induced microtubule bundling (Fig. 5C, middle). Unlike the intense colocalization of DCLK1 with microtubules in DCLK1-overexpressing hepatoma cells (Fig. 2F), we observed mostly punctate staining of DCLK1 in H1975 probably due to alternatively spliced or post-translationally modified forms that do not preferentially bind microtubules (Fig. 5C).

To understand the mechanism of Z-TMS–induced cell death, H1975 cells were cultured in a 96-well plate and treated with Z-TMS (1 μmol/L) in quadruplets. After fixing the cells, nuclei were stained and the extent of nuclear fragmentation was assessed. The data suggest that Z-TMS induces nuclear fragmentation in nearly 70% to 80% cells as compared with the untreated or DMSO or cisplatin (50 μmol/L) treated controls (Fig. 5D). Total lysates of H1975 cells treated with the drugs as described above showed activation of autophagy markers, LC3B-I and LC3B-II proteins in Z-TMS–treated cell lysate (Fig. 5E, lane 3), which were much higher than the corresponding controls (lanes 1, 2 and 4). Similar Western blot analysis suggested that only cisplatin, but not Z-TMS, induces cleaved caspase-3 (Asp175), which marks cell death by apoptosis (Fig. 5F). Thus, Z-TMS appears to induce H1975 cell death by autophagy and nuclear fragmentation.

The scheduled administration of DEN (one injection) followed by twice-weekly injections of carbon tetrachloride (CCl₄) induces liver injury that further progresses to fibrosis and HCC within 8 to 10 months (40–42). We treated immunocompetent C57BL/6 mice with two DEN (25 mg/kg BW) injections once a week followed by administration of CCl₄ (1 mL/kg BW) alone or with Z-TMS (20 or 40 mg/kg BW) twice per week for 7 weeks (groups 1 through 3 in Table 1; Fig. 6A). As expected, the DEN/CCl₄–treated mice developed liver injury characterized by histologic perturbations and appearance of nonepithelial cells in the parenchyma (Fig. 6A, extreme left panels). These regions also exhibited unusually high numbers of DCLK1⁺ cells with intense staining in epithelial and nonepithelial compartments at the injury sites (only two liver sections from different mice of the same group are shown in top and bottom). This observation suggests that DEN/CCl₄–led injury significantly induces DCLK1 expression in liver (highlighted in Fig. 6B). The untreated control (normal) mice livers were largely negative for DCLK1 stain or occasionally revealed a fewer DCLK1⁺ cells within portal triads (extreme right). The mice treated with Z-TMS (groups 2 and 3), which were simultaneously receiving DEN/CCL4 doses, exhibited improved histologic appearance with significantly reduced number of DCLK1⁺ cells as compared with group 1 (DEN/CCl₄ treated, Fig. 6A). Both histologic and liver function panel (ALB, ALT, AST, ALKP, and TBIL in Table 1) data clearly suggest that Z-TMS alone (group 4) does not exhibit hepatotoxicity under these treatment conditions. Instead, Z-TMS successfully protected the liver from toxic agents and improved injury most likely by eliminating or suppressing DCLK1⁺ cells. Mice in any group did not lose their body weight.

Viral hepatitis, steatohepatitis, and metabolic disorders substantially contribute to the development of cirrhosis and primary HCC. End-stage liver disease due to chronic HCV infection is expected to remain a major health care issue in most countries partly due to lack of effective treatment options or inaccessibility to newly discovered drugs. Current HCC antineoplastic drugs, sorafenib alone or in combination with other drugs, cause considerable toxicity and only improve median overall survival by 3 months. Here, we have reported novel properties of Z-TMS that exerts its antiviral and antitumor activities by interference with the CSC marker DCLK1, microtubule dynamics, induction of autophagy, G₂–M arrest and nuclear fragmentation. The combined effects reflect in vitro efficacy of Z-TMS against the drug-resistant tumor cells such as NSCLC. Z-TMS also promotes hepatic recovery from extensive DEN (genotoxic)/CCl₄ (hepatotoxic) induced liver injury in immunocompetent C57BL mice. Thus, Z-TMS may potentially block initiation and progression of HCC by targeting DCLK1. This notion is also supported by our observation that DCLK1 downregulation results in reduction of hepatoma-like tumor growth (43). Although Z-TMS at doses 20 and 40 mg/Kg BW did not exhibit hepatic toxicity, 20% mice showed complications in gastrointestinal emptying. These observations warrant us for optimizing drug dosages and duration of treatment to gain its maximum health benefits.

Using subgenomic replicons and full-length JFH1 infection models of HCV, we have demonstrated the efficacy of Z-TMS against widely prevalent HCV genotypes 1 and 2. The magnetic levitation experiments revealed that growth of HCV-expressing spheroids/aggregates could also be effectively targeted by Z-TMS. The heterogeneity of this cellular mass and cell-to-cell contacts in the spheroids were revealed by immunohistochemical staining showing membranous CK19. It further showed expected DCLK1⁺ subpopulations (usually 1%–2% of total GS5 cells; ref. 15). Our in vivo experiments suggest that Z-TMS can diminish the DCLK1⁺ population in injured livers. Because DCLK1 appears to be involved in HCV replication and hepatic malignancy (15, 16, 43), it is likely that Z-TMS will be able to eliminate HCV-expressing cancer stem–like cells to prevent progression of liver diseases.

Although normal liver parenchyma lacks detectable DCLK1 expression, DCLK1 expression and number of DCLK1⁺ cells are increased significantly in the stroma, regenerative cirrhotic nodules, and HCC. Z-TMS clearly targeted DCLK1-microtubule dynamics that promotes HCV replication, cell migration, and cell-cycle progression. Microtubule-stabilizing drugs (paclitaxel and docetaxel) and destabilizing drugs (vinblastine and nocodazole) have been extensively used for the treatment of cancer (44, 45). A few of these drugs have also been shown to hamper movement of the HCV replication complexes required for viral replication (46). By sequestering DCLK1 with bundled microtubules, Z-TMS will block survival, proliferation, and division of DCLK1⁺ CSC cells.

Cancer/tumor stem cells (CSC) are key contributors of drug resistance, metastasis, and recurrence of tumors. At present, effectively targeting CSCs for treatment is challenging (18, 19). We previously demonstrated that HCV induces several CSC markers (CD133, Lgr5, and c-Myc), including DCLK1 (47, 48). Our current data revealed that DCLK1 overexpression in HCC-positive patients considerably reduces survival rate. Z-TMS causes sequestration of DCLK1 with microtubule bundles accompanied by a dramatic decrease in the NS5B polymerase level in HCV⁺DCLK1⁺ hepatoma cells. Z-TMS appears to be effective against DCLK1-positive CSCs. DCLK1 has been shown to play important roles in the development of intestinal polyps, intestinal response to radiation injury, cell survival, and control of metastasis. In addition, we have shown that DCLK1 positively regulates HCV replication in knockdown and overexpression systems. Thus, Z-TMS–targeted elimination of HCV-positive cells bearing CSCs footprints is likely to benefit patients with advanced liver diseases, including HCC.

The molecular mechanisms of Z-TMS's antiproliferative effects were also studied. We observed that Z-TMS treatment resulted in G₂–M arrest (95%) of hepatoma cells independent of HCV within 48 hours. Cell-cycle progression is orchestrated by cyclins and cyclin-dependent kinases (CDK). CyclinB1/CDK1 complex specifically regulates cell entry into mitosis, and enhances mitochondrial respiration and ATP generation for G₂–M transition. In accordance with these observations, we observed a considerable decrease in total CDK1 and a concomitant increase in the cell-cycle inhibitor, p21^Cip/Waf1, following Z-TMS treatment. The observed p21^Cip/Waf1 induction after Z-TMS treatments resulted in inhibition of CDK1 activities and cell-cycle arrest. Akt activation downstream of the PI3K/Akt/mTOR pathway regulates multiple cellular processes such as cell growth, CSC survival, metabolism, angiogenesis, and proliferation. Phosphorylation of Akt at Ser⁴⁷³ by mTORC2 is required for its hyperactivation and stabilization of its active conformation. We observed a considerable inhibition of Akt (Ser⁴⁷³) phosphorylation following Z-TMS treatment. In addition, Z-TMS induced autophagy that was accompanied by nuclear fragmentation. These properties are especially desirable for the treatment of hard-to-treat drug-resistant cancers.

In conclusion, the results presented here collectively support Z-TMS as a unique potent anti-HCV and antitumor drug that exerts its effects by targeting multiple pathways. Z-TMS significantly reduces DCLK1⁺ CSC populations during hepatic injury, interferes with DCLK1-microtubule dynamics, arrests cell cycle at G₂–M phase, promotes autophagy, and causes nuclear fragmentation. Thus, Z-TMS potentially overrides NSCLC resistance to erlotinib because of its unique mechanisms of action.

S. Sureban has ownership interest (including patents) in patents. M.S. Bronze is a consultant/advisory board member for Genentech/Roche. C.W. Houchen has ownership interest (including patents) and is a consultant/advisory board member for COARE Biotechnology Inc. No potential conflicts of interest were disclosed by the other authors.

Conception and design: C.B. Nguyen, H. Kotturi, G. Waris, A. Mohammed, C.W. Houchen, N. Ali

Development of methodology: C.B. Nguyen, H. Kotturi, A. Mohammed, C.W. Houchen, N. Ali

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): C.B. Nguyen, H. Kotturi, G. Waris, A. Mohammed, N. Ali

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): C.B. Nguyen, H. Kotturi, G. Waris, A. Mohammed, P. Chandrakesan, S. Sureban, D. Qu, C.V. Rao, N. Ali

Writing, review, and/or revision of the manuscript: C.B. Nguyen, H. Kotturi, A. Mohammed, M.S. Bronze, C.W. Houchen, N. Ali

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): H. Kotturi, R. May, N. Weygant, D. Qu, C.V. Rao, D.N. Dhanasekaran, N. Ali

Study supervision: H. Kotturi, A. Mohammed, C.W. Houchen, N. Ali

The authors are thankful to the Liver Tissue Cell Distribution System, (Minneapolis, MN). We thank Stephenson Cancer Center Pathology Core Laboratory for immunohistochemistry.

This work was partially supported by VA Merit Award and the Frances and Malcolm Robinson Endowed Chair (C.W. Houchen). N. Ali is partially supported by an Institutional Development Award (IDeA) from NIH (1P20GM103639NIH) and COMAA Research Fund Seed Grant (OUHSC).

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.