Stroma-dependent development of two dendritic-like cell types with distinct antigen presenting capability

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1 Experimental Hematology 2013;41: Stroma-dependent development of two dendritic-like cell types with distinct antigen presenting capability Pravin Periasamy and Helen C. O Neill Research School of Biology, Australian National University, Canberra, Australia (Received 26 August 2012; revised 9 November 2012; accepted 16 November 2012) Novel antigen presenting cells (APCs) have been described in the murine spleen. Cells have a distinct CD11c lo CD11b hi MHC-II L CD8a L phenotype as highly endocytic dendritic-like cells that cross-present antigen to CD8 + T cells but fail to activate CD4 + T cells. These cells are named L-DCs because they reflect dendritic cells (DCs) produced in long-term spleen cultures (LTC). Similar cells were produced when bone marrow progenitors were cocultured over the splenic stromal line 5G3. Cocultures continuously produced a majority of L-DCs and a transient population of cells reflecting conventional dendritic cells (cdcs). Both the L-DC and cdc-like subsets cross-present antigen to CD8 + T cells, inducing their activation and proliferation. However, as MHC-II L cells, L-DCs are unable to activate CD4 + T cells, while MHC-II + cdc-like cells present antigen for CD4 + T cell activation. These results distinguish two APC subsets produced in vitro: a transient population of cdc-like cells and L-DCs that are continuously produced, presumably from self-renewing progenitors. These subsets are not developmentally linked via a precursor or progeny relationship. L-DCs and cdc-like cells are also distinct in terms of cytokine expression, with 65 of 84 tested genes displaying greater than a twofold difference by quantitative reverse-transcriptase polymerase chain reaction. Splenic stroma supports production of two APC subsets reflecting different lineage origins. Ó 2013 ISEH - Society for Hematology and Stem Cells. Published by Elsevier Inc. Bone marrow (BM) is a primary lymphoid organ that contains progenitors and precursors that differentiate to give all blood cell lineages. In terms of dendritic cell (DC) development, the immediate conventional DC (cdc) precursors (pre-cdc) [1,2] and plasmacytoid (p)-pre DC [3], traffic through blood and enter the spleen, where they differentiate further to produce mature cdcs and plasmacytoid DCs (pdcs). Similarly, Ly6C hi monocytes can differentiate under inflammatory conditions within the spleen to give monocyte-derived DCs (modcs) [4]. When longterm cultures (LTCs) were established from whole murine spleens [5,6], they were shown to support continuous hematopoiesis of a homogenous population of phenotypically distinct, large, immature, dendritic-like cells termed LTC-DC [6]; it was hypothesized that spleen may reflect an independent site for hematopoiesis and contain an endogenous self-renewing progenitor [7 9]. When the in vivo counterpart cell to LTC-DC was discovered in the spleen (namely L-DC ), this reflected further support for the claim that these cells represent a different lineage Offprint requests to: Helen C. O Neill, Research School of Biology, Australian National University, Acton ACT 0200, Australia; Helen.ONeill@anu.edu.au of dendritic-like cells to previously described cdcs and pdcs [10]. Phenotypic and functional characterization of the in vivo L-DC subset has defined it as a unique dendritic-like subset restricted to the spleen that differs from the commonly described splenic cdc, pdc, and modc subsets. Splenic stromal lines derived from LTCs [11,12] have been found to support the long-term development of cells resembling L-DC in cocultures established with spleen and BM as a source of hematopoietic progenitors [7,13]. The cloned splenic stromal line 5G3 has been selected for further study based on its superior capacity to support L-DC development of progenitors in Lin BM [13], and the 5G3 coculture system has been developed as a microenvironment providing optimal conditions for L-DC development. 5G3 cocultures compare favorably with splenic LTCs in their capacity to support myelopoiesis [13]. Dendriticlike cells produced in 5G3 cocultures have been studied by antibody staining to determine the dynamic production of dendritic and myeloid cells over time. Cocultures have been shown to produce two distinct subsets: cdc-like cells, which are transient, and a continuous population of L-DC. These subsets have been distinguished here by their distinct antigen-presenting functions X/$ - see front matter. Copyright Ó 2013 ISEH - Society for Hematology and Stem Cells. Published by Elsevier Inc.

2 282 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: Methods Animals Specific pathogen-free C57BL/6J (H-2 b ), CBA/H (H-2 k ), C57BL/ 6.Tg(TcraTcrb)1100Mjb (OT-I), and C57BL/6.SJL/J.OT-II. CD45.1 (OT-II) mice aged 8 days or 6 weeks were obtained from the John Curtin School of Medical Research (Canberra, Australia). Mice were housed and handled according to protocols approved by the Animal Experimentation Ethics Committee at the Australian National University (Canberra, Australia). Cell culture Cells were cultured at 37 Cin5%CO 2 in air in Dulbecco modified Eagle medium (DMEM) supplemented with 10% FCS, mol/l 2-mercaptoethanol, 10 mmol/l 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 100 U/mL penicillin, 100 mg/ ml streptomycin, 4 g/l glucose, 6 mg/l folic acid, 36 mg/l L-asparagine, 116 mg/l L-arganine HCl (sdmem). The establishment of LTC from neonatal spleens has been described previously [11,14,15]. LTCs show continuous production of dendritic-like cells for more than 1 year [5]. Splenic stromal lines were established from LTCs of B10.A(2R) mice that had ceased production of DCs over time because of a loss of hematopoietic cells [11,15]. Stroma were maintained by scraping attached cells for passage into a new flask. The cloned 5G3 stromal line, which supports in vitro hematopoiesis, has been described previously [12,13]. To maintain the stability of cloned stromal lines, frozen stocks were established and cell cultures were discarded after five passages. In controlled coculture experiments, stromal cells were dissociated and harvested using 0.25% trypsin ethylenediaminetetraacetic acid treatment before plating a given number of cells. Preparation of cell suspensions BM and spleen cells were dissociated by forcing tissue through a fine wire sieve, followed by washing through centrifugation at 300 g for 5 min. For lysis of red blood cells, supernatant was decanted and cells resuspended in 5 ml red blood cell lysis buffer (140 mmol/l NH 4 Cl, 17 mmol/l Tris Base, ph 7.5) followed by incubation for 5 min at room temperature. Five milliliters of phosphate-buffered saline (PBS) were then added, and the cell suspension was centrifuged for 5 minutes at 300 g. Cells were washed twice with 10 ml of PBS to remove cell debris and then resuspended in 5 ml of sdmem. Trypan blue staining (4% in 0.85% normal saline) was used to count live cells. Establishment of coculture assays The capacity of 5G3 to support hematopoiesis was assessed by overlay of Lin BM or T cell and B cell (T/B) depleted spleen cells above stromal monolayers followed by coculture for several weeks. Stromal cell lines were grown to 80% 90% confluency, and Lin BM or T/B depleted spleen, were plated at cells/ml above stromal monolayers. At 7-day intervals, nonadherent cells were collected by gently shaking the flask, with removal and replacement of supernatant. Antibody staining and flow cytometry Fluorochrome-labeled antibodies specific for murine markers were purchased from three suppliers: CD86 (GL1), CD172a (SIRPa; P84), and Ly6C (AL-21) from BD Pharmingen (San Diego, CA, USA); CD69 (H12F3) and F4/80 (C1:A3-1) from Bio- Legend (San Gabriel, CA, USA); and B220 (RA3-6B2), CD4 (H129.19), CD8a (53-6.7), CD11b (M1/70), CD11c (N418), CD24 (M1/69), CD45RB (C363.16A), CD80 (16-10A1), Gr-1 (Ly-6C/G) (RB6-8C5), I-A b (MHC-II) (AF ), TCR-Va2 (B20.1), and Thy1.2 (30-H12) from ebiosciences (San Diego, CA, USA). These antibodies were used at minimal saturating concentrations in multicolor staining experiments according to previously described methods [13]. Cells were plated in the wells of a flexible 96-well microtiter plate on ice for 15 min in 25 ml of FACS buffer (DMEM with 0.1% sodium azide and 1% FCS) containing 40 mg/ml of Fc block specific for FcgII/IIIR (CD32/CD16; ebiosciences) [16]. Cells were centrifuged for 5 min, and supernatant was discarded before resuspension in 25 ml of primary antibody and incubation on ice for 30 minutes. Cells were washed twice with 150 ml of fluorescence-activated cell sorting (FACS) buffer by centrifugation and resuspended in 20 ml of diluted secondary antibody or streptavidin conjugate as required, followed by incubation on ice for 30 min. Following two washes with 150 ml FACS buffer, cells were resuspended in 50 ml of FACS buffer before flow cytometric analysis. In some experiments, 5 ml of 10 mg/ml propidium iodide (PI) was added to cells for flow cytometric discrimination of dead cells. For multicolor staining, the same incubation and washing procedures were used with up to four different primary antibodies added together in the first incubation step. The specificity of antibody binding was monitored using isotype-matched control antibodies, second stage reagents alone, or FACS buffer only when isotype control antibodies were unavailable. Flow cytometry was performed on an LSRII FACS machine (Becton Dickinson; Franklin Lakes, NJ, USA). FACSDIVA software (Becton Dickinson) was used to set voltage parameters and event counts while running samples. For multicolor analysis, single-color compensation controls were used to set compensation on the machine. FlowJo software (FlowJo, Ashland, OR, USA) was used to analyze data. Cell debris was gated out using a forward scatter (FSC) threshold of 100. Cells were gated further on the basis of side scatter (SSC) and the absence of PI staining to detect live PI cells. Post-acquisition gating was used to obtain information on cell subsets, and staining with isotype controls was used to set gates to distinguish specific antibody staining. Cell sorting by FACS was used to isolate hematopoietic cell subsets based on marker expression. BM and spleen cells were prepared and stained with antibodies as described earlier. Sorting was performed on a BD FACSAriaII cell sorter (Becton Dickinson). Sorted cells were reanalyzed with flow cytometry to check purity of subsets. Endocytosis was measured flow cytometrically to assess the capacity of cells to take up antigen by the addition of 100 mg/ ml ovalbumin conjugated to fluorescein isothiocyanate (OVA- FITC) (Molecular Probes, Eugene, OR, USA) in a total volume of 100 ml sdmem. Cells were then incubated at 37 C for 45 min before endocytosis was halted by the addition of 100 ml of chilled PBS/0.1%NaN 3. Cells were washed three times using centrifugation at 300 g for 5 min and then resuspended in FACS buffer for analysis with flow cytometry. Fractionation of cells using magnetic bead methodology Lin BM was prepared using an antibody cocktail specific for hematopoietic lineage cells and containing biotin-labeled antibodies specific for CD5, CD45R, CD11b, Gr-1 (Ly-6G/C), 7-4,

3 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: and Ter-119 (Lineage Depletion Kit; Miltenyi Biotec, North Ryde, Australia) according to the manufacturer s protocol. Following antibody binding, 30 ml of labeling buffer containing 20 ml of MACS anti-biotin microbeads (Miltenyi Biotec) were added and incubated on ice for an additional 15 min. After washing and resuspension in 500 ml labeling buffer, cells were transferred to a MACS MS column (Miltenyi Biotec) placed in the permanent magnet of a SuperMACS II Separator (Miltenyi Biotec). Cells binding the superparamagnetic antibiotin microbeads are retained in the MACS MS column (Miltenyi Biotec). The MACS MS column was then washed three times with 500 ml of labeling buffer, and the flow-through cells were collected and washed twice with sdmem. An aliquot of the Lin cell population was tested with FACS analysis for the presence of Lin þ cells to determine the efficiency of depletion. Over multiple experiments, efficiency of depletion was shown to be w95%. T cells were purified from the spleen through depletion of macrophages, B cells, and MHC-II þ antigen presenting cells (APCs). The procedure used has been described previously [10,13]. The antibody cocktail contained antibodies specific for CD11b (clone M1/70), B220 (clone RA3-6B3), and IA b/k (clone TIB120; ebiosciences). For depletion of CD4 þ or CD8 þ T cells, either anti-cd4 (GK1.5) or anti-cd8 (53-6.7) were included in the antibody cocktail (ebiosciences). Cells were incubated with antibodies for 25 min on ice and washed twice with MACS labeling buffer. Cells were then incubated with sheep anti-rat Ig Dynabeads (Invitrogen Dynal, Oslo, Norway; 50 ml beads per 10 7 cells) at 4 C for 25 min with rotation before placing cells in a Dynal magnetic particle separator for 2 min. Supernatant containing unbound T cell subsets was then transferred into a new tube. Over multiple experiments, the efficiency of depletion was shown to average 90%. The procedure to isolate a mixed population of splenic CD11c þ DCs from T and B cell depleted splenocytes has been described previously [10]. T and B cell depleted splenocytes (10 8 cells/ml) were incubated for 25 min on ice with 20 ml of anti-cd11c magnetic MACS microbeads (Miltenyi Biotec) per 10 8 cells and washed once with 5 ml MACS labeling buffer and resuspended in 500 ml MACS labeling buffer. The cell suspension was run into an MACS MS column, and the column washed to deplete unbound cells. After the final wash, the column was removed from the SuperMACS magnet and placed over a fresh tube for elution of CD11c þ labeled cells. The depletion efficiency of this procedure was shown by flow cytometry to average 90% over many experiments. Microscopy The morphology of stromal cells was photographed under bright field microscopy using an inverted Fluovert FS microscope (Leica, North Ryde, Australia) equipped with a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI, USA). Images were processed using SPOT RT software v3.5.1 (Diagnostic Instruments). A DM IRE2 inverted research microscope (Leica) equipped with a DFC digital camera (Leica) was used to obtain phase contrast photomicrographs. Images were processed using Leica IM software version 4.0. For Giemsa staining, cells ( in 200 ml) were pelleted on to a glass slide using a cytospin, fixed in methanol, and stained in a two-step procedure using 0.25% Eosin YO Sorensen buffer (ph 6.8), followed by 0.25% methylene blue polychrome Sorensen buffer (ph 6.8; CliniPure Staining Kit; HD Scientific, Wetherill Park, Australia). Slides were washed, dried, and mounted with a nonaqueous mounting agent (Depex; Fluka Analytical, Buchs, Switzerland). Photography used a DFC digital camera (Leica) connected to a bright field inverted microscope. T cell activation assays A mixed lymphocyte reaction (MLR) was used to investigate the ability of cells produced in cocultures to induce an allogeneic T cell response using syngeneic T cells as control group. Responders were prepared as the T cell fraction of C57BL/6J and CBA/H spleen. The cross-presenting capacity of coculture-produced APC was measured by the ability of antigen-pulsed cells to induce proliferation of CD8 þ T cells purified from spleens of OT-I T cell receptor (TCR)-transgenic (Tg) mice specific for OVA /H- 2K b. The ability of isolated coculture-produced APC subsets to present antigen to CD4 þ T cells was measured according to the ability of antigen-pulsed cells to induce proliferation of purified CD4 þ T cells isolated from OT-II TCR-Tg mice specific for OVA /H-2IA b. APC included nonadherent cells collected from cocultures established from C57BL/6J BM or from LTCs established from C57BL/6J spleen. Freshly isolated CD11c þ splenic DCs from C57BL/6J or CBA/H mice served as control APC. In some experiments, APC were pulsed with 10 mg/ml of OVA or control antigen hen egg lysozyme (HEL) overnight (8 12 hours), then washed by centrifugation (300 g, 5 min) twice. Some cultures were given lipopolysaccharide (LPS; 10 mg/ml) as an activator of DCs for 12 hours. The time for treatment was optimized in preliminary experiments. To measure T cell activation, cells were cultured at cells per well in a 96-well plate together with graded numbers of APC or alone as a control. Cells were plated in a total volume of 200 ml of sdmem, and plates incubated at 37 C for 4 days. Separate plates were stained with antibody to CD4 or CD8 to determine T cell proliferation in terms of reduced levels of 5- (and 6-) carboxyfluorescein diacetate succinimidyl esters (CFSE) for each of the CD4 þ and CD8 þ T cell subsets. Fractionated T cells were labeled with CFSE for flow cytometric analysis of their proliferation. For labeling, CFSE (Molecular Probes) was added to cells (0.5 ml/ml) to give a final concentration of 10 mg/ml, and samples were vortexed and then incubated at room temperature for 5 min. Cells were then washed twice by adding 1 ml of ice-cold sdmem followed by centrifugation at 300 g and 4 C for 5 min. After culture for various periods of time, cell division was assessed flow cytometrically in terms of reduction in CFSE level as cells divided. Quantitative polymerase chain reaction L-DC and cdc-like cells were sorted from 14-day cocultures of Lin BM over 5G3. Extraction of RNA involved the RNeasy mini kit following the supplier s instructions (Qiagen, Clifton Hill, Australia). RNA purity and quality were determined by loading material on to RNA 6000 Pico Chips (Agilent Technologies, Waldbronn, Germany) with analysis on an Agilent 2100 Bioanalyser (Agilent Technologies). RNA was converted to cdna using the RT 2 First Strand Synthesis Kit following the manufacturer s instructions (Qiagen). In real-time, quantitative reversetranscriptase polymerase chain reactions (qrt-pcr), cdna, RT 2 SYBR Green Mastermix, and RNase-free water were added

4 284 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: directly to PCR arrays already loaded with primers for 84 genes related to a common pathway (e.g., cytokine expression; PCR array #PAMM-021, SABioscience, Frederick, MD, USA), and the plate was loaded on to a LightCycler 480 (Roche, Castle Hill, Australia). Cycling conditions were set according to manufacturer s recommendation: 1 cycle of 10 min at 95 C, followed by 45 cycles of 15 sec at 95 C, and 1 min at 60 C. Data analysis was performed using Roche LightCycler 480 software version to calculate the cycle number at which the maximal increase in fluorescence emission occurs in the log-linear phase Figure 1. Phenotype of cells produced in BM cocultures. Cocultures were established for 28 days by overlay of Lin BM over 5G3 stroma. LTCs were established from dissociated spleen of C57BL/6J mice for up to 18 months. (A) Nonadherent cells were collected from cocultures at 7-day intervals and at a single time point from LTCs. Cell surface marker expression was analyzed flow cytometrically by staining cells for CD11c, CD11b, MHC-II, CD45RB, CD86, CD80, CD24, CD8a, F4/80, B220, Ly6G, Ly6C, and CD172a. Propidium iodide staining allowed gating of PI live cells for analysis. FSC and SSC plots were used to gate large cells for multichannel analysis. Isotype control antibodies were used to indicate background binding, shown as a red overlay on LTC-DC plots, and to set gates on bivariate L-DC plots. Numbers shown in quadrants represent percent positive cells. (B) Photomicroscopy of L-DCs and LTC-DCs. Electron microscopy revealed similar ultrastructure morphology (original magnification 3000; scale bar, 10 mm). Giemsa staining and photography using bright field illumination revealed similar cell types (original magnification 600; scale bar, 30 mm). LTCs and 28 day cocultures were photographed under phase contrast to show cells developing above stroma (original magnification 200; scale bar, 100 mm).

5 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: (threshold cycle C t ). C t values were calculated for genes of interest (GOI), the b-actin housekeeping gene (HKG): DC t 5 C t (GOI) C t (HKG), and the average DC t taken from duplicate experiments. The fold change in mrna quantity between two samples was calculated as 2 DCt (sample 1)/2 DCt (sample 2). The production of an amplified product was validated through gel electrophoresis. Statistical analysis The Wilcoxon rank sum test was used to test significance (p # 0.05) for sample sizes of n # 5 when a normal distribution could not be assumed. The Student t test was used to assess significance (p # 0.05) when a normal distribution could be assumed. Results Cells produced in BM cocultures over splenic stroma Nonadherent cells produced in cocultures of Lin BM over 5G3 stroma were compared with cells produced in splenic LTCs in terms of morphology, phenotype, and function. Flow cytometric analysis of nonadherent cells produced in well-established LTCs revealed a distinct dendritic-like cell population with a CD11b þ CD11c þ MHC-II CD80 þ CD86 þ phenotype as described previously [6] (Fig. 1A). Cells produced did not express known markers of cdc and pdc, like CD24, F4/80, CD8a, Ly6C, Ly6G, B220, CD172a, and CD45RB. Cell production in cocultures of lineage (Lin) BM over 5G3 was monitored over time by staining for the same markers. At 14 days, both CD11b þ CD11c and CD11b þ CD11c þ subsets were detected. The relative abundance of CD11b þ CD11c þ MHC-II þ cdc-like cells is dependent on the number of precursors present in Lin BM preparations, which cannot be controlled. As a result, numbers of cdc-like cells can vary in early cocultures (!28 days). At this early stage of coculture establishment, MHC-II þ cells were also detected, although these were lost over time. After 28 days of coculture, a CD11b þ CD11c þ CD80 þ CD86 þ dendritic-like cell subset was seen to predominate as the cdc-like population disappeared (Fig. 1A). The majority of cells were negative for MHC-II, F4/80, CD8a, B220, Ly6G, Ly6C, CD45RB, and CD172a, with a 37% subset of cells expressing CD24. Morphologically, the CD11b þ CD11c þ MHC-II dendritic-like cells produced in LTCs and in cocultures appear as similarly large, highly granulated cells, with a large lobulated nucleus and fingerlike membrane projections or dendrites (Fig. 1B). In both cocultures and LTCs, cells developed as large round cells above stroma. As a result, the coculture-derived population was termed L- DC to indicate resemblance with previously described LTC-DCs. Although 28-day cocultures were enriched for CD11b þ CD11c þ MHC-II L-DCs, earlier stages during coculture revealed a transient appearance of CD11b þ CD11c þ MHC-II þ cdc-like cells. By day 28, the population of L-DCs had reached w62%, and the population of cdc-like cells was w37% (Fig. 1A). At 7 days of Figure 2. Endocytic function of cells produced in cocultures. Nonadherent cells were collected from Lin BM cocultures, produced as in Figure 1 at 28 days, and sorted to isolate L-DCs (CD11b þ CD11c þ MHC-II ) and cdc-like (CD11b þ CD11c þ MHC-II þ ) subsets. (A) Cells were stained for CD11b, CD11c, and MHC-II and then incubated with propidium iodide to allow gating of PI live cells. CD24 expression distinguishes the two subsets. (B) Sorted L-DCs and cdc-like cells were compared with LTC-DCs for their capacity to endocytose soluble FITC-OVA (100 mg/ml) for 45 min at 37 C, or at 4 C as a background control. Uptake was assessed flow cytometrically to give a percent of endocytic cells. Data reflect mean 6 SE from three separate experiments. *Endocytosis by cdc-like cells is significantly different from L-DCs and LTC-DCs (p # 0.05, Wilcoxon Rank Sum Test).

6 286 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: coculture, outgrowth of a distinctly large population of CD11b þ CD11c myeloid-like cells was evident, but the outgrowth disappeared by 21 days (all data not shown). Lin BM appears to contain a mixture of precursors and progenitors that are supported in the early stages of stromal cocultures. Although the relative numbers of these cells can vary between cocultures, their transient nature is a true characteristic. Over time, however, the outgrowth of L- DCs surpassed other transient subsets. Outgrowth of similar cells has been reported in LTCs [6]. Characterization of MHC-II distinct dendritic-like cells produced in co-cultures To compare the subsets produced in cocultures after 28 days of co-culture, nonadherent cells were collected and sorted as CD11b þ CD11c þ MHC-II L-DCs and CD11b þ CD11c þ MHC-II þ cdc-like cells. In addition to their disparity in MHC-II expression, cdc-like cells were shown to be distinct by expressing CD24, consistent with previous evidence for CD24 expression on cdcs [17] (Fig. 2A). These subsets were compared with isolated LTC-DCs for their capacity to endocytose FITCconjugated ovalbumin (OVA-FITC) as soluble antigen. Both coculture-produced L-DCs and LTC-DCs were superior to cdc-like cells, having 100% endocytic cells compared with 25% for cdc-like cells (Fig. 2B). The ability of L-DCs and cdc-like cells to stimulate T cell proliferation in an MLR was compared with LTC-DCs and freshly isolated splenic CD11c þ DCs (Fig. 3). In an allogeneic MLR of CBA/H (H-2 k ) T cells against C57BL/ 6J (H-2 b ) APCs, all APCs stimulated CD8 þ T cell proliferation measured flow cytometrically by reduction in CFSE staining (Fig. 3A). However, LTC-DCs and splenic DCs gave stronger responses. Although control splenic DCs were strong CD4 þ T cell activators, neither coculture produced L-DCs and cdc-like cells, nor LTC-DCs, activated CD4 þ T cell proliferation. In a control syngeneic B6 anti-b6 MLR, none of the APCs showed activation or proliferation of syngeneic CD4 þ or CD8 þ T cells, confirming that T cell activation is antigen-specific (Fig. 3B). Figure 3. Cells produced in cocultures activate allogeneic T cells. Cocultures of Lin BM isolated from C57BL/6J (H-2 b ) (B6) mice were established over 5G3 stroma. Nonadherent cells were collected after 28 days and sorted to give L-DCs (CD11b þ CD11c þ MHC-II ) and cdc-like cells (CD11b þ CD11c þ MHC-II þ ) as described in Figure 2. LTC-DCs were collected as nonadherent cells from LTCs established 6 months previously from C57BL/6J mice. Freshly isolated CD11c þ DCs (f-dc) were prepared from C57BL/6J (H-2 b ) spleens as control APC using MACS magnetic bead isolation. T cells were isolated from spleens of CBA/H (H-2 k ) (CBA) and C57BL/6J (H-2 b ) (B6) mice depleted of B cells and myeloid cells using Dynabead technology. These were labeled with CFSE and cocultured with either f-dcs, L-DCs, cdc-like cells or LTC-DCs at a T cell:apc ratio of 3:1, 15:1, and 75:1. Microtiter plates were incubated at 37 C and analyzed after 4 days by flow cytometry to detect the CFSE profile of CD4 þ and CD8 þ T cell subsets. Data are presented as percent of divided cells for (A), an allogeneic CBA anti-b6 MLR, and (B) a syngeneic B6 anti-b6 MLR. Controls included T cells only. Data are presented as mean 6 SE for two replicate experiments. L-DCs produced in cocultures are cross-presenting To compare their function further, L-DCs and cdc-like cells were sorted from non-adherent cells produced in 28-day Lin BM cocultures and compared for ability to cross-present antigen to CD8 þ T cells purified from OT-I TCR anti-h-2k b /OVA transgenic mice. Also compared were LTC-DCs and splenic DCs. All four subsets were able to cross-present OVA, but not control antigen HEL. Activation of CD8 þ T cells after 24 hours was identified by upregulation of CD69 in both the presence and absence of LPS (Fig. 4A). After 72 hours, all subsets induced antigen-specific proliferation of CD8 þ T cells, measured by reduction in CFSE as cells divided (Fig. 4B). The ability to cross-present antigen was specific for OVA, and no HELspecific response was detected. The addition of LPS caused a significant increase in CD8 þ T cell proliferation at T cell:apc ratios of 2:1 and 10:1 for all APC tested, including

7 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: Figure 4. Cross-presentation capacity of coculture-produced cells. L-DCs, cdc-like cells, and LTC-DCs were prepared along with freshly isolated splenic CD11c þ DCs (f-dc) control cells as described in Figures 1 and 3. These cells were compared for their capacity to cross-present soluble antigen OVA, or control antigen HEL, in the presence and absence of LPS, to CD8 þ T cells isolated from OT-I TCR-tg anti-h-2k b /OVA mice. All APCs were pulsed for 12 hours with OVA, HEL, OVA þ LPS, or HEL þ LPS (each at 10 mg/ml). CD8 þ T cells were purified from OT-I spleen by depletion of B cells, myeloid cells, DCs, and CD4 þ T cells using magnetic bead protocols and then labeled with CFSE. Cells were cocultured in T cell:apc ratios of 2:1, 10:1, and 100:1. (A) CD8 þ T cell activation was analyzed after 24 hours as a percentage of cells staining for CD69. (B) CD8 þ T cell proliferation was measured after 4 days as a percentage of cells showing a reduction in CFSE staining. Data reflect mean 6 SE (n 5 3) collected in three replicate experiments. CD8 þ T cells were gated as live (PI ) CD11c Thy1.2 þ Va2 þ CD8 þ cells using flow cytometry. Response to OVA þ LPS is significantly different to the response to OVA for all APC at T cell:apc ratios of 2:1 and 10:1 (onetailed t test; p # 0.05). fresh cdcs isolated from spleen (Fig. 4B). In previous studies, LPS treatment of L-DCs generated in cocultures [18] and LTC-DCs [6,10] increased their capacity to activate CD8 T cells. This finding is consistent with the effect of LPS on LTC-DCs [6] and L-DCs [18] to upregulate MHC-I and CD80/86, although not MHC-II. Figure 5. Inability of L-DCs to activate CD4 þ T cells. L-DCs, cdc-like cells, and LTC-DCs were prepared along with freshly isolated splenic CD11c þ DCs (f-dc) control cells as described in Figures 1 and 3. Cells were compared for their capacity to present soluble antigen OVA, or control antigen HEL, in the presence and absence of LPS to CD4 þ T cells from OT-II TCR-tg anti-h-2ia b /OVA mice. All APCs were pulsed for 12 hours with OVA, HEL, OVA þ LPS, or HELþLPS (each at 10 mg/ml). CD4 þ T cells were purified from OT-II spleen through depletion of B cells, myeloid cells, DCs, and CD8 þ T cells using magnetic bead protocols and then labeled with CFSE. Cells were cocultured in T cell:apc ratios of 2:1, 10:1, and 100:1. (A) CD4 þ T cell activation was analyzed after 24 hours as a percentage of cells staining for CD69. (B) CD4 þ T cell proliferation was measured after 4 days as a percentage of cells showing a reduction in CFSE staining. Data reflect mean 6 SE (n 5 3) collected in three replicate experiments. CD4 þ T cells were gated as live (PI ) CD11c Thy1.2 þ Va2 þ CD4 þ cells using flow cytometry. OT-II response to f-dcs with OVA þ LPS is significantly greater than the response to OVA alone (one-tailed t test; p # 0.05). L-DCs are weak activators of CD4 þ T cells The ability of L-DCs and cdc-like cells to present OVA to OT-II TCR anti-ova/h-2ia b CD4 þ T cells was compared with freshly isolated CD11c þ splenic DCs and LTC-DCs. All APCs were pulsed with specific-antigen OVA and control-antigen HEL in the presence and absence of LPS.

8 288 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: Figure 6. Differential potential of cells produced in cocultures. Nonadherent cells were collected on day 14 from Lin BM cocultures established as described in Figure 1. (A) Cells were stained for CD11c and MHC-II, and PI staining was used to gate PI live cells for sorting. Sorted subsets of CD11c MHC-II, CD11c þ MHC-II þ, and CD11c þ MHC-II cells were then recultured over 5G3 stroma. Progeny cells were collected at 3, 7, and 14 days for phenotypic analysis. (B) Nonadherent cells were collected and stained with fluorochrome-conjugated antibodies specific for CD11c, CD11b, and MHC-II, or with isotype control antibodies. FSC versus SSC analysis was used to gate large (FSC hi ) cells for subsequent multicolor analysis. Isotype controls were used to indicate background binding and to set gates on bivariate antibody plots. Numbers shown in quadrants represent the percentage of positivestaining cells. (C) Percentage of live cell recovery relative to total input cell number was estimated using trypan blue exclusion for cell counting. Three replicate cocultures were established, and data reflect mean 6 SE (n 5 3). (D) The percent yield of L-DCs (CD11c þ CD11b þ MHC-II ) and c-dc-like cells (CD11c þ CD11b þ MHC-II þ ) in cocultures was estimated relative to input cell number. Data represent mean 6 SE (n 5 3). Control splenic DCs showed antigen-specific, LPS-responsive activation of CD4 þ T cells at 12 hours, with T cell proliferation at 4 days (Fig. 5). This was significantly enhanced by the presence of LPS at T cell:apc ratios of 2:1 and 10:1. Both coculture-produced L-DCs and LTC- DCs were unable to activate or stimulate the proliferation of CD4 þ T cells. Interestingly, coculture-derived cdc-like cells were unable to stimulate CD4 þ T cell proliferation after 4 days, although they did show strong, antigenspecific, LPS-responsive activation of CD4 þ T cells at 12 hours. The inability of L-DCs and LTC-DCs to activate CD4 þ T cells is not surprising because both of these subsets do not express MHC-II. However, cdc-like cells grown in vitro are MHC-II þ, and their capacity to activate CD4 þ T cells without inducing proliferation is suggestive of immature DCs or DCs that are not immunogenic. Furthermore, the addition of LPS did not activate these cells to make them immunogenic.

9 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: Figure 7. Expression of common cytokines by L-DCs and cdc-like cells. Real-time PCR was performed using an RT 2 Profiler PCR Array (SABioscience) to measure the expression levels of genes for 84 common cytokines on sorted L-DCs and cdc-like cells isolated from 28 day cocultures. (A) The expression of each gene in L-DCs and cdc-like cells is shown as a single point on a log 10 plot of 2 DCt indicating gene expression. Gates are drawn to reflect a window of a twofold difference. Points lying outside these gates distinguish genes upregulated (twofold or greater) in L-DCs or cdc-like cells. (B) Genes upregulated twofold or greater in L-DCs. (C) Genes upregulated twofold or greater in cdc-like cells. MHC-II distinct subsets produced in cocultures are developmentally distinct Three distinct subsets of cells can be identified in Lin BM cocultures by 14 days based on expression of MHC-II and CD11c (Fig. 6A). To investigate the developmental relationship between cells produced in cocultures, the CD11c MHC-II, CD11c þ MHC-II þ, and CD11c þ MHC- II subsets were sorted and recultured over 5G3 stroma. Cell productivity was monitored, and progeny cells were identified by marker expression at 3, 7, and 14 days after reculture over 5G3 stroma (Fig. 6B). Only the reculture of CD11c þ MHC-II cells yielded an increase in cell production over a 14-day period, and cells produced resembled CD11c þ CD11b þ MHC-II L-DCs with no production of cdc-like cells (Fig. 6B, C). The CD11c þ MHC-II þ subset did not proliferate after reculture and declined in numbers over 14 days, suggesting a nonproliferating population (Fig. 6C). Similarly, in CD11c MHC-II recultures, cell number reduced significantly after 14 days, despite a weak proliferative peak in CD11c þ MHC-II þ cdc-like cell production at 7 days. Yields were reproducible across three separate flasks for each subset analyzed. Although it is always possible that low numbers of contaminants exist in the sorted populations, the data refute this. For example, the absence of CD11c þ MHC-II contaminants amongst CD11c MHC-II cells is confirmed because cocultures produce no CD11c þ MHC-II cells. Similarly, the absence of CD11c MHC-II cells among the CD11c þ MHC-II

10 290 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: population used to seed cocultures is confirmed because no CD11c þ MHC-II þ cells are produced. This experiment indicated no developmental relationship between L-DCs and cdc-like cells produced in cocultures. For example, L-DCs did not upregulate MHC-II expression on reculture to become cdc-like cells, and cdc-like cells were not precursors of L-DCs. Furthermore, CD11c MHC-II cells contained precursors of cdc-like cells but not L-DCs. This finding suggested that distinct precursors of L-DCs must exist among the CD11c þ MHC-II subset that is maintained in 5G3 stromal cultures. Cytokine gene expression by L-DCs and cdc-like cell subsets Gene expression for 84 common cytokines was quantified for the L-DCs and cdc-like cell subsets produced in cocultures using real-time PCR and an RT2 Profiler PCR Array (SABioscience). Cells were sorted out of cocultures established for 28 days. Results were collated from two separate experiments. Sixty-five of 84 genes were differentially expressed at a twofold difference or greater between L-DCs and cdc-like cells (Fig. 7A). This difference in gene expression supports the derivation of distinct DC types. The majority of genes (55) were upregulated twofold or greater in cdc-like cells compared with L-DCs (Fig. 7C), whereas 10 genes were upregulated twofold or greater in L-DCs (Fig. 7B). The genes most highly upregulated in L-DCs reflect their prominent role as activators of CD8 þ T cells. These genes include Tnfsf8 (tumor necrosis factor [ligand] superfamily, member 8) which encodes CD30L, and Tnfsf9 (tumor necrosis factor [ligand] superfamily, member 9) which encodes 4-1BBL. 4-1BBL is a co-stimulator that activates cytotoxic T cell function [19], and CD30L is a cytokine that supports the generation of long-lived memory CD8 þ T cells [20]. Gene expression by cdc-like cells reflects a mature DC phenotype with expression of cytokine genes naturally produced by mature cdcs and reflective of their functional capacity. These genes include IL12b (interleukin 12b), Ifng (interferon gamma), Ifna/Ifnb (interferon alpha/ beta) commonly expressed by cdcs [21 23]. Expression of a number of cytokines in a high level relative to L-DCs suggests two distinct cell types produced in cocultures. Discussion Previously it was shown that splenic LTCs support continuous production of a unique DC subset arising from selfrenewing progenitors supported by a stromal cell niche [6]. However, the heterogeneity of stromal cells and progenitor subsets in LTCs has made the study of cell differentiation in that system highly complex. In this study, we show that cocultures of Lin BM involving the 5G3 stromal line can also produce dendritic-like cells that are phenotypically and functionally similar to those produced in LTCs. Cocultures can replace LTCs as a way to produce L-DCs in vitro. The coculture system comprises three essential components: stroma, progenitors, and differentiated large nonadherent cells. By analyzing each component of the coculture system, it will be possible to gain an understanding of how a splenic stroma supports myelopoiesis. Lin BM preparations do, however, vary as a source of precursors and progenitors, which leads to some variability in the number and type of cells produced. While the time course for production of cdc-like cells is transient, it is also variable in length. An overriding feature of cocultures, however, is their continuous production of L-DCs, which becomes the only cell type produced once cdc-like cell production ceases. The possibility that contaminating mesenchymal and endothelial progenitors enriched in Lin BM contributes to hematopoiesis in 5G3 cocultures is refuted on two grounds. First, Lin BM alone does not undergo hematopoiesis. Second, highly purified hematopoietic stem cells sorted out of BM represent an adequate source of L-DC progenitors in cocultures over 5G3 [18]. In this study, the focus has been on characterizing the differentiated large nonadherent cells that resemble DCs. The cloned stromal cell line 5G3 provides a uniform splenic stromal environment for hematopoiesis. The use of BM as a source of progenitors in 5G3 cocultures and the study of L-DCs is justified by evidence that L-DCs produced resemble splenic LTC-DCs as large mononucleate cells with fingerlike membrane projections and a common phenotype as CD11c þ CD11b þ MHC- II CD80 þ CD86 þ cells with no expression of CD24, F4/ 80, CD8a, B220, Ly6C, Ly6G, CD172a, or CD45RB. Like LTC-DCs, coculture-produced L-DCs are highly endocytic and are able to activate and stimulate the proliferation of CD8 þ T cells via cross-presentation, but are unable to activate CD4 þ T cells. In a previous study, we characterized a functional and phenotypic in vivo equivalent to LTC-DCs [10]. This in vivo subset was present in the steady-state and shown to be highly endocytic and capable of presenting antigen to CD8 þ T cells, resulting in T cell activation, but it was unable to activate CD4 þ T cells. This finding is significant because it emphasizes the importance of L-DCs as a unique APC in the spleen. Using 5G3 cocultures established with Lin BM, it has been possible to map the development of two distinct dendritic-like cell types over time, including the transient, early CD11c þ CD11b þ MHC-II þ CD24 þ cdc-like population, and the predominant, continuously produced CD11c þ CD11b þ MHC-II CD24 L-DC subset. The L-DC subset is distinct from cdc-like cells and continues to be produced for as many as 5 weeks or more in cocultures, whereas the cdc-like cells are produced in lower numbers and only transiently, disappearing by 3 4 weeks. L-DCs and cdc-like cells differ phenotypically in terms of CD24 and MHC-II expression. Functionally, cdc-like

11 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: cells are able to activate but not induce proliferation of naive CD4 þ T cells. This fact distinguishes them from L-DC, which can neither activate nor induce proliferation of CD4 þ T cells. cdc-like cells are not highly endocytic like L-DCs, but are equally capable of cross-presentation for CD8 þ T cell activation. Phenotypically, the cdc-like subset resembles described subsets of splenic CD8a cdcs, and the expression of MHC-II by cdc-like cells is consistent with a mature DC. Further evidence is provided by the expression of cytokine genes associated with DCs. One example is the 10-fold upregulation of the IL2 gene in cdc-like cells over L-DCs, which is associated with DC maturity. Mature DCs secrete interleukin (IL) 2 to activate and proliferate T cells during antigen presentation [24]. In contrast to L-DCs, the cdc-like subset expresses many other genes associated with mature DC function, including Ifng, Ifna, IL12b, IL27, and Cd40lg [25 27]. In contrast to the cdc-like subset, L- DCs show several features that are reflective of immature DCs. The absence of MHC-II expression and high endocyticcapacityarecommonpropertiesofimmaturedcs. L-DCs also show an approximately ninefold upregulation of IL16 gene expression over cdc-like cells. Production of IL-16 has been shown to distinguish immature DC, since addition of inflammatory activators to DC like TNFa (tumor necrosis factor-alpha) and PGE2 (prostaglandine2)canresultinlossofil16 mrna and protein expression [28]. The cdc-like subset has been named as such on the basis of phenotype and mature status as antigen presenting DC. Functionally, however, cdc-like cells are distinct from described mature CD8a cdcs because they do not stimulate the proliferation of CD4 þ T cells, although they have a strong capacity to cross-present antigen to CD8 þ T cells. Cross-presentation is a property more commonly associated with CD8a þ cdcs than with CD8a cdcs in the spleen [29]. The cross-presentation capacity of cdc-like cells would be consistent with expression of CD24, a costimulator molecule known to facilitate antigen presentation to CD8 þ T cells by CD24 þ CD8a þ cdc [30].AninvivoCD8a CD24 þ cdc subset has been characterized as highly crosspresenting and a precursor of CD8a þ cdc [31]. However, cdc-like cells produced in cocultures express CD11b and reflect a cell type distinct to CD8a þ cdcs. Cocultureproduced cdc-like cells can also be distinguished from modcs, because precursors of modcs are Ly6C þ cells [4]. In addition, after more than 28 days of coculture, dendritic-like cells produced in cocultures showed no expression of Ly6C. Although it is not yet possible to equate the in vitro derived cdc-like subset with an in vivo cell counterpart, these cells have many characteristics of cdcs and are distinct from the dominant L-DC subset, including high expression of CD11b and MHC-II, weak endocytic capacity, and an ability to activate both CD4 þ and CD8 þ T cells. In comparison with L-DCs, the cdc-like subset also produced in cocultures was shown to have approximately 10-fold upregulated IL10 gene expression and upregulated expression of other IL-10 family immunoregulatory cytokines, such as IL-19 and IL-24. IL-10 is also produced by regulatory DCs, which also have a CD11c lo CD11b hi MHC- II þ phenotype similar to the cdc-like subset [32,33]. Regulatory DCs, however, reflect a loosely related, heterogeneous cell subset described in a number of in vivo and in vitro studies. Multiple regulatory DC subsets have been described, linked by an ability to inhibit T cell function [32 36]. In one study, in vitro coculture of BM-derived hematopoietic stem cells above endothelial stroma was found to result in production of regulatory DC [34]. Upregulation of IL-10, a CD11c þ CD11b þ MHC-II þ phenotype, and derivation above splenic stroma would all be consistent with cdc-like cells reflecting regulatory DCs. However, crosspresentation capacity has not been described for regulatory DC. Furthermore, the ability to activate CD4 þ T cells without induction of their proliferation would also represent a distinctive property of cdc-like cells that could relate to the ability to induce tolerance or suppression rather than activation of CD4 þ T cells. Further investigation is required to categorize further the cdc-like subset produced transiently in 5G3 cocultures. This study has clearly demonstrated the independent lineage origin and in vitro developmental capacity of the L-DC subset in comparison with cdc-like cells. When the L-DC subset of CD11c þ MHC-II cells was isolated and recultured over 5G3, a steady rise in L-DC numbers was observed over 14 days, suggesting that either the CD11c þ MHC-II population contains a precursor that differentiates to give L-DCs, or that 5G3 stroma supports L-DC proliferation. In contrast, reculture of isolated cdclike cells revealed a short-lived population, unable to replenish itself over 14 days in stromal cultures. This finding is consistent with fully differentiated cells or an absence of precursors within the isolated CD11c þ MHC- II þ subset. The CD11c MHC-II subset was found to contain a small number of precursors of cdc-like cells, because low numbers of cdc-like cells were observed transiently upon reculture. In terms of phenotype and functional capacity, L-DCs are distinct both in relation to known subsets of cdcs and pdcs, as well as the cdc-like cells produced in cocultures. They are also distinct from monocytes, macrophages, and granulocytes, both in terms of markers studied here and recent data in preparation. Increasing evidence now distinguishes L- DCs from regulatory DCs and modcs and indicates their lineage origin as a spleen-endogenous APC derived from self-renewing progenitors resident in the spleen and dependent on a splenic stromal cell type for their development [7 9]. Future studies are being directed at defining the L- DC progenitor in the spleen and the functional capacity of L-DCs, which makes them unique as a tissue-specific APC.

12 292 P. Periasamy and H.C. O Neill/ Experimental Hematology 2013;41: The 5G3 splenic stromal cell line is also being studied in terms of its lineage, ability to support hematopoiesis, and localization within the spleen environment. Acknowledgments This work was supported by project Grant No (to H.C.O.) from the National Health and Medical Research Council of Australia. P.P. was supported by an Australian National University Graduate Scholarship. Author contributions: P.P. designed and performed experiments and wrote the paper. H.C.O. designed experiments, interpreted data, and wrote the paper. Conflict of interest disclosure No financial interest/relationships with financial interest relating to the topic of this article have been declared. References 1. Naik SH, Metcalf D, van Nieuwenhuijze A, et al. Intrasplenic steadystate dendritic cell precursors that are distinct from monocytes. Nat Immunol. 2006;7: Onai N, Obata-Onai A, Schmid MA, et al. 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Splenic stroma drives mature dendritic cells to differentiate into regulatory dendritic cells. Nat Immunol. 2004;5:

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