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JOURNAL OF INTERFERON & CYTOKINE RESEARCH Volume 00, Number 00, 2015 ª Mary Ann Liebert, Inc. DOI: 10.1089/jir.2015.0029

Soluble Mediators in Platelet Concentrates Modulate Dendritic Cell Inflammatory Responses in an Experimental Model of Transfusion Alexis J. Perros,1,2 Anne-Marie Christensen,1,2 Robert L. Flower,1,2 and Melinda M. Dean1,2

The transfusion of platelet concentrates (PCs) is widely used to treat thrombocytopenia and severe trauma. Ex vivo storage of PCs is associated with a storage lesion characterized by partial platelet activation and the release of soluble mediators, such as soluble CD40 ligand (sCD40L), RANTES, and interleukin (IL)-8. An in vitro whole blood culture transfusion model was employed to assess whether mediators present in PC supernatants (PC-SNs) modulated dendritic cell (DC)-specific inflammatory responses (intracellular staining) and the overall inflammatory response (cytometric bead array). Lipopolysaccharide (LPS) was included in parallel cultures to model the impact of PC-SNs on cell responses following toll-like receptor-mediated pathogen recognition. The impact of both the PC dose (10%, 25%) and ex vivo storage period was investigated [day 2 (D2), day 5 (D5), day 7 (D7)]. PC-SNs alone had minimal impact on DC-specific inflammatory responses and the overall inflammatory response. However, in the presence of LPS, exposure to PC-SNs resulted in a significant doseassociated suppression of the production of DC IL-12, IL-6, IL-1a, tumor necrosis factor-a (TNF-a), and macrophage inflammatory protein (MIP)-1b and storage-associated suppression of the production of DC IL-10, TNF-a, and IL-8. For the overall inflammatory response, IL-6, TNF-a, MIP-1a, MIP-1b, and inflammatory protein (IP)-10 were significantly suppressed and IL-8, IL-10, and IL-1b significantly increased following exposure to PC-SNs in the presence of LPS. These data suggest that soluble mediators present in PCs significantly suppress DC function and modulate the overall inflammatory response, particularly in the presence of an infectious stimulus. Given the central role of DCs in the initiation and regulation of the immune response, these results suggest that modulation of the DC inflammatory profile is a probable mechanism contributing to transfusion-related complications.

others 1993; Spiess and others 2004; Aslam and others 2008; Kiefel 2008; Bilgin and others 2011; Refaai and others 2011; Sahler and others 2011). Poorer clinical outcomes in patients post-transfusion, associated with modulation of the recipient’s inflammatory profile and immune response, are referred to as transfusionrelated immunomodulation (TRIM) (Aslam and others 2008; Bilgin and Brand 2008). The mechanisms underlying TRIM as a result of PC transfusion remain largely unknown. A few mechanisms have been proposed, including the presence of residual leukocytes in PCs (Slichter and others 2005) and the release of major histocompatibility complex (MHC) class I molecules associated with PC storage (Ghio and others 1999). Despite these hypotheses, the underlying mechanisms, key cells, and mediators of adverse transfusion complications post-PC transfusion remain largely undefined.

Introduction

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latelets (PLTs) are key contributors to coagulation and wound healing, as well as providing a significant contribution to inflammation and innate immune responses ( Jurk and Kehrel 2005). Transfusion of platelet concentrates (PCs) is standard in the treatment of thrombocytopenia and severe trauma (Hunt 1998; Sahler and others 2012). In addition, the transfusion of PCs is used prophylactically to maintain hemostasis in patients undergoing ablative chemotherapy before bone marrow transplantation (Hunt 1998; Sahler and others 2012). While it is clear that PC transfusions are an effective therapy, they have the potential to modulate the immune system, contributing to poor clinical outcomes, including increased length of stay in intensive care, prolonged ventilation, infection, mortality, and severe trauma (Heddle and

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Research and Development, Australian Red Cross Blood Service, Brisbane, Australia. Faculty of Health, School of Biomedical Sciences, Queensland University of Technology, Brisbane, Australia.

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PCs are routinely stored for up to 7 days before transfusion, although country-specific guidelines exist [eg, 7-day expiry in Ireland, Denmark, the Netherlands, and the United Kingdom (Korte 2011; Sto¨rmer and Vollmer 2014) and 5-day expiry in Australia (National Health and Medical Research Council and the Australasian Society of Blood Transfusion 2002)]. The 7-day expiry was introduced in some countries following the introduction of bacterial screening for all PLT products and extensive quality testing comparing day 5 (D5) and day 7 (D7) PCs. While Australia has introduced bacterial screening, the current shelf life remains at 5 days. Significant biochemical and biological changes occur during ex vivo storage of PCs (Sparrow 2010; Spiess 2010; Sahler and others 2011). During storage, PLTs become partially activated, lose metabolic function, and PLT matrix adhesion properties are altered (Boomgaard and others 1994; Rosenfeld and others 1995; Seghatchian 2006; Shrivastava 2009; Sahler and others 2011). In addition, the release of biological mediators, including histamine (Konca and others 2006), soluble CD40 ligand (sCD40L) (Kaufman and others 2007), P-selectin, RANTES, PLT factor-4 (PF4), transforming growth factor-b (TGF-b), and interleukin (IL)8 (Apelseth and others 2006; Sahler and others 2011), have widely been reported to increase in the PC during routine storage. Despite these biochemical and biological changes, the relationship between blood product storage before transfusion and poorer clinical outcomes in patients remains controversial (Boehlen and Clemetson 2001; Karkouti and others 2006; Koch and others 2008; Sparrow 2010; Spiess 2010; Sahler and others 2011). The accumulation of sCD40L, in particular, could have a detrimental impact on clinical outcomes by inducing inflammation, impacting other mediators downstream, inducing cytokine and chemokine release and adhesion molecule surface expression, and driving dendritic cell (DC) maturation (Martinson and others 2004; Kaufman and others 2007; Hamzeh-Cognasse and others 2008; Sahler and others 2012). The presence of biological mediators in PCs may affect the function of important immune cells such as DCs. DCs play a central role at the interface of the innate and adaptive immune response as they are the only antigen-presenting cells that can present antigens to naı¨ve T cells (Lee and Iwasaki 2007). Therefore, dysfunction in the DC, a key immunoregulatory cell, may contribute to adverse complications of transfusion contributing to TRIM. In humans, there are 2 main subtypes of blood DCs, the myeloid DCs; CD11c+ with a monocytoid appearance, which produce tumor necrosis factor-a (TNF-a), and plasmacytoid DCs; CD11c-, which are morphologically similar to plasma cells and produce interferons, in particular interferon-a (IFN-a) (Reid and others 1992; Robinson and others 1999; Adams and others 2005). While these cells have a similar capacity to stimulate an immune response (Banchereau and others 2000), they differ in terms of tissue distribution, cytokine production, and growth requirements (Banchereau and others 2000; Adams and others 2005). DCs express a number of pattern recognition receptors (PRRs) that interact with pathogen-associated molecular patterns or damage-associated molecular patterns. PRRs expressed by DCs include C-type lectins, mannose receptors, and toll-like receptors (TLRs) (Lee and Iwasaki 2007).

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Upon activation of PRRs and receptor/ligand binding, DCs undergo differentiation and antigens are processed for presentation on MHC class I and/or MHC class II molecules. Together with changes in expression of adhesion molecules and TLRs, antigen presentation mediates DC migration from the tissue to the draining lymph node through the lymphatic system (Sallusto and others 1998; Lee and Iwasaki 2007). During this process, costimulatory molecules and cytokines (such as IFN-a or TNF-a) are released, assisting in the maturation process and activation of naı¨ve T cells, eosinophils, macrophages, and natural killer (NK) cells (Banchereau and others 2000; Lee and Iwasaki 2007). Despite a clear understanding of the basic functions of DCs, little is known about transfusion-associated modulation of these key immunological processes. Given the central role of DCs in the initiation and regulation of the immune response, the potential for dysfunction in these cells needs to be considered as a potential mechanism contributing to transfusion-related complications. Using in vitro models, transfusion-mediated modulation of DC costimulatory molecules and inflammatory responses has recently been described following exposure to packed red blood cells (PRBCs) (Dean and others 2013). Inflammatory cytokines, IL-6, TNF-a, IL-1a, chemokine macrophage inflammatory protein (MIP)-1a, and activation molecule CD83, were proposed as potential biomarkers associated with PRBC transfusion-related modulation of DC function (Dean and others 2013). Based on similar panels and established assays, this study aimed to investigate transfusion-associated changes in the DC inflammatory profile following exposure to PCs. Given that it is well accepted that PLTs modulate DC responses (Mazzoni and others 2001; Martinson and others 2004; Hamzeh-Cognasse and others 2008; Gudbrandsdottir and others 2013; Hottz and others 2014), the soluble rather than cellular component of PCs was investigated in this study. Furthermore, this study was designed to provide insight into the extent to which immunomodulation is affected by PC dose and/or ex vivo storage period.

Materials and Methods Platelet concentrates Leukodepleted buffy coat-derived PCs (pooled from the buffy coat of 4 whole blood donations) were obtained from the Australian Red Cross Blood Service (Blood Service, Brisbane). The PCs were prepared according to standard Blood Service protocols based on the Council of Europe Guidelines for the preparation and quality assurance of blood components (Council of Europe 2008). PCs were stored under routine conditions (20–24C with gentle agitation) at 326 mL (–14 mL) at a concentration of 284 · 109 PLTs per pool (–40 · 109 PLTs per pool). In Australia, PCs are routinely stored for up to 5 days before transfusion, while a number of European countries, including Denmark, Ireland, The Netherlands, and the United Kingdom, routinely store PCs for up to 7 days before transfusion. Therefore, we considered whether a 7-day expiry would further impact on the immunomodulatory capacity of soluble mediators in PCs. Thus, following 2 (fresh, n = 2), 5 (expiry, n = 2), or 7 (extended expiry, n = 3) days of storage, PC supernatant (PC-SN) was aseptically collected

SOLUBLE MEDIATORS IN PCs MODULATE DC RESPONSES

following 2 subsequent centrifugation steps at 1,500 g for 15 min and at 3,000 g for 10 min. PC-SN was aliquoted and stored at -80C until use in the in vitro assay.

Ethics This study was approved by the Australian Red Cross Blood Service Human Research Ethics Committee (2012#05). Whole blood from consenting volunteers was collected in ethylenediaminetetraacetic acid (EDTA)-anticoagulated tubes [Becton Dickinson (BD)] and utilized as the recipient in the in vitro whole blood culture transfusion model. EDTA-anticoagulated whole blood has been demonstrated to be reliable and consistent for cytokine analysis (Thavasu and others 1992; Flower and others 2000). PCSNs were used at a 10% replacement volume and a 25% replacement volume approximating a 1 and 2–3 unit transfusion, respectively.

In vitro whole blood culture transfusion model PC-SNs [day 2 (D2), D5, or D7] and EDTA-anticoagulated recipient whole blood from consenting volunteers were cultured in RPMI 1640 media (containing 2 mM L-glutamine; Gibco by Life Technologies) in 24-well plates (Costar, Corning Life Sciences). The total volume was 1.5 mL/well and a 10% and 25% replacement volume transfusion was set up (10% replacement: 150 mL PC-SN, 600 mL RPMI, 750 mL recipient whole blood; 25% replacement: 375 mL PC-SN, 375 mL RPMI, 750 mL recipient whole blood). Lipopolysaccharide (LPS, 1 mg/mL; Sigma Aldrich) was added to duplicate wells to model an infectious complication. GolgiPlug (containing 0.1% brefeldin A, 1 mg/mL; BD Biosciences) was added after the first hour to facilitate intracellular detection of inflammatory markers. For assessment of the overall inflammatory response, the GolgiPlug was omitted. A no transfusion (no tx) control (containing recipient blood and RPMI – LPS) was run in parallel for each recipient to establish an individual baseline for donor variation and facilitate analysis of immunomodulation. Following a total of 6 h of incubation (37C, 5% CO2), cells were then harvested and centrifuged (1,000 g, 2 min). The supernatant (SN) from wells without the GolgiPlug was collected and stored at -80C for assessment of the overall inflammatory response using the cytometric bead array (CBA; see ‘‘Assessment of the overall inflammatory response from the whole blood culture transfusion model through CBA’’). The cells harvested from wells containing the GolgiPlug were assessed for DC-specific cytokine production. (see ‘‘Assessment of DC inflammatory response from the whole blood culture transfusion model through intracellular staining’’). Data represent the following combinations (D2: 4 recipients, each cocultured with SN derived from 2 independent PCs stored for 2 days, total n = 8; D5: 4 recipients, each cocultured with SN derived from 2 independent PCs stored for 5 days, total n = 8; D7: 3 recipients, 2 recipients cocultured with SN derived from 2 independent PCs stored for 7 days and 1 recipient cocultured with SN derived from 3 independent PCs stored for 7 days, total n = 7). The same recipients were used for no LPS and LPS data within each time point.

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Assessment of DC inflammatory response from the whole blood culture transfusion model through intracellular staining For wells with the GolgiPlug included, harvested cells were surface stained [room temperature (RT), 15 min, dark] with a panel of fluorescently labeled monoclonal antibodies to identify DC populations as follows: lineage [CD3, CD19, CD20, CD34, and CD56, all fluorescein isothiocyanate (FITC) conjugated], CD45-peridinin chlorophyll (PerCP), CD11c-allophycocyanin (APC), human leukocyte antigen DR (HLA-DR)-violet 450 (V450), and CD14-violet 500 (V500) (all from BD Biosciences). Following surface staining, erythrocytes were then lysed (FACS Lyse, RT, 10 min, dark; BD Biosciences) and leukocytes permeabilized (FACS Perm, RT, 10 min, dark; BD Biosciences). Leukocytes were then stained with phycoerythrin (PE)-conjugated anti-human IL-6, IL-8, IL-10, IL-12, IL-1a, TNF-a, MIP-a, MIP-1b, monocyte chemoattractant protein (MCP)-1, and inflammatory protein (IP)-10 (30 min, RT, dark, all BD Biosciences), washed with 3% fetal calf serum (Gibco by Life Technologies)/phosphate-buffered saline (PBS; Gibco by Life Technologies), and resuspended in 1% paraformaldehyde (PFA; Sigma Aldrich)/PBS (Gibco by Life Technologies) before analysis using flow cytometry. DCs were identified on flow cytometry dot plots as lineage- CD14- HLA-DR+ CD11c + and median fluorescent intensity (MFI) was determined for each inflammatory marker. A ratio of MFI was then determined (sample MFI/ matched no tx control MFI) for each inflammatory marker. Log2 of the ratio was then calculated equalizing the no tx control to zero, which represents the baseline and takes into account biological variation between different individual responses. In this way, a -1 result indicates a 2-fold suppression compared with the matched no tx control, and a + 1 result indicates a 2-fold increase.

Assessment of the overall inflammatory response from the whole blood culture transfusion model through CBA CBA (BD Biosciences) was used to assess the inflammatory response in the culture SNs from the in vitro whole blood culture transfusion model (wells incubated without GolgiPlug). For the purpose of this study, the overall immune response refers to quantification of the panel of markers outlined below and represents the combined inflammatory response for all the leukocyte populations as these are present in the whole blood model. CBA is a multiplex flow cytometrybased sandwich immunoassay that utilizes a plex of capture beads each with a defined fluorescence and each coated with a different specific capture antibody. The CBA was performed according to the manufacturer’s specifications (BD Biosciences). Briefly, culture SN and capture beads were mixed (1 h, RT, dark), and PE-labeled detection reagent was then added (2 h, RT, dark). Capture beads were then washed with wash buffer, centrifuged (200 g, 5 min), and resuspended in wash buffer before flow cytometric analysis. Unknowns were interpolated from standard curves run in parallel. The target panel consisted of IL-4, IL-6, IL-8, IL-10, IL-12, IL-1a, IL1b, TNF-a, MCP-1, MIP-1a, MIP-1b, IP-10, IFN-a, and IFN-g (all BD Biosciences). Culture SNs were all tested in

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one batch with no freeze/thaw of samples to minimize assay-to-assay variation. The limit of detection for this assay was 10 pg/mL. For statistical analysis, the marker had to be detected in at least 6 of 8 biological replicates (different individual responses).

Flow cytometry A 3-laser BD FACSCanto II flow cytometer (BD Biosciences), along with FACS Diva, and FCAP Array (both from BD Biosciences) were used for all flow cytometry and associated analyses. For intracellular staining, a minimum of 500 DC events were analyzed per treatment, per inflammatory marker.

Statistical analyses A repeated measure one-way analysis of variance (ANOVA) with Tukey’s post-test was used on paired data for analysis of the MFI of inflammatory markers detected by flow cytometry. A separate repeated measure one-way ANOVA with Tukey’s post-test was performed on data generated from each time point (D2, D5, D7) as different recipients were used at each time point. A dose response was considered present when the post hoc analyses indicated a significant difference between a 10% and 25% dose, or, between the no tx and 25% dose, but not between the no tx and 10% dose. To assess storage for DC-specific responses, the log2 MFI was compared across D2, D5, and D7 for each dose (eg, D2 10%, D5 10%, D7 10%) using ANOVA with Tukey’s posttest. To assess storage-associated modulation of the overall immune response, the concentration (pg/mL) of each marker was compared across D2, D5, and D7 within each dose using ANOVA with Tukey’s post-test. A storage-associated modulation was considered present when the post hoc analyses determined that there was a significant difference between D2, D5, and D7 for the 10% dose and/or the 25% dose. For all statistical analyses, P values < 0.05 were considered significant. GraphPad Prism 5 (GraphPad software) was used for all analyses and graphics.

Results Assessment of the DC-specific inflammatory profile in response to PC-SN in an in vitro whole blood culture transfusion model In the first instance, a panel of markers was used to assess changes in DC inflammatory response following exposure to SN derived from different buffy coat PCs that had been stored for 2, 5, or 7 days. In the absence of LPS, there was minimal modulation of the DC inflammatory profile. Of the inflammatory markers tested, only production of IL-10 changed significantly, with suppression evident following exposure to D5 PC-SN (ANOVA, P = 0.032, Fig. 1A). Doseassociated suppression was also evident for DC IL-10 production following exposure to D5 PC-SN ( post hoc 10% vs. 25%, P < 0.05, Fig. 1A). IL-10 production was also suppressed when LPS was included as a model of infection, with D2, D5, and D7 PCSNs significantly suppressing IL-10 production compared with the matched LPS no tx control (ANOVA, P = 0.025

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(D2), P = 0.007 (D5), P = 0.042 (D7), Fig. 1A). A doseassociated response was evident at each of the time points assessed, and, a storage-associated modulation of IL-10 was also observed for both doses [Dose: post hoc LPS 10% vs. LPS 25%, P < 0.05 (D2 and D5); post hoc no tx LPS vs. LPS 25%, P < 0.05 (D7). Storage: post hoc LPS 10% D2 vs. LPS 10% D7, P < 0.05; post hoc LPS 25% D2 vs. LPS 25% D7, P < 0.05, Fig. 1A]. Significant modulation of a number of other key proinflammatory cytokines and chemokines was also observed following exposure to PC-SNs in coculture with LPS. Significant suppression of LPS-induced DC IL-8 production was evident following exposure to D5 and D7 PC-SNs [ANOVA, P = 0.034 (D5), P = 0.02 (D7), Fig. 1B] with a significant dose-associated modulation at both D5 and D7 and storage-associated modulation evident for the 25% dose [Dose: post hoc LPS 10% vs. LPS 25%, P < 0.05 (D5); post hoc no tx LPS vs. LPS 25%, P < 0.05 (D7). Storage: post hoc LPS 25%, P < 0.05 (D7), Fig. 1B]. Similar modulation of IL-8 was also observed following exposure to D2 PC-SN, but this did not reach significance. For IL-6, IL-12, and MIP-1b, a significant dose-associated suppression of LPS-induced DC responses was evident following exposure to D2, D5, and D7 PC-SNs [IL-6 ANOVA, P = 0.001 (D2), P = 0.027 (D5), P = 0.002 (D7); IL-12 ANOVA, P = 0.003 (D2), P = 0.013 (D5), P = 0.008 (D7); MIP-1b ANOVA, P = 0.0001 (D2), P = 0.004 (D5), P < 0.0001 (D7): for post hoc analysis, see Fig. 1C–E]. DC production of MIP-a, MCP-1, and IP-10 was not significantly modulated by D2, D5, or D7 PC-SNs in the presence of LPS. A clear dose-associated modulation of LPS-induced DC production of IL-1a was observed following exposure to D2, D5, and D7 PC-SNs [ANOVA, P = 0.022 (D2), P = 0.03 (D5), P = 0.030 (D7): for post hoc analysis, see Fig. 1F], with a substantially different profile evident following exposure to D5 and D7 PC-SNs (Fig. 1F). LPS-induced DC production of TNF-a was significantly suppressed following exposure to D7 PC-SN (ANOVA, P < 0.0001, Fig. 1G) with both a significant dose- ( post hoc LPS 10% vs. LPS 25%, P < 0.05) and storage-associated response ( post hoc LPS 25% D2 vs. LPS 25% D7, P < 0.05) observed. Suppression of LPS-induced TNF-a production in DCs was evident following exposure to D2 and D5 PC-SNs, but did not reach significance. These results suggest that soluble mediators present in buffy coat-derived PCs significantly modulate the DC inflammatory profile, particularly when an infectious complication was modeled. While both dose and storage modulated the DC immune profile, the dose more than the length of ex vivo storage affected DC function.

Assessment of the overall inflammatory response to PC-SNs in an in vitro whole blood culture transfusion model In addition to the measurement of DC-specific responses, the study design facilitated assessment of the overall inflammatory response in the culture SN from the in vitro whole blood culture transfusion model. Using multiplex CBA to quantify a panel of inflammatory markers, these data represent the inflammatory response of the entire leukocyte population present in the in vitro whole blood culture

SOLUBLE MEDIATORS IN PCs MODULATE DC RESPONSES

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transfusion model. Using this approach, IL-4, IL-12, IL-1a, IFN-a, and IFN-g were all below the limit of detection and no further analysis could be undertaken (data not shown). As observed for the DC-specific inflammatory response, minimal PC-SN transfusion-associated modulation of the overall inflammatory response was evident in the absence of LPS. A significant induction in IP-10 production was observed following exposure to PC-SN derived from PCs stored for 2, 5, and 7 days, with a dose-associated response evident following exposure to D2, D5, and D7 PC-SNs [ANOVA, P < 0.0001 (D2), P = 0.016 (D5), P = 0.003 (D7), post hoc no tx vs. 25%, P < 0.05 (D2, D5, and D7), Fig. 2A]. Significant suppression of IL-6 was observed following exposure to D5 PC-SN (ANOVA, P = 0.015, post hoc no tx LPS vs. LPS 25%, P < 0.05, Fig. 2B). A significant induction was also observed for MCP-1 following exposure to PC-SN derived from PCs stored for 7 days with a dose-associated response evident following exposure to D7 PC-SN (ANOVA, P = 0.025, post hoc no tx vs. 25%, P < 0.05, Fig. 2C). In the presence of LPS as a model of infection, significant suppression of IP-10, IL-6, TNF-a, MIP-1a, and MIP-1b production was evident [IP-10 ANOVA, P < 0.0001 (D2), P < 0.0001 (D5), P = 0.001 (D7); IL-6 ANOVA, P < 0.0001 (D2), P < 0.0001 (D5), P < 0.0001 (D7); TNF-a ANOVA, P = 0.003 (D2), P < 0.0001 (D5), P = 0.001 (D7); MIP-1a ANOVA, P < 0.0001 (D2), P < 0.0001 (D5), P < 0.0001 (D7); MIP-1b ANOVA, P < 0.0001 (D2), P < 0.0001 (D5), P < 0.0001 (D7); Fig. 2A, B, D–F]. For IL-6, TNF-a, MIP1a, and MIP-1b, neither the PC storage period nor dose further impacted on the production of these inflammatory markers.

‰ FIG. 1. Assessment of the DC-specific inflammatory profile following exposure to D2, D5, and D7 PC-SNs. Flow cytometry was used to measure DC-specific inflammatory markers in cell cultures of an in vitro whole blood culture transfusion model following exposure to SN derived from D2, D5, and D7 PCs (6 h culture, 5% CO2, 37C). Minimal PC-SN-associated modulation of the DC inflammatory response was evident in the absence of LPS, with only modulation of the DC inflammatory response evident for IL-10 (A). In the presence of LPS, significant modulation of the DC inflammatory response was observed for IL-10 (A), IL-8 (B), IL-6 (C), IL-12 (D), MIP-1b (E), IL-1a (F), and TNF-a (G) following exposure to PC-SN. Dose rather than ex vivo storage period predominantly had a more profound effect on the inflammatory response. Open symbols indicate culture without LPS, and the solid symbols indicate culture with LPS (1 mg/mL). Circle, square, and diamond symbols represent D2, D5, and D7 data, respectively. Y axis indicates log2-fold change, normalized to the matched no tx control, indicated by the dotted line at zero. X axis indicates dose (10%, 25%) and PC storage period (D2, n = 8; D5, n = 8; D7, n = 7). Bars represent mean – standard error of the mean (SEM). ANOVA indicated by an asterisk: *P < 0.050, **P < 0.010, ***P < 0.001. Tukey’s post-test (dose): no tx versus 25%, a P < 0.050, aaP < 0.010. Tukey’s post-test: 10% versus 25%, b P < 0.050, bbP < 0.010, bbbP < 0.001. LTukey’s post-test (storage). ANOVA, analysis of variance; D2, day 2; D5, day 5; D7, day 7; DC, dendritic cell; IL, interleukin; LPS, lipopolysaccharide; MIP, macrophage inflammatory protein; no tx, no transfusion; PC, platelet concentrates; PC-SN, PC supernatants; TNF-a, tumor necrosis factor-a.

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For IP-10, a dose-associated response was evident following exposure to D2, D5, and D7 PC-SNs [post hoc no tx LPS vs. LPS 25%, P < 0.05 (D2, D5 and D7), Fig. 2A]. Exposure to PC-SNs and LPS resulted in induction of MCP1 [ANOVA, P = 0.014 (D2), P = 0.001 (D7)] and IL-8 [ANOVA, P < 0.0001 (D2, D5, D7)] (Fig. 2C, G) production, particularly at the 25% dose. This pattern was seen across the storage time points tested, suggesting that the PCSN dose was important for modulation of MCP-1 and especially IL-8 production (For post hoc analysis, see Fig. 2C). For IL-10 and IL-1b, a variable dose response was observed in the presence of LPS (Fig. 2H, I). IL-10 production was significantly increased following exposure to D2, D5, and D7 PC-SNs, with a 10% dose inducing more IL-10 than the 25% dose at D5 [ANOVA, P = 0.036 (D2), P = 0.001 (D5), P = 0.002 (D7), post hoc LPS 10% vs. LPS 25%, P < 0.05 (D5), Fig. 2H]. For IL-1b, the 10% dose reduced production and the 25% dose induced production [ANOVA, P = 0.001 (D2), P = 0.002 (D5), P = 0.001 (D7), Fig. 2I]. These results demonstrate that soluble mediators present in PCs mediate significant dysregulation of the overall inflammatory response, particularly in the presence of an infectious stimulus.

Discussion The transfusion of PCs is standard for the management of thrombocytopenia and severe trauma to maintain coagulation and hemostasis (Hunt 1998; Sahler and others 2012). Unfortunately, this form of treatment also has the potential to contribute to the development of transfusion-associated complications, including infection, multiple organ dysfunction syndrome, and mortality (Heddle and others 1993; Spiess and others 2004; Aslam and others 2008; Kiefel 2008; Bilgin and others 2011; Refaai and others 2011; Sahler and others 2011); yet, the underlying mechanisms of TRIM associated with transfusion of PCs remain largely unknown.

‰ FIG. 2. Assessment of the overall inflammatory response to D2, D5, and D7 PC-SNs. Cytometric bead array was used to quantitate inflammatory markers in culture SNs from the in vitro whole blood culture transfusion model following exposure to PC-SN derived from D2, D5, and D7 PCs (6 h culture, 5% CO2, 37C). Minimal PC-SN-associated modulation of the inflammatory response was evident in the absence of LPS with significant modulation only observed for IP-10 (A) and IL-6 (B). In the presence of LPS as a model of infection, significant modulation of IP-10 (A), IL-6 (B), MCP-1 (C), TNF-a (D), MIP-1a (E), MIP-1b (F), IL-8 (G), IL-10 (H), and IL-1b (I) was evident following exposure to PC-SN. Importantly, dose rather than storage period had a more profound effect on the inflammatory response. The open symbols indicate culture without LPS, and the solid symbols indicate culture with LPS (1 mg/mL). Circle, square, and diamond symbols represent D2, D5, and D7 data, respectively. Y axis indicates pg/mL. X axis indicates dose (10%, 25%) and ex vivo PC storage period (D2, n = 8; D5, n = 8; D7, n = 7). Bars represent mean – SEM. ANOVA indicated by an asterisk: *P < 0.050, **P < 0.010, ***P < 0.001. Tukey’s post-test: no tx versus 25%, aP < 0.050, aaP < 0.010, aaaP < 0.001. Tukey’s post-test: 10% versus 25%, bP < 0.050, bbP < 0.010, bbbP < 0.001. LTukey’s post-test (storage). IP, inflammatory protein; MCP, monocyte chemoattractant protein.

SOLUBLE MEDIATORS IN PCs MODULATE DC RESPONSES

Human immune responses are regulated by a broad range of cytokines and chemokines, which target and regulate the multitude of cellular responses (Borish and Steinke 2003). In this study, the impact of PC-SNs on a panel of proinflammatory, regulatory/anti-inflammatory, and differentiation/ proliferation cytokines and a number of chemokines was evaluated in an in vitro whole blood culture transfusion model. DCs were the focus of this study due to their central role at the interface of the innate and the adaptive immune response, and we hypothesized that the transfusion of PCs would modulate DC cytokine and chemokine production, potentially impacting on clinical outcomes due to deregulated cytokine and chemokine signaling. Buffy coat-derived PCSNs were used in this study rather than apheresis PC-SN as buffy coat-derived PCs represent the majority of PCs used in transfusion practice. We assessed the potential for both dose and ex vivo storage period of PCs to impact on DC function. This is the first report showing that soluble factors present in PCs modulate DC immune responses as well as the overall inflammatory response in a dose-associated manner. The immune profile of DCs was investigated by intracellular staining of inflammatory markers following culture with PC-SNs in an in vitro whole blood culture transfusion model. The use of this model facilitates cell-to-cell interactions without the need for cell isolation procedures involving density gradients or magnetic-assisted cell separation. Using this model, there was minimal modulation of the DC profile following exposure to PC-SN in the absence of LPS. In contrast, when LPS was added as a model of infection, significant suppression of DC cytokine and chemokine production was observed following culture with PC-SNs. A dose-associated suppression of DC production of IL-10, IL8, IL-6, IL-12, and MIP-1b was evident. Both dose- and storage-associated modulations of DC production of TNF-a were evident. We are unable to directly compare these findings with other literature as, to date, there have been no previous reports of the modulation of the blood myeloid DC inflammatory profile following exposure to PC-SNs, highlighting the uniqueness of this study. Monocyte-derived DCs (MoDCs) are one of the closest comparable models to blood myeloid DCs, and modulation of the immune profile of MoDCs has, however, previously been reported following exposure to PCs that contained both the PLTs and soluble components of the blood product (Kissel and others 2006; Hamzeh-Cognasse and others 2008; Cognasse and others 2009). In a model using cultured MoDCs, a reduction in levels of IL-12 and TNF-a and an increase in IL-10 production were observed in culture SNs following exposure of LPS-activated MoDCs to both resting and thrombin receptor-activating peptide-activated PLTs (Kissel and others 2006). These results, based on MoDCs, support the observations of this study in a number of ways; first, the reduction in IL12 and TNF-a production is similar to that observed in the current study, and second, PLTs become partially activated during routine ex vivo storage, so the nonactivated and activated PLTs used in the aforementioned MoDC model also approximate changes that may be associated with fresh and stored PCs (Kissel and others 2006). Another study investigated PLT-associated changes in the MoDC profile and reported a significant increase in DC surface receptor expression of CD80, CD83, and CD86 and, using a transwell approach, indicated that the DC profile of activation was

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different, depending on direct cell-to-cell interaction versus soluble factors (Hamzeh-Cognasse and others 2008). In a recent study (Loh and others 2015), significant suppression of LPS-induced IL-12 production from monocytes cultured with PCs or PC-SNs was reported in a whole blood culture model of transfusion similar to that utilized in this study. Taken together with the dose-associated suppression of IL-12 observed in this study, the suppression of IL-12 in monocytes and in SNs of MoDC cultures in separate independent studies suggests that administration of PCs has the potential to significantly impact on the outcome of the immune response due to a lack of IL-12 production required for differentiation and proliferation of DCs, NK cells, and T cells (Borish and Steinke 2003). In addition to the investigation of DC-specific modulation, the in vitro whole blood culture transfusion model was also used to assess changes in the overall inflammatory response following exposure to PC-SNs. The overall inflammatory response represents the combined inflammatory response of multiple cell subsets found in circulation that would interact in the context of transfusion. Overall, while some variation was evident between PC-SNs derived from PCs stored for 2, 5, or 7 days, largely the same inflammatory mediators were modulated to similar levels regardless of the ex vivo storage period. In the absence of LPS, IP-10 production was consistently increased at all time points investigated, with a dose response evident following exposure to D2, D5, and D7 PCSNs. Another primarily monocyte-derived chemokine, MCP-1, was also increased following exposure to PC-SNs, with a significant induction observed following exposure to D7 PC-SN. IP-10 and MCP-1 have not been reported to increase during routine storage of buffy coat-derived PCs (Muylle and others 1993; Stack and Snyder 1994; Ferrer and others 2000; Wadhwa and others 2000; Cognasse and others 2006; Costa and others 2012; Tung and others 2012; Kaur and others 2015), and these data suggest that monocytedriven chemotaxis may be induced following exposure to soluble mediators in PC-SNs. With coculture of PC-SNs with LPS, quite a different profile of modulation of the overall response was observed. While MCP-1 was increased following exposure to D2 and D7 PC-SNs, IP-10 production was reduced, suggesting divergent pathways of response to soluble mediators present in PCs. In addition to IP-10, the overall production of IL-6, TNF-a, MIP-1a, and MIP-1b was significantly reduced following coculture of PC-SNs and LPS. Of note, IP-10 and IL-6 production was severely perturbed at the 10% dose, indicating that even a 1 unit PC transfusion can result in a significant suppression of these mediators and a dose response was not observed due to the profound suppression observed at the lowest dose studied. Like MCP-1, PC-SNs significantly enhanced IL-8 production in the presence of LPS, with a very clear doseassociated response evident, which was independent of the storage period of the PC-SNs. IL-8 is produced by a broad range of inflammatory cells, including neutrophils and monocytes (Harada and others 1994), and these data suggest that PC-SNs contain soluble mediators that target IL-8mediated pathways associated with cell recruitment. In our study, modulation of the overall inflammatory response was largely dose-associated, not storage associated. In addition, we observed a heterogeneous response, with

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suppression of proinflammatory mediators (IL-6 and TNFa) and simultaneous suppression and activation of mediators driving cell recruitment. Our data highlights the complex nature of biological therapeutics where multiple cell receptors and pathways can be engaged simultaneously and the output is the result of balance of multiple factors. While there has not been another in vitro study as comprehensive as this, these data are in line with some previous observations. A study using an in vitro whole blood culture transfusion model observed a significant reduction in LPSstimulated TNF-a and significant induction of IL-10 production in the SN of 24-h cell cultures following incubation with PCs (Schneider and others 2009). These findings differ from our study as IL-10 production was found to be reduced following exposure to PC-SNs. These conflicting outcomes for IL-10 may be due to the different cell culture periods (12/24 vs. 6 h) employed in the 2 in vitro models. PCs are complex biological components, thus it is likely that multiple mechanisms are involved in driving the immunomodulation detected in this study. While this study has not specifically analyzed the biological mediators in the PC-SN, the PLT storage lesion has previously been well characterized. Ex vivo storage of PCs is associated with significant increases in levels of histamine (Konca and others 2006), sCD40L (Kaufman and others 2007), P-selectin, RANTES, PF4, TGF-b, and IL-8 (Slichter and others 2005; Sahler and others 2011). Histamine has been reported to regulate the expression and secretion of cytokines and chemokines during LPSdriven maturation of DCs by downregulating IL-12, IL-6, IP10, MIP-3a, IL-18, and RANTES, while increasing levels of IL-8 and IL-10 (Mazzoni and others 2001). Further study into the specific role of histamine in PC-associated suppression of immune responses would be warranted. sCD40L is abundant in PCs and has the potential to induce an inflammatory response and it has been reported to drive DC activation and maturation following stimulation with activated PLTs (Martinson and others 2004; Blumberg and others 2006; Hamzeh-Cognasse and others 2008). The accumulation of sCD40L, in particular, may have a detrimental impact on DC function by inducing inflammation, impacting downstream mediators, inducing cytokine and chemokine release (eg, MCP-1), inducing adhesion molecule surface expression, and driving DC maturation (Martinson and others 2004; Sahler and others 2012). PF4, stored in PLT alpha granules, is released during routine ex vivo storage and has been reported to mediate a reduction in IL-12 and TNF-a production from LPS-stimulated MoDCs. Further experiments utilizing blocking studies or selective filtration approaches of these known DC modulators may be useful to define more specific mechanisms. Proinflammatory cytokines, such as IL-6, IL-1a, and TNF-a, promote the activation of the immune response, leading to inflammation (Borish and Steinke 2003). These responses are moderated by anti-inflammatory or regulatory cytokines such as IL-10, which inhibit the production of inflammatory mediators produced by key immune regulators, such as T cells, B cells, and monocytes (Borish and Steinke 2003). IL-12 is a key differentiation/proliferation cytokine mainly produced by monocytes, macrophages, and DCs. It is a key cytokine that is essential for differentiation, cytotoxicity, cytokine production of NK cells, and proliferation of T-helper and cytotoxic lymphocytes and plays a

PERROS ET AL.

key counter-regulatory role in an allergic inflammatory response (Borish and Steinke 2003). Chemokines are key mediators required for recruitment of lymphocytes to the sites of inflammation (Borish and Steinke 2003). Chemotactic signaling molecules, including IL-8, MIP-1a, MIP-1b, MCP-1, and IP-10, are derived from mononuclear phagocytes, T cells, neutrophils, and eosinophils and have the capacity to activate polymorphonuclear cells and drive chemotaxis and cell recruitment to the sites of inflammation (Borish and Steinke 2003). Given the significant role these cytokines and chemokines play in the immune response, modulation of these as a result of PC transfusion and/or ex vivo storage can significantly impact upon the patient, leading to poor outcomes. Such modulation was clearly evident in this study, providing evidence of the impact of PC transfusion on recipients’ immune profile. PC-SN-associated modulation of inflammatory signals (observed in this study) produced by DCs is likely to result in significant perturbation of other detrimental downstream immune processes. Monocytes act as myeloid precursors for the differentiation of macrophages and DCs (Auffray and others 2009; Hettinger and others 2013), and modulation of key cytokines required for differentiation, as observed in this study (eg, IL-12 and TNF-a) (Scheuerer and others 2000; Xia and Kao 2003), would significantly impact upon the functions of these downstream mediators by impacting the release of inflammatory mediators from other immune cell subsets with potentially detrimental consequences to the patient immune response post-transfusion. DC-driven cytokine secretion (eg, TNF-a) is required as an essential signal to activate T and B cells, naı¨ve T cells, eosinophils, macrophages, and NK cells (Banchereau and others 2000; Lee and Iwasaki 2007). Without activation of these cell types as a result of PC-associated modulation of cytokine and chemokine signaling, as observed in this study, the recipient immune system would not be able to mount an appropriate immune response, subsequently leading to increased risk of infection, a common transfusion-associated complication (Aslam and others 2008; Kiefel 2008). Therefore, if DC function (eg, cytokine and chemokine signaling) is compromised by soluble mediators in PCs, as observed in this study, host immune responses would also be compromised, leading to poor patient outcomes. In this study, there was clear evidence of dose-associated suppression of the DC-specific as well as the overall inflammatory responses. Therefore, it can be postulated that higher doses of PC transfusions would be associated with more significant modulation in the patient, increasing the likelihood of transfusion-associated complications. In summary, these novel findings demonstrate that soluble mediators present in PCs modulate both the DC-specific and overall inflammatory responses. These results suggest that DC immunosuppression associated with PC transfusion may contribute to poor patient outcomes, such as morbidity, and increased rates of infection through a failure to induce an appropriate adaptive immune response due to a lack of cytokine and chemokine signaling. In addition, our data suggest that in the absence of infectious complications, the soluble mediators present in PCs have minimal impact on the immune profile, but in patients with infectious complications, DC function may be significantly impaired following PC transfusion, particularly for patients receiving multiple PC transfusions (>2–3 unit).

SOLUBLE MEDIATORS IN PCs MODULATE DC RESPONSES

Acknowledgment The Australian government funds the Australian Red Cross Blood Service to provide blood, blood products, and services to the Australian community.

Author Disclosure Statement A.J.P. carried out laboratory work, data analysis, and wrote the manuscript. M.M.D. designed the project, assisted with data analysis, and reviewed the manuscript. R.L.F. and A.-M.C. contributed to the design of the project and reviewed data analysis and the manuscript. No competing financial interests exist.

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Address correspondence to: Ms. Alexis J. Perros Research and Development Australian Red Cross Blood Service PO Box 145 Kelvin Grove Brisbane Queensland 4059 Australia E-mail: [email protected] Received 11 February 2015/Accepted 27 April 2015