Our group previously demonstrated that inhibition of PRMT5 enzymatic activity with a PRMT5 small molecule inhibitor called CMP5 or through shRNA-mediated PRMT5 protein knockdown resulted in significantly decreased cellular proliferation and viability in HTLV-1-transformed T-cell lines (Panfil et al., 2015). CMP5 is a novel, first-generation selective PRMT5 inhibitor designed to selectively block the symmetric di-methylation of H4R3 (S2Me-H4R3) by PRMT5 (Alinari et al., 2015). CMP5 was identified through virtual docking of candidate small molecules with low binding energy in SAM cofactor and arginine-binding pockets of PRMT5. In our studies, we found that higher concentrations of CMP5 (25–50 μM) were required to achieve a significant decrease in the viability and proliferation of HTLV-1-transformed T-cells. To both improve and expand upon our previous studies, we assessed the capacity of a next generation orally bioavailable PRMT5 inhibitor, called EPZ015666, to selectively induce cellular toxicity in HTLV-1-transformed T-cell lines in vitro. EPZ01566 is a potent peptide-competitive and SAM-cooperative inhibitor that displays more than 10,000-fold specificity for PRMT5 over other protein arginine methyltransferases (Chan-Penebre et al., 2015). Treatment of mantle cell lymphoma (MCL) cell lines with EPZ015666 led to cell death with biochemical half-maximal enzyme-inhibition concentrations (IC₅₀) in the nanomolar range (Chan-Penebre et al., 2015). Of note, this same study found that EPZ015666 had dose-dependent antitumor activity in multiple MCL xenograft mouse models.

In order to evaluate the selective toxicity of EPZ015666 in HTLV-1-infected T-cells, a panel of HTLV-1-transformed, ATL-derived, and newly immortalized T-cell lines were treated with titrating amounts of EPZ015666 (0.1–10 μM) for 12 days. The antiproliferative effects of the inhibitor developed over several days, therefore a long-term proliferation assay was required. HTLV-1-transformed T-cell lines included SLB-1 and MT-2 (Figures 1A,B), PBL-HTLV-1 represents an infected and immortalized primary T-cell line (Figure 1C), and TL-Om1 and ATL-ED are patient ATL-derived T-cell lines (Figures 1D,E). EPZ015666 treatment of each HTLV-1-infected cell line led to a dose- and time-dependent decrease in cell viability. The effects of EPZ015666 treatment on total cellular symmetric arginine methylation (sdme-RG) in each cell line were examined by immunoblot after 4 days of treatment (Figures 1A–E, right panel). EPZ015666 treatment resulted in a concentration-dependent decrease in the intensity of multiple bands, indicating the effectiveness of the PRMT5 inhibitor. Cellular toxicity of EPZ015666 was next evaluated in normal resting CD4⁺ T-cells (Figure 1F), activated primary CD4⁺ T-cells (Figure 1G), and the HTLV-1-negative transformed T-cell lines, Jurkat (Figure 1H), and Hut-78 (Figure 1I). CD4⁺ T-cells represent the primary and preferential target of HTLV-1 infection (Richardson et al., 1990). EPZ015666 treatment of activated CD4⁺ T-cells, Jurkat, and Hut-78 cells led to a dose- and time-dependent decrease in cell viability. Interestingly, the inhibitor did not significantly affect the viability of normal resting CD4⁺ T-cells. The EPZ015666 IC₅₀ concentrations for resting CD4⁺ T-cells were approximately 100-1,000-fold higher than that of HTLV-1-infected T-cell lines (Figure 1K). Collectively, these data demonstrate that PRMT5 inhibition with EPZ015666 is selectively toxic to activated, rapidly proliferating CD4⁺ T-cells (including HTLV-1-infected T-cell lines), while demonstrating limited toxicity to normal resting T-cells even after prolonged incubation (Figure 1J).

To further characterize the mechanism of EPZ015666-mediated cellular toxicity in HTLV-1-transformed T-cell lines, we examined the level of apoptosis in Jurkat (HTLV-1-negative), SLB-1 (HTLV-1-transformed), and ATL-ED (ATL patient-derived) cell lines. Cells were treated with titrating doses of EPZ015666 for 12-days. The level of apoptosis was examined in each cell population using flow cytometry and Annexin V staining for early apoptosis and Annexin V/propidium iodide (PI) staining for late apoptosis (Figure 2A). Total cellular symmetric arginine methylation (sdme-RG) in each cell line was examined by immunoblot after 4 days of treatment to confirm PRMT5 enzymatic inhibition (Figure 2A, right panel). We found no significant difference in the level of apoptosis in vehicle-treated compared to EPZ015666-treated Jurkat cells. In contrast, both SLB-1 and ATL-ED cell lines displayed a significant dose-dependent increase in apoptosis in EPZ015666-treated cells. To further confirm induction of apoptosis, we measured the expression of PARP in cells treated with EPZ015666 after 4 days (Figure 2B). PARP is a nuclear poly (ADP-ribose) polymerase that is involved in DNA repair in response to environmental stress. PARP helps cells to maintain their viability, such that cleavage of PARP facilitates cellular disassembly and thereby serves as a marker of cellular apoptosis. We found that EPZ015666 treatment of ATL-ED and SLB-1 cells led to a dose-dependent increase in PARP cleavage. We next compared the levels of early and late apoptosis in ATL-ED cells following a 4-day treatment with EPZ015666 (Figure 2C). We found that while EPZ015666 treatment led to an overall increase in apoptosis, it was the amount of early apoptosis that increased in an EPZ015666 dose-dependent manner.

Our group previously demonstrated that PRMT5 becomes dysregulated and overexpressed during the HTLV-1-driven T-cell transformation process (Panfil et al., 2015). Given these findings, we hypothesized that PRMT5 likely represents a key cellular factor that is required for the transformation of HTLV-1-infected T-cells. To test this hypothesis, we asked if enzymatic inhibition of PRMT5 would alter HTLV-1-driven T-cell transformation. Freshly isolated human PBMCs co-cultured with lethally irradiated HTLV-1 producer cells showed progressive growth over time, consistent with HTLV-1-mediated immortalization (Figure 3A). As a control, PBMCs co-cultured with lethally irradiated HTLV-1-negative cells were unable to sustain progressive growth (data not shown). However, PBMCs co-cultured with HTLV-1 producer cells in the presence of EPZ015666 were unable to sustain progressive cell growth over time. We further quantified the number of wells with cellular outgrowth in both the vehicle control and EPZ015666-treated co-culture groups at the end of the study (Figure 3B). Approximately 50% of our wells had sustained cellular outgrowth and became newly immortalized HTLV-1 T-cell lines, while 0% of the EPZ015666-treated wells were immortalized. This suggests that PRMT5 enzymatic activity is required for HTLV-1-mediated T-cell immortalization. Earlier results demonstrated that EPZ015666 was effective against an immortalized primary T-cell line called PBL-HTLV-1 (Figure 1C). To further characterize EPZ015666, we next examined the impact of PRMT5 inhibition on four separate newly immortalized (i.e., less than 9 months post-infection/immortalization) T-cell lines (Figures 3C–F). Each newly immortalized PBL cell line had dose-dependent sensitivity to EPZ015666, similar to results obtained with well-established HTLV-1-transformed T-cell lines (Figure 1). In addition, the IC50 values for each of the newly immortalized cell lines were similar to results obtained with HTLV-1-transformed T-cell lines (Figure 3G).

HTLV-1 is unable to infect and replicate in murine cells (Delebecque et al., 2002). Therefore, the use of humanized immune system (HIS) mice is necessary to evaluate HTLV-1-mediated disease progression. HIS mice are created by intra-hepatic injection of CD34⁺ umbilical cord stem cells in NSG (NOD.Cg-Prkdc^scid Il2rg^tm1Wjl/SzJ) mice. Subsequent analysis by flow cytometry and histology demonstrates normal circulation and population of lymphoid organs by human lymphocytes. Infection of these mice with HTLV-1 results in integration of the virus into the genome of human lymphocytes, followed by activation and proliferation of CD4⁺ T-cells. Subsequently, a CD4⁺ T-cell leukemia and lymphoma develops, which leads to eventual death of the mice (Maksimova et al., 2022). HTLV-1-infected HIS mice typically start to become moribund after 6–7 weeks. Using this accelerated HIS model of HTLV-1-mediated disease, we investigated if EPZ015666 treatment of infected HIS mice could impact disease development or survival outcomes. EPZ015666 has a favorable pharmacokinetic profile in mice with a reported oral bioavailability of 69% following oral administration of 10 mg kg⁻¹ (Chan-Penebre et al., 2015). Twenty-one days after infection, HTLV-1-infected HIS mice were treated twice daily orally with of EPZ015666 (50 mg per kilogram of body weight [mg kg⁻¹]). EPZ015666 was previously shown to be well tolerated in immune-deficient mice up to 200 mg kg⁻¹ (Chan-Penebre et al., 2015) and we also detected minimal bodyweight loss in control mice (data not shown). EPZ015666-treated HTLV-1-infected HIS mice had significantly enhanced survival at study endpoint (46 days) compared to vehicle-treated mice (Figure 4A). At necropsy, splenic PBMCs were collected from each mouse, and proviral load (Figure 4B), viral sense transcript (tax; Figure 4C), and viral antisense transcript (hbz; Figure 4D) were measured using qPCR. Although proviral load and hbz transcript levels were slightly elevated in EPZ015666-treated mice, we found no significant differences in either proviral load or viral gene expression in vehicle compared to EPZ015666-treated mice. Serum was collected from each mouse throughout the course of study to measure human IL-2Rα levels. IL-2Rα is secreted by actively proliferating HTLV-1-infected cells and therefore, can be used as a biomarker for cellular proliferation in vivo. Using an IL-2Rα ELISA, we found no significant difference in the level of HTLV-1-infected cellular proliferation in vehicle vs. EPZ015666-treated mice (Figure 4E). To ensure in vivo target inhibition, we measured total cellular symmetric arginine methylation (sdme-RG) in each mouse spleen by immunoblot (Figure 4F). Although EPZ015666 treatment in HTLV-1-infected HIS mice increased survival, there was no significant difference in the proviral load, viral gene expression, or infected cell proliferation.

NSG mice characteristically lack mature T, B, and natural killer (NK) cells, resulting in impaired cytokine signaling and several deficiencies in their adaptive and innate immune response. This particular immune phenotype makes these mice an excellent model for the engraftment of human cells (Panfil et al., 2013). In an effort to further characterize PRMT5 as a viable therapeutic target for ATL patients, we next evaluated the efficacy of EPZ015666 treatment in NSG mice bearing subcutaneous SLB-1 (Figure 5) and ATL-ED (Figure 6) xenografts. NSG mice inoculated subcutaneously with HTLV-1-infected cell lines such as SLB-1 and ATL-ED will develop solid tumors (Dewan et al., 2003; Ohsugi et al., 2005; Arnold et al., 2008). SLB-1 and ATL-ED xenograft mice were treated with bidaily oral dosing of 25 or 50 mg kg⁻¹ EPZ015666. Importantly, PRMT5 inhibitor was administered after the formation of small measurable tumors. EPZ015666-treated HTLV-1 xenograft mice had significantly enhanced survival compared to vehicle-treated mice (Figures 5A, 6A). 3-D tumor dimensions and tumor weight were measured at time of necropsy in each mouse. As expected, we found that EPZ015666-treated ATL-ED xenograft mice had decreased overall tumor burden (Figures 6B,C). We did not record a measurable difference in the overall tumor burden in SLB-1 xenograft mice at time of necropsy (Figures 5B,C). Serum was collected from each mouse throughout the course of study to measure human IL-2Rα levels. Using an IL-2Rα ELISA, we found no significant difference in the level of cellular proliferation in vehicle or EPZ015666-treated SLB-1 xenograft mice (Figure 5D). However, we observed a dose-dependent decrease in IL-2Rα levels in EPZ015666-treated ATL-ED xenograft mice (Figure 6D). To ensure in vivo target inhibition within the solid tumor, we measured total cellular symmetric arginine methylation (sdme-RG) in each mouse tumor by immunoblot (Figures 5E, 6E). Taken together, our results suggest that EPZ015666 can significantly decrease HTLV-1 solid tumor burden and increases overall survival in HTLV-1 xenograft mice.

Human PBMCs (hPBMCs) were isolated from whole blood freshly collected from healthy donors using Ficoll-Paque PLUS (Cytiva, Marlborough, MA). Protocols for blood collection from human donors were approved by the Ohio State University Institutional Review Board. Naïve resting T-cells were enriched for CD4⁺ T-cells using a Human CD4 T Lymphocyte Enrichment Set (BD Biosciences, Franklin Lakes, NJ) according to the manufacturer’s instructions. CD4 T-cells were activated using the Dynabeads™ Human T-Activator CD3/CD28 kit (Thermo Fisher). Primary T-cells and early passage, HTLV-1-immortalized primary human T-cell lines (PBL-HTLV-1) were cultured in RPMI 1640 supplemented with 20% FBS, 100 U/ml penicillin, 100 μg/ml streptomycin, 2 mM L-glutamine, and 20 U/ml recombinant human interleukin-2 (hIL-2; Roche Diagnostics GmbH, Mannheim, Germany). Jurkat (HTLV-1-negative transformed T-cell line), Hut-78 (HTLV-1-negative transformed T-cell line), MT-2 (HTLV-1-transformed T-cell line), ATL-ED (ATL-derived T-cell line), and TL-Om1 (ATL-derived T-cell line) cells were maintained in RPMI 1640 supplemented with 10% FBS, 2 mM glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. SLB-1 cells (HTLV-1-transformed T-cell line) and 729.ACH cells (HTLV-1 producer cell line) were maintained in Iscove’s medium supplemented with 10% FBS, 2 mM glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. All cells were grown at 37°C in a humidified atmosphere of 5% CO₂ and air.

Cell lysates were harvested in NP-40 lysis buffer containing Complete Mini protease inhibitor cocktail (Roche Diagnostics). A small portion of each solid tumor (from NSG mice) or mouse spleen (from HIS mice) was collected in NP-40 lysis buffer containing Complete Mini protease inhibitor cocktail and homogenized using a BioMasher® II Micro Tissue Homogenizer (VWR). Protein quantification was carried out using the Pierce™ BCA Protein Assay Kit (Thermo Fisher) and a FilterMax F5 Multi-Mode Microplate Reader (San Jose, CA, United SA). Equal concentrations of protein were separated in Mini-PROTEAN® TGX™ Precast 4–20% Gels (Bio-Rad, Hercules, CA, USA) and transferred to nitrocellulose membranes. Membranes were blocked in PBS containing 5% milk and 0.1% Tween-20 and incubated with primary antibody overnight at 4°C. The following antibodies were used: anti-Symmetric Di-Methyl Arginine Motif [sdme-RG] (1:1000, Cell Signaling, Danvers, MA, USA), anti-PARP (1:1000, Cell Signaling), anti-β-actin (1:5000, Sigma, St. Louis, MO, USA), and anti-α-tubulin (1:250, Santa Cruz Biotechnology). Secondary antibodies used include IgG HRP anti-rabbit and anti-mouse (1:5000, Promega). Blots were developed using Pierce™ ECL Western chemiluminescence substrate (Thermo Scientific). Images were captured using the Amersham Imager 600 (GE Healthcare Life Sciences).

Activated and naïve T-cells were seeded in 12-well plates at 5 × 10⁵ cells/mL and all other lymphoid cells were seeded in 12-well plates at 1 × 10⁵ cells/mL. EPZ015666 (MedChemExpress) or vehicle (DMSO) were added to duplicate wells at the indicated concentrations. Cells were incubated at 37°C. Cell viability was measured on days 4, 8, and 12 using Trypan blue dye exclusion. Fresh EPZ015666 or vehicle were added to the appropriate wells at the indicated concentrations every 4 days.

Lymphoid cells were seeded in 6-well plates at 0.5 × 10⁵ cells/mL. EPZ015666 or vehicle (DMSO) were added to duplicate wells at the indicated concentrations on days 0, 4, and 8. Cells were incubated at 37°C for 12 days. Following incubation, cells were collected by slow centrifugation (5 min, 3,000 rpm) for apoptosis analysis via flow cytometry. Collected cells were stained using the FITC Annexin V Apoptosis Detection Kit I (BD Biosciences, Franklin Lakes, NJ, USA) according to the manufacturer’s instructions. Early apoptosis was defined was cells which stained positive for Annexin V (FITC) and negative for PI. Late apoptosis was defined as cells which stained positive for Annexin V (FITC) and PI.

Long-term immortalization assays were performed as detailed previously (Green et al., 1995). Briefly, 2 × 10⁶ freshly isolated human PBMCs were co-cultivated at a 2:1 ratio with lethally irradiated 729.ACH cells (HTLV-1-producer cell line) in 24-well culture plates (media was supplemented with 20 U/ml rhIL-2). At 1-week post-infection (co-culture), cells were treated weekly with 1 μM EPZ015666 or vehicle (DMSO). Viable cells were enumerated weekly using Trypan blue dye exclusion.

Breeding pairs of NOD.Cg-Prkdc^scid Il2rg^tm1Wjl/SzJ mice (NSG strain, specific pathogen free) were purchased from The Jackson Laboratory (Bar Harbor, ME). These mice lack mature T-cells, B-cells, or functional NK or dendritic cells, and are deficient in cytokine signaling. Animals were housed in individually ventilated microisolation cages with corncob bedding and provided with commercial pelleted rodent chow and reverse-osmosis–purified water without restriction. Cages containing autoclaved bedding were used for cage changes, which were performed in a ventilated biosafety cabinet. Mice were maintained in a room with constant temperature (20 ± 2°C) and relative humidity (50% ± 20%) under a 12:12-h light–dark cycle. The animal use protocol received prior approval by the Institutional Animal Care and Use Committee of The Ohio State University.

Shortly (24 to 48 h) after birth, pups were removed temporarily from the dam and treated with whole-body irradiation at 1 Gy (RS 2000, Rad Source, Suwanee, GA). Each mouse then was injected into the liver with 3 × 10⁴ to 1 × 10⁵ CD34⁺ HUSC (Lonza, Allendale, NJ) in 50 μl PBS (pH 7.4) by using a sterile 26-gauge hypodermic needle. After recovery, pups were returned to their dams, allowed to mature normally, weaned at 21 days, and then housed in groups (maximum, 5 mice per cage). At 10 weeks, after HUSC engraftment, mice were tested for the presence of human peripheral blood cells. Mice with at least 15% human CD45-positive lymphocytes were infected by intraperitoneal inoculation of 10⁶ lethally irradiated (100 Gy) 729.ACH producer cells; an aliquot of cells was maintained in culture to control for irradiation treatment. Mice began bidaily administration of vehicle (0.5% methylcellulose) or PRMT5 inhibitor (50 mg kg⁻¹ EPZ015666) via oral gavage (n = 8–9 per treatment group) at day 21 post-infection. Mouse serum was collected at weekly time points by submandibular bleeds. Serum was used for quantification of human IL-2Rα levels using a human CD25/IL-2Rα Quantikine ELISA kit (R&D Systems, Inc., Minneapolis, MN). Animals were euthanized when they lost more than 20% of their body weight within 48 h.

The NOD.Cg-Prkdc^scidIl2rg^tm1Wjl /SzJ (NSG) xenograft mice were purchased from Charles River. NSG mice were subcutaneously inoculated with 1 × 10⁷ SLB-1 or 1 × 10⁶ ATL-ED cells. Mice were treated through bidaily administration of vehicle (0.5% methylcellulose) or PRMT5 inhibitor (25, 50 mg kg⁻¹ EPZ015666) via oral gavage (n = 9 per treatment group) once palpable tumors were observed (8 days post-injection for SLB-1, 11 days post-injection for ATL-ED). Mouse serum was collected at weekly time points by submandibular bleeds. Serum was used for quantification of human IL-2Rα levels using a human CD25/IL-2Rα Quantikine ELISA kit (R&D Systems, Inc., Minneapolis, MN). At 24 days post-infection for SLB-1 and 33 days post-infection for ATL-ED, all remaining mice were euthanized. Tumors were extracted for assessment of weight (g) and dimensions (mm³). Animals were euthanized when they lost more than 20% of their body weight within 48 h.

Genomic DNA and RNA were isolated from HIS mouse spleens using the AllPrep DNA/RNA Mini Kit (QIAGEN, Hilden, Germany) according to manufacturer’s instructions. RNA samples were subjected to on-column DNase digestion, and 250 ng of RNA was used for cDNA synthesis, as described above. Approximately 1–2 μl cDNA was used per qPCR reaction with iQ™ SYBR® Green Supermix (Bio-Rad, Hercules, CA) and 300 nM of each sense and antisense primer (20 μl final reaction volume). Primer pairs to specifically detect viral mRNA species (tax, hbz) and gapdh were described previously (Li and Green, 2007). The reaction conditions were 95°C for 5 min followed by 40 cycles of 94°C for 30 s and 60°C for 45 s. Total copy numbers of each gene target in HIS mice were determined by log₁₀ dilutions of plasmid DNA to generate a standard curve. Copy numbers were normalized to 1 × 10⁴ human GAPDH. Samples and standards were run in duplicate with no-RT and no-template controls included on each plate. To determine proviral load, 250 ng genomic DNA was used for qPCR with a primer set specific to HTLV-1 gag/pol. The 20 μl final reaction volume included iQ™ SYBR® Green Supermix and 300 nM each of 5′ primer (#20) and 3′ primer (#19). The reaction conditions were as follows: 95°C for 3 min followed by 40 cycles of 94°C for 30 s, 60°C for 45 s, and 72°C for 60 s. Total copy number was calculated using a standard curve generated by duplicate log₁₀ dilutions of ACHneo plasmid DNA. Proviral copies per cell in HIS mice was calculated based on the approximation that 6 pg. human DNA is equivalent to 1 cell.