Targeting xCT, a cystine-glutamate transporter induces apoptosis and tumor regression for KSHV/HIV-associated lymphoma
© Dai et al.; licensee BioMed Central Ltd. 2014
Received: 25 February 2014
Accepted: 30 March 2014
Published: 4 April 2014
Kaposi’s sarcoma-associated herpesvirus (KSHV) is the etiological agent of primary effusion lymphoma (PEL), which represents a rapidly progressing malignancy arising in HIV-infected patients. Conventional chemotherapy for PEL treatment induces unwanted toxicity and is ineffective — PEL continues to portend nearly 100% mortality within a period of months, which requires novel therapeutic strategies. The amino acid transporter, xCT, is essential for the uptake of cystine required for intracellular glutathione (GSH) synthesis and for maintaining the intracellular redox balance. Inhibition of xCT induces growth arrest in a variety of cancer cells, although its role in virus-associated malignancies including PEL remains unclear. In the current study, we identify that xCT is expressed on the surface of patient-derived KSHV+ PEL cells, and targeting xCT induces caspase-dependent cell apoptosis. Further experiments demonstrate the underlying mechanisms including host and viral factors: reducing intracellular GSH while increasing reactive oxygen species (ROS), repressing cell-proliferation-related signaling, and inducing viral lytic genes. Using an immune-deficient xenograft model, we demonstrate that an xCT selective inhibitor, Sulfasalazine (SASP), prevents PEL tumor progression in vivo. Together, our data provide innovative and mechanistic insights into the role of xCT in PEL pathogenesis, and the framework for xCT-focused therapies for AIDS-related lymphoma in future.
The oncogenic γ-herpesvirus known as the Kaposi’s sarcoma-associated herpesvirus (KSHV) is a principal causative agent of cancer arising in patients with compromised immune systems . One of these cancers, primary effusion lymphoma (PEL), comprises transformed B cells harboring KSHV episome and arises preferentially within the pleural or peritoneal cavities of patients infected with HIV . PEL is a rapidly progressing malignancy with a median survival time of approximately 6 months . Currently, combinational chemotherapy is the standard of care for PEL, and cyclophosphamide, doxorubicin, vincristine, and prednisone (CHOP) regimens are considered first-line therapy [4, 5]. However, the myelosuppressive effects of systemic cytotoxic chemotherapy synergize with those caused by antiretroviral therapy or immune suppression [3, 4, 6]. Several novel approaches for PEL therapy have been reported in recent studies and increase survival for some patients, but a lack of sufficient safety and efficacy data have precluded their routine use. The proteasome inhibitor bortezomib and the combination of arsenic trioxide and interferon both suppress the NF-κB activation and may work synergistically with cytotoxic chemotherapy to reduce PEL viability [7, 8]. Unfortunately, proteasome inhibition and arsenic incur significant toxicities limiting their clinical application. The mammalian target of rapamycin (mTOR) inhibitor, sirolimus inhibits PEL cell growth in a murine xenograft model , but which paradoxically induces expression of the serine/threonine kinase Akt and tumor cell growth, resulting in treatment failures . Recently, we report that inhibition of sphingosine kinase 2 (SphK2) by a novel compound, ABC294640, effectively prevents and represses PEL tumor progression in vivo. Even though, novel targeted, safer and more effective strategies are urgently needed for PEL treatment.
The xc− antiporter, consisting of xCT (also named as SLC7A11) and its chaperone CD98, functions as a Na+-independent electroneutral exchange system for cystine/glutamate . Expression of xCT on the cell membrane is essential for the uptake of cystine required for intracellular glutathione (GSH) synthesis, which plays an important role in maintaining the intracellular redox balance [11, 12]. Cystine/cysteine represents an essential amino acid for many cancer cells and its uptake from the microenvironment is crucial for their growth and viability. Therefore, xCT is highly expressed by a variety of malignant tumors [13–16], and also contributes to multidrug resistance for cancer cells [17, 18].
Interestingly, xCT has been also identified as a fusion-entry receptor for KSHV and mediated KSHV entry either in isolation or as part of a complex with other receptors for the virus [19, 20]. Recently, our study has demonstrated that xCT is upregulated within more advanced Kaposi’s sarcoma (KS, another KSHV-caused malignancy) lesions containing a greater number of KSHV-infected cells . Moreover, xCT can be upregulated by KSHV-microRNAs by directly targeting BACH-1, one of the negative transcription regulators of xCT, thereby facilitating viral dissemination and persistence in the host . In the same study, we also report that xCT protects KSHV-infected endothelial cells from death induced by reactive oxygen species (ROS) . In another our recent study, we report that xCT is able to activate intracellular signaling pathways especially MAPK, cytokine release and cell invasiveness through induction of 14-3-3β protein . Even though these understandings about xCT and KSHV pathogenesis, it remains unclear whether xCT is also expressed on KSHV-infected PEL tumor cells and its functions in PEL pathogenesis especially tumor cells growth/survival and underlying complex mechanisms. More importantly, it is interested to know whether targeting xCT may represent a promising therapeutic strategy against PEL tumor progression in vivo.
xCT is highly expressed on KSHV-infected PEL cell-lines
Targeting xCT induces KSHV-infected PEL cell death/apoptosis in vitro
Inhibition of xCT reduces intracellular glutathione (GSH) but increasing reactive oxygen species (ROS) from KSHV-infected PEL cells
Akt complex activities are reduced by xCT inihibitors
Inhibition of xCT induces viral lytic gene expression from KSHV-infected PEL cells
xCT inhibitor suppresses PEL tumor progression in vivo
xCT expression is differentially regulated during oxidative stress through transcription factors binding to the cis-acting “Antioxidant Response Element” (ARE) in its promoter [35, 36]. Transcription factors that bind to the ARE include a positive regulator known as Nuclear factor erythroid 2-related factor-2 (Nrf-2)  and negative regulators, including BACH-1 and c-Maf, which competitively reduce Nrf-2 binding to the ARE thereby repressing ARE-mediated gene expression [37, 38]. We and others have confirmed that at least one KSHV-microRNA, miR-K12-11, can directly target BACH-1 to increase xCT expression [21, 39, 40]. In addition, c-Maf can also be directly targeted by several KSHV-microRNAs to promote endothelial cell reprogramming . Therefore, it is interested to understand whether these KSHV-microRNAs expressed within PEL cells represent the major contributor to xCT expression and its functions in future studies.
Our data indicate that induction of PEL apoptosis by targeting xCT is potentially through repressing intracellular GSH, increasing ROS and viral lytic gene expression. In accordance with this, Li et al. have reported that depletion of GSH and upregulation of ROS strongly induce KSHV reactivation and final cell death for KSHV-infected PEL in vitro. The authors also demonstrate that ROS is upregulated by NF-κB inhibition and is required for subsequent KSHV reactivation. However, we do not observe apparent NF-κB inhibition in MSG- or SASP-treated PEL cells when compared with vehicle-treated control (data not shown), implying NF-κB signaling is not responsible for disruption of redox balance by targeting xCT. On the other hand, it has been reported that ROS-induction by KSHV plays a causal role in KS oncogenesis by promoting proliferation and angiogenesis . Furthermore, Rac1 is overexpressed in AIDS-KS lesions and in KSHV-infected mECK36 tumors, and the antioxidant NAC was able to completely suppress Rac1-induced tumor formation in RacCA transgenic mice . Interestingly, published literature indicates a close association between ROS upregulation and Akt signaling activation in a variety of cells [44–46]. In the contrary, we have found here that Akt signaling is greatly impaired while ROS production increased in MSG- or SASP-treated PEL cells. We assume the presence of other compensatory mechanisms for ROS upregulation when Akt signaling is repressed by inhibition of xCT, although which requires further experimental validation.
In fact, xCT is involved in more cellular functions including multidrug resistance for cancer cells. The xc− transporter can mediate cellular uptake of cystine to enhance biosynthesis of glutathione, which has a major role in the protection of cells from drug-induced oxidative stress by mediating detoxification of drugs and their extrusion via multidrug resistance proteins [17, 18, 47–50]. For instance, glutathione induces a conformational change within the multidrug resistance-associated protein-1 (MRP1) and impairs its interaction with a drug and subsequent extrusion function . In a pharmacogenomics approach, Huang et al. reported that linking expression of xCT with potency of 1,400 candidate anticancer drugs identified 39 showing positive correlations, and 296 with negative correlations . Interestingly, we recently identify a membrane-protein-complex including Emmprin (CD147), LYVE-1 (a hyaluronan receptor) and BCRP (a drug-efflux pump protein), responsible for multidrug resistance of KSHV-infected PEL cells [52, 53]. Therefore, future work will focus on determining whether xCT is also involved in this protein-complex to mediate multidrug resistance for PEL. Finally, it is interested to identify more cellular genes within PEL cells potentially regulated by xCT, through analysis of the global gene profile changed due to inhibition of xCT using “-omics” technologies.
Materials and methods
Cell culture and reagents
Body cavity-based lymphoma cells (BCBL-1, KSHV+/EBVneg) and a Burkitt’s lymphoma cell line BL-41 (KSHVneg/EBVneg) were kindly provided by Dr. Dean Kedes (University of Virginia) and maintained in RPMI 1640 medium (Gibco) with supplements as described previously . The Burkitt’s lymphoma cell line BJAB (KSHVneg/EBVneg), RAMOS (KSHVneg/EBVneg), AKATA (KSHVneg/EBV+) were kindly provided by Dr. Erik Flemington (Tulane University) and cultured as described elsewhere . PEL cell line BC-1 (KSHV+/EBV+), BC-3 (KSHV+/EBVneg), and BCP-1 (KSHV+/EBVneg) cells were purchased from American Type Culture Collection (ATCC) and maintained in complete RPMI 1640 medium (ATCC) supplemented with 20% FBS. A diffuse large cell lymphoma (DLCL) cell line CRL2631 (KSHVneg/EBVneg) was purchased from ATCC and maintained in complete RPMI 1640 medium (ATCC) supplemented with 10% FBS. Primary human umbilical vein endothelial cells (HUVEC) were cultured as described previously . KSHV infection was verified for all cell lines using immunofluorescence assays for detection of the KSHV latency-associated nuclear antigen (LANA) . All cells were cultured at 37°C in 5% CO2. All experiments were carried out using cells harvested at low (<20) passages. Monosodium glutamate (MSG), Sulfasalazine (SASP) and Bay11-7082 were purchased from Sigma.
Cell viability assays
Cell viability was assessed using MTT assays for assessment of proliferative capacity, and flow cytometry for quantitative assessment of apoptosis. The standard MTT assays were performed as described previously . For flow cytometry, the FITC-Annexin V/propidium iodide (PI) Apoptosis Detection Kit I (BD Pharmingen) were used according to the manufacturer’s instructions.
Cells were lysed in buffer containing 20 mM Tris (pH 7.5), 150 mM NaCl, 1% NP40, 1 mM EDTA, 5 mM NaF and 5 mM Na3VO4. Total cell lysates (30 μg) were resolved by 10% SDS–PAGE, transferred to nitrocellulose membranes, and immunoblotted using 100–200 μg/mL antibodies, including cleaved-caspase 3/9, p-Akt/t-Akt, p-GSKα/t-GSKα, p-P70S6/t-P70S6, p-S6/t-S6, XIAP, Rac1 (cell signaling), Nox1 (Abcam), xCT, p22phox, p47phox, Nox2, Nox4 (Santa Cruz), KSHV-K8.1 (ABI). For loading controls, blots were reacted with antibodies detecting β-Actin (Sigma). Immunoreactive bands were developed using an enhanced chemiluminescence reaction (Perkin-Elmer) and visualized by autoradiography.
Immunofluorescence Assays (IFA)
Cells were incubated in 1:1 methanol-acetone at −20°C for fixation and permeabilization, then with a blocking reagent (10% normal goat serum, 3% bovine serum albumin, and 1% glycine) for an additional 30 minutes. Cells were then incubated for 1 h at 25°C with 1:2000 dilution of a mouse anti-K8.1 monoclonal antibody (ABI) followed by 1:200 dilution of a goat anti-mouse secondary antibody conjugated to Texas Red (Invitrogen). For identification of nuclei, cells were subsequently counterstained with 0.5 μg/mL 4′,6-diamidino-2-phenylindole (DAPI; Sigma) in 180 mM Tris–HCl (pH 7.5). Cells were washed once in 180 mM Tris–HCl for 15 minutes and prepared for visualization using a Leica TCPS SP5 AOBS confocal microscope.
For RNA interference assays, xCT ON-TARGET plus SMART pool siRNA (Dharmacon), or negative control siRNA, were delivered using the DharmaFECT transfection reagent according to the manufacturer’s instructions. To confirm initial transfection efficiency for siRNA experiments, PEL cells were transfected with green fluorescent protein (GFP)-tagged siRNA, and GFP expression determined by flow cytometry 24 h later as described previously . Three independent transfections were performed for each experiment, and all samples were analyzed in triplicate for each transfection.
Total RNA was isolated using the RNeasy Mini kit according to the manufacturer’s instructions (QIAGEN). cDNA was synthesized from equivalent total RNA using SuperScript III First-Strand Synthesis SuperMix Kit (Invitrogen) according to the manufacturer’s procedures. Primers used for amplification of target genes are displayed in Additional file 2: Table S1. Amplification was carried out using an iCycler IQ Real-Time PCR Detection System, and cycle threshold (Ct) values were tabulated in duplicate for each gene of interest in each experiment. “No template” (water) controls were used to ensure minimal background contamination. Using mean Ct values tabulated for each gene, and paired Ct values for β-actin as an internal control, fold changes for experimental groups relative to assigned controls were calculated using automated iQ5 2.0 software (Bio-rad).
Measurement of virion production
BCBL-1 cells (10 mL) were treated by vehicle, MSG (20 mM), SASP (0.5 mM) and valproic acid (1.5 mM as a positive control), respectively, for 48 h, then followed by centrifugation at 1200 rpm for 5 min and 3000 rpm for 20 min. The clear supernatants were collected followed by ultracentrifugation at 30000 g for 2 h, and virion pellet were resuspended at 100 μL of fresh medium. The HUVEC cells were infected with these virion suspensions as described previously  and followed by qRT-PCR measurement of Lana transcripts mentioned above.
Vehicle-, MSG- and SASP-treated PEL cells were loaded with 10 μM of the ROS dye c-H2DCFDA (Invitrogen) for 30 min at 37°C in Hanks’ Balanced Salt Solution (HBSS) containing calcium and magnesium (HBSS/Ca/Mg). Cells will be washed once with HBSS/Ca/Mg to remove dye, resuspended in HBSS/Ca/Mg and subjected to flow cytometry analyses as described elsewhere . To block ROS production, PEL cells were treated with MSG, SASP or vehicle in the presence or absence of the antioxidant N-acetylcysteine (NAC) 2.5 mM for 48 h, then cell apoptosis was assessed as described above.
Cells were resuspended in staining buffer (3 BSA in 1× PBS) for 20 minutes, then incubated on ice for 30 min with 1:20 dilution of primary antibody xCT (Santa Cruz). Following two subsequent wash steps, cells were incubated for an additional 30 min with 1:200 dilution of secondary antibodies (Invitrogen) including Donkey anti-goat IgG Alexa Fluor 647. Controls included cells incubated with the secondary antibody only. Cells were resuspended in 1× PBS and analyzed using a FACS Calibur 4-color flow cytometer (BD) and FlowJo software (TreeStar) to quantify cell surface localization of target proteins.
NADPH oxidase activities assays
The chemiluminescence-based NADPH oxidase activities assays were performed as described previously with modifications . After drug-treatment, cells were centrifuged at 500 g for 10 min at 4°C. The cell pellet was resuspended with 35 μL ice-cold lysis buffer and kept on ice for 20 min. To a final 200 μl of HBSS/Ca/Mg buffer containing NADPH (1 μM, Sigma) and lucigenin (20 μM, Sigma), 5 μl of cell lysates was added to initiate the reaction for 5 min at 37°C. Chemiluminescence was measured immediately using a Synergy HT microplate reader (BioTek Instruments).
Intracellular GSH measurement
The intracellular GSH levels in vehicle-, MSG- and SASP-treated PEL cells were quantified using the GSH-Glo™ Glutathione Assay Kit (Promega), according to the manufacturer’s instructions.
PEL xenograft model
BCBL-1 cells maintained at early passage number in cell culture were washed twice in sterile-filtered PBS prior to performance of trypan blue and flow cytometry assays for verification of their viability. Aliquots of 107 viable cells were diluted in 200 μL sterile PBS, and 6–8 week-old male non-obese diabetic/severe-combined immunodeficient (NOD/SCID) mice (Jackson Laboratory, Taconic Inc.) received intraperitoneal (i.p.) injections with a single cell aliquot. SASP solutions were prepared at 20 mg/mL, dissolved in 0.1 N NaOH in PBS at pH 7.2, and sterile-filtered prior to in vivo administration. The SASP (150 mg/kg body weight), or vehicle alone, was administered using an insulin syringe for i.p. injection. Drug was administered 24 h after BCBL-1 injection, once daily for 5 days/week. Two experiments, with 8 mice per group for each experiment, were performed. The PEL expansion in vivo was confirmed by testing the expression of cell-surface markers including CD45, CD138, EMA and viral protein LANA in nuclear within ascites tumor cells, using IFA and flow cytometry as described in our previous publications . Weights were recorded weekly as a surrogate measure of tumor progression, and ascites fluid volumes were tabulated for individual mice at the completion of each experiment. All protocols were approved by the Louisiana State University Health Science Center Animal Care and Use Committee in accordance with national guidelines.
Significance for differences between experimental and control groups was determined using the two-tailed Student’s t-test (Excel 8.0).
This work was supported by grants from the National Institutes of Health (R01-CA142362), a Center for Biomedical Research Excellence Award (P20-RR021970), the SOM Research Enhancement Funding (5497400038), the Ladies Leukemia League Grant (2014-2015), the National Natural Science Foundation (81272191), the NNSF for Young Scientists of China (81101791) and the Foundation for Innovative Research Groups of the National Natural Science Foundation of China (81221001). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
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