Immunosuppressive potential of astemizole against LPS activated T cell proliferation and cytokine secretion in RAW macrophages, zebrafish larvae and mouse splenocytes by modulating MAPK signaling pathway

International Immunopharmacology 65 (2018) 268–278

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Immunosuppressive potential of astemizole against LPS activated T cell proliferation and cytokine secretion in RAW macrophages, zebrafish larvae and mouse splenocytes by modulating MAPK signaling pathway Rekha Jakhar1, Chanchal Sharma1, Souren Paul, Sun Chul Kang

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Department of Biotechnology, Daegu University, Jillyang, Naeri-ri, Gyeongsan, Gyeongbuk 38453, Republic of Korea

A R T I C LE I N FO

A B S T R A C T

Keywords: Immunomodulatory Astemizole LPS RAW macrophages Zebrafish larvae MAPK

In this study, the immunomodulatory effects of astemizole (AST) against lipopolysaccharide (LPS) mediated T cell proliferation and induction of inflammation in RAW macrophages (in vitro), and zebrafish larvae (in vivo) were determined. AST significantly suppressed the phagocytic activity of macrophages (3.303 ± 0.115) and inhibited lysosomal enzyme secretion (13.27 ± 2.52) induced by LPS (100 ng/ml). Moreover, AST subdued the morphological deformities such as yolk sac edema (YSE) and spinal curvature curving (SC) by inhibiting ROS generation in zebrafish larvae 24 h after microinjection of LPS (0.5 mg/ml). AST was also shown to inhibit the production of the major cytokines TNF-α (150.8 ± 0.6), IL-1β (276.5 ± 1.6), and PGE2 (194.6 ± 0.6) pg/ml in RAW macrophages. It also subdued the ROS induced iNOS and COX-2 generated in response to LPS mediated immune dysfunctions in zebrafish larvae. These results suggested the immunosuppression effect of AST. Furthermore, induction of immune-suppression due to AST resulted in significant down-regulation of innate immunity directed by MAPK (p38, ERK and JNK), which was found to be associated with decreased production of acute inflammatory mediators both in vitro and in vivo. To confirm its activity, splenocytes were prepared using BALB/c mice and a mitogen activated splenocyte proliferation assay was also performed. Our findings suggest that AST has the ability to inhibit T cell proliferation and cytokine secretion both in vitro and in vivo by interfering with MAPK signaling pathway. Taken together, our results showed the potential of AST as a countermeasure to immune dysfunction and suggest its use as immunosuppressant compound in inflammatory disease.

1. Introduction Immunomodulation is a process that mainly consists of biological and pharmacological effects of various metabolites on humoral and cellular aspects of immune response [1]. As the immune system is involved in pathophysiological mechanisms of every disease, modulation of the immune system requires either immuno-enhancers or immunosuppressors to alleviate various diseases. Sometimes in response to foreign attack excess inflammation initiated by immune suppressor cells, various cytokines, chemokines and angiogenic factors becomes toxic which can be suppressed to enable immune recovery [2]. Hence, there is a need to identify new immunosuppressants for the success of immune therapies. A large number of clinically applicable immunosuppressant drugs are available such as cyclosporine A which binds to cyclophilin, FK506 and rapamycin, they binds with FK506

related binding proteins, silymarin and cyclophosphamide. They have been reported for significantly improving the patient first-year survival with up to 90% for renal transplant [2–5]. Additionally, they have also been reported to have neuroprotective effect against brain injury and neurodegenerative disorder such as Alzheimer and Parkinson [6]. However these drugs have a narrow therapeutic range and cause serious adverse effects including headaches, insomnia, hepatotoxicity, nephrotoxicity, neurotoxicity, and induction of diabetes. As a result, it is essential to search for new, potential immunosuppressant drugs with low toxicity. Identification of immunosuppressants without any side effects is still a challenge to the medical system. Recently, several synthetic compounds were reported to exert immunosuppressing effects such as the augmentation of macrophage function inhibitors of calcineurin, and mitogen-activated protein kinase kinase kinases (MAPKKK) [2,7]. There

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Corresponding author at: Department of Biotechnology, College of Engineering, Daegu University, Jillyang, Naeri-ri, Gyeongsan, Gyeongbuk 38453, Republic of Korea. E-mail addresses: [email protected] (C. Sharma), [email protected] (S.C. Kang). 1 Authors contributed equally. https://doi.org/10.1016/j.intimp.2018.10.014 Received 16 June 2018; Received in revised form 27 September 2018; Accepted 10 October 2018 1567-5769/ © 2018 Elsevier B.V. All rights reserved.

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Fig. 1. In vitro effect of AST treatment. (A) Chemical structure of AST. (B) AST effect on the viability of macrophage cells based on the MTT assay. (C) Predicted IC50 of AST on RAW macrophage cells. Significant differences from the control were designated as *p < 0.02, **p < 0.005, ***p < 0.0002, and ****p < 0.0001.

All other required chemicals used were of highest quality and purchased from Sigma Aldrich, St. Louis, MO, USA.

is now a convincing evidence that T cells also play an important role in immune regulation and that their secreted cytokines are involved in the inflammatory process. Hence, the blockade of T cell activation and proliferation as well as consequential cytokine production is one of the main immunosuppression principles [8]. Alterations of important intracellular signaling pathways such as NF-kB and the mitogen-activated protein kinase (MAPK) pathway have been suggested to play a critical role during the process of immunosuppression. Therefore, in an effort to search for new and clinically safe immunosuppressant, we analyzed the immuno-suppressive effects of astemizole (AST) (Fig. 1a), a potent histamine H1 receptor antagonist, and elucidated its effects on key immunomodulatory properties that could be used to treat several inflammatory and autoimmune diseases. AST exerts both anticholinergic and antipruritic effects. In addition, it is a well-known second generation drug which is absorbed rapidly from the gastrointestinal tract and competitively binds to histamine H1 receptor by suppressing the formation of edema and pruritus in the gastrointestinal tract, uterus, blood vessels, and bronchial muscle regions. AST has also been studied for treatment of malaria, hERG and hEAG channel function in cancer and as a second generation antihistamine H1 antagonist [9,10]. However, the immunosuppressive effects of AST on the immune responses to RAW macrophage cells and on zebrafish larvae have not been reported yet. Zebrafish (Danio rerio) is an important animal model to study innate immune systems due to the development of immunity immediately after 1 day post fertilization. And larval transparency (during first 2 weeks of development) is a unique feature of zebrafish that allows direct assessment of drug effect. In the present study we aimed to investigate the immunosuppressive effect of AST on LPS (Escherichia coli 055: B5) stimulated inflammatory response in vitro (RAW macrophages 264.7) and in vivo (zebrafish larvae) model systems. In addition, we characterized the role of p38 MAPK during the immunosuppressive effects of AST in both systems.

2.2. In silico pharmacokinetics and drug-likeness assessment of astemizole Chemical properties and biological activity spectrum were determined by using PASS professional 2010 & Pharma Expert 2010 Evaluation (v. 10.1) respectively. ADMETx (absorption, distribution, metabolism, elimination and toxicity) properties, were determined by using SwissADME ((http://www.swissadme.ch) and GUSAR (http:www.pharmaexpert.ru/GUSAR/qsar.html), freely accessible web tool. The drug likeness test was carried out using the criteria used by major pharmaceutical companies; MDL drug data report, Pfizer, Pharmacia and GSK [11–14]. 2.3. Cell culture and treatment

2. Materials and methods

RAW 264.7, a macrophage cell line, was purchased from the American Type Culture Collection (ATCC Manassas, VA). Cells were maintained in DMEM (Sigma, St. Louis, MO, USA) and supplemented with 10% (v/v) fetal calf serum and 1% penicillin/streptomycin (pen/ strep) (Invitrogen, Carlsbad, CA). Cells were seeded in a 96-well plate at a density of 1 × 106 cells/well and allowed to adhere at 37 °C in a 5% CO2 incubator for next 24 h. Cells with 80–90% confluence were treated with varying concentrations of AST (0.2, 0.5, 2, and 10 μM) for 24 h, after which the cell viability (~80–90%) was measured using MTT (3(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay as reported by Mosmann, 1983 with some modifications [15]. 10 μl of MTT working solution (5 mg/ml) dissolved in phosphate buffer saline (PBS) was added to each well for 4 h. The medium was then aspirated, after which the generated formazan crystals were dissolved completely in 50 μl dimethyl sulfoxide (DMSO) per well for 30 min and absorbance was read at 540 nm using ELISA plate reader. Experiment was performed in triplicates and mean value was used to calculate the percentage of cell proliferation using the following equation

2.1. Reagents and antibodies

Cell viability (%) =

Astemizole (AST), Lipopolysaccharide (LPS) (Escherichia coli 055: B5), Concanavalin A (ConA), Griess reagent, Indomethacin (Indo), Neutral Red, Triton X-100, and nitroblue tetrazolium (NBT) were purchased from Sigma-Aldrich (St. Louis, MO, USA). Rabbit monoclonal primary antibodies, β-Actin, p-P38, p-ERK, JNK, iNOS and COX-2 were purchased from Abcam (Cambridge, MA, USA). c-Fos, c-Jun and COX-1 were purchased from Santa Cruz Biotechnology Inc. (Santa Cruz, CA, USA). Horseradish peroxidase labeled goat anti-rabbit immunoglobulin was purchased from Bethyl Laboratories, USA. Supplementary Tables 1 and 2 provide the details of primary and secondary antibody sources.

Absorbance of treated cells × 100 Absorbance of control cells

2.4. In vitro phagocytic assay of cellular lysosomal enzyme activity The cellular lysosomal enzyme activity was evaluated by measuring the acid phosphatase activity in macrophages as per the protocol described by Rainard (1986) [16,17]. A mix of 20 μl RAW macrophages 264.7 (1 × 106 cells/well), 40 μl of DMEM media (supplemented with 10% (v/v) fetal calf serum and 1% penicillin/streptomycin) and 20 μl AST (0.35 μM and LD50 0.74 μM) each were added in triplicate into a 96 well culture plate and incubated for next 24 h in a CO2 (5%) dependent 269

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140 μl of DMSO were added, after which the absorbance was measured at 570 nm using a microplate reader. Finally, the percentage of NBT reduction was calculated by the following equation:

atmosphere at 37 °C. After treatment with AST (0.35 μM and LD50 0.74 μM), the medium was removed by aspiration and 20 μl of 0.1% Triton X-100 were added to each well. Next, 100 μl of 10 mM p-nitro phenyl phosphate (p-NPP) solution and 50 μl of 0.1 M citrate buffer (pH 5.0) were added. The plate was then further incubated for 30 min, after which 150 μl of 0.2 M borate buffer (pH 9.8) was added and the absorbance was measured at 405 nm. The percentage activity was then calculated according to the following equation:

NBT reduction (%) =

OD sample − OD negative control × 100 OD negative control

2.8. NO measurement using Griess reagent

OD sample − OD negative control Lysosomal enzyme activity (%) = OD negative control

Total nitrite concentration was measured in the medium using Griess reagent and the concentration thus estimated was taken as the production of NO. To this aim, RAW macrophages 264.7 (1 × 106 cells/ well) were seeded in a 96 well plate (triplicates per group). Cells after reaching confluency (~80–90%) were treated with PBS (blank control), 100 ng/ml LPS (negative control), 0.35 μM AST and LD50 0.74 μM of AST in presence of 100 ng/mL LPS. After incubation for 24 h, equal amount (100 μl) of supernatant and Griess reagent (1% sulfanilamide in 5% phosphoric acid and 0.1% naphthylethylenediamine dihydrochloride in water) was mixed and incubated for 10 min at room temperature. Absorbance was measured using a microplate reader at 540 nm. Standard cure was generated using sodium nitrite.

× 100

2.5. Neutral red phagocytosis assay of macrophages RAW 264.7 macrophages with the density of 1 × 106 cells/well in a 96 well plate were used to examine the influence of AST on phagocytic activity by a neutral red phagocytosis assay system, as previously described [17,18] with some modifications. Entire experiment was carried out in triplicates for each group. Cells with 80–85% confluence were treated with 0.35 μM AST and LD50 0.74 μM of AST for next 24 h. Next, supernatant was removed and cells were treated with 200 μl of 0.075% neutral red and incubated for 30 min at 37 °C followed by thrice aspiration and rinsing thoroughly with 200 μl PBS. Treated cells were lysed in 200 μl per well of 100 mmol/l acetic acid:dehydrated alcohol (1:1, v/v), and were incubated for next 8 h at 4 °C for sufficient schizolysis of cells and the release of phagotrophic neutral red. The optical density was determined at 490 nm and the phagocytosis was expressed as OD values. Control cells were treated with phosphate buffer saline, and LPS (100 ng/ml) was used as a positive control.

2.9. Zebrafish maintenance, embryo collection, and larval maintenance Mature zebrafish of wild type strain were commercially purchased from a local shop in Daegu, South Korea and maintained at 28 °C under a light-dark cycle of 14:10 h in a circulating system at Daegu University, South Korea [19–21]. Embryos obtained after 12 h of natural pair-wise mating were collected in fresh unchlorinated water, then maintained in E3 medium (0.2 mM Ca (NO3)2, 0.13 mM MgSO4, 19.3 mM NaCl, 0.23 mM KCl, and 1.67 mM HEPES) supplemented with a drop of methylene blue to prevent contamination. Morphologically distorted and dead eggs were removed 1 day post-fertilization (dpf). At 36 h postfertilization (hpf), larvae were used for experiments [22].

2.6. ELISA for cytokine determination The TNF-α and IL-1β levels in cell supernatants were determined using an enzyme-linked immunosorbent assay (ELISA) kit by Invitrogen, Frederick, MD, USA, while the prostaglandin levels were measured using a Prostaglandin E2 Competitive Elisa (PGE2) Kit by Enzo Life Sciences Co., USA. Experiment was conducted in triplicates where 1 × 106 cells/well were seeded in a 96-well plate. Cells treated with phosphate buffer saline was taken as blank control and cells treated with LPS (100 ng/ml) was taken as positive control. Cells were treated with 0.35 μM AST and LD50 0.74 μM of AST in the presence of LPS. Further procedures were done according to the manufacturer's protocols. Monoclonal anti-mouse TNF α/IL-1β and PGE2 antibodies were used as capture; and specific biotin conjugates were added as secondary antibodies. Recombinant mouse TNF α/IL-1β and PGE2 were used as protein standard. Finally, the absorbance at 450 nm was read on microtiter plate reader (Bio-Tek Instrument Co., WA, USA).

2.10. Experimental design Decorticated eggs were maintained under optimum conditions for 24 h before treatment. The LD50 of AST was determined based on the mortality, morphological changes, and heart-beat per minute of larvae (n = 10) using different doses. A mixture of (0.5 mg/ml) LPS dissolved in sterilized physiological saline was infused via a microinjection cannula connected to a motor-driven injector nanolitre 2010 (World precision instruments, Sarasota USA) at a rate of 13.8 nl to induce immune dysfunction and inflammation. The injection cannula was retained for another 5 min after injection with optimized dip switch setting. Control larvae were injected with the same amount of sterile physiological saline alone. Larvae at 2 dpf were randomly divided into four groups (10 larvae/group) in 24 well plates maintained in E3 medium. At 24 h after LPS induction, larvae were treated with the IC50 concentration of AST for 24 h. Group 1 served as control larvae, Group II was the LPS control, Group III was treated with LPS + AST (10 μM) and Group IV was treated with LPS + AST (20 μM).

2.7. Detection of superoxide anions To determine the superoxide anions produced by phagocytic cells that were reduced by the redox dye nitroblue tetrazolium (NBT), an NBT dye reduction assay was conducted according to Rainard (1986), with few modifications [17,18]. 20 μl RAW macrophages 264.7 (with density of 1 × 106 cells/well) dissolved in 40 μl of DMEM were seeded in triplicates/group in a 96-well plate and kept at 37 °C in humidified 5% CO2 incubator. Next, macrophage cells were treated with 20 μl of 0.35 μM AST and LD50 0.74 μM of AST respectively, and incubated for 24 h under the same conditions. After incubation, 20 μl of phorbol myristate acetate (PMA, 10 ng/ml) and 20 μl of NBT solution in PBS (1.5 mg/ml) were added and the mixture was further incubated for 1 h under the same conditions. After incubation, the adherent macrophages were rinsed vigorously with RPMI medium, then washed four times with 200 μl methanol. Following air-drying, 120 μl of 2 M KOH and

2.11. Determination of ROS production and measurement of cell death marker The ROS production was monitored using live larvae. Larvae were anesthetized with a 1:100 dilution of tricaine (4 mg/ml). All groups were incubated with DCFH-DA (5.0 μM/ml) for 30 min at 28 °C in the dark, after which images were captured using a fluorescent microscope and the representative fluorescent intensities were quantified and analyzed for statistical significance. To identify any apoptotic characteristics, all groups were treated with acridine orange (AO) (7.5 μM/ ml) for 30 min in dark. After subsequent washing, samples were 270

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observed under a fluorescent microscope (Nikon Eclipse TS100 Epifluorescence microscope, Japan) using an excitation wavelength of 488 nm.

Cell proliferation (%) =

Absorbance of treated cells × 100 Absorbance of control cells

2.12. Whole mount immunofluorescence staining

2.16. Statistical analysis

Whole mount staining was performed for COX-2 (substrate for prostaglandin) and iNOS (one of the isoform of NO system). Treated larvae were fixed overnight in 4% paraformaldehyde, after which methanol permeabilization was conducted overnight in the dark. Blocking was accomplished using 10% heat inactivated fetal bovine serum and 3% BSA prepared using PBS-0.1% Tween 20 mixture. Larvae were further incubated overnight with COX-2 and iNOS (1:500 dilution) primary antibodies at 4 °C with mild shaking. Following incubation, the larvae were washed with PBST and incubated again with FITC-conjugated (for COX-2) and Texa-red (for iNOS) anti-rabbit secondary antibody (1:400 dilutions) for 2 h at room temperature. Counter staining was done using Hoechst dye. The intensity of signals was observed using a Nikon Eclipse TS100 Epi-fluorescence microscope (Japan).

Statistical analysis was conducted using GraphPad Prism (version 7.01). Differences were identified by ANOVA and the Student's t-test. Results were considered significant at *p < 0.01, **p < 0.001 and ***p < 0.0001. All data were calculated from three individual sets of experiments, and the data are expressed as the means ± the standard deviation. 3. Results 3.1. In silico pharmacokinetics and drug-likeness assessment of AST Astemizole (C28H31FN4O) is a synthetic piperidinyl-benzimidazol derivative having antiallergic properties. Its chemistry includes a total of four H-bond acceptors, one H-bond donor, eight rotatable bonds, total polar surface areas (TPSA) of 42.32 Ǻ, and MLOGP of 4.46. It has 0.32 fraction of carbons in sp3 hybridization, lipophilicity (iLOGP) of 4.48 units, corrective factor (XLOGP3) of 5.97 and log S of −9.15. Support vector machine algorithm (SVM) was applied to determine the pharmacokinetics parameter. With the prediction of drug-likeness assessment for AST, we found that it exactly followed the rule of Veber (Rotable bonds ≤ 10 & TPSA ≤ 140) and MDDR (Rings ≥ 3, rigid bonds ≥ 18 & Rotable bonds ≥ 6). Moreover, it also followed the Lipinski rule of five (MW ≤ 500, MLOGP ≤ 4.15, N or O ≤ 10 and NH or OH ≤ 5) and Egan's rule (WLOGP ≤ 5.88 and TPSA ≤ 131.6).

2.13. Western blotting After treatment with LPS and AST, both cells and zebrafish larvae were harvested, lysed with RIPA buffer (Sigma, St. Louis, MO, USA). The protein concentrations were determined on the supernatant harvested using Bradford Reagent (Sigma, St. Louis, MO, USA). For western blot analysis, an equal amount of protein (50 μg in each lane) was subjected to electrophoresis on SDS-polyacrylamide gel and transferred to a polyvinyldifluoridine (PVDF) membrane (Roche Diagnostics, Indianapolis, IN, USA) in 25 mM Tris buffer containing 192 mM glycine and 20% (vol/vol) methanol by electro-blotting. The membranes were then blocked with 5% non-fat milk for 2 h at room temperature. Next, blots were probed with the primary antibodies overnight at 4 °C and then incubated with an appropriate dilution of secondary antibody conjugated with horse radish peroxidase (HRP). Finally, the protein bands were detected using WEST-ZOL™ Plus enhanced chemiluminescence (ECL) according to the recommended procedure (Amersham Pharmacia, Piscataway, NJ, USA).

3.2. In vitro toxicity assessment of AST The cytotoxicity of AST was determined by MTT on RAW 264.7 macrophages (Fig. 1B), which are important immune cells that are primarily active in the host body after foreign invasion. To determine the cytotoxicity of AST toward macrophages, the effects of four different doses from lower to higher concentration (1 μM, 5 μM, 10 μM and 50 μM) were compared with those of control. The cell reducing potential was found to decrease as the AST dose increased from 0.2 μM (72.88 ± 2.12) to 10 μM (20.97 ± 0.79). The IC50 (0.74 ± 0.07) was calculated by using a non-linear regression curve (Fig. 1C); hence, the IC50 and ~half 0.35 μM) was used during the further in vitro experiments.

2.14. Preparation of mouse splenocytes BALB/c Mice obtained from Orient Bio, Inc. Seoul, South Korea were sacrificed, after which their spleens were removed aseptically. Cell suspensions were then prepared by flushing. Next, 1 × 105 cells were placed in a 16-mm well and incubated under 5% CO2 at 37 °C for 3 h to remove adherent cells such as macrophages. The supernatant together with the non-adherent cells were collected by centrifugation at 1000 rpm at 37 °C for 10 min. The cell pellets were then re-suspended in CRPMI medium and adjusted to 1 × 105 cells/ml. Finally, cell numbers and viability were determined using a hematocytometer and trypanblue dye exclusion, respectively.

3.3. AST subdues the phagocytosis and lysosomal activity against LPS in RAW macrophages We next determined the anti-phagocytic potential of AST and its effects on the release of lysosomal enzyme. A significant increase (p < 0.0001) between the control (1.507 ± 0.28) and LPS treatment (9.937 ± 0.08) was found in the phagocytic activity of macrophages (Fig. 2A). Treatment with AST at 0.35 μM (4.933 ± 0.202) and 0.74 μM (3.303 ± 0.115) after LPS treatment showed a significant dose dependent decrease (p < 0.0001) relative to the LPS treated group; therefore, we concluded that AST elicited a significant decrease in the phagocytic activity that was related to the inhibition of macrophage proliferation. To confirm this, we evaluated the release of lysosomal enzyme in response to LPS (100 ng/ml) and after AST treatment. As shown in Fig. 2B, LPS exhibited the maximum potency, approximately 60%; however, this was found to be significantly reduced (p < 0.002) in response to treatment with 0.35 μM AST (24 ± 3.57) and 0.74 μM AST (13.27 ± 2.52).

2.15. Mitogenic and comitogenic activity assay The comitogenic activity was tested by incubating mouse splenocytes (1 × 105 cells/ml) with different concentrations of AST (0.05, 0.35 and 0.74 μM) in the presence of mitogen (i.e., ConA at 2 μg/ml) for 24 h in 96-well plates [23,24]. Plates were pre-incubated for 24 h before adding ConA and AST in various concentrations. Control cells were treated with phosphate buffer saline. Splenocytes proliferation activity was then subjected to MTT assay. Absorbance at 570 nm was read with an ELISA reader (Bio-Tek Instrument Co., WA, USA). The percentage of cell proliferation was calculated using following equation 271

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Fig. 2. Effect of AST treatment on in vitro phagocytosis response of RAW macrophages. (A) AST induced macrophage phagocytosis. Control cells were treated with PBS only and LPS (100 ng/ml) was used as a positive control. (B) % lysosomal enzyme activity. (C) % NBT reduction. (D) Nitric oxide production (E) Cytokine (TNF α, IL-1β, and PGE2) secretion from LPS-activated (1 μg/mL) RAW macrophages cells. 20 μM Indo was used as a positive control. Values are presented as the means ± S.D. (n = 4). (F) Effects of AST on iNOS protein expression. Significant differences from the control were designated as *p < 0.01, **p < 0.001 and ***p < 0.0001.

3.5. In-vivo toxicity assessment of AST in Zebrafish larvae

3.4. AST inhibits LPS mediated phagocytosis by suppressing superoxide anion, release of nitric oxide and secretion of cytokines in RAW macrophages

To determine the effects of AST in vivo, zebrafish larvae model was developed. We evaluated the effects of six AST doses (0.1, 1, 5, 10, 25, and 50 μM) on survival percentage recorded after 12 and 24 h of AST treatment. As shown in Fig. 3A, no significant sign of death was observed in response to treatment with AST (0.1, 1, 5, 10 μM) after 24 h, whereas only 21.65% and 17.1% survival was observed in response to treatment with 25 μM and 50 μM after 12 h, which was significant (p < 0.0003) based on the Logrank test for trend. The LD50 (20.37 μM) was determined by non-linear regression curve. Dose dependent effects of AST on the morphology of larvae were determined by microscopic analysis. No significant morphological deformities were found among the control and 1, 5, and 10 μM AST treatment groups after 24 h. Moreover, changes such as yolk sac edema (YSE), spine curvature curving (SC), opaque yolk (OC), and pericardial edema (PE) were observed in response to treatment with higher doses of 25 μM and 50 μM (Fig. 3B). Next, the heart beat rate per min (60 s) after 24 h of AST treatment was calculated to determine the effects of AST over cardiac output (Fig. 3C). Non-significant variation between the control (166 beats per min (bpm)) and AST 0.1 μM (163.2 bpm) and AST 1.0 μM (148.8 bpm) were found. In addition, significant differences (p < 0.01) were found for the 5.0 μM (130.8 bpm), 10 μM (112.8 bpm) and 25 μM (98.4 bpm) treatments. These finding provided sufficient evidence of direct effects of AST on zebrafish larvae. Hence, further in vivo studies were conducted using two doses of AST 10 μM and 20 μM (LD50).

Fig. 2C shows that treatment with AST at 0.35 μM (68.02 ± 1.04) and at its IC50 of 0.74 μM (27.96 ± 1.63) produced a significant dosedependent decrease (p < 0.0003) in NBT reduction stimulated by phorbol myristate acetate (PMA) (81.85 ± 1.04). We next evaluated the effect of AST on the NO production that subsequently occur when macrophages are activated and caused the accumulation of nitrite in the cultured medium. In the presence of LPS (77.08 ± 2.50), AST significantly (p < 0.0001) suppressed the NO production in a dose dependent manner (0.35 μM AST - 66.42 ± 0.166; 0.74 μM AST 54.79 ± 1.04) (Fig. 2D). The protein concentrations of all investigated pro-inflammatory cytokines (TNF-α, IL-1β, and PGE2) decreased dose dependently in LPS (100 ng/ml) treated macrophage culture supernatants after the addition of AST (Fig. 2E). The amounts of TNF-α, IL-1β, and PGE2 in the control were found to be 13.74 ± 0.08, 28.7 ± 0.15, and 10.06 ± 0.06, respectively, whereas elevated levels were of 279.5 ± 1.2, 455.3 ± 1.5, and 699.5 ± 0.9 pg/ml, respectively, were recorded for LPS-treated supernatants. The amounts of TNF-α, IL-1β, and PGE2 decreased significantly (p < 0.0001) after 0.74 μM AST treatment and were found to be 150.8 ± 0.6, 276.5 ± 1.6 and 194.6 ± 0.6 pg/ml for TNF-α, IL1β, and PGE2, respectively. Based on these results, we concluded that AST inhibited the respiratory burst and phagocytic activity through superoxide anion suppression and that it controlled the secretion of TNF-α, IL-1β, and PGE2 generated against LPS invasion in host (macrophages) cells. Fig. 2F represents the fold change after LPS treatment in the iNOS protein expression. With iNOS results, it was clear that AST even in a low dose of 0.35 μM (3.15 ± 0.08) was capable to cope with the LPS (4.39 ± 0.09) at significance of p < 0.0001. Reduction in the expression of iNOS due to AST - IC50 was reported to be 2.36 ± 0.06.

3.6. Immunosuppressant effect of AST on morphology of zebrafish larvae Morphological changes were monitored 24 h after AST treatment (hat) over 24 h of LPS induction (Fig. 4A). Significant changes were observed between the control and LPS treated group. The presence of pericardial edema (PE) and spinal column curving (SC) were observed in the group treated with LPS. However, after treatment with 10 μM and 272

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Fig. 3. In vivo effect of AST treatment on zebrafish larvae. (A) Kaplan-Meier analysis of zebrafish larval survival rate after AST treatment. (B) Phenotypic changes that occurs after AST treatment in increased dose dependent manner. (C) Effect on heartbeat rate per min after AST treatment in dose dependent manner.

20 μM AST these morphological deformities were not seen, demonstrating the immunosuppressant activity of AST.

We next checked how AST regulates apoptosis in response to increased oxidative stress. To accomplish this, we conducted AO staining to detect any apoptotic markers or bodies. As expected, high fluorescence signals (assumed to be due to apoptosis) were recorded for LPS relative to the control (Fig. 4D). These signals were relatively low after AST treatment in a dose dependent manner when compared with LPS generated signals. Hence, we concluded that AST inhibited the LPS mediated apoptosis.

3.7. AST reduced intra-cellular ROS production by inhibiting apoptosis Larvae generated a significant (p < 0.001) amount of reactive oxygen species in response to LPS (2.759 ± 0.11) attack in comparison with the absence of LPS (control 1 ± 0), which was evident upon live imaging of larvae for ROS using DCFH-DA (Fig. 4B). The imaging data provided the significant fold change after treatment with AST at the LD50 dose (1.297 ± 0.10) (Fig. 4C).

Fig. 4. AST treatment normalizes the (A) morphology, (B) ROS, (C) ROS densitometry and (D) inhibited apoptosis in vivo in zebrafish larvae after LPS exposure. Significant differences were designated as *p < 0.01, **p < 0.001 and ***p < 0.0001. 273

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Fig. 5. Photomicrograph representing the immunofluorescence expression for (A) COX-2 and (B) iNOS for zebrafish larvae treated with AST and counterstained with Hoechst (Blue). The scale bar indicates 200 μm (magnification 4×). Significance was calculated by Tukey's test with significant differences from the control were designated as *p < 0.01, **p < 0.001 and ***p < 0.0001. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

suppression of JNK and p38 was associated with the downregulation of c-fos (LPS, 2.31 ± 0.05; 20 μM-AST, 1.08 ± 0.01) and c-jun (LPS, 14.15 ± 0.02; 20 μM-AST, 2.47 ± 0.005) respectively. Level of COX-1 (LPS, 3.03 ± 0.06; 20 μM-AST, 1.17 ± 0.01) and COX-2 (LPS, 2.75 ± 0.01; 20 μM-AST, 1.63 ± 0.005) were also found to be significantly (p < 0.001) reduced after AST treatment (Fig. 6C and D).

3.8. AST suppressed LPS-driven inflammatory response in zebrafish larvae To assess the impact of AST protection against LPS induced cytokine secretion and increased inflammation we performed whole mount staining and expression analysis of iNOS and COX-2 levels. As shown in Fig. 5, the intensity of both COX-2 (Fig. 5A) and iNOS (Fig. 5B) were significantly upregulated in LPS treated larvae when compared to the control. Conversely, treatment of zebrafish larvae with AST significantly (P < 0.01) reduced the expression of iNOS and COX-2, providing evidence of its inhibition capacity against inflammation.

3.10. Confirming the immunosuppressant effect of AST in a mouse model To confirm the effects of AST on the immune system, we next conducted a mitogen-stimulated splenocyte proliferation assay in a mouse model as shown in Fig. 7. The ConA treatment significantly (p < 0.0001) enhanced the proliferation rate of mouse splenocytes (T cells), which was then reduced significantly (p < 0.0001) in a concentration-dependent manner by AST treatment. Moreover, the maximum suppressing effect was observed in response to 0.74 μM (0.151 ± 0.004) AST. Under these conditions, we found that the AST induced an important suppression of ConA triggered splenocyte proliferation.

3.9. AST impairs LPS-mediated activation of the p38-MAPK signaling pathway both in vitro and in vivo To determine if the inhibition of cytokine production, ROS and RNS mediated inflammation by AST is mediated through modulation of signaling pathways involving p38 MAPK, we examined the effects of AST on the activation of various MAPK pathways both in vitro (RAW macrophages 264.7) (Fig. 6A) and in vivo (zebrafish larvae) (Fig. 6C) through immunoblotting assay. In vitro results revealed a significant (p < 0.01) downregulation of p-p38 (LPS, 1.48 ± 0.02; 0.75 μM-AST, 1.24 ± 0.004) and JNK activation (LPS, 1.26 ± 0.02; 0.75 μM-AST, 1.14 ± 0.004) in macrophage cells upon AST treatment (Fig. 6A and B). Suppression of JNK and p38 were also found to be associated with subsequent downregulation of c-fos (LPS, 1.27 ± 0.01; 0.75 μM-AST, 1.03 ± 0.01) and c-jun (LPS, 1.17 ± 0.007; 0.75 μM-AST, 1.13 ± 0.01). Moreover, the expression of COX-1 (LPS, 3.03 ± 0.06; 0.75 μM-AST, 2.62 ± 0.03) and COX-2 (LPS, 16.35 ± 0.06; 0.75 μMAST, 13.21 ± 0.02) were significantly (p < 0.001) reduced after AST treatment. Similar results were found in case of in vivo experiment conducted using zebrafish larvae where LPS mediated activation of p-p38 (LPS, 10.41 ± 0.14; 20 μM-AST, 0.32 ± 0.03) and JNK (LPS, 3.27 ± 0.08; 20 μM-AST, 1.52 ± 0.07) was suppressed upon AST exposure in a dose-dependent manner (Fig. 6C and D). Unlike macrophages, the

4. Discussion Immune regulation is a complex and fundamental process to any immune response that involves multiple mechanisms to ensure that it is appropriate for the perceived threat to the host. Regulatory T cells (Tregs) play an important role in modulating immune response to selfantigens and infectious agents, but the method by which they modulate discrete immune functions has not been well defined. In pharmacologic doses, immunosuppressive drugs are used to suppress various allergic, inflammatory, and autoimmune disorders. To address the consequences, the present investigation focused on the immunosuppressive activity of AST as a potential candidate for development of immunomodulatory drugs. Hence first we made efforts to predict various unknown physicochemical (structure and chemistry), chemo-informatics (QSAR and molecular descriptors), and pharmacokinetics 274

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Fig. 6. AST exerted immunosuppressant activity by activating the MAPK signaling pathway both in vitro and in vivo. (A) RAW macrophage cells were treated with LPS and AST and the activation of p38, JNK, ERK, c-Fos, c-Jun, COX-1 and COX-2 was detected by Western blotting. (B) All Western blots were analyzed by densitometry analysis using the ImageJ software. (C) Zebrafish larvae were treated with LPS and AST and the activation of p38, JNK, ERK, c-Fos, c-Jun, COX-1 and COX-2 was detected by Western blotting. (D) Relative fold change expressed in bar graph using ImageJ software. β-actin was used as an internal control. Values are presented as the means ± S.D. (n = 4). Significant differences from the control were designated as *p < 0.05, **p < 0.01 and ***p < 0.0001.

Macrophages are multifunctional cells that play a distinct role in inflammation and immune response because of their ability to undergo phagocytosis, a process that is accompanied by the formation of several phagosomes that undergo sequential fusion events with early, then late endosomes and finally with lysosomes to yield a phagolysosome, an acidic compartment full of hydrolases leading to the generation of reactive oxygen species (ROS) [23,25–27]. In association with the antiphagocytic effects, our results also showed the inhibition of phagocytosis and a significant decrease in the release of lysosomal enzymes and NBT reduction in macrophage cells (Fig. 2). It has been reported that macrophages are indispensable accessory cell lines in the in vitro generation of cytotoxic T cells and are involved in antigen presentation through TCR and CD28 and thus required for the induction of cytotoxic T cell response [28,29]. Similar results were observed in cytotoxic T cell activation after AST treatment, leading to a decline in the process of phagocytosis because of their incapability in antigen presentation [30,31]. The process of inhibition of phagocytosis was rapid, but the mode of action of AST is at present not understood. Possible mechanism for inhibition of phagocytosis could be the increase in membrane fluidity and the activation of some membrane proteins [32]. Apparently, little alteration in membrane function and permeability was observed after AST treatment, as indicated by neutral red uptake and spontaneous lysosomal enzyme release, both of which are metabolically active processes. Thus, we assumed that the rapid inhibitory effect on phagocytosis could be because of changes in physicochemical properties, especial the fluidity or membrane protein mobility within the bilayers of the macrophage membrane. These specific processes of immunosuppression through alteration of the macrophage activity were assumed to trigger a cascade of signal transduction to maintain the host's immune response. ROS has been reported to play a major role in immune system

Fig. 7. Effects of AST on the proliferative response of mouse splenocytes induced by ConA. Mouse splenocytes were induced to proliferate by the addition of 2 μg/ml of ConA over a 24 h culture period. AST was applied at concentrations of 0.05–0.74 μM. The values are presented as the means ± S.D. (n = 4). Significant differences from the control were designated as *p < 0.02, **p < 0.005, ***p < 0.0002, and ****p < 0.0001.

(ADMETox) of AST using system biology approach. Being flexible due to the presence of 8 rotational bonds, AST fulfils an additional criterion of drug metabolism and pharmacokinetics (DMPK). Major advantage using system biology resists in its lower cost, easy availability, handiness and less time investment. 275

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Fig. 8. Overview of the mechanisms involved in the regulation of AST-host crosstalk by immunosuppression. AST can act at three different levels by: (1) inhibiting T cell activation by directly or indirectly affecting the splenocyte proliferation and inhibiting cytokine release; (2) phagocytosis inhibition by suppressing lysosomal enzyme release and NBT reduction; (3) signal transduction by interfering with upstream and downstream kinases as well as the transcription factors involved in the inflammatory and immune response activation.

responses are triggered through both the TCR and CD28 co-stimulatory receptor, and are thought to function primarily in various stress responses, and are sensitive to immunosuppressive drugs [41–43]. Cooperative activation of JNK and p38 leads to activation of transcription factors including AP-1, a heterodimer composed of fos and jun family members, which further stimulate the production of the chronic inflammatory mediators TNF-α, IL-1β, IL-10, TGF-β, and IL-6 [44]. In accordance with these reports, we found a significant decrease in c-Fos and c-Jun protein levels after AST treatment both in vitro and in vivo, which led to decreased TNF-α and IL-1β level. Indeed, as in other reports, the results of this study also revealed that down-regulation of the p38 MAPK signaling was strongly associated with reduction of TNF-α production and decreased levels of the above mentioned factors in the acute inflammatory microenvironment [45–49]. In activated macrophage cells, p38-MAPK leads to the production of NO by activating inducible NO synthase (iNOS) [50,51]. In agreement with this, our findings demonstrated that AST-mediated abrogation of p38 MAPK in macrophage cells resulted in decreased capacity to produce NO (Fig. 2D) by suppressing the iNOS expression level (Fig. 2F). Similar results were confirmed in zebrafish larvae (Fig. 5B). Furthermore, treatment with AST markedly inhibited LPS-mediated induction of two cyclooxygenase (COX) isoforms, COX-1 and COX-2, consistent with our previous finding that p38 activates COX-1 and COX2 by increasing production of IL-1β, which metabolizes arachidonic acid to PGH2, the common substrate for thromboxane A2 (TXA2), prostacyclin (PGI2), and PGE2 synthesis [52,53]. We observed suppression of COX-1 and COX-2 after AST treatment, which further led to a significant decrease in PGE2 secretion from LPS treated macrophage cells. COX-2 mediates prostaglandin synthesis (PGE2), which in turn promotes cell proliferation and cytokine secretion, while suppressing immune surveillance by mechanisms such as inhibition of Th-1 activation, IL-2 secretion, and DC suppression through IL-10 [54,55]. Finally, we validated the immunosuppressant potential of AST using in vitro proliferation of splenocytes prepared from mouse and observed that AST significantly suppressed the ConA stimulated splenocytes in a concentration dependent manner (Fig. 7). ConA was found to be

against any pathogen attack. Increase in ROS production is always a result of dysregulated immune function. For in vivo experiments, we used zebrafish larvae and checked the immunosuppressant effects of AST by measuring ROS levels and corresponding changes in the level of iNOS and COX-2. LPS stimulation increased the production of ROS in zebrafish larvae as evidenced in Fig. 4B. Increased ROS functions as a signal to induce inducible nitric oxide synthase (iNOS), which in result enhances the NO generation [33,34], and which in turn is associated with activation of inflammatory response against the pathogen attack. ROS increase also results in the binding of c-Jun complex at cAMP responsive element sites. During the present study, we analyzed the ROS intensity using DCFH-DA in live zebrafish larvae. Increased intensity after LPS treatment was correlated with increase expression of inflammatory marker COX-2 and iNOS (Fig. 5) and also with the protein expression of c-Jun and c-Fox (Fig. 6). The ROS intensity gradually decreased in a dose dependent manner after AST treatment. The process of apoptosis was determined using live staining of zebrafish larvae using acridine orange stain (AO). LPS resulted in increased fluorescence due to the presence of dead apoptotic bodies which was decreased after AST treatment (Fig. 4D). MAP kinase pathways and NF-kB are major pathways induced by TC stimulation that play a key role in T cell response [35]. The MAPK pathway is a conserved eukaryotic signaling cascade activated by TCR response that participates in a diverse array of biological processes and consists of three distinct subgroups of the MAPK superfamily: ERK, JNK and p38 [36–40]. Hence, next we addressed MAPK signaling pathways affected by the AST-based immunosuppression. We treated cells with AST after LPS treatment to suppress the LPS generated acute inflammatory microenvironment that activates the MAPK pathway [39]. We observed that the phosphorylation of p38 and JNK-MAPK in macrophage cells was strongly downregulated upon the AST treatment when compared with the LPS treated group, but that the phosphorylation of ERK was slightly increased, suggesting that AST had no effect on ERK activation (Fig. 6A and B). However p-ERK was also found to be downregulated in larvae (Fig. 6C). According to a previous report, among these three subgroups, JNK and p38 are activated when T cell 276

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associated with induction of T cell proliferation; thus, our results suggested that AST suppressed the activation of T cells, including cytotoxic T cells and helper T cells. Thus, we hypothesized that ConA-stimulated mouse splenocyte cells treated with AST also lost their ability to induce an allo-reactive or MHC-restricted cytotoxic T cell response.

[10]

[11]

5. Conclusion [12]

In conclusion, we demonstrated that AST has the ability to cause significant induction of immunosuppression in macrophage and splenocyte cells because of decrease T cell proliferation and Th1-related cytokine secretion in vitro. These effects were strongly related to the down-regulation of p38 and JNK-MAPK signaling in macrophage cells associated with a decreased production of acute inflammatory mediators (TNF-α, IL-1β, and PGE2). AST suppresses the activation of JNK, p38 and ERK by inhibiting COX-2 and iNOS in zebrafish larvae. Our findings also indicate that AST-based immunosuppression could efficiently condition the cell's microenvironment, suggesting that administration of AST in non-cytotoxic doses can be considered a novel therapeutic approach for decreasing the inflammatory potential of macrophage cells (Fig. 8).

[13] [14]

[15]

[16]

[17]

[18]

Acknowledgments

[19]

Not applicable.

[20]

Funding [21]

This research was supported by a NRF Research Grant from the Republic of Korea in 2016. [22]

Compliance with ethical standards [23]

All animals were approved by the committee for Laboratory Animal Care and use of Daegu University. All procedures were conducted in accordance with the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health.

[24] [25]

Appendix A. Supplementary data [26]

Supplementary data to this article can be found online at https:// doi.org/10.1016/j.intimp.2018.10.014.

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