Piperine impairs the migration and T cell-activating function of dendritic cells

Accepted Manuscript Title: Piperine impairs the migration and T cell-activating function of dendritic cells Author: Gemma Rodgers Carolyn D. Doucette David A. Soutar Robert S. Liwski David W. Hoskin PII: DOI: Reference:

S0378-4274(15)30120-X http://dx.doi.org/doi:10.1016/j.toxlet.2015.11.025 TOXLET 9279

To appear in:

Toxicology Letters

Received date: Revised date: Accepted date:

16-7-2015 23-11-2015 25-11-2015

Please cite this article as: Rodgers, Gemma, Doucette, Carolyn D., Soutar, David A., Liwski, Robert S., Hoskin, David W., Piperine impairs the migration and T cell-activating function of dendritic cells.Toxicology Letters http://dx.doi.org/10.1016/j.toxlet.2015.11.025 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Piperine impairs the migration and T cell-activating function of dendritic cells Gemma Rodgersa, Carolyn D. Doucettea, David A. Soutara, Robert S. Liwskia,b, and David W. Hoskina,b,c a

Department of Pathology, b Department of Microbiology and Immunology, c

Department of Surgery, Faculty of Medicine, Dalhousie University Halifax, Nova Scotia B3H 4R2, Canada

Correspondence to:

Dr. David W. Hoskin Department of Microbiology and Immunology Dalhousie University 5850 College Street, P.O. Box 15000 Halifax, Nova Scotia, Canada B3H 4R2 Telephone: 902-494-6509 E-mail: [email protected]

Running title: Piperine inhibits dendritic cell migration and function

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Highlights: 

Piperine is an alkaloid found in black pepper

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Piperine inhibits dendritic cell migration

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Piperine causes dendritic cells to retain an immature phenotype

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Piperine-treated dendritic cells are deficient in T cell-activating function

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Abstract Piperine, a major alkaloid found in the fruits of black and long pepper plants, has antiinflammatory properties; however, piperine’s effect on dendritic cell (DC) migration and T cell-activating function has not been investigated. Bone marrow-derived mouse DC that were matured in the presence of 100 µM piperine showed reduced in vitro migration in response to CCL21, as well as reduced in vivo migration to lymph nodes. In addition, piperine-treated DC had reduced CCR7 expression and elevated CCR5 expression, as well as reduced expression of CD40 and class II major histocompatibility complex molecules and decreased nuclear accumulation of RelB. DC production of interleukin (IL)-6, tumor necrosis factor, and monocyte chemoattractant protein-1 in response to lipopolysaccharide stimulation was also reduced following piperine treatment. Exposure to piperine during maturation therefore caused DC to retain an immature phenotype, which was associated with a reduced capacity to promote T cell activation since coculture of ovalbumin (OVA323–339)-specific T cells with OVA323–339-pulsed DC that were previously matured in the presence of piperine showed reduced interferon-γ and IL-2 expression. OVA323–339-specific T cell proliferation was also reduced in vivo in the presence of piperine-treated DC. Inhibition of DC migration and function by piperine may therefore be a useful strategy to down-regulate potentially harmful DC-driven T cell responses to self-antigens and transplantation antigens.

Keywords Cytokine; Dendritic cell; Migration; Piperine; T lymphocyte

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1. Introduction Dendritic cells (DCs) are antigen-presenting cells that are notable for their ability to efficiently activate naïve T cells (Banchereau and Steinman, 1998). This process begins with the stimulation of tissue-resident immature DC by pathogen-associated molecular pattern molecules, or other danger signals, and the subsequent capture of foreign antigens by DC, which results in a co-ordinately regulated process of DC maturation and migration to regional draining lymph nodes where the interaction with naïve T cells takes place (Tan and O’Neill, 2005). The resulting adaptive immune response is an important component of host defence against pathogens and other threats. Immature DCs that reside in peripheral tissues express high levels of the chemokine receptors CCR1, CCR2, and CCR5, which orchestrate the homing of DC to sites of inflammation (Sallusto et al., 1998; Sozzani et al., 1998). In peripheral tissues, DCassociated pattern recognition receptors that include Toll-like receptor (TLR) 3, TLR4, and TLR9, bind double-stranded RNA, lipopolysaccharide (LPS), and unmethylated CpG oligodeoxynucleotides, respectively (Trinchieri and Sher, 2007), resulting in DC activation and maturation with concomitant up-regulation of peptide-presenting major histocompatibility complex (MHC) class II and CD40, CD86, and CD80 co-stimulatory molecules (Chow et al., 2002; Granucci et al., 1999). Importantly, during the activation and maturation process, DCs down-regulate CCR5 expression and up-regulate CCR7 expression, which promotes CCL19- and CCL21-driven DC mobilization to lymph nodes (Sallusto et al., 1999; Sozzani et al., 1998). In this regard, CCL21 is the critical CCR7 ligand in regulating tissue-resident DC migration (Britschgi et al., 2010). CCL21 is produced constitutively by endothelial cells of lymphatic vessels, high endothelial

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venules and stromal cells present within the T cell zone of the lymph node (Gunn et al., 1998; Luther et al., 2000), and thus acts as homing beacon for lymph node access by DCs and DC-T cell interactions (Gunn et al., 1999). Within the lymph node, DCs process and present foreign peptide in the context of MHC I and II, which is recognized by CD8+ T cells and CD4+T cells, respectively, via the T cell receptor (Corse et al., 2011). A critical second signal is provided to T cells as a result of interactions between CD28 on T cells and the co-stimulatory molecules CD80 and CD86 on DCs (Chen and Flies, 2013). Finally, DCs help to create the cytokine environment that drives the differentiation of naïve T cells into different types of effector T cells (Satpathy et al., 2012). A number of bioactive phytochemicals are capable of inhibiting pattern recognition receptor-mediated inflammation, suggesting that a diet rich in these antiinflammatory phytochemicals may reduce the risk of developing chronic diseases associated with dysregulated inflammation caused by pattern recognition receptor stimulation (Zhao et al., 2011). Piperine, the major alkaloid present in the fruits of long pepper plants (Piper longum Linn) and black pepper plants (Piper nigrum Linn) (Madhavi et al., 2009), is responsible for the characteristic pungent smell and taste of black pepper, which has a long history of use in Ayurvedic medicine (Johri and Zutshi, 1992). As reviewed by Srinivasan (2007), piperine has diverse physiological effects on many cell types, including in vitro and in vivo anti-inflammatory activities (Meghwal and Goswami, 2013). For example, piperine inhibits the LPS-induced synthesis of proinflammatory tumor necrosis factor α (TNFα) by mouse macrophages (Bae et al., 2010), as well as constitutive expression of TNFα, interleukin (IL)-1β, and IL-6 by B16-F10 melanoma cells (Pradeep and Kuttan, 2004). Piperine also inhibits the TNFα-induced

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expression of intercellular adhesion molecule-1 (ICAM-1), vascular cell adhesion molecule-1 (VCAM-1) and E-selectin, which are involved in leukocyte extravasation into inflamed tissues (Kumar et al., 2007). In addition, piperine suppresses LPS-induced activation of extracellular signal-regulated and c-Jun N-terminal kinases in mouse DCs (Bae et al., 2012), as well as inhibiting signal transduction pathways that mediate T cell activation and clonal expansion (Doucette et al., 2015a; 2015b). In this study, we explored the effect of piperine on the ability of DCs to migrate and promote T cell activation. We show for the first time that piperine-treated DC failed to stimulate T cells in vivo, which we attribute to reduced DC migration to lymph nodes, as well as reduced expression of cell-surface molecules involved in DC-T cell interactions. Synthesis of T cells cytokines characteristic of a Th1 response was also diminished in the presence of piperine-treated DCs, suggesting selective inhibition of cell-mediated immunity.

2. Materials and methods 2.1. Mice Female C57BL/6 wild-type and OT-II mice were purchased from Charles River Laboratories (Lasalle, QC). All mice were housed at Dalhousie University’s Carleton Animal Care Facility and were fed a standard diet of mouse chow and water. All animal protocols were approved by Dalhousie Committee on Laboratory Animals and were in accordance with the Canadian Council on Animal Care Guidelines.

2.2. Generation of bone marrow-derived DCs Mature DCs were cultured from the bone marrow of C57BL/6 wild-type mice 6

essentially as previously described (Lutz et al., 1999). Briefly, bone marrow cells were isolated under aseptic conditions from the femur and tibia, then cultured in RPMI 1640 medium (Invitrogen Canada Inc., Burlington, ON) supplemented with 10% fetal calf serum (Wisent Inc., St-Bruno, QC), 2% penicillin/streptomycin, 1% HEPES (both from Invitrogen Canada Inc.), 5 mM 2-mercaptoethanol (Sigma-Aldrich Canada, Oakville, ON), and 20 ng/ml recombinant murine granulocyte-macrophage colony-stimulating factor (R&D Systems Inc., Minneapolis, MN). Fresh medium was added on day 3, and on day 6 non-adherent cells were transferred to fresh 6-well plates and cultured in fresh medium containing 10 ng/ml granulocyte-macrophage colony-stimulating factor. On days 3 and 6, DCs were treated with either 100 μM piperine (> 97%, Sigma-Aldrich Canada) or 0.1% ethanol (vehicle for piperine). On day 8, piperine-treated and control DCs were matured with 500 ng LPS (Sigma-Aldrich Canada) for 2 h and non-adherent cells were harvested. In some experiments, DCs were matured with 5 μg CpG oligodeoxynucleotide or 5 μg poly I:C (both from Sigma-Aldrich Canada) in place of LPS.

2.3. In vitro DC migration assay DC migration assays were performed in 24-well transwell plates (Corning Inc., Tewksbury, MA) containing inserts with 8 µm pores. The top and bottom of the membranes were coated for 1 h at 4°C with either phosphate-buffered saline (PBS) or 0.5% bovine serum albumin ([w/v]; fraction V; Sigma-Aldrich Canada). Wells were washed and 0.6 ml serum-free RPMI-1640 medium was added to each of the lower wells. In some wells, 500 ng CCL21 (R&D Systems, Inc.) was added to the lower chamber to serve as a chemoattractant. Piperine-treated or control DCs (4 x 105) were added to the top chamber of each insert and incubated for 5 h at 37°C. Cells that migrated to the 7

bottom chamber were harvested and counted using a hemocytometer. All migration conditions were done in duplicate and each experiment was repeated 3 times.

2.4. In vivo DC migration assay Piperine-treated or control DCs were harvested on day 8 and labelled with 5 µM CellTracker Green CMFDA (5-chloromethylfluorescein diacetate) dye (Invitrogen Canada Inc.), according to the manufacturer’s protocol. Briefly, DCs were labelled for 5 min, washed and incubated with warm RPMI-1640 medium for 30 min, washed 3 times, and then resuspended in PBS. DCs (1 x 106) in 20 µl of PBS were placed into the right forelimb of C57BL/6 wild-type mice by subcutaneous injection. After 48 h, the draining axial LNs were harvested and frozen in optimum cutting temperature solution containing 20% sucrose. Lymph nodes were cut in 10 µm sections using a cryostat (Leica Microsystems Inc., Concord, ON) and transferred onto sialinated slides. Lymph node sections were blocked with 5% goat serum (v/v) and incubated overnight with rat antimouse pan-reticular fibroblast marker IgG2a antibody (Ab) (Cedarlane Laboratories, Burlington, ON), followed by incubation with goat anti-rat Cy3 Ab (Jackson ImmunoResearch Laboratories, Inc., West Grove, PA). Whole lymph node sections containing high endothelial venules, where DCs predominately cluster (Luther et al., 2000), were photographed using a Nikon E-600 fluorescent microscope and DCs in each field were counted in a blinded manner.

2.5. Flow cytometry In order to assess the phenotype of piperine-treated and control DCs, 5 x 105 cells were suspended in flow cytometry buffer (2% fetal calf serum [v/v], 0.5% EDTA [w/v],

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0.1% sodium azide [w/v] in PBS) and incubated with 1:100 anti-CD16.1/32.1 Ab for 30 min to block Fc receptors. cells were then washed and incubated with an optimal dilution of the desired Ab for 30 min at 4°C for 30 min. The following Abs were used for DC phenotyping by flow cytometry: allophycocyanin (APC)-conjugated anti-CD11c Ab, phycoerythrin (PE)-conjugated anti-CD195 (CCR5) Ab, PE-conjugated rat IgG2a, APCconjugated Armenian hamster IgG1, and PE-conjugated Armenian Hamster IgG (eBioscience Inc., San Diego, CA); Alexa Fluor 488-conjugated anti-MHC II (I-A/I-E) Ab, Alexa Fluor 488-conjugated anti-CD54 (ICAM-1) Ab, PE-conjugated anti-CD197 (CCR7) Ab, and Alexa Fluor 488-conjugated rat IgG2b (BioLegend, San Diego, CA); PE-conjugated anti-CD86 Ab, PE-conjugated anti-CD40 Ab, fluorescein isothiocyanate (FITC)-anti-CD11b/CD18 (Mac-1) Ab ( Cedarlane Laboratories Ltd.); and PEconjugated anti-CD11a (LFA-1) Ab (BD Biosciences). Following additional washes, Ablabelled cells were fixed in 2% paraformaldehyde and fluorescence was determined using a FACSCalibur flow cytometer (BD Biosciences, Mississauga, ON). In some experiments, cell viability was determined by flow cytometric analysis of cells stained with 7-amino-actinomycin D (7-AAD) Viability Staining Solution (eBioscience Inc.), according to the manufacturer’s instructions. Fluorescence data was analyzed using FCS Express V3 software (DeNovo Software, Thornhill, ON). Analysis was performed by gating on CD11c-expressing cells, which comprised >90% of the population, to ensure that findings were DC-specific.

2.6. Cytokine analysis

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For analysis of DC cytokines, piperine-treated and control DCs were harvested on day 7 and plated at 4 x 105 cells/ml, then treated with 500 ng LPS for 18 h. Cell-free supernatants were harvested and cytokine analysis was performed using a Cytometric Bead Array Mouse Inflammation Kit (BD Biosciences), according to the manufacturer's instructions. Briefly, 50 μl supernatant was incubated with fluorescent capture beads coated with Abs against IL-6, monocyte chemoattractant protein (MCP)-1, and TNF-α. After 2 h incubation with PE-conjugated secondary Abs, fluorescence intensity was determined by flow cytometry and quantified relative to a standard curve.

For analysis of T cell cytokines, CD4+ T cells specific for chicken ovalbumin (OVA323–339) peptide were isolated from the spleens of OT-II mice using the MACS CD4+ negative isolation system (Miltenyi Biotech, Auburn, CA), according to the manufacturer’s protocol. Piperine-treated and control DCs were harvested on day 8 and loaded with 300 nM OVA323–339 peptide (Sigma-Aldrich Canada), then matured with LPS for 2 h at 37°C. CD4+ T cells were seeded into a 96-well round-bottom plate (1 x 105 cells/well) and co-cultured with peptide-pulsed DCs at a 10:1 ratio. As a negative control, CD4+ T cells were co-cultured with DCs that were not loaded with OVA323–339 peptide. After 72 h culture, cell-free supernatants were harvested and cytokine content was determined using enzyme-linked immunosorbent assay (ELISA) kits for IL-2 and interferon (IFN)-γ (R&D Systems, Inc.) or a Cytometric Bead Array Mouse Inflammation Kit for IL-4 and IL-17 (BD Biosciences), according to the manufacturer's instructions.

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2.7. Western blot analysis Piperine-treated and control DCs from day 8 cultures were matured with LPS for 2 h. Cells were then washed twice with PBS, resuspended in 1.5 mL of ice-cold lysis buffer (10 mM HEPES [pH 7.9], 10 mM potassium chloride, 1 mM dithiothreitol, 0.5 mM EDTA, 0.5 mM EGTA, 0.1 mM phenylmethylsulfonyl fluoride, 0.1% nonyl phenoxylpolyethoxylethanol, and 1x protease inhibitor cocktail) (all from Sigma-Aldrich Canada) and incubated on ice for 15 min. Cells were gently homogenized with a Dounce homogenizer and centrifuged at 1000 g for 10 min at 4°C. Supernatants comprising the cytoplasmic fraction were collected. The pellet was resuspended in 2 volumes of ice-cold saline buffer (20 mM HEPES [pH 7.9], 420 mM sodium chloride, 1 mM EDTA, 1 mM EGTA, 10% glycerol, 1 mM dithiothreitol, and 1x protease inhibitor cocktail) and incubated at 4°C for 30 min on rocking platform to lyse nuclei. Lysates were centrifuged at maximum speed for 10 min at 4°C and supernatants comprising the nuclear fraction were collected. Proteins in cytoplasmic and nuclear lysates were resolved by sodium dodecyl sulphate-polyacrylamide gel electrophoresis and transferred to polyvinylidene difluoride membranes (Millipore, Billerica, MA), which were incubated in blocking solution containing Tris-buffered saline (20 mM Tris–HCl [pH 7.6], 200 mM NaCl) and 0.05% Tween-20 (v/v) (TBST) with 5% fat-free milk (w/v) for 1 h at room temperature. To confirm proper fractionation of cytoplasmic and nuclear compartments, the cytoplasmic protein cathepsin D was probed for using anti-mouse cathepsin D Ab (R&D Systems Inc.) and the nuclear protein lamin was probed for using anti-mouse lamin Ab (BD Biosciences). RelB was probed for using anti-rabbit RelB Ab (Santa Cruz Biotechnology Inc., Santa Cruz, CA). Membranes were incubated with an optimal

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dilution of primary Abs in blocking buffer for 1 h at room temperature on a rocker, then washed extensively with TBST. Membranes were next incubated with an optimal dilution of the appropriate horseradish peroxidase-conjugated secondary Ab (mouse anti-rabbit IgG, donkey anti-goat IgG, or goat anti-mouse IgG) (all from Jackson ImmunoResearch Laboratories Inc.) for 1 h at room temperature on a rocker, then washed extensively with TBST. Protein bands were detected using an enhanced chemiluminescent kit (GE Healthcare, Baie d’Urfe, QC) and exposure to X-ray film (Sci-Med Inc., Truro, NS). Densitometry was assessed using NIH ImageJ software.

2.8. In vivo T cell proliferation assay Piperine-treated and control DCs were harvested on day 8 and loaded with 300 nM OVA323–339 peptide, then matured with LPS for 2 h at 37°C. DCs (1 x 106 cells in 20 µl PBS) were placed into the right forelimb of C57BL/6 wild-type mice by subcutaneous injection. The following day, CD4+ T cells specific for OVA323–339 peptide were isolated from the spleens of OT-II mice using the MACS CD4+ negative isolation system (Miltenyi Biotech, Auburn, CA) and labelled with 5 μM carboxyfluorescein diacetate succinimidyl ester (CFSE). Mice were inoculated with CSFE-labeled CD4+ T cells (5 x 106 cells in 0.1 ml PBS) by tail vein injection. After 72 h, the draining axial and brachial lymph nodes were harvested and a single cell suspension was created using 30G needles and a cell strainer. The proliferation of the CFSE-labeled OT II T cells within the lymph nodes was determined by flow cytometry. Contralateral brachial and axial lymph nodes from the same mouse were harvested and used as a negative control. The percentage of the original T cell population that underwent proliferation (percent responders) was calculated as previously described (Lyons, 2000). Briefly, the number of daughter cells 12

within each division was divided by the precursor extrapolation (2n) to estimate the number of precursor T cells from the original population that proliferated. Following extrapolation, precursor numbers were summed and the number of precursors that proliferated was divided by the total number of precursors and multiplied by 100 to generate the percent responders, also known as the division precursor frequency, of the original T cell population.

2.9. Statistical analysis Variability between replicate experiments is expressed as standard error of the mean (SEM). Multiple datasets was compared using one-way analysis of variance (ANOVA) with a Tukey-Kramer post-test. A paired Student’s t-test was used to compare paired and normally distributed datasets. A 1-sample t-test was used when a dataset was being compared to a normalized control. Results were considered to be statistically significant when p < 0.05. NS indicates p > 0.05 and were not statistically significant.

3. Results 3.1. Piperine inhibits DC migration Given that DC migration to lymph nodes is crucial for antigen-specific activation of naïve T cells, we investigated the impact of DC maturation in the presence of 100 µM piperine on the subsequent ability of DCs to migrate in response to CCL21, which is a ligand for CCR7 (Britschgi et al., 2010). As shown in Fig. 1, the migration of piperinetreated DC in an in vitro transwell assay was significantly reduced in comparison to untreated control DCs. Furthermore, there was no significant difference between the

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migration of piperine-treated and control DCs in the absence of CCL21, suggesting that piperine-treated DCs were refractory to CCL21. An initial dose-response study established that an optimal inhibitory effect on DC migration was achieved with 100 µM piperine, which was shown by flow cytometric analysis of 7-AAD-stained DCs to have no adverse effect on DC viability (Supplemental Fig. S1). To determine whether piperine treatment also inhibited DC migration in vivo, piperine-treated and control DCs were activated with LPS, then labelled with CMFDA and injected into the forelimb skin of syngeneic mice. As shown in Fig. 2, draining lymph nodes from mice injected with piperine-treated DCs contained significantly fewer migrated DCs than lymph nodes from mice injected with control DCs (28 ± 2 cells/field versus 130 ± 22 cells/field, respectively). 3.2. Piperine alters maturation-associated chemokine receptor expression by DCs Since piperine-treated DCs were refractory to CCL21 stimulation in vitro and exhibited decreased migration in vivo, we assessed the effect of piperine on chemokine receptor expression by LPS-matured DCs. Flow cytometric analysis of piperine-treated DCs demonstrated a significant decrease in CCR7 expression and significantly higher levels of CCR5 expression in comparison to control DCs (Fig. 3), indicating that piperine inhibited the maturation-associated DC chemokine receptor switch from CCR5 to CCR7 involved in DC mobilization from peripheral tissues to lymph nodes (Sallusto et al., 1998; Sozzani et al., 1998). The piperine-induced increase in CCR5 expression was dosedependent (Supplemental Fig. S2). 3.3. Piperine inhibits pathogen-associated molecular pattern-induced expression of CD40 and MHC II by DCs

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We next determined the effect of piperine on LPS-induced expression of additional maturation markers by DCs. As shown in Fig. 4, in comparison to vehicletreated DCs, piperine-treated DCs had significantly decreased expression of CD40 and MHC class II. The piperine-induced decrease in CD40 and MHC class II expression was dose-dependent (Supplemental Fig. S3). In contrast, CD86 expression was not affected by piperine. Interestingly, the MHC class II high population was dramatically reduced following exposure to piperine, which was in line with a decrease in the number of highly mature DCs. Flow cytometric analysis showed that piperine by itself, i.e., without LPS, did not affect CD40, CD86 or MHC class II expression by DCs (Supplemental Fig. S4), indicating that piperine alone did not activate DCs. In addition, piperine did not affect TLR4 expression (Supplemental Fig. S5), suggesting that piperine-treated DCs remained capable of binding LPS. In addition, piperine treatment did not alter expression of the adhesion molecules ICAM-1, MAC-1, and LFA-1 by LPS-matured DCs (Supplemental Fig. S6). As an alternative to LPS-induced maturation, DCs were matured with CpG (TLR9 agonist) or poly I:C (TLR3 agonist) in the absence or presence of 100 µM piperine. As with LPS-matured DCs, DCs matured with CpG or poly I:C in the presence of piperine had decreased expression of MHC class II and CD40 in comparison to control DCs (Fig. 5A and 5B, respectively), indicating that the inhibitory effect of piperine on DC maturation was not restricted to the LPS/TLR4 axis. 3.4. Piperine inhibits LPS-induced cytokine synthesis by DCs DCs are an important source of chemokines and pro-inflammatory cytokines (Blanco et al., 2008). We therefore determined the effect of piperine on LPS-induced production of TNFα and IL-6, as well as MCP-1, by DCs. In comparison to control DCs,

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piperine-treated DCs synthesized significantly less TNFα (Fig. 6A), IL-6 (Fig. 6B), and MCP-1 (Fig. 6C) in response to LPS stimulation. 3.5. Piperine inhibits nuclear translocalization of RelB Since the NF-κB subunit RelB is present at a high level in the nuclei of mature DCs relative to immature DCs (Neumann et al., 2000), we predicted that treatment with piperine might impact nuclear translocalization of RelB. As shown in Fig. 7, western blot analysis of cytoplasmic and nuclear fractions from DCs that were matured in the presence of piperine showed a significant reduction in the amount of nuclear RelB in comparison to control DCs. In contrast, there was no significant difference between cytoplasmic RelB in control and piperine-treated DCs. 3.6. Differential effect of piperine-treated DCs on cytokine production by T cells Given that piperine inhibited DC secretion of cytokines and expression of CD40 and MHC class II, which are important for antigen-specific T cell activation by DCs (Banchereau and Steinman, 1998), we compared the ability of control and piperinetreated DCs to promote cytokine synthesis by OVA323–339-specific CD4+ T cells. Coculture of OVA323–339 -specific CD4+ T cells with OVA323–339 peptide-pulsed DCs resulted in reduced production of IFN-γ (Fig. 8A) and IL-2 (Fig. 8B) when DCs were matured in the presence of piperine; however, there was no significant effect of piperinetreated DCs on T cell synthesis of IL-4 (Fig. 8C) and IL-17A (Fig. 8D). 3.7. Piperine ablates DC-driven T cell responses in vivo Finally, we determined whether piperine treatment of DCs impacted their ability to activate T cells in vivo. Control and piperine-treated DCs were pulsed with OVA323–339 peptide, and injected into the forearm skin of syngeneic mice, followed by intravenous

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injection of CFSE-labeled OVA323–339 -specific T cells. T cell proliferation was assessed by flow cytometric analysis of draining lymph node cells. As shown in Fig. 9, in comparison to control DCs, piperine-treated DCs failed to induce T cell proliferation.

4. Discussion The anti-inflammatory and immunomodulatory properties of piperine suggest that this dietary phytochemical may be useful in controlling chronic inflammation associated with pattern recognition receptor stimulation (Meghwal and Goswami, 2013; Srinivasan, 2007); however, little is known about the effect of piperine on the function of DCs, which play a central role in the pathogenesis of a number of human inflammatory diseases (Blanco et al., 2008). In this study, we report for the first time that piperine inhibited the in vitro and in vivo migration of DCs, as well as DC-induced T cell proliferation in vivo. We attribute piperine-mediated inhibition of DC migration to impairment of the chemokine receptor switch from CCR5 to CCR7 since CCL21 causes DC migration via stimulation of CCR7 (Yoshida, 1998). Reduced in vivo migration of piperine-treated DCs was likely also caused by reduced expression of CCR7. In addition, piperine-treated DCs exhibited decreased LPS-induced expression of CD40 and MHC II molecules, as well as reduced synthesis of the pro-inflammatory cytokines, TNF-α, IL-6, MCP-1. Furthermore, exposure to piperine caused a significant decrease in MHC II high-expressing DC, which is indicative of a reduction in highly mature DCs (Lutz et al., 1999). Taken together, these findings suggest that exposure to piperine caused DCs to retain an immature or semi-mature DC phenotype in spite of TLR4 stimulation with LPS. Interestingly,

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piperine-treated DCs have increased phagocytic activity (Bae et al., 2012), which is also consistent with an immature DC phenotype. Our findings are in general agreement with two previous studies that showed reduced expression of maturation markers and proinflammatory cytokines following DC exposure to piperine (Bae et al., 2012; Cavalcanti et al., 2014); however, unlike these earlier reports, we observed reduced synthesis of IL-6 but no effect on CD86 expression by piperine-treated DCs. Reduced production of proinflammatory cytokines has also been reported in piperine-treated B16-F10 melanoma cells, adipocytes, and macrophages (Pradeep and Kuttan, 2003, 2004; Woo et al., 2007), indicating that this effect is not cell type-specific. Moreover, normal expression of ICAM-1, MAC-1, and LFA-1 adhesion molecules by piperine-treated DCs shows selective inhibition of DC surface marker expression, which may reflect targeting of specific signal transduction pathways. Indeed, piperine is a known inhibitor of extracellular signal-regulated and c-Jun N-terminal kinases that are activated in mouse DCs as a result of LPS stimulation (Bae et al., 2012). Piperine-mediated inhibition of DC maturation marker expression induced by LPS, CpG, and poly I:C indicates that the inhibitory effect of piperine on DC maturation caused by pattern recognition receptor stimulation was not restricted to the LPS/TLR4 axis. Furthermore, this finding suggests that piperine modulates signaling pathways associated with the TLR adaptor molecule Toll/IL-1R domain-containing adaptor inducing interferon (TRIF) and the TLR adaptor protein MyD88 since TLR3 agonists such as poly I:C act via TRIF, TLR9 agonists such as CpG act via MyD88, and TLR4 agonists such as LPS activate both TRIF and MyD88 signaling pathways (Trinchieri and Sher, 2007). It therefore seems likely that piperine acts downstream of TRIF and MyD88

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by targeting the transcription factor NF-κB, which promotes the expression of genes involved in the inflammatory response (Gasparini and Feldmann, 2012). Other investigators have also reported piperine-mediated inhibition of the NF-κB pathway. For example, nuclear localization of the NF-κB subunits p65, p50, and cRel is inhibited in piperine-treated B16-F10 melanoma cells (Pradeep and Kuttan, 2004), while piperine interferes with NF-κB transcriptional activation in HT-1080 fibrosarcoma cells (Hwang et al., 2011). Piperine also inhibits the degradation of IκBα in TNFα-stimulated endothelial cells and PMA-stimulated HT-1080 cells ( Kumar et al., 2007; Hwang et al., 2011). However, piperine does not affect the IL-1β-induced transcriptional activity of NF-κB in synoviocytes (Bang et al., 2009). Piperine-mediated inhibition of the NF-κB pathway therefore appears to be cell type-specific. We report here that piperine-treated DCs had decreased nuclear levels of the NFκB subunit RelB, which is normally abundant in the nuclei of DCs matured by stimulation with pathogen-associated molecular pattern molecules (Neumann et al., 2000). Since RelB is critical for DC maturation and antigen-presenting function (Zanetti et al., 2003) and NF-κB regulates CCR7 transcription (Sanchez-Sanchez et al., 2006), it is reasonable to conclude that decreased RelB expression in the nuclei of piperine-treated DCs accounts at least in part for the inhibitory effect of piperine on DC maturation, migration, and T cell stimulation observed in the present study. However, it is important to note that piperine also inhibits mitogen activated protein kinase signaling in various cell types (Hwang et al., 2011; Bang et al., 2009), including DCs (Bae et al., 2012). We suggest that piperine-mediated inhibition of mitogen activated protein kinases also contributes to the failure of DCs to migrate in response to CCL21 since CCR7-induced

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chemotaxis of DCs involves the activation of extracellular signal-regulated kinase and p38 mitogen-activated protein kinase (Riol-Blanco et al., 2005). Antigen-specific stimulation of OVA323–339 -specific CD4+ T cells with piperinetreated DCs resulted in decreased production of IL-2 and IFN-γ, indicating that T helper cells were directed away from the Th1 phenotype that is typically induced by DCs differentiated with granulocyte-macrophage colony-stimulating factor (Eksioglu et al., 2007). In contrast, there was no reduction in CD4+ T cell production of IL-4 and IL-17, suggesting that T helper cells had not been alternatively directed to a Th2 or Th17 phenotype. A similar decrease in CD4+ T cell synthesis of IFN-γ with no change in IL-4 production occurs when CD4+ T cells are stimulated with DCs that were treated with dexamethasone prior to LPS maturation (Matyszak et al., 2000). Interestingly, multiple restimulation of CD4+ T cells with dexamethasone-treated DCs resulted in the selective induction of T regulatory cells. Although we did not determine whether piperine-treated DCs also induced T regulatory cells, such an outcome seems unlikely given that T regulatory cells require IL-2 for their development and function (de la Rosa et al., 2004; Buchill et al., 2007). We also observed that the in vivo proliferation of OVA323–339 specific CD4+ T cells was virtually ablated when piperine-treated DCs were used as antigen-presenting cells. This profound inhibitory effect on CD4+ T cell activation was likely due to reduced migration of piperine-treated DCs to lymph nodes and impaired interactions with CD4+ T cells as a result of decreased DC expression of CD40 and MHC II molecules. In conclusion, we have demonstrated a direct inhibitory effect of piperine on DC maturation, chemokine receptor expression and migration, as well as an alter T helper

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cell cytokine profile and impaired T helper cell proliferation following stimulation with piperine-treated DCs. It is important to note that the 100 µM concentration of piperine used throughout our study is substantially higher than the concentration that could be reasonably achieved in extravascular spaces where DCs are resident by diet alone; however, it may be feasible to achieve an effective systemic concentration of piperine by oral administration of the purified phytochemical as a pharmacologic intervention since oral gavage of rats with piperine at 54 mg/kg results in a peak plasma concentration of 20 µM (Liu et al., 2011). Since DCs play an important role in the pathogenesis human inflammatory diseases (Blanco et al., 2008), including of inflammatory bowel disease (Rutella and Locatelli, 2011), we suggest that the inhibitory effect of piperine on DC maturation and function may have clinical application. Acknowledgments This work was supported by a grant from the Natural Sciences and Engineering Research Council of Canada (NSERC) and a regional partnership grant from the Canadian Institutes of Health Research/Crohn's and Colitis Foundation of Canada/Nova Scotia Health Research Foundation (NSHRF). C.D. was supported by a NSERC Postgraduate Scholarship and a NSHRF Student Research Award.

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Figure legends Fig. 1. Piperine inhibits DC migration in vitro. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with LPS, then added to transwell plates to measure CCL21-induced migration. Migrated cells were counted and baseline migration was determined. Fold increases over baseline migration by control DCs (black) and piperine-treated DCs (white) were determined and results are expressed as mean ± SEM of 3 independent experiments; * indicates p < 0.05, as calculated by one way ANOVA with Tukey-Kramer multiple comparisons post-test.

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Fig. 2. Piperine inhibits DC migration in vivo. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with LPS, then stained with CMFDA dye and injected into the forearm of C57BL/6 mice. After 48 h, DC migration to axial lymph nodes was determined by fluorescence microscopy of lymph node sections stained with anti-mouse pan-reticular fibroblast marker Ab or an isotype control Ab. DCs in 5 high powered photographs from each lymph node were counted in a blinded manner. Images shown are from a representative experiment. The bar graph shows the mean number of control DCs (black) or piperine-treated DCs (white) per mouse ± SEM of 4 independent experiments; * indicates p < 0.05 compared to vehicle control, as calculated by paired Student’s t test.

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Fig. 3. Piperine modulates DC chemokine receptor expression. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with LPS, then stained for CCR7 or CCR5. Representative flow cytometry histograms are shown. In all histograms, grey lines to the left represent cells stained with an isotype control Ab, while black lines represent cells stained with Ab against CCR7 or CCR5. Bar graphs show the mean % control cells (black) or piperine-treated cells (white) positive for CCR7 or CCR5 ± SEM of data obtained from the analysis of single samples in 5 and 3 independent experiments, respectively; * indicates p < 0.05 compared to vehicle control, as calculated by paired Student's t test.

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Fig. 4. Piperine down-regulates CD40 and MHC II expression by LPS-matured DCs. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with LPS, then expression of CD40, CD86, and MHC II was determined by flow cytometry. Representative flow cytometry histograms are shown. In all histograms, grey lines to the left represent cells stained with an isotype control Ab, while black lines represent cells stained with Ab against CD40, CD86 or MHC II. Bar graphs show the mean % control cells (black) or piperine-treated cells (white) positive for CD40, CD86 or MHC II ± SEM of 3 independent experiments; * indicates p < 0.05, NS indicates p > 0.05, compared to vehicle control, as calculated by paired Student's t test.

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Fig. 5. Piperine down-regulates CD40 and MHC II expression by CpG- or poly I:Cmatured DCs. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with (A) CpG or (B) poly I:C. Expression of CD40 and MHC II was determined by flow cytometry. The bar graphs show (A) the mean fluorescence intensity (MFI) of anti-CD40 Ab-stained control cells (black) or piperine-treated cells (white) ± SEM of 3 independent experiments and (B) the mean % control cells (black) or piperinetreated cells (white) stained for MHC II ± SEM of 3 independent experiments; * indicates p < 0.05, NS indicates p > 0.05, compared to vehicle control, as calculated by paired Student's t test.

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Fig. 6. Piperine inhibits DC cytokine synthesis. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with LPS. Supernatants were collected 24 h later and TNFα, IL-6, and MCP-1 levels were determined by cytometric bead array. Data are expressed as amount of (A) TNFα, (B) IL-6, and (C) MCP-1 produced by piperinetreated DCs (white) relative to control DCs (black) ± SEM from at least 3 independent experiments; * indicates p < 0.05 compared to vehicle control, as calculated by 1-sample t test.

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Figure 7 Piperine inhibits RelB translocation to the DC nucleus. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with LPS. Lysates of cytoplasmic and nuclear fractions were then isolated and subjected to western blot analysis for RelB. Equal loading and proper nuclear and cytoplasmic fractionation was confirmed by probing for lamin and cathepsin D. A representative blot is shown. Bar graphs show the mean cytoplasmic and nuclear RelB localization in control DCs (black) or piperine-treated DCs (white) relative to the corresponding cytoplasmic or nuclear housekeeping genes ± SEM of 3 independent experiments, as determined by densitometry; * indicates p < 0.05, NS indicates p > 0.05 compared to vehicle control, as calculated by paired Student's t test.

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Figure 8 Piperine-treated DCs alter the cytokine secretion profile of T cells. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with LPS. DCs were then loaded with OVA323–339 peptide, and co-cultured with naïve OT-II CD4+ T cells for 72 h. Levels of (A) IFN-γ, (B) IL-2, (C) IL-4, and (D) IL-17 in cell-free supernatants were determined by (A, B) ELISA or (C, D) cytokine bead. Bar graphs show the mean cytokine levels in culture containing control DCs (black) or piperinetreated DCs (white) ± SEM of 3 independent experiments; * indicates p < 0.05, NS indicates p > 0.05 compared to vehicle control, as calculated by paired Student's t test.

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Fig. 9. Piperine-treated DCs have a reduced capacity to activate T cells in vivo. DCs were exposed to 100 μM piperine or the ethanol vehicle prior to maturation with LPS. DCs were then loaded with OVA323–339 peptide, and injected into the forearm of C57BL/6 mice, which then received CFSE-labeled OT-II CD4+ T cells by tail vein injection. After 72 h axial and brachial lymph nodes were harvested for assessment of T cell proliferation by flow cytometry. A representative flow cytometry histogram is shown. The bar graph shows the mean % responder T cells ± SEM from 4 independent experiments, where % responders is the percentage of the original T cell population that underwent proliferation and was calculated by extrapolation of the number of cells in each division peak relative to the number of cell divisions associated with that peak; * indicates p < 0.05, as calculated by paired Student's t test.

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