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The Journal of Immunology, 2004, 172: 7408-7416.
Copyright © 2004 by The American Association of Immunologists

Increased and Long-Term Generation of Dendritic Cells with Reduced Function from IL-6-Deficient Bone Marrow1

Joshua I. Bleier, Venu G. Pillarisetty, Alaap B. Shah and Ronald P. DeMatteo2

Hepatobiliary Service, Memorial Sloan-Kettering Cancer Center, New York, NY 10024


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The importance of IL-6 in dendritic cell (DC) development and function has not been well defined. To establish the role of IL-6, we studied bone marrow-derived DC (BMDC) and freshly isolated splenic DC from IL-6–/–-transgenic mice. We found that although IL-6–/– bone marrow had a similar composition to that of wild-type (WT) mice, it generated up to 10 times more DC when cultured in GM-CSF. The difference persisted even when IL-6–/– and WT bone marrow were cultured together, excluding the possibility that the effects were simply due to different cytokine microenvironments. In comparison to WT BMDC, IL-6–/– BMDC captured at least as much Ag, had an equivalent surface phenotype, and matured similarly in response to LPS or CpG. However, IL-6–/– BMDC induced less T cell allostimulation and Ag-specific T cell activation, but only the former was related to their inability to generate IL-6. Although WT bone marrow cultures died within 4 wk, IL-6–/– cultures continued to generate BMDC for >120 days, although the BMDC became immature and less functional. In vivo, we found that IL-6–/– mice had similar numbers and types of splenic DC as WT mice, both normally and after treatment with either Flt-3 ligand or GM-CSF. These findings demonstrate that IL-6 has profound effects on DC development in vitro, although the number and subtype composition of DC are unaffected by the absence of IL-6 in vivo. Furthermore, secretion of IL-6 is critical to certain DC functions.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Interleukin 6 is a pleiotropic cytokine involved in immunity, liver regeneration, and hemopoiesis (1, 2). It is produced by macrophages, dendritic cells (DC),3 T cells, B cells, fibroblasts, endothelial cells, and keratinocytes (3). IL-6 exerts variable effects in innate and acquired immunity. IL-6 secretion is an early response to LPS and TNF-{alpha} and stimulates the production of acute phase proteins in the liver (4). However, in other circumstances, IL-6 reduces inflammation by suppressing IL-1 and TNF-{alpha} synthesis while leaving the inhibitory cytokines IL-10 and TGF-{beta} unaffected (5). IL-6 provides a "second-signal" to T cells, especially in conjunction with accessory cytokines such as IL-1 (6). IL-6 promotes Th2 responses in two ways. It induces CD4+ T cells to make IL-4 and inhibits Th1 responses by activating suppressor of cytokine signaling 1, which interferes with IFN-{gamma} signaling (7, 8). IL-6 can also convert resting T cells into an IL-2-responsive state by promoting their entry into the G1 phase of the cell cycle (9, 10).

DC are the most potent APCs in the immune system. A number of cytokines have been implicated in DC differentiation and function. However, there are discrepant reports of the effects of IL-6 on DC development. Treatment of CD34+ human DC progenitors with medium from renal cell carcinomas containing IL-6 and G-CSF (11) or IL-6 alone (12) inhibited DC generation and promoted monocytic differentiation. Similarly, Mitani et al. (13) found that IL-6 skewed human monocytes toward a macrophage lineage and away from DC. Chomarat et al. (14) determined that the mechanism was via up-regulation of monocyte M-CSF receptors which then bind autocrine M-CSF. In contrast, there are two reports showing that IL-6 promotes DC differentiation. Brasel et al. (15) established that blocking IL-6 function reduced the number of DC generated in Flt-3 ligand-supplemented murine wild-type (WT) bone marrow cultures. In another report, higher IL-6 levels correlated with greater DC yield in cultures of CD34+ cells (16).

Although DC are known to produce IL-6, data regarding the contribution of IL-6 secretion to DC function are sparse. Initial experiments demonstrated that the addition of supraphysiologic doses of IL-6 increased allogeneic T cell proliferation in a MLC with human monocytes (6). In contrast, others showed that adding IL-6 to human peripheral blood monocytes (17) or blocking IL-6 in lung DC (18) had no effect on alloproliferation. Grohmann et al. (19) have shown that treating "tolerogenic" CD8{alpha}+ DC with supraphysiologic doses of IL-6 enabled them to induce immunity against a tumor Ag while blockade of IL-6 abrogated the immunogenic effects of CD40 ligation in CD8{alpha}+ DC. A recent report (20) demonstrated that DC production of IL-6 abrogates the suppressive effects of regulatory T cells.

To resolve the conflicting data on the contribution of IL-6 to DC development and establish the importance of IL-6 secretion to their function, we studied bone marrow DC (BMDC) and splenic DC from IL-6–/– mice. We found that IL-6–/– bone marrow cultured in GM-CSF generated a far greater quantity of DC than WT bone marrow. Strikingly, IL-6–/– bone marrow generated DC for as long as it was cultured (>120 days), although the DC became immature and less functional. In contrast, IL-6–/– and WT mice had similar numbers of splenic DC, both normally and after treatment with GM-CSF or Flt-3 ligand. Although IL-6–/– BMDC and splenic DC had a normal phenotype, their inability to produce IL-6 significantly altered some of their functions. These data establish that IL-6 plays a critical role in normal DC function and in vitro development, but is redundant to achieve normal numbers of DC in vivo.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals

Male 4- to 8-wk-old C57BL/6 (B6, H-2Kb, CD45.2) mice and BALB/c (H-2Kd) mice were purchased from Taconic Farms (Germantown, NY). IL-6–/– (H-2Kb, CD45.2) mice (backcrossed for 22 generations with B6 mice) (21), OT-II mice, and B6 mice carrying the CD45.1 Ag were purchased from The Jackson Laboratory (Bar Harbor, ME). The Institutional Animal Care and Use Committee approved all procedures. Mice were euthanized by CO2 inhalation.

DC isolation and culture

DC were generated from bone marrow according to the method of Inaba (22) with modifications (23). Briefly, bone marrow from the femurs and tibiae of mice was grown in RPMI 1640 with 10% heat-inactivated FBS, 2 mM L-glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin, 0.05 mM 2-ME, and 20 ng/ml GM-CSF (J558 supernatant, gift from R. Steinman, The Rockefeller University, New York, NY). Cultures were initiated by placing 7.5 x 106 bone marrow cells in 10 ml of medium onto 100-mm petri dishes (Falcon 1029 plates; BD Labware, Franklin Lakes, NJ). On day 3, another 10 ml of medium was added. Starting on day 6, 10 ml of medium was replaced every 3–4 days. DC were purified from cultures by using CD11c immunomagnetic beads according to the manufacturer’s protocol (Miltenyi Biotec, Auburn, CA). IL-6 (10 ng/ml; R&D Systems, Minneapolis, MN) or anti-IL-6 Ab (10 µg/ml, clone MP5–20F3; BD PharMingen, San Diego, CA) or its isotype (BET-2, rat IgG1, mAb Core; Sloan-Kettering Institute, New York, NY) was added in some experiments. Cultures that survived for longer than 45 days were considered to be long term.

Splenic DC were isolated as described previously (24). Liver DC were isolated as detailed elsewhere (25), with modifications. Briefly, after animals were euthanized by CO2 inhalation, the portal vein was cannulated and injected with 2 ml of 1% (w/v) collagenase D (Sigma-Aldrich, St. Louis, MO) in PBS. After mechanical disruption, the cell suspension was passed through a 100-µm filter and then centrifuged three times at 30 x g to remove hepatocytes. The remaining nonparenchymal cells were pelleted and after RBC lysis, CD45+ cells were purified using immunomagnetic beads according to the manufacturer’s protocol (Miltenyi Biotec).

Recombinant adenovirus

Recombinant adenoviruses were propagated, purified, and stored as previously described (26). AdGM-CSF, AdFlt3L, and AdGFP (Quantum Biotechnologies, Montreal, Quebec, Canada) encode murine GM-CSF, murine Flt-3 ligand, and green fluorescent protein, respectively, under the control of the CMV promoter. Mice (two per group) were treated with a single tail vein injection of 6 x 1010 particles of virus. Ten days later, they were sacrificed and splenic DC were isolated.

Flow cytometry

Flow cytometry was performed on a FACSCalibur flow cytometer (BD Biosciences, Mountain View, CA). We stained 1 x 106 cells/tube with 1 µg FITC, PE, Per-CP, or allophycocyanin-conjugated Ab (BD PharMingen, except as indicated) after blocking with 1 µg anti-Fc{gamma}RIII/II Ab (clone 2.4G2, mAb Core; Sloan-Kettering Institute). Cells were stained for the DC marker CD11c (HL3), lineage markers (CD3{epsilon} (145-2C11)), Ter-119, CD11b (M1/70), Gr-1 (RB6-8C5), CD45R (RA3-6B2), CD4 (RPA-T4), CD8{alpha} (53-6.7), NK1.1 (PK136), Mac-3 (M3/84), F4/80 (CI:A3-1; Serotec, Oxford, U.K.), Sca-1 (E13-161.7), Thy1.1 (HIS51), and KIT (2B8)); MHC class II (I-Ab); intercellular adhesion marker CD54 (3E2); costimulatory molecules CD40 (3/23), CD80 (B7-1 (1G10)), and CD86 (B7-2 (GL1)); IL-6R (CD126); and the apoptosis markers Fas (JO2), Fas ligand (MFL3), TRAIL (R&D Systems), TRAIL-R4 (R&D Systems), and annexin V.

DC assays

To determine the ability of DC to mature, we cultured 1 x 105 DC in 24-well plates with LPS (1 µg/ml), CpG (oligodeoxynucleotide 1826, 10 µg/ml), CpG control (oligodeoxynucleotide 1982, 10 µg/ml] (Oligos, Etc., Wilsonville, OR), anti-CD40 Ab (FGK45, 25 µg/ml; Memorial-Sloan Kettering Cancer Center Monoclonal Core Facility) (27), TNF-{alpha} (100 ng/ml), IFN-{gamma} (100 ng/ml), or IL-4 (20 ng/ml; R&D Systems). After 24 h, cells were harvested for flow cytometric analysis and supernatant was collected for ELISA. IL-4, IL-6, IL-10, IL-12, IFN-{gamma}, TNF-{alpha} (R&D Systems), and IL-2 (BD PharMingen) levels were measured according to the manufacturers’ protocol. To quantitate proliferation, we plated in triplicate on a 96-well plate (2 x 104 cells/well) WT BMDC from day 13 of culture, IL-6–/– BMDC from day 13 of culture, and IL-6–/– BMDC from day 87 of culture. The number of DC was measured 1, 2, and 4 days later using the CellTiter 96 Proliferation Assay (Promega, Madison, WI). In other experiments, we labeled adherent cells from day 10 bone marrow cultures with 5 µM CFSE (Molecular Probes, Eugene, OR) for 5 min and then measured their fluorescence 3 and 4 days later.

To measure DC Ag uptake, we incubated 1–5 x 105 DC in 100 µl with FITC-albumin or FITC-dextran (1 mg/ml; Sigma-Aldrich) and measured mean fluorescence at various time points using flow cytometry. MLR was performed as previously described (24). CD11c-beaded DC were irradiated (3000 rad) and mixed in various amounts with 3 x 105 allogeneic T cells in a 96-well U-bottom plate (Falcon; BD Labware). T cells were isolated from BALB/c mouse spleens with CD90 (Thy1.2) immunomagnetic beads according to the manufacturer’s protocol (Miltenyi Biotec). LPS (1 µg/ml), CpG (10 µg/ml), TNF-{alpha} (20 ng/ml), or IL-12 (20 ng/ml) was added in some experiments. Cultures were pulsed with [3H]thymidine (1 µCi/well) at 72 h and radioactive uptake was measured 20 h later. We measured Ag-specific CD8+ T cell stimulation as we have previously (24). Briefly, we incubated purified DC with 10 µg/ml OVA257–264 peptide (SIINFEKL) for 1 h and then mixed them in various amounts with 5 x 104 GB10 cells, which are H-2Kb-restricted OVA-specific CD8+ T cells. After 72 h, supernatant was harvested and assayed for IL-2 production with ELISA. To assess CD4+ T cell-specific stimulation, we incubated purified DC with 10 µg/ml OVA323–339 peptide for 1 h and then mixed them in various amounts with 2.4 x 103 OT-II T cells. In some wells, IL-6 (10 ng/ml) was added. Cultures were pulsed with [3H]thymidine (1 µCi/well) at 96 h and radioactive uptake was measured 20 h later.

Statistical analysis

Flow cytometry results were assessed using the {chi}2 test. The Student’s t test was used to compare two groups. All other comparisons were performed using ANOVA. SPSS statistical software (version 11.5; SPSS, Chicago, IL) was used.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
IL-6–/– bone marrow generates greater numbers of DC than WT bone marrow

To determine the effect of IL-6 on DC development in vitro, we cultured 7.5 x 106 bone marrow cells from IL-6–/– and WT mice separately. By day 10, both cultures had a confluent layer of adherent cells, but IL-6–/– bone marrow cultures contained three to four times more floating cells. We discovered that nonadherent IL-6–/– cells had a significantly higher percentage of DC based on CD11c expression (Fig. 1A). In fact, IL-6–/– cultures were nearly all CD11c+ by day 15, while WT cultures contained only 65% DC at most. The majority of the CD11c– WT cells proved to be granulocytes since they were Ly6-G+ and did not stain for T cell, B cell, or macrophage markers (Fig. 1B). Because of both the increased total number of cells and the higher percentage of DC, IL-6–/– bone marrow cultures generated up to 10 times more DC than WT cultures. This effect was most pronounced by days 11 and 12 of culture, when IL-6–/– cultures contained ~1.2 x 107 BMDC per 100-mm plate compared with 1–2 x 106 BMDC in WT cultures (Fig. 1C). To determine whether the difference was due to increased production or longevity of IL-6–/– BMDC, we labeled adherent cells from day 10 of bone marrow cultures with CFSE and then measured their fluorescence over time. We found equivalent mean fluorescence between adherent IL-6–/– and WT BMDC 3 (200 vs 173) and 4 (114 vs 103) days later. Furthermore, adherent IL-6–/– and WT cells had a similar percentage of cells in S phase (4.0 vs 3.4%). These findings suggest that IL-6–/– BMDC are longer-lived than WT BMDC.



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  		FIGURE 1. IL-6–/– (knockout (KO)) bone marrow generate more DC than WT bone marrow. On day 0, 7.5 x 106 bone marrow cells were placed on 100-mm plates. Nonadherent cells were harvested, counted, and stained for CD11c on various days thereafter. Values shown are a mean of three plates per time point and results are representative of five experiments. A, IL-6–/– bone marrow cultures contained a higher percentage of CD11c+ cells at each time point tested (p < 0.001). B, WT bone marrow cultures contained a higher proportion of granulocytes than did IL-6–/– cultures on day 10. C, IL-6–/– bone marrow produced up to 10 times more DC than WT bone marrow (p < 0.001).

 
IL-6–/– bone marrow generates increased numbers of DC despite physiologic levels of IL-6

Using ELISA, we could identify only minimal amounts (6 pg/ml) of IL-6 in WT bone marrow cultures on day 10. To determine whether physiologic levels of IL-6 from WT bone marrow would prevent the increased generation of BMDC from IL-6–/– bone marrow, we cultured equal numbers of WT (CD45.1) and IL-6–/– (CD45.2) bone marrow together in the same plate. After 7 days of culture, IL-6–/– BMDC began to outgrow WT BMDC and by day 17 almost all of the cells were. IL-6–/– (CD45.2+) BMDC (Fig. 2A). The physiologic levels of IL-6 from WT cells did appear to exert some effect though, as there was an ~33% decrease in the yield of IL-6–/– BMDC when IL-6–/– bone marrow was cultured with WT bone marrow compared with when it was cultured alone. In contrast, a supraphysiologic dose (10 ng/ml) of IL-6 added to culture on day 0 was able to substantially reduce the yield of BMDC from IL-6–/– and, to a lesser extent, WT bone marrow (Fig. 2B). Next, we wanted to determine whether there was a specific time period during which the absence of IL-6 was critical to enhancing IL-6–/– BMDC development. We added IL-6 (10 ng/ml) to IL-6–/– bone marrow cultures starting at various time points and then measured the number of BMDC on day 12. We found that the inclusion of IL-6 at any time reduced the yield of IL-6–/– BMDC. In particular, the addition of IL-6 starting on days 2 or 4 had the most profound effect (Fig. 2C).



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  		FIGURE 2. The growth advantage of IL-6–/– bone marrow DC is not accounted for by a lack of physiologic amounts of IL-6 and is not due to a different starting composition. Values shown are a mean of three plates per time point and results are representative of two experiments. A, 3.75 x 106 WT (CD45.1) and IL-6–/– (CD45.2) bone marrow cells were cultured together on the same 100-mm plate. Using flow cytometric analysis, we determined the relative percentage of CD45.1+ and CD45.2+CD11c+ DC various days later. IL-6–/– cells became the dominant lineage by day 12 of culture (p < 0.01). B, To determine whether supraphysiologic amounts of IL-6 could alter BMDC generation, we added 10 ng/ml IL-6 on day 0 to separate cultures of WT or IL-6–/– bone marrow. Addition of IL-6 to the bone marrow cultures of either group resulted in a decrease in the yield of DC (p < 0.001). C, A supraphysiologic amount of IL-6 (10 ng/ml) was added to IL-6–/– bone marrow cultures starting on various days and then the number of DC on day 12 were counted. Results with WT and IL-6–/– bone marrow cultured without IL-6 are shown on the left. D, Flow cytometric analysis of freshly isolated IL-6–/– and WT bone marrow showed similar expression of the lineage markers Sca-1, Kit, F4/80, CD3, NK 1.1, and B220. KO, Knockout.

 
IL-6–/– and WT bone marrow have similar composition

We then postulated that differences in the composition of IL-6–/– and WT bone marrow could account for the disparity in the yield of BMDC. We found that age-matched IL-6–/– and WT mice had equal numbers of marrow cells (~7.5 x 106/hind limb). By flow cytometry, we found that there was equal bone marrow expression of Sca-1, Kit, F4/80, CD3, NK 1.1, and B220 (Fig. 2D). Furthermore, there was a similar percentage of Lin–c-kit+Thy1.1lowSca-1high hemopoietic stem cells (28) in IL-6–/– and WT bone marrow (0.11 vs 0.16%, respectively). Lin– was defined as the absence of CD3{epsilon}, Ter-1, B220, CD11b, Gr-1, CD4, and CD8{alpha} staining.

IL-6–/– BMDC have a similar phenotype and maturation as WT BMDC

We next determined the phenotype of IL-6–/– BMDC. Similar to WT BMDC, IL-6–/– BMDC had high CD11b expression but lacked CD4 and CD8{alpha} staining (Fig. 3A) (22, 26). Nevertheless, microscopic analysis of IL-6–/– and WT BMDC demonstrated that the former were larger and contained more extensive cytoplasmic veils and dendritic processes (Fig. 3B). To test whether there were differences between the maturation level of IL-6–/– and WT BMDC, we examined their surface expression of MHC class II, CD40, and CD86. IL-6–/– and WT BMDC had similar maturation by flow cytometry at baseline and after a 24-h incubation with LPS, CpG, anti-CD40, TNF-{alpha}, IFN-{gamma}, or IL-4 (Fig. 3C). Neither IL-6–/– nor WT BMDC underwent maturation in response to bead purification and replating alone.



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  		FIGURE 3. IL-6–/– BMDC express similar phenotypic markers as WT BMDC. Nonadherent cells from day 10 of bone marrow culture were harvested and tested by flow cytometry or processed for cytospin. A, CD11c+ IL-6–/– and WT BMDC had similar staining for the dendritic subset markers CD11b, CD8{alpha}, and CD4. Representative of five experiments. B, By microscopic analysis, IL-6–/– BMDC were significantly larger and had more extensive veils and dendritic processes after 10 or 24 days of culture (magnification, x400). C, IL-6–/– (open histograms) and WT (filled histograms) BMDC from day 10 of culture had similar expression of MHC class II, CD40, and CD86 at baseline. IL-6–/– and WT BMDC had similar maturation after overnight incubation with LPS (1 µg/ml), CpG (10 µg/ml), anti-CD40 (25 µg/ml), TNF-{alpha} (100 ng/ml), IFN-{gamma} (100 ng/ml), or IL-4 (20 ng/ml). Representative of two experiments. All isotypes fell within the first decade of histograms (data not shown). KO, Knockout.

 
IL-6–/– BMDC are functionally distinct from WT BMDC

Given that IL-6–/– BMDC had a similar surface marker phenotype as WT BMDC, we then determined whether they had similar function. We tested Ag uptake and found that although both took up dextran equally, IL-6–/– BMDC had a greater ability to capture OVA than did WT BMDC (Fig. 4A). However, this difference involved only the magnitude of OVA uptake since nearly all DC in both groups contained fluorescent Ag within 5 min (data not shown).



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  		FIGURE 4. IL-6–/– BMDC are functionally distinct from WT BMDC. A, To determine Ag uptake ability, BMDC were mixed with either FITC-OVA or FITC-dextran (1 mg/ml). The reaction was stopped and mean fluorescence was determined at various time points by flow cytometry. IL-6–/– BMDC captured similar amounts of dextran, but took up more OVA on a per cell basis than did WT BMDC. B, A MLR was performed by mixing various numbers of immunomagnetic bead-purified, irradiated WT, or IL-6–/– BMDC with 2 x 105 allogeneic (BALB/c) T cells. In some groups, 10 ng/ml IL-6 were added to the wells on day 0. T cells incubated alone proliferated minimally (~7900 cpm). This was increased ~2- to 3-fold when IL-6 was added, but remained unchanged when anti-IL-6 was added. The MLR was repeated by first stimulating the DC for 2 h with either CpG (10 µg/ml, C) or LPS (1 µg/ml, D). Ag-specific CD8+ T cell activation was tested by loading DC with OVA257–263 peptide and then mixing them with 5 x 104 OVA257–263- specific GB10 T cells in a 96-well dish. Supernatant IL-2 was measured at 72 h. In some cases, the BMDC were stimulated with either CpG (10 µg/ml, E) or LPS (1 µg/ml, F). IL-6 (10 ng/ml) or an IL-6 blocking Ab (10 µg/ml) was added to some wells. G, Ag-specific CD4+ T cell activation was tested by loading DC with OVA323–339 peptide and then mixing them with 2.4 x 103 OVA323–339 TCR-transgenic T cells in a 96-well dish. In one group, 10 ng/ml IL-6 was also added. H, The cytokine profiles of day 10 IL-6–/– and WT BMDC were determined by assaying supernatants with ELISA 24 h after culture with either medium, LPS (1 µg/ml), CpG (10 µg/ml), anti-CD40 (25 µg/ml), IL-4 (20 ng/ml), or IFN-{gamma} (100 ng/ml). Results of triplicate wells are shown. IL-6–/– BMDC made more IL-12 than WT BMDC after stimulation with LPS and TNF-{alpha} (p < 0.01). IL-6–/– BMDC made more TNF-{alpha} than WT BMDC in all conditions tested (p < 0.05). Neither group made measurable amounts of IL-4 or IFN-{gamma}. IL-10 secretion was ~25 pg/ml after LPS stimulation, 150 pg/ml with CpG, and 275 pg/ml with CD40 ligand. In each case, IL-6–/– BMDC produced slightly less IL-10 (data not shown). CpG control peptide did not produce any effect (data not shown). Culture with either LPS or CpG resulted in dramatic IL-6 production by WT BMDC. Data are presented as mean ± SEM.

 
Because the hallmark of DC function is their activation of T lymphocytes, we tested the ability of IL-6–/– and WT BMDC to stimulate allogeneic and Ag-specific T cells. In a MLR, IL-6–/– BMDC induced less proliferation of allogeneic T cells than did WT BMDC (Fig. 4B). This difference was only partially due to the inability of IL-6–/– BMDC to secrete IL-6. Although adding an excess (10 ng/ml) of IL-6 increased the MLR of both types of BMDC, WT BMDC still induced substantially more allogeneic T cell proliferation (Fig. 4B). To determine whether other stimulatory cytokines would synergize with IL-6 to increase the function of IL-6–/– BMDC, we cultured IL-6–/– BMDC in IL-6 plus either TNF-{alpha} or IL-12. We found that TNF-{alpha} had no effect and, consistent with our previous findings (29) and those of Kelleher et al. (30), IL-12 actually decreased the magnitude of allostimulation induced by IL-6–/– BMDC (data not shown). To determine whether microbial stimulation allowed IL-6–/– BMDC to "catch up" to WT BMDC, we incubated IL-6–/– BMDC with CpG or LPS for 2 h before using them in a MLR. Stimulated IL-6–/– BMDC treated with either CpG or LPS induced equal alloproliferation as unstimulated WT BMDC but less than stimulated WT BMDC (Fig. 4, C and D). However, the addition of IL-6 to stimulated IL-6–/– BMDC allowed them to achieve equal MLR as stimulated WT BMDC. This was consistent with our finding that stimulated WT BMDC that had been incubated with CpG or LPS for 2 h made enormous amounts (86 and 135 ng/ml, respectively) of IL-6 after subsequent overnight culture. To determine whether the enhanced MLR ability seen by stimulated IL-6–/– BMDC was due to other cytokines being secreted, we blocked either TNF-{alpha} or IL-12, but this had no effect (data not shown).

When we tested DC interaction with Ag-specific T cells using an H-2Kb-restricted OVA-specific CD8+ T cell line, we again found that IL-6–/– BMDC were less potent (Fig. 4, E and F). However, in this case, IL-6 did not contribute to T cell activation since adding it to either unstimulated or stimulated (with LPS or CpG) IL-6–/– BMDC or blocking IL-6 in WT BMDC had no effect. This result was not specific to this cell line since we obtained similar results with freshly isolated CD8+ T cells from an OVA257–264 TCR (OT-I)-transgenic mouse (data not shown) as well as with freshly isolated CD4+ T cells from an OVA323–339 TCR (OT-II)-transgenic mouse (Fig. 4G).

Because IL-6–/– BMDC had decreased T cell stimulatory capacity despite their similar costimulatory molecule expression as WT BMDC, we questioned whether other differences in cytokine expression besides IL-6 explained their disparate function. We analyzed the supernatants of BMDC from day 10 of culture and found that unstimulated IL-6–/– and WT BMDC had similar cytokine profiles. Both had undetectable levels of IL-4, IL-6, IL-10, IL-12, and IFN-{gamma} while IL-6–/– BMDC and, to a lesser extent, WT BMDC secreted TNF-{alpha}. Therefore, the cytokine profiles did not account for the functional differences. We did detect differences after stimulation with either LPS or CpG as IL-6–/– BMDC made more IL-12 (p70) and TNF-{alpha} (Fig. 4H), but less IL-10 (data not shown). These data are consistent with recent findings (31) showing that IL-6 inhibits IL-12 production by CD11b+ pulmonary DC and that IL-6–/– pulmonary DC secrete increased amounts of IL-12.

IL-6–/– BMDC persist in long-term culture but become immature and less functional

To determine whether there were differences in the longevity of IL-6–/– and WT bone marrow cultures beyond 2 wk (Fig. 1C), we continued adding fresh medium to the cultures every 3–4 days. As expected, WT bone marrow cultures died out by 20–30 days. Surprisingly, IL-6–/– bone marrow cultures remained viable and continued to proliferate for as long as we changed the medium (>120 days, Fig. 5A). Both the adherent and floating cells were able to perpetuate the cultures. Adding fresh medium to the IL-6–/–, adherent cells generated 3–4 x 106 DC per 100-mm plate within 4 days. Alternatively, when we replated the floating BMDC, they quickly adhered and generated nonadherent DC over the course of 2–3 wk (data not shown). Flow cytometric analysis revealed that both the adherent and nonadherent cells subsequently generated floating cells that were uniformly CD11c+. Additionally, we found that nonadherent long-term IL-6–/– BMDC had similar growth kinetics as young IL-6–/– BMDC (Fig. 5B).



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  		FIGURE 5. IL-6–/– bone marrow cultures live much longer than WT bone marrow cultures. A, When 10 ml of medium was replaced every 3 days, WT cultures died out by ~4 wk. In contrast, IL-6–/– bone marrow cultures continued to generate DC for as long as they were cultured (>120 days). Data shown are from an average of three plates per day. B, We plated WT or IL-6–/– BMDC from day 10 of culture or IL-6–/– BMDC from day 87 of culture. Living cells were counted on days 1, 2, and 4. IL-6–/– BMDC from day 10 and day 87 of culture had similar growth kinetics. Results shown are from an average of triplicate wells. Data are presented as mean ± SEM. C, By flow cytometry, day 87 IL-6–/– BMDC were more immature than day 10 IL-6–/– BMDC and had reduced expression of MHC class II, CD40, CD80, and CD86. Isotypes fell within the first decade (data not shown). Long-term IL-6–/– BMDC induced less allostimulation in a MLR (D) and less Ag-specific T cell activation (E) in an Ag-specific T cell assay than younger IL-6–/– BMDC. Representative of at least three experiments except for D, which was performed twice. Over 10 separate long-term cultures were generated, all with comparable characteristics. KO, Knockout.

 
We then examined the phenotype and function of the long-term IL-6–/– BMDC. They were immature with decreased expression of MHC class II, CD40, CD80, and CD86 compared with IL-6–/– BMDC from day 10 of culture (Fig. 5C). However, 30% of long-term IL-6–/– BMDC expressed the IL-6R, which we were unable to detect on either WT or IL-6–/– BMDC from day 10 of culture. Long-term IL-6–/– BMDC were able to mature in response to LPS or CpG stimulation (as determined by class II, CD40, and CD86 expression), although to a lesser degree than day 10 IL-6–/– BMDC (data not shown). Consistent with their immature state, the long-term IL-6–/– BMDC had decreased ability to stimulate allogeneic (Fig. 5D) or Ag-specific GB10 CD8+ T cells (Fig. 5E). However, we cannot conclude that the observed decrease in maturation and function in long-term IL-6–/– BMDC is truly specific, since there is no appropriate control for comparison.

IL-6 affects splenic DC function but not in vivo development

We next sought to determine whether IL-6 also affected DC development in vivo as it had in vitro. We found that naive WT and IL-6–/– mice had an equal number of splenic DC (Table I). Just as we had seen with BMDC, splenic DC from IL-6–/– and WT mice had similar maturation with equal expression of MHC class II and CD40 (data not shown). In addition, spleens from WT and IL-6–/– mice had a similar percentage of myeloid (85.4 vs 83.3%), lymphoid (9.5 vs 12.1%), and plasmacytoid (0.9 vs 1.0%) DC. Liver DC subset composition was also comparable (Fig. 6). Because GM-CSF led to increased numbers of BMDC from IL-6–/– bone marrow, we then tested whether systemic treatment with GM-CSF led to greater expansion of DC in IL-6–/– mice than in WT mice. We found equivalent splenic DC expansion 10 days after i.v. treatment with an adenoviral vector that we have used previously (26) which encodes the murine GM-CSF transgene. DC expansion was also equal between IL-6–/– and WT mice when we treated them with murine Flt-3 ligand.


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  		Table I. Comparison of splenic DC from IL-6–/– and WT micea

 


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  		FIGURE 6. WT and IL-6–/– mice have similar splenic and liver DC subset composition. Freshly isolated splenic DC and liver DC from WT and IL-6–/– mice contain similar numbers of myeloid (CD11b+CD8{alpha}–), lymphoid (CD11b–CD8+, and plasmacytoid (CD11c+B220+) DC. In the upper row, cells are gated on CD11c+ splenocytes or CD45+ hepatic nonparenchymal cells. In the lower row, cells are gated on viable splenocytes determined by forward scatter vs side scatter of CD45+ hepatic nonparenchymal cells. Data are representative of three separate analyses. KO, Knockout.

 
Having established that IL-6 was not essential for achieving normal numbers of DC in vivo, we next assessed whether DC secretion of IL-6 contributed to their function. As we found with BMDC, IL-6–/– splenic DC captured more OVA but equal dextran when compared to WT splenic DC (Fig. 7A). Additionally, we showed that like BMDC, IL-6–/– splenic DC had a decreased ability to stimulate allogeneic T cells. This deficit was due to their inability to produce IL-6. When exogenous IL-6 was added, WT and IL-6–/– splenic DC caused similar alloproliferation. Conversely, anti-IL-6 greatly reduced the immunostimulatory capacity of WT splenic DC (Fig. 7B).



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  		FIGURE 7. Splenic DC from IL-6–/– mice have altered function. A, Freshly isolated, immunomagnetic bead-purified CD11c+ splenic DC from WT and IL-6–/– mice were tested for Ag uptake ability. IL-6–/– splenic DC captured similar amounts of dextran but took up more OVA on a per cell basis than did WT BMDC. B, A MLR was performed with freshly isolated splenic DC from WT and IL-6–/– mice. IL-6–/– splenic DC induced less allogeneic T cell proliferation than did WT splenic DC. The addition of IL-6 (10 ng/ml) enabled WT and IL-6–/– splenic DC to achieve similar alloproliferation which was increased in both instances. Blocking IL-6 (10 µg/ml) reduced WT splenic DC MLR ability to below that of IL-6–/– splenic DC. T cells incubated alone had ~1500 cpm and this was increased ~3- to 4-fold when IL-6 was added, but remained unchanged when anti-IL-6 was added. Representative of two experiments. Data are presented as mean ± SEM. KO, Knockout.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We have shown that IL-6 has a profound influence on DC development in vitro. IL-6–/– bone marrow generated up to 10 times more DC than WT bone marrow. Furthermore, IL-6–/– bone marrow cultures became nearly 100% CD11c+ by day 15 of culture compared with 65% purity reached in WT cultures. Although IL-6–/– BMDC generation was only slightly reduced in the presence of physiologic levels of IL-6, excessive (10 ng/ml) IL-6 reduced BMDC generation in both IL-6–/– and WT bone marrow cultures. Furthermore, we established that the IL-6-mediated inhibition of DC development from IL-6–/– bone marrow was time dependent, with inhibition being most potent when IL-6 was added starting between days 2 and 4 of culture. Our data demonstrating that IL-6 inhibits BMDC generation are consistent with several previous findings that IL-6 prevents monocytes from differentiating into DC (11, 14). However, Santiago-Schwarz et al. (16) showed that when CD34+ progenitor cells were cultured with GM-CSF, TNF-{alpha}, and stem cell factor, the yield of DC was increased in cultures with higher supernatant IL-6 levels and reduced by the addition of an anti-IL-6 Ab. In contrast to our findings of increased BMDC generation from IL-6–/– bone marrow culture in GM-CSF, Brasel et al. (15) demonstrated that IL-6–/– bone marrow cultured solely in Flt-3 ligand produced much fewer BMDC than WT bone marrow. Thus, the effect of IL-6 on DC generation appears to depend on the cytokine microenvironment as IL-6 generally inhibits DC differentiation in the presence of GM-CSF but promotes DC development with Flt-3 ligand.

We examined other possibilities besides the absence of IL-6 to explain the higher yield of IL-6–/– BMDC compared with WT BMDC. We excluded that other cytokine differences between IL-6–/– and WT bone marrow were responsible. We proved this by mixing IL-6–/– and WT bone marrow together and demonstrating that IL-6–/– BMDC still predominated, although the absolute number of IL-6–/– BMDC produced was less, presumably due to the physiologic amounts of IL-6 released by WT bone marrow (Fig. 2A). We also showed that the composition of IL-6–/– bone marrow was similar to that of WT mice. Specifically, IL-6–/– mice had equivalent numbers of total bone marrow cells as well as Lin–c-kit+Thy1.1lowSca-1high hemopoietic stem cells. Nevertheless, Bernad et al. (32) have previously reported that IL-6–/– mice contain one-half the normal amount of bone marrow granulocyte/monocyte precursors. Furthermore, they found fewer than normal day 12 spleen (CFU-S) in IL-6–/– bone marrow and spleen, as well as fewer pre-CFU-S in IL-6–/– bone marrow. The ability of the IL-6–/– long-term repopulating stem cell compartment (which reflects the activity of hemopoietic stem cells) to reconstitute irradiated mice was adequate initially, but diminished with successive transplantation, implying a defect in its self-renewal ability (32). Given that we found increased and sustained BMDC generation from IL-6–/– bone marrow, it appears that any potential deficits in the quantity of committed progenitors or function of stem cells in IL-6–/– mice are overcome by culture in GM-CSF.

To our surprise, IL-6–/– bone marrow cultures generated DC for as long as we cultured them (>120 days). Both the floating and adherent cells were each capable of perpetuating the cultures. Meanwhile, WT bone marrow cultures generally died within 1 mo. We were unable to successfully culture WT BMDC long-term even when we added a blocking anti-IL-6 Ab at the outset of culture. Instead, we found that this reduced the yield of WT BMDC as much as 30% by day 11 of culture (data not shown). There are only a few other reports of long-term DC cultures. Bone marrow from TNFR1–/– mice has also been shown to produce BMDC continuously and this depended on GM-CSF, just as it did for IL-6–/– BMDC in our experiments (33). However, there are numerous differences between IL-6–/– and TNFR1–/– BMDC. TNFR1–/– bone marrow cultures produced similar numbers of BMDC as WT bone marrow in the first few weeks of culture and achieved only ~50% purity of DC by CD11c staining compared with nearly 100% purity in IL-6–/– cultures. Although TNFR1–/– BMDC contained both mature and immature DC, we found long-term IL-6–/– BMDC to be uniformly immature. TNFR1–/– BMDC were MHC class IIhigh, CD40high, and CD86high while IL-6–/– BMDC had low expression of these markers. Consistent with their relative phenotypic immaturity, both types of BMDC had less ability to stimulate T cells. Nevertheless, TNFR1–/– BMDC matured in response to LPS but, unlike IL-6–/– BMDC, not to anti-CD40 or TNF-{alpha}. The mechanism of continual DC generation in TNFR1–/– bone marrow was postulated to be a lack of proapoptotic signaling via TNFR1 and a down-regulation of CD95. In contrast, Fas expression was normal in IL-6–/– BMDC (data not shown). In addition, we found that the expression of Fas ligand, TRAIL, and TRAIL-R4 was absent in both WT and IL-6–/– BMDC (data not shown). Nevertheless, we did find evidence to suggest that IL-6–/– BMDC live longer. The rate of BMDC generation was comparable between IL-6–/– and WT bone marrow since CFSE-labeled adherent cells from each group had a similar dissipation of fluorescence during 4 days of subsequent culture. However, IL-6–/– cultures contained more cells, implying enhanced longevity. We only found differences in apoptosis in the long-term cultures, which had almost 50% less annexin staining than did day 10 IL-6–/– or WT BMDC (data not shown). Other long-term DC have been generated from splenocyte cultures, but they required the addition of unidentified stromal growth factors (34, 35, 36). Winzler et al. (36) were able to continuously propagate DC from splenocytes cultured in GM-CSF and fibroblast growth factors. These DC had an immature phenotype and generated less T cell allostimulation but equal Ag-specific activation as WT DC. Ni and O’Neill (34) reported a technique using spleen-derived stromal cells to support long-term DC cultures from bone marrow, splenocytes, and thymocytes.

Although IL-6–/– mice are known to have a normal splenic lymphocyte composition (37), their spleen and liver DC have not been characterized. Despite the dramatic effects we observed of IL-6 on DC development in vitro, we found that IL-6 was unnecessary to achieve normal numbers of DC in vivo. IL-6–/– mice had a similar number and composition of DC subtypes (myeloid, lymphoid, and plasmacytoid) in their spleens as well as in their livers as compared with WT (Fig. 6). Although GM-CSF and Flt-3 ligand have striking and opposing effects on IL-6–/– BMDC development in vitro based on our findings and those of Brasel et al. (15), endogenous overexpression of these cytokines in vivo had similar effects on splenic DC expansion in IL-6–/– mice compared to WT mice. Moreover, although Funk et al. (33) reported that TNFR1–/– bone marrow cultures continuously generated BMDC, we found that spleens of TNFR1–/– mice contained normal numbers of DC (our unpublished data), further supporting that the regulation of DC development in vivo is redundant.

Unstimulated IL-6–/– BMDC and splenic DC each induced less allogeneic T cell stimulation than their WT counterparts. Exogenous IL-6 partially rescued IL-6–/– BMDC and completely rescued IL-6–/– splenic DC in a MLR. That IL-6 alone could restore at least some function to IL-6–/– DC was consistent with the fact that IL-6–/– BMDC and splenic DC were otherwise normal with a similar phenotype to WT DC and a comparable maturation after stimulation (Fig. 3, A and C). We also observed that stimulation of WT BMDC with LPS or CpG causes massive IL-6 secretion. Despite our findings with allogeneic T cell stimulation, IL-6 was not essential for all DC functions as it did not influence Ag-specific T cell activation in either IL-6–/– or WT BMDC. In addition to our findings with T cells, recent reports have demonstrated the importance of IL-6 on the interaction of DC with B cells. IL-6 produced by DC has been shown to promote B cell differentiation to plasma cells (38) and induce mucosal IgA secretion (39).

Thus, IL-6 has profound effects on DC development in vitro as IL-6–/– bone marrow generates high numbers of BMDC for an extended time. Nevertheless, the absence of IL-6 does not alter the number of DC in vivo. Our findings have application to the function of normal DC since we have shown that DC secretion of IL-6 is critical to some of their functions.


   Acknowledgments
 
We gratefully acknowledge Jan Hendrikx, Vinod Sahi, and the staff of the Memorial Sloan-Kettering Flow Cytometry Core for their advice and assistance.


   Footnotes
 
1 This work was supported in part by Grant CA94503 (to R.P.D.). Back

2 Address correspondence and reprint requests to Dr. Ronald P. DeMatteo, Memorial Sloan-Kettering Cancer Center, Box 203, 1275 York Avenue, New York, NY 10021. E-mail address: dematter{at}mskcc.org Back

3 Abbreviations used in this paper: DC, dendritic cell; WT, wild type; BMDC, bone marrow DC. Back

Received for publication September 9, 2003. Accepted for publication March 12, 2004.


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 Results
 Discussion
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