Ochratoxin a mediates MAPK activation, modulates IL-2 and TNF-α mRNA expression and induces apoptosis by mitochondria-dependent and mitochondria-independent pathways in human H9 T cells

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Correspondence: Youssef Darif (E-mail: darifyoussef2000@yahoo.fr)

Ochratoxin A mediates MAPK activation, modulates IL-2 and TNF-α mRNA expression and induces apoptosis by mitochondria-dependent and mitochondria-independent

pathways in human H9 T cells

Youssef Darif¹, Driss Mountassif², Abdelkarim Belkebir¹, Younes Zaid³, Kaustuv Basu², Walid Mourad⁴ and Mounia Oudghiri¹

1Laboratory of Physiology and Molecular Genetics, Immunology Unit, Faculty of Sciences, Hassan II Ain Chock University, KM 8, Route El Jadida BP., 5366, Casablanca, Morocco

2Department of Anatomy and Cell Biology, McGill University, 3640 University Street, Montreal, Quebec, H3A 0C7, Canada

3Laboratory of Thrombosis and Hemostasis, Montreal Heart Institute, Montreal, Quebec, Canada

4Laboratoire d’immunologie cellulaire et moléculaire, Centre de Recherche-Centre Hospitalier de l’Université de Montréal (CR-CHUM), 900, rue St-Denis, Montréal, Quebec, H2X 0A9, Canada

(Received December 23, 2015; Accepted March 25, 2016)

ABSTRACT — Ochratoxin A (OTA) is a natural fungal secondary metabolite that contaminates food and animal feed. Human exposure and involvement of this mycotoxin in several pathologies have been demonstrated worldwide. We investigated OTA immunotoxicity on H9 cells, a human cutaneous CD4+

T lymphoma cell line. Cells were treated with 0, 1, 5, 10, and 20 μM OTA for up to 24 hr. Western blotting revealed increased phosphorylation of all three major mitogen-activated protein kinases (extracellular signal–regulated kinase, c-Jun amino-terminal kinase, p38). OTA triggered mitochondrial transmembrane potential loss and caspase-3 activation. The 24-hr OTA treatment caused marked changes in cell morphology and DNA fragmentation, suggesting the occurrence of apoptotic events that involved a mitochondria- dependent pathway. Moreover, OTA triggered significant modulation of survivin, interleukin 2 (IL-2) and tumor necrosis factor α (TNF-α): mRNA expression of survivin and IL-2 were decreased, while TNF-α was increased. OTA also caused caspase-8 activation in a time-dependent manner, which evokes the death receptor pathway activation; we suspect that this occurred via the autocrine pro-apoptotic effect of TNF-α on H9 cells.

Key words: OTA, TCD4+, Apoptosis, Mitochondrial pathway, Cytokines modulation, TNF-α

INTRODUCTION

Ochratoxin A (OTA) is a natural mycotoxin produced mainly by Aspergillus ochraceus and Penicillium ver- rucosum; it was described as the major agent in Bal- kan endemic nephropathy (Kuiper-Goodman and Scott, 1989). Following the evidence of OTA toxicity in mammalian species, the International Agency for Research on Cancer has classified OTA as a possible carcinogen (Group 2B) for humans (IARC, 1993; Malir et al., 2013).

OTA consists of a chlorinated dihydroisocoumarin moi- ety linked to one l-phenylalanine molecule by a peptide bond through its 7-carboxyl group. This structure inhib-

its protein synthesis by competing with phenylalanine in the phenylalanine-tRNA aminoacylation reaction (Baudrimont et al., 1997).

The ubiquitous occurrence of OTA as a food contami- nant and its high stability make avoidance in dietary intake very difficult (El Khoury and Atoui, 2010), the high- est level of contamination reported was 80 mg OTA/kg in mouldy bread intended for animal feeding. Human exposure via food and beverages has been estimated to be 15-60 ng OTA/kg b.w. per week for adult consumer (Heussner and Bingle, 2015): however dietary intake of 1.21 μg per day of OTA has been associated with endemic nephropathy in Balkan countries, with serum concen-

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tration of OTA ranged between 5 and 50 ng/mL. Exposed persons develop DNA adducts in renal tissue, karyome- galy in proximal and distal tubular epithelial cells and tumors. In Italy a case of acute renal failure was reported in a person who worked for 8 hr in a farm granary that had been closed for several months; the A. ochraceus producing OTA was isolated from the wheat and intox- ication by OTA inhalation was suspected (Reddy and Bhoola, 2010).

OTA is the most frequently detected mycotoxin in human plasma, has a long half-life of 35 days and is considered a cumulative toxin in which chronic exposure is more frequent and more hazardous than acute exposure. In fact, upon its absorption from the gastrointesti- nal tract, OTA binds rapidly to serum proteins, due to the high affinity of the human serum albumin to OTA with a Kb ~5 × 10⁶ M^-1. In addition the reabsorption during the enterohepatic circulation reduces its elimination and promotes bio-accumulation in the organs (Bow et al., 2006).

Following oral intake of low to moderate amounts of OTA, the mucosal immunity which acts as the first obsta- cle for pathogens will be adversely affected, due to the decrease in the percentage of lymphocytes, macrophages and the modulation of cytokine secretion (Müller et al., 1999). In vivo studies suggest that low to moderate concentrations of OTA could also influence human susceptibility to infectious diseases (O’Brien and Dietrich, 2005).

Recently, in Khayelitsha, South Africa, it has been shown the risk of HIV transmission to newborns increases by up to 11-fold in infants receiving a mixed diet (breast milk with formula/cereals). The presence of OTA in the mixed diet was suspected to induce mucosal inflammation and a positive correlation was found between OTA plasma levels and the importance of CD4+ T lymphocyte population in oral mucosa (Wood et al., 2013).

To date, only a few studies have investigated the effect of OTA on the human immune system; Lea et al. (1989) reported that 12.5 μM of OTA is sufficient to complete- ly abolish the peripheral blood lymphocyte proliferation, in response to the mitogen-stimulator. The decreased proliferation was correlated with the impairment of IL-2 production and IL-2 receptor expression in activated T cells (Lea et al., 1989). Later the same researchers showed that DNA synthesis inhibition occurs when T cells are exposed to 6.4 μM of OTA while protein synthesis inhibition occurs at 12.5 μM, which is suspected as the OTA immunosuppresor mechanism (Størmer and Lea, 1995).

Other studies indicate that OTA-immunomodulation is instead responsible for lymphoproliferation decrease. It has been shown that low concentrations of OTA stimu- late the expression of CD69 and CD25 markers on the

cell surface and IL-6 production on lymphocytes after mitogenic stimulation, while higher concentrations inhib- it the expression of CD69, CD25, CD71 and TNF-α production (Marin and Taranu, 2015; Köhler et al., 2002).

Other studies have shown that the incubation of human peripheral blood lymphocytes with OTA triggers apoptosis through the mitochondrial pathway and decreases expression of the pro-survival protein Bcl-xL, (Assaf et al., 2004; Hsuuw et al., 2013).

Human blood represents a reservoir in which the T4 lymphocytes are chronically exposed to OTA; these cells play a pivotal role in both cellular and humoral immune responses. However despite the strong immunosuppressive potential of OTA, there are only a few papers describing the events following the exposure of human T4 lymphocytes to OTA and the mode of action remains unclear. To further understand the mechanism of OTA-induced apoptosis in human T lymphocytes, we investigated the succession of apoptotic events in H9 cells, a human cutaneous CD4+ T lymphoma cell line, by evaluating the effects of OTA on mitochondrial transmembrane potential (∆Ψm), changes in cell morphology, caspase-3 activation and DNA degradation. We further examined the effects on the stimulation of mitogen-activated protein kinases (MAPKs), modulation of survivin, IL-2 and tumor necrosis factor α (TNF-α) expression, and caspase-8 activation.

MATERIALS AND METHODS Cell line and culture conditions

The H9 cell line (HTB-176™, ATCC; Manassas, VA, USA) was grown in 75-cm² tissue culture flasks with RPMI 1640-glutamine medium (Invitrogen, Burlington, ON, Canada) supplemented with 10% fetal bovine serum (inactivated at 56°C for 30 min) and 1% penicillin (100 IU/mL)/streptomycin (100 μg/mL). Cells were grown at 37°C in 5% CO₂ and 95% air; cultures were split 1:3 every 4 days.

Reagents and antibodies

OTA (Cat. No. 01877, Sigma-Aldrich, St. Louis, MO, USA) was dissolved in sterile 0.1 M NaHCO₃ solution (pH 7.4) to yield a stock solution of 1 mM OTA and added to cultures at the indicated times and concentrations.

Unless stated otherwise, all chemicals and reagents were of analytical grade and purchased from Sigma-Aldrich.

RPMI 1640 medium, fetal bovine serum and penicillin/

streptomycin were obtained from Gibco Laboratories (Grand Island, NY, USA).

The primary antibodies (Abs) used were rabbit antibodies against phosphorylated (phospho)-p38, p38, phospho-

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extracellular signal-regulated kinase (ERK)1/2, ERK1/2, phospho-c-Jun amino-terminal kinase (JNK), JNK, rabbit monoclonal anti-active caspase-8 clone 18C8 (Cell Signaling Technology, Beverly, MA, USA), and rabbit monoclonal fluorescein isothiocyanate (FITC)-conjugated anti-active caspase-3 clone C92-605 (BD Biosciences, San Jose, CA, USA). The secondary Ab was goat anti-rabbit horseradish peroxidase-conjugated immunoglobulin G (Santa Cruz Biotechnology, Santa Cruz, CA, USA).

Cell viability measurement

Cell viability was assessed with propidium iodide (PI) staining. PI penetrates dead and damaged cells; in contrast, live cells with intact membranes exclude PI and appear less fluorescent than dead cells under 488 nm exci- tation. H9 cells were incubated with 0 (control), 1, 5, 10 and 20 μM OTA for 3, 6, 12 and 24 hr. At the end of each indicated incubation period, 5 × 10⁵ cells were collected, washed with phosphate-buffered saline (PBS) and stained with PI at a final concentration of 5 μg/mL for 10 min in the dark. PI fluorescence of individual cells was measured by flow cytometric analysis using an LSR II flow cytometer and FACS Diva software (Becton Dickinson, San Jose, CA, USA).

Transmission electron microscopy analysis of morphological apoptosis

Treated and control H9 cells were centrifuged in 0.4-mL microcentrifuge tubes at 1,500 rpm at room temperature for 15 min to obtain a moderately firm pellet.

The supernatant was replaced with buffered 2.5% glutar- aldehyde (pH, 7.2-7.4), resuspended and fixed at room temperature for at least 60 min. The pellets were further treated with 1% osmium tetroxide for 1 hr, followed by three washes in cacodylate buffer to remove the osmium tetroxide. Graded alcohol dehydration (50% to 100%) was performed, with each step lasting 10 min. The pellets underwent two more washes with propylene oxide, followed by centrifuging each time. Finally, the pellets were mixed with 50% propylene oxide and 50% Epon 812 for a minimum of 1 hr, followed by 1 hr in 100% Epon 812.

The pellets were further embedded in fresh Epon and polymerized in an oven at 60°C for at least 48 hr. The blocks were put in chuck and mounted on an ultrami- crotome (Leica Microsystems, Richmond Hill, Canada).

Ultrathin sections (80 nm) were cut using a diamond knife (Diatome AG, Biel, Switzerland). The sections were collected in 200-mesh Formvar-coated grids and stained with 2% uranyl acetate and lead citrate (5 min each) for contrast. Micrographs were acquired at 120 kV in a Philips Tecnai 12 transmission electron microscope (FEI

Company, Eindhoven, the Netherlands) using a Gatan CCD 4Kx4K camera (Gatan, Inc., Pleasanton, CA, USA).

Detection of apoptotic DNA ladder

After OTA treatment, cell pellets were collected, resuspended in 0.5 mL lysis buffer (5 mM Tris, 20 mM EDTA [pH 8], 0.5% Triton X-100) and centrifuged for 20 min at 12,000 rpm (Eppendorf F-45-24-11 Fixed Angle Rotor, r =83 mm) at 4°C. The supernatant was collected and incubated with proteinase K and RNase A (DNase-free) at 56°C for 2 hr. After DNA was extracted using phenol/

chloroform/isoamyl alcohol and centrifuged for 30 min at 6,000 rpm, it was precipitated overnight with 0.1 volume 3 M sodium acetate (pH 5.2) and 2 volumes 100%

ethanol at -20°C. After 30-min centrifugation at 12,000 rpm, the pellet was washed twice with 0.1 mL 70% ethanol, dried and resuspended in sterile deionized water. The quality and quantity of extracted DNA were determined using a Synergy Microplate Reader with Gen5 Data Analysis Software (Bio-Tek Instruments, Inc., Winooski, VT, USA). Five micrograms from 10 μL of each treatment were loaded onto 1.5% agarose gels containing 350 μg/mL ethidium bromide. After separation, bands were visualized using a Universal Hood II Gel Imager and Quantity One software, version 4.5.2 (Bio-Rad, Montreal, QC, Canada).

Annexin V/PI staining

The percentage of early apoptotic cells was determined through analysis of phosphatidylserine (PS) translocation and plasma membrane integrity. H9 cells were seed- ed in 12-well plates at 1 × 10⁶ cells/well and incubated with 0, 1, 5, 10 or 20 μM OTA for 12 hr. Double staining with FITC-conjugated annexin V (AV) and PI was performed as follows: after being washed with PBS, pellets were resuspended in 200 μL binding buffer (10 mM HEPES/NaOH [pH 7.4], 140 mM NaCl, 2.5 mM KCl).

AV was added at a final concentration of 1 μg/mL for 15 min in the dark. Cells were then washed, centrifuged at 300 rpm for 5 min, and resuspended in 200 μL binding buffer. Before flow cytometric analysis, 10 μL PI (100 μg/

mL in binding buffer) was added to each sample. AV⁺/PI^- cells were early apoptotic cells. As a positive control for apoptosis, cells were incubated with 1 μM staurosporine for 12 hr.

Mitochondrial transmembrane potential

Variation of ∆Ψm was studied by evaluating the changes in fluorescence intensity of cells stained with the cationic dye JC-1 (5,5′, 6,6′-tetrachloro-1,1′,3,3′ tetraethylbenzimidazolo-carbocyanine iodide). H9 cells

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(3 × 10⁵ cells/mL) were incubated with 10 μg/mL JC-1 for 15 min at 37°C in the dark and analyzed by flow cytometry. Changes from red (JC-1 aggregate, emission:

590 nm) to green fluorescence (JC-1 monomer, emission:

525 nm) indicated mitochondrial membrane depolarization. Decreases in the red/green fluorescence intensity ratio were evaluated on histograms.

Intracytoplasmic staining of active caspase-3 Active caspase-3 was measured by flow cytometry with an FITC-conjugated monoclonal active caspase-3 Ab apoptosis kit (BD Biosciences) according to the manufacturer’s instructions. Briefly, after washing with cold PBS, 1 × 10⁶ cells were fixed and permeabilized in Cyto- fix/Cytoperm buffer for 20 min on ice. Then, the cells were washed and incubated with anti-active caspase-3 Ab for 30 min at room temperature, washed and analyzed by flow cytometry.

Polyacrylamide gel electrophoresis and immunoblot analysis

After OTA treatment, cells were lysed using lysis buffer (10 mM Tris [pH 7.5], 150 mM NaCl, 5 mM EDTA) with 1% Triton X-100, 2 mM Na₃VO₄ and 1 mM protease inhibitors (Roche Diagnostics, Montreal, QC, Canada).

Lysates from each sample were adjusted to equal protein concentrations and added (v/v) to 2× Laemmli buffer.

After boiling for 5 min at 95°C, proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (4%-12% polyacrylamide gels) and transferred to polyvinylidene difluoride membranes (Immobilon-P;

Millipore, Ontario, Canada). Subsequently, the membranes were blocked with 5% skim milk in Tris-buffered saline with 0.05% Tween 20 for 30 min.

Procaspase-8 cleavage and p38, ERK1/2 and JNK phosphorylation were assessed by immunoblotting using 18C8 (anti–active caspase-8, Asp391) and phospho-specific Abs, according to the manufacturer’s instructions, followed by washing and labeling with the secondary Ab at room temperature. The blots were visualized using enhanced chemiluminescence (Amersham/GE Healthcare Biosciences, Piscataway, NJ, USA). Each phosphorylation blot was stripped and reprobed with a total Ab (p38, ERK1/2 or JNK) to verify the endogenous protein levels.

Semi-quantitative and real-time quantitative RT-PCR

Total RNA from each treatment was extracted from 5 × 10⁵ cells using TRIzol reagent (Invitrogen). After quantification with spectrophotometry, complementary DNA was synthesized by reverse transcription from 1 μg total RNA using oligo(dT)_12-18 primers, RNaseOUT and Moloney murine leukemia virus reverse transcriptase in 20 μL reactions according to the manufacturer’s protocol (Invitrogen). The level of survivin mRNA was assessed with semi-quantitative reverse transcription-polymerase chain reaction (RT-PCR) as described previously (Granziero et al., 2001); Table 1 lists the sequences of the forward and reverse primers for survivin and β-actin (internal control gene). IL-2 and TNF-α mRNA levels were assessed with real-time RT-PCR using a Fast SYBR Green kit (Roche) and a Rotor-Gene 3000 real-time ther-

Table 1. Primers used to monitor the expression of survivin and cytokine genes in reverse transcription-polymerase chain reaction or real-time polymerase chain reaction analyses.

Gene Primer sequence (sense, antisense) PCR product size (bp)

Survivin^a 5′-CTCTACATTCAAGAACTGGCC-3′,

5′-TTGGCTCTTTCTCTGTCCAG-3′ 342

β-Actin^{a, c} 5′-AATCTGGCACCACACCTTCT-3′,

5′-TAATGTCACGCACGATTTCC-3′ 386

IL-2^b 5′-TACAACTGGAGCATTTACTG-3′,

5′-GTTTCAGATCCCTTTAGTTC-3′ 268

TNF-α^b 5′-CAGAGGGCCTGTACCTCATC -3′,

5′-GGAAGACCCCTCCCAGATAG-3′ 219

S9^{b, c} 5′-CGTCTCGACCAAGAGCTGA-3′,

5′-GGTCCTTCTCATCAAGCGTC-3′ 133

aAnalyzed with RT-PCR.

bAnalyzed with real-time PCR.

cInternal control.

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mal cycler (Montreal Biotech, Montreal, QC, Canada).

Table 1 lists the specific primers for IL-2, TNF-α and the ribosomal protein S9 (S9, internal control gene). Rela- tive fold expression values were determined using the comparative threshold cycle (∆∆Ct) method and normal- ized to S9. Standard curves were established with ampli- fications of serial dilutions of pCR2.1-TOPO-S9 plasmid (Soucy-Faulkner et al., 2010).

Statistical analysis

Data from at least three independent experiments were analyzed using Prism-6 software (version 6.04). For cell viability, LC₅₀ values were calculated by linear regression from experimental data, using trendline equation, where 50 = aX + b. Statistical analysis involving two means was performed by an unpaired Student’s t test, whereas com- parisons of more than two means was made using one- way ANOVA followed by Tukey’s multiple comparison

test. Results are presented as the mean ± S.E. A p-value

< 0.05 was considered significant, and p < 0.01 was considered highly significant when compared to the corresponding control.

RESULTS OTA affected H9 cell viability

We investigated the cytotoxicity of OTA on H9 cells using the PI assay (Fig. 1). The effect of OTA on loss cell viability was most obvious after 24 hr of treatment, whereas 1 μM OTA did not exert a significant loss of cell viability at any time point (p < 0.05). There was an early decrease in relative cell viability after 3 hr and 6 hr of treatment at 10 μM OTA. However, after 12 hr of treatment, the decrease became more evident in cells treated with 5, 10 and 20 μM OTA. The loss in H9 cell viability was OTA dose- and time-dependent, which sug- gests OTA toxicity through apoptosis and/or necrosis (see Supplementary Fig. 1).

Morphological changes in OTA-treated H9 cells Based on the results of cell viability, ultrastructural study was performed using transmission electron microscopy (TEM) on normal control H9 cells and treated cells with 1, 5, 10 and 20 μM OTA for 24 hr, when the cell mortali- ty was more visible. Control cells were irregularly round- ed with well-defined outlines and contained abundant cytoplasm, many mitochondria with clear transverse cristae and uniform nuclei (Fig. 2A). While in treated cells, TEM revealed nuclei deformation like crescent (Fig. 2B), shrinkage, smaller size, dense cytoplasm and more tightly packed organelles in cells that were incubated with OTA (Fig. 2C).

The mitochondria were slightly swollen, while the nuclei underwent the karyopyknosis phenomenon, eventually followed by several fragmentations (Fig. 2D). There was extensive plasma membrane blebbing, followed by pyk- nosis, which is the result of chromatin condensation (Fig.

2E), and karyorrhexis with separation of cell fragments into apoptotic bodies in a process called budding. Electron- dense nuclear material aggregates peripherally under the nuclear membrane fragment (Fig. 2F). The ultrastructural examination of OTA-treated H9 cells revealed many hallmarks of the apoptotic process (Wyllie et al., 1980).

DNA damage in OTA-treated H9 cells

The induction of apoptosis led to nuclease activation, which cleaved chromosomal DNA into oligonucleosomal fragments of approximately 200 bp, forming the characteristic “stepladder” visible following agarose gel separation. Figure 3 depicts the DNA ladder in OTA-treated Fig. 1. Effect of OTA on H9 cell viability was dose-and time-

dependent. (A) H9 cells were incubated with 0, 1, 5, 10 and 20 μM OTA for different durations. Data are presented as the mean ± S.D. from three independent experiments and each sample were run in duplicate.

The overall comparison of the slopes (3, 6, 12 and 24 hr) vs. T0 shows a significant difference (p < 0.05), P: 0.0379; 0.0137; 0.0011; 0.0053 respectively. (B) Corresponding LC₅₀ of each incubation time was determined as the concentration inducing 50% of cell viability loss, using linear regression and trendline equation.

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(B)

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Fig. 2. Electron micrographs of ultrastructural damage in H9 cells caused by different concentrations of OTA. After 24-hr incu- bation with 0, 1, 5, 10 or 20 μM OTA, cells were fixed and observed under electron microscopy. (A) Control (× 7200).

(B) Marginalization of organelles; nuclei-like crescent moon body (× 7200). (C) Cell shrinkage and nuclear fragmentation forming two nuclei with condensed chromatin structure (× 7800). (D) Apoptotic body derived from membrane blebbing and swollen mitochondria (× 6500). (E) Pronounced shrinkage and chromatin condensation (× 9500). (F) Breakdown of cytoplasmic organelles, destruction of the nucleus, and appearance of undefined clumps of nuclear chromatin (× 8000).

Bar = 1 μm.

Fig. 3. Oligonucleosomal DNA fragmentation in OTA-treated H9 cells. Cells were incubated with 1, 5, 10 or 20 μM OTA for 24 hr.

The cells were lysed and DNA was extracted. Control: H9 cells incubated in the same conditions but without OTA. For each sample, 5 μg DNA was electrophoresed in a 1.5% agarose gel. DNA size marker: 1 Kb Plus DNA Ladder (Life Technologies). Data are representative of three independent experiments. OTA, ochratoxin A.

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H9 cells. After 24-hr incubation with OTA, the DNA ladder was slightly visible in cells treated with 1 μM OTA, whereas it was clearly visible in cells treated with 5, 10 and 20 μM OTA, and there was uniform distribution of the 200-1000 bp oligonucleosomal DNA fragments. Fur- thermore, there was a marked increase in DNA laddering following treatment with 20 μM OTA, with complete dis- appearance of high-molecular weight DNA corresponding to intact genomic DNA. In contrast, control cells did not yield the characteristic oligonucleosomes, only intact genomic DNA. The DNA “stepladder” is commonly recognized as a consequence of apoptosis (Peitsch et al., 1993). In the present study, OTA caused DNA fragmentation in H9 cells, confirming the ability of OTA to induce

apoptosis in this cell line.

Discrimination between apoptotic and necrotic cells

Translocation of PS to the exterior of the plasma membrane is one of the earliest events in apoptosis. AV, which has a high specific affinity for PS, tracks its translocation;

PI enables verification of plasma membrane integrity because it is excluded by viable cells. Figure 4 illustrates AV/PI double staining of H9 cells after 12 hr of treatment with 0, 1, 5, 10 and 20 μM OTA. The percentage of early apoptotic cells (AV⁺/PI^-) following treatment with 0, 1, 5, 10 and 20 μM OTA was 1.7 ± 0.38%, 2.7 ± 0.67%, 5.0

± 0.99%, 6.7 ± 0.51% and 3.2 ± 0.61% respectively; that

Fig. 4. Annexin V-FITC/PI staining in H9 cells at 12 hr post-treatment with different OTA concentration (0, 1, 5, 10 and 20 μM).

(A) Dot plot for flow cytometric analysis of apoptotic cells. , AV^-/PI^-: viable cells (lower left quadrant), AV^-/PI⁺: necrotic cells (upper left quadrant), AV⁺/PI⁺: late apoptotic cells (upper right quadrant), AV⁺/PI^-: early apoptotic cells (lower right quadrant). (B) Bar graph quantifying the percentage of, early apoptotic, late apoptotic and necrotic cells according to OTA treatment. The values shown are the mean of three independent experiments ± S.E. and samples were run in duplicate. Con- trol: treatment without OTA (negative control), Stauro: treatment with 1 μM staurosporine for 12 hr (positive control) of apoptosis. AV, FITC-conjugated annexin V; PI, propidium iodide.

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of late apoptotic cells (AV⁺/PI⁺) was 10.7 ± 0.07%, 10.6

± 0.07%, 18.0 ± 1.23%, 23.8 ± 0.83% and 20.8 ± 0.60%, respectively, while that of necrotic cells (AV^-/PI⁺) was 2.0

± 0.66%, 2.1 ± 0.98%, 3.8 ± 0.93%, 12.5 ± 1.31% and 23.7 ± 5.67%, respectively. As compared to the control, 1 μM OTA induce slight increase in early apoptotic cells (2.7 ± 0.67% vs 1.7 ± 0.38%), whereas the percentages of late apoptotic and necrotic cells were similar to that of the control cells, which may be explained by the presence of spontaneous apoptotic cells in the population. In cells treated with 5 and 10 μM OTA, there was a significant (p < 0.05) increase in the percentages of early apoptotic/late apoptotic/necrotic cells, confirming the apop-

totic effect of OTA on H9 cells. However, there was an increased percentage of necrotic cells in cells incubated with 20 μM OTA for 12 hr versus a decrease in early apoptotic and late apoptotic cells. This suggested that at high OTA concentrations (i.e., 20 μM), necrosis takes precedence over apoptosis in terms of cell death.

OTA induces mitochondrial membrane permeabilization and caspase-3 activation

To investigate whether OTA-induced apoptosis in H9 cells takes place through the mitochondrial apoptotic pathway, we measured the loss of ∆Ψm using the potential sensor JC-1 and flow cytometry analysis. JC-1 stains

Fig. 5. Loss of mitochondrial membrane potential. H9 cell ∆Ψm was determined after incubation with 0, 1, 5 and 10 μM OTA for the in- dicated periods by JC-1 staining. (A) Representative dot plot of JC-1 aggregates and JC-1 monomers from incubation with 0 and 5 μM OTA. (B) JC-1 aggregate (red fluorescence) alterations to JC-1 monomers (green fluorescence) are presented as the percentage of JC-1 ratio (red/green fluorescence) relative to control cells at 0 hr (treatment without OTA). Stauro: cells treated with 1 μM staurosporine. Data are the means ± S.E. from three independent experiments, and each experiment was carried out in duplicate. *p < 0.05; **p < 0.01 (versus control at indicated time). OTA, ochratoxin A; JC-1, 5,5′, 6,6′- tetrachloro-1,1′,3,3′ tetraethylbenzimidazolo-carbocyanine iodide.

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high-∆Ψm mitochondria by emitting orange-red fluorescence as a consequence of the presence of JC-1 aggregates. In cells with depolarized or damaged mitochondria, the ∆Ψm collapses and JC-1 remains in monomeric form, which emits green fluorescence. Figure 5B illustrates the loss of ∆Ψm in H9 cells as presented by the decreased relative JC-1 ratio (aggregate/monomer). Increas- ing the OTA concentration led to a significant decrease (p 0.014) in the relative ratio only after 6-hr incubation (1 μM OTA, 90.2% ± 0.94% versus 5 μM OTA, 65.5%

± 2.01%). The decrease of the relative ratio continued over time for all OTA concentrations. There was a slight decrease following treatment with 1 μM OTA (76.9% ± 5.89%) and a near complete loss of ∆Ψm following treatment with 5 μM (11.6% ± 2.45%) and 10 μM OTA (7.5%

± 4.91%), which resembled that obtained following 24 hr of treatment with 1 μM staurosporine (2.6% ± 2.02%).

These results indicate that OTA can induce loss of ∆Ψm in a time- and dose-dependent manner; the loss of ∆Ψm reflects the disruption of mitochondrial membrane perme- ability and the subsequent release of apoptogenic factors into the cytosol and activation of caspases.

Activation of caspase-3 is one of the critical steps in the execution of programmed cell death; it commonly occurs in apoptosis of different cell types, resulting

from specific cleavage of procaspase-3 and release of the active fragment into the cytosol (Peterson et al., 2010). It appeared that OTA activated caspase-3 in a dose-dependent manner following 24-hr incubation (Fig. 6). Cells treated with 1 μM OTA exhibited active caspase-3 levels similar to that of untreated control cells, while there was a 5.6-fold increase in active caspase-3 levels in cells treated with 5 μM OTA. Treatment with 10 μM and 20 μM OTA led to 7.8- and 8.2-fold increases, respectively, in active caspase-3 levels. The decreased rate of caspase-3 activation following treatment with 10 μM and 20 μM OTA confirmed the results of the AV/PI assay (between 10 μM and 20 μM OTA), where the mode of cell death shifted from apoptosis to necrosis. In fact, based on the results, it appears that the mitochondria play an important role in OTA-induced apoptosis in H9 cells. Incubation for 24 hr to 1 μM OTA did not result in the hallmarks of apoptosis, indicating that at low doses, OTA requires more time to generate the hallmarks of apoptosis in H9 cells.

OTA led to MAPK activation in H9 cells

To gain insight into the early events of OTA-induced apoptosis in H9 cells, we evaluated the response of the signaling pathways of three major MAPKs. Cells were treated with 1 μM OTA for 1; 3; 6; 9; 12 and 24 hr at this

Fig. 6. Active caspase-3 in H9 cells after 24-hr incubation with OTA. Intracytoplasmic staining was performed with mAb against active caspase-3 after 24-hr incubation with 1, 5, 10 and 20 μM OTA; the percentage of active caspase-3-positive cells was measured by flow cytometry. Control: treatment without OTA. Stauro: treatment with 1 μM staurosporine for 24 hr. Results shown are from a representative experiment; an additional experiment produced similar results.OTA, ochratoxin A.

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concentration the major apoptotic events did not yet take place. We performed immunoblot analysis following different periods (Fig. 7) to identify the duration required to activate p38, ERK1/2 and JNK. Incubating H9 cells with 1 μM OTA induced p38, ERK1/2 and JNK phosphorylation that peaked after 6 hr and was subsequently main- tained for the next 18 hr (Fig. 7).

Inhibition of survivin mRNA expression

We next investigated the resistance of H9 cells to OTA- induced apoptosis using survivin, a member of the human IAP (inhibitor of apoptosis protein) family; After 12 hr of incubation, 10 μM OTA decreased the level of survivin mRNA in H9 cells, whereas at 1 μM and 5 μM OTA, survivin mRNA expression levels were apparently unaffect- ed and remained similar to that of untreated cells (Fig. 8).

After 24 hr, there was an obvious decrease in survivin mRNA levels in cells treated with 5 μM and 10 μM OTA.

This indicated that OTA could also promote apoptosis in H9 cells by inhibiting survivin expression in a time- and dose-dependent manner.

Modulation of IL-2 and TNF-α mRNA expression We then analyzed the ability of OTA to modulate the expression of the cytokines IL-2 and TNF-α in H9 cells.

Real-time PCR revealed decreased IL-2 mRNA expression (Fig. 9A): in H9 cells incubated with 1, 5 and 10 μM

OTA, IL-2 mRNA levels decreased by 11%, 8% and 72%

(p < 0.05), respectively, when compared to control cells incubated for 12 hr in medium only. The decrease was more significant between 12 hr and 24 hr in cells incubated with 1, 5 and 10 μM OTA (p < 0.01), decreasing by 73%, 84% and 81%, respectively (Fig. 9A). In contrast, OTA stimulated TNF-α mRNA expression (Fig.

9B). There was a 2-fold induction of TNF-α promoter as compared to control cells after 12 hr of incubation with 10 μM OTA, whereas 24 hr of incubation with 10 μM OTA led to a strong, 8-fold induction of TNF-α promoter (p < 0.05). However, both 12 hr and 24 hr incubation with 1 μM and 5 μM OTA did not lead to a significant effect on TNF-α expression (Fig. 9B).

TNF-α activate caspase-8

In view of previous findings and our results, we used western blotting to evaluate active caspase-8, considered a key factor in activation of the death receptor pathway.

Figure 10 illustrates the cleavage of caspase-8 in H9 cells incubated with 10 μM OTA for various intervals up to 24 hr. Caspase-8 activation by 10 μM OTA became appar- ent after 3 hr and steadily increased during the next 24 hr in a time-dependent manner. These results suggested that OTA might also induce apoptosis in H9 cells by the death receptor pathway, likely via TNF-α release.

Fig. 7. OTA triggers p38, ERK1/2 and JNK phosphorylation.

H9 cells were incubated with 1 μM OTA for the indicated durations and analyzed by immunoblotting using phospho-specific antibodies: P-p38 (Thr180/Tyr182), P-ERK1/2 (Thr202/Tyr204) and P-JNK (Thr183/

Tyr185). Membranes were stripped and rehybridized with total (T) specific Abs. Blots are representative of three independent experiments. OTA, ochratoxin A.

Fig. 8. OTA decreases survivin mRNA levels in H9 cells.

Cells were incubated with 0, 1, 5 and 10 μM OTA for 12 hr and 24 hr. Total RNA was extracted and analyzed by RT-PCR to determine the mRNA level of survivin and β-actin (for equal loading). Gels are representative of two independent experiments. OTA, ochratoxin A.

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DISCUSSION

OTA has been recognized as an immunosuppressive mycotoxin that acts at different levels of the immune system: it inhibits humoral and cellular immunity and depletes lymphoid organs. OTA has also been identified as an immunomodulatory molecule for cytokines secretion, leading to several immune function alterations (Marin and Taranu, 2014). Various mechanisms of immunotoxicity have been reported in immune cell lines from different species, including protein synthesis inhibition, oxidative stress, cell signaling pathway alteration and apoptosis.

Despite the fact that the half-life of OTA is very high in humans (35 days), there have been few reports regard- ing its mechanisms in the human immune system (Marin- Kuan et al., 2011). In the present study, we attempted to elucidate the molecular events that occurred during OTA- induced apoptosis in H9 cells which is a derivative clone

of human cutaneous CD4+ T lymphoma isolated from a Caucasian patient with leukemia. H9 cell line expresses naturally high level of CD4 receptor and is permissive to the HIV infection/replication in vitro (Mann et al., 1989).

They provide the best approach for understanding the reaction and the succession of apoptotic events in human activated CD4+ T cells once they are exposed to OTA.

Generally, in vitro treatment with relatively high OTA doses provides similar effects to those observed in vivo after chronic low doses exposure, especially in case of DNA-adducts caused by OTA (Pfohl-Leszkowicz and Castegnaro, 2005; Mantle et al., 2015), recent studies carried on human peripheral blood lymphocytes, showed a viability decrease of 20-30% following incubation with OTA concentrations less than 20 μM, (Gonzàlez-Arias et al., 2014; Ali et al., 2011). In the light of these data we have investigated cytotoxicity of OTA on H9 cells with concentrations up to 20 μM, to evaluate the sensibility of Fig. 10. OTA induced caspase-8 activation. Cells were incubated with 10 μM OTA for the indicated durations, lysed and total cell lysates were analyzed by immunoblotting using anti-active caspase-8 mAb. Medium without OTA was used as the control.

Immunoblot is representative of two independent experiments. OTA, ochratoxin A.

Fig. 9. OTA modulates IL-2 and TNF-α mRNA expression in H9 cells. Cells were incubated with 0, 1, 5 and 10 μM OTA for 12 hr and 24 hr. IL-2 (A) and TNF-α (B) mRNA levels were analyzed by real-time PCR using specific primers. IL-2 or TNF-α mRNA expression is presented as fold increases versus control cells (black bars) after normalization to S9 mRNA.

Each bar represents the mean ± S.E. from three independent experiments, and each experiment was performed in duplicate.

*p < 0.05; **p < 0.01. OTA, ochratoxin A; IL-2, interleukin 2; TNF-α, tumor necrosis factor α.

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this cells line to OTA and the mechanism of OTA cytotoxicity. Basically we found that OTA affects negatively cells viability in time-dose-manner (Fig.1). To determine whether the death is caused by necrosis or apoptosis we have performed TEM analysis of ultrastructural changes following OTA treatment and as suspected, micrographs revealed all the major hallmarks of apoptosis (Fig. 2). However higher doses (20 μM) meanly cause death by necrosis without morphological changes. Next, we explored the events by which OTA induced apoptosis in H9 cells. There are two major pathways of apoptosis:

death receptor and mitochondria-dependent. In the latter, the mitochondria play a critical role in the early events of apoptosis by both mitochondrial depolarization (∆Ψm) and the release of cytochrome c from the mitochondrial intermembrane space into the cytosol; the two phenome- na are a consequence of mitochondrial membrane permeabilization and have been considered as early irreversible events during apoptosis (Vander Heiden et al., 1997). We investigated the mitochondrial potential (Fig. 5), which exhibited significant loss following 6-hr incubation of H9 cells with 5 μM and 10 μM OTA (p < 0.001), and near-total ∆Ψm collapse after 18 hr (p < 0.001). OTA provoked the loss of ∆Ψm in a time- and dose-dependent manner.

These results confirm previous studies on the capacity of OTA to induce apoptosis by disrupting mitochondrial function in human T lymphocytes (Assaf et al., 2004). It has been well documented that mitochondrial damage is a major event that leads to the formation of the apoptosome and activation of caspase-9 and subsequently caspase-3.

At least two endogenous mitochondrial proteins, apoptosis-initiating factor and cytochrome c, were implicated in this activation of caspases (Li et al., 1998). Activation of procaspase-3 (executioner caspase) appears to be one of the most common steps involved in the execution of apoptosis. In different cell types, active caspase-3 leads to morphological changes and cell death. Studies have demonstrated that OTA induces ∆Ψm loss and activation of caspase-9 and caspase-3 in peripheral blood mononuclear cells (PBMCs) and Kit 225 cells (Assaf et al., 2004).In our experiments, we examined procaspase-3 activation in H9 cells following 24-hr OTA treatment (Fig. 6). The percentage of caspase-3-positive H9 cells increased with the OTA dose. However, we did not observe an increase in active caspase-3 at the low 1 μM dose, which may explain the absence of the major hallmarks of apoptosis.

At low doses, OTA required more time to induce apoptosis in H9 cells (data not shown).

Several mechanisms are associated with OTA-induced apoptosis, including protein and mRNA synthesis inhibition, oxidative stress, cell signaling pathway alteration

and disruption of the balance of the pro- and anti-apoptotic Bcl-2 protein family (Chopra et al., 2010). Oxida- tive stress is often cited as a key mechanism in OTA-induced apoptosis in different cell types (Marin-Kuan et al., 2011): it has been reported that OTA accelerates reac- tive oxygen species (ROS) formation by lipid peroxidation and the inhibition of antioxidant enzymes such as superoxide dismutase. Increased ROS induce Ca²⁺ release from intracellular organelles, affecting MAPK phosphorylation and the intracellular signaling cascade (Liang et al., 2015). In vitro OTA stimulated the phosphorylation of three major MAPKs identified in mammalian cells:

ERK1/2, JNK and p38. Generally, ERK1/2 is involved in cell survival, whereas JNK and p38 are often reported as stress kinases implicated in apoptosis. The pivotal balance between the ERK1/2 and JNK/p38 pathways determines whether a cell will survive or undergo apoptosis. MDCK cells illustrate the importance of this balance: OTA induces apoptosis in MDCK-C7 cells but not in MDCK-C11 cells. At nanomolar concentrations, OTA induces JNK and ERK1/2 phosphorylation in MDCK- C7 cells (ERK1/2 phosphorylation is induced after 2 hr).

The strong activation of these two MAPKs is responsible for the pronounced sensitivity to OTA, Phospho-ERK1/2 leads to stable and irreversible dedifferentiated MDCK- C7 cells with morphology distinct from the parent cell line, whereas phospho-JNK promotes apoptosis by a 14-fold increase in caspase-3 activation and DNA ladder formation (Gekle et al., 2000). In this study, 1 μM OTA induced ERK1/2, JNK and p38 phosphorylation in H9 cells after 3 hr (Fig. 7). The MAPK phosphorylation level remained constant over the next 3 hr; these results are in concordance with those of previous studies and confirm the time-dependent OTA activation of these three MAPKs (Sauvant et al., 2005).

OTA promotes apoptosis by acting on both pro- and anti-apoptotic Bcl-2 family members, which are considered among the principal regulators of mitochondrial apoptosis pathway. In vivo, it was demonstrated that topical application of OTA on mouse skin resulted in Bax upregulation (pro-apoptotic) and Bcl-2 downregu- lation (anti-apoptotic), thus promoting the pro-apoptotic signal and release of cytochrome c (Kumar et al., 2012).

Assaf et al. reported that OTA triggered a decrease in Bcl-xL protein but not Bcl-2 protein in PBMCs and Kit 225 cells (Assaf et al., 2004). The IAP family is anoth- er family of apoptotic regulator proteins that represent the last line of cellular defense against apoptosis; they act by directly binding to caspases, but few studies have been carried out to investigate OTA effect on this family.

Among the IAP family members, we studied surviv-

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ing, which binds directly to caspase-3, and its deregula- tion has been suspected in chemoresistance and tumor development. Typically, survivin is not expressed in adult tissues; however, it is highly expressed in embryonic tissues and dedifferentiated cells (Vegran et al., 2005). OTA treatment greatly affected the level of survivin mRNA in H9 cells (Fig. 8). Chopra et al. (2010) reported that 1 μM OTA triggered apoptosis in primary rat hepatocytes, inducing the expression of both pro- and anti-apoptotic genes. Survivin was one of the examined genes; it was induced 3-fold. In our experiments, 5 μM or 10 μM OTA greatly decreased the level of survivin expression in H9 cells. Despite these results, OTA could promote cell death downstream of the apoptotic cascade via the inhibition of IAPs.

OTA plays an important role in cytokine modulation.

Some studies have demonstrated that mycotoxins inhib- it cytokines such as circulating IL-2, IL-5 and TNF-α, while others have demonstrated that OTA stimulates IL-1, IL-2, IL-6 and TNF-α secretion. The modulatory effect of OTA may depend on the cell type and environment in which the secretion occurs (Al-Anati and Petzinger, 2006;

Bondy and Petska, 2000).

IL-2 and TNF-α play an important role in CD4 T lymphocyte activation, differentiation and proliferation. CD4 T lymphocytes may secrete the two cytokines, and dereg- ulation of their production affects the immune response considerably. In our cellular model, 24-hr incubation with 10 μM OTA significantly decreased IL-2 mRNA expression and increased that of TNF-α (Fig. 9). TNF-α has been described as a powerful pro-inflammatory molecule and has been reported to mediate apoptosis in H9 cells through binding to TNF-α receptor-1 (TNFR1) (Chuang et al., 2007). Additionally, two apoptotic pathways have been described: the mitochondria-dependent pathway that responds to extra- and intracellular stress, and the death receptor pathway mediated by activation of death receptor superfamily members such as TNFR1, which involves caspase-8 activation (Micheau and Tschopp, 2003;

Schmitz et al., 2000). We investigated the possible role of TNF-α in death receptor pathway activation through caspase-8 activation (Fig. 10) and illustrated caspase-8 activation in H9 cells following incubation with 5 μM OTA for different durations. Interestingly, OTA activation of caspase-8 was time-dependent and occurred more con- sistently at 3 hr. We suggest that besides the mitochondria-dependent apoptotic pathway, OTA can also indi- rectly trigger the death receptor apoptotic pathway in H9 cells through a possible autocrine effect of TNF-α (Sup- plementary Fig. 2).

In conclusion, we have demonstrated the signifi-

cant apoptotic potential of OTA on human CD4+ T lymphocytes. OTA led to MAPK activation and apoptosis by disrupting mitochondrial functions in H9 cells, and modulated survivin, IL-2 and TNF-α expression. These mechanisms clarify the pronounced immunotoxicity of OTA. Our findings prove that OTA activates caspase-8 in CD4+ T lymphocytes, which is a specific event of the death receptor apoptotic pathway, suggesting that OTA can additionally trigger the death receptor pathway in H9 cells through the autocrine effect of TNF-α.

ACKNOWLEDGMENTS

This work was supported by the Moroccan Society of Mycotoxicology. We are grateful to Karin Fink for her assistance with the realtime PCR and Laurence Lejeune for his technical assistance with the flow cytometry.

Conflict of interest---- The authors declare that there is no conflict of interest.

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