Liver Dendritic Cells Are Less Immunogenic Than Spleen Dendritic Cells because of Differences in Subtype Composition

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1 This information is current as of July 13, Liver Dendritic Cells Are Less Immunogenic Than Spleen Dendritic Cells because of Differences in Subtype Composition Venu G. Pillarisetty, Alaap B. Shah, George Miller, Joshua I. Bleier and Ronald P. DeMatteo J Immunol 2004; 172: ; ; doi: /jimmunol References Subscription Permissions Alerts This article cites 21 articles, 15 of which you can access for free at: Why The JI? Submit online. Rapid Reviews! 30 days* from submission to initial decision No Triage! Every submission reviewed by practicing scientists Fast Publication! 4 weeks from acceptance to publication *average Information about subscribing to The Journal of Immunology is online at: Submit copyright permission requests at: Receive free -alerts when new articles cite this article. Sign up at: Downloaded from by guest on July 13, 2018 The Journal of Immunology is published twice each month by The American Association of Immunologists, Inc., 1451 Rockville Pike, Suite 650, Rockville, MD Copyright 2004 by The American Association of Immunologists All rights reserved. Print ISSN: Online ISSN:

2 The Journal of Immunology Liver Dendritic Cells Are Less Immunogenic Than Spleen Dendritic Cells because of Differences in Subtype Composition 1 Venu G. Pillarisetty, Alaap B. Shah, George Miller, Joshua I. Bleier, and Ronald P. DeMatteo 2 The unique immunological properties of the liver may be due to the function of hepatic dendritic cells (DC). However, liver DC have not been well characterized because of the difficulty in isolating adequate numbers of cells for analysis. Using immunomagnetic bead and flow cytometric cell sorting, we compared freshly isolated murine liver and spleen CD11c DC. We found that liver DC are less mature, capture less Ag, and induce less T cell stimulation than spleen DC. Nevertheless, liver DC were able to generate high levels of IL-12 in response to CpG stimulation. We identified four distinct subtypes of liver DC based on the widely used DC subset markers CD8 and CD11b. Lymphoid (CD8 CD11b ) and myeloid (CD8 CD11b ) liver DC activated T cells to a similar degree as did their splenic DC counterparts but comprised only 20% of all liver DC. In contrast, the two more prevalent liver DC subsets were only weakly immunostimulatory. Plasmacytoid DC (B220 ) accounted for 19% of liver DC, but only 5% of spleen DC. Our findings support the widely held notion that liver DC are generally weak activators of immunity, although they are capable of producing inflammatory cytokines, and certain subtypes potently activate T cells. The Journal of Immunology, 2004, 172: The liver is believed to play a critical role in the regulation of both tolerance and immunity as evidenced by the observations that 1) liver allografts are commonly accepted without the need for immunosuppression, 2) Ags administered orally or directly into the portal vein can induce tolerance, 3) metastatic cancer frequently involves the liver, and 4) the liver may become the target of autoimmune disease. Because of their central role in Ag processing and presentation, it is likely that dendritic cells (DC) 3 are major determinants of immunity within the liver (1, 2). A complete understanding of the phenotype and function of liver DC is therefore desirable. Due to the relative scarcity of hepatic DC in normal mice, most of our knowledge has been extrapolated from studies of liver-derived DC progenitors generated by in vitro culture of hepatic nonparenchymal cells (NPC) in GM- CSF (3). To circumvent the difficulty in procuring adequate numbers of freshly isolated DC for functional analysis, we and others have used cytokine treatment to recruit liver DC in vivo. Overexpression of GM-CSF leads to the expansion of a predominantly myeloid (CD8 CD11b ) DC subtype as well as a population of myeloid DC progenitors in the liver (4), whereas Flt3 ligand treatment leads to the nearly equal expansion of both lymphoid (CD8 CD11b ) and myeloid DC subtypes (5, 6). These studies provided important clues to the in vivo signals that drive DC development and differentiation. However, it remains difficult to apply data from these artificially expanded DC to the function and phenotype of normal liver DC under steady state conditions. In contrast to murine splenic DC, which have been well characterized (7), considerable controversy exists with regard to the phenotype and function of normal murine liver DC. Early studies of the murine liver using in situ immunohistochemical techniques to identify DC based on their expression of MHC class II molecules showed that these cells did not display measurable levels of the T cell-costimulatory molecules CD40, CD80, and CD86, implying that they would have low immunostimulatory capacity (8). The initial isolation of liver NPC provided similar phenotypic data but also showed that these cells are indeed capable of triggering allogeneic T cell proliferation in vitro (9). Additionally, we have previously shown that freshly isolated CD11c liver DC are potent T cell stimulators (4). Lian et al. recently defined four subtypes of murine liver DC based on the relative expression of B220, CD11b, and CD4 but notably failed to identify CD8 DC, which are thought to play a crucial role in the physiologic maintenance of peripheral tolerance (10 12). To resolve the conflicting data regarding hepatic DC phenotype and function, we studied freshly isolated hepatic CD11c DC from normal untreated mice. To have a reference point from which to interpret the findings, we compared liver DC with spleen DC. We show that liver DC are immature relative to spleen DC and are heterogeneous with respect to maturational state as well as CD8 and CD11b expression. We also demonstrate that liver DC are less immunostimulatory in vitro, in part due to the predominance of two DC subtypes not found in the spleen. Hepatobiliary Service, Memorial Sloan-Kettering Cancer Center, New York, NY Received for publication June 12, Accepted for publication November 6, The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. 1 This work was supported in part by Grants CA (to V.P.) and CA (to R.D.). 2 Address correspondence and reprint requests to Dr. Ronald P. DeMatteo, Memorial Sloan-Kettering Cancer Center, Box 203, 1275 York Avenue, New York, NY address: dematter@mskcc.org 3 Abbreviations used in this paper: DC, dendritic cell; NPC, nonparenchymal cell; pdc, plasmacytoid DC. Materials and Methods Mice Adult 6- to 10-wk-old male C57BL/6 (H-2K b ) and BALB/c (H-2K d ) mice were purchased from Taconic Farms (Germantown, NY) and maintained in a pathogen-free animal housing facility at Memorial Sloan-Kettering Cancer Center. All procedures were approved by the Institutional Animal Care and Use Committee. Liver and spleen DC isolation Liver and spleen DC were isolated as previously described, with minor modifications (4). Animals were euthanized by CO 2 inhalation before laparotomy. In all experiments, the portal vein was cannulated in situ with a 24-gauge catheter (BD Biosciences, Sandy, UT) and perfused with 2 ml of Copyright 2004 by The American Association of Immunologists, Inc /04/$02.00

3 1010 LIVER DC SUBTYPES 1% (w/v) collagenase D (Sigma-Aldrich, St. Louis, MO) in PBS. To maximize the yield of DC for flow cytometry experiments, we then cut the liver into small pieces before incubation in 20 ml of collagenase solution for 40 min at 37 C. For experiments testing liver DC function, we omitted the in vitro collagenase incubation step because we previously found it to impair DC viability (4). Performance of the additional collagenase incubation in vitro did not affect the distribution of DC subtypes released or their expression of surface markers, merely the magnitude of the yield (not shown). We then mechanically disrupted the perfused liver using the flat portion of a plunger from a 10-ml syringe. The cell suspension was then passed through a sterile 100- m pore size nylon cell strainer (Falcon; BD Biosciences, Franklin Lakes, NJ) and spun three times at 30 g for 4 min to remove hepatocytes. The cells were pelleted (300 g for 10 min), and the hepatic NPC were resuspended in 1.5 ml of medium and added to 2.8 ml of 30% (w/v) metrizamide (Sigma-Aldrich) in PBS. This suspension was layered under 5 ml of PBS in a 15-ml Falcon centrifuge tube and spun at 1500 g for 20 min. The layer of low density cells at the interface, containing NPC, was harvested. The cells were fractionated based on CD11c expression with immunomagnetic beads (Miltenyi Biotech, Auburn, CA) after blocking Fc III/II receptors with mab 2.4G2 (Fc block; mab Core Facility, Sloan-Kettering Institute, NY). Typically, 95% of positively selected cells were CD11c DC after two rounds of positive selection with LS columns (Miltenyi Biotec). For assays in which individual liver and spleen DC subtypes were isolated for functional studies, metrizamide gradient-enriched cell suspensions were positively selected for CD45 expression with immunomagnetic beads (Miltenyi Biotec). This functioned to eliminate CD45 liver sinusoidal endothelial cells that are highly autofluorescent. Cells were then stained with fluorescently conjugated Abs to CD8, CD11b, and CD11c for subsequent sorting using a MoFlo cell sorter (DakoCytomation, Fort Collins, CO). Similarly, liver plasmacytoid DC were isolated by staining with Abs to B220 and CD11c and sorting for the B220 CD11c int cells using the MoFlo cell sorter (95 98% purity). Typical experiments starting with 10 livers yielded only total liver DC, which contained cells of each DC subtype. Spleen DC were isolated simultaneously from the same donors. Spleens were pooled, mechanically disrupted, pelleted, and run on a metrizamide gradient; and low density cells were harvested and sorted as described above. DC were cultured in RPMI 1640 with 10% heat-inactivated FBS, 2 mm L-glutamine, 100 U/ml penicillin, 100 g/ml streptomycin, and 0.05 mm 2-ME (complete medium). Flow cytometry Flow cytometry was performed on a FACSCalibur flow cytometer (BD Biosciences) after blocking cells per tube with 0.1 g of Fc block and then staining with FITC, PE, allophycocyanin, or biotin-conjugated Ab (BD PharMingen, San Diego, CA, except as indicated). Biotinylated Abs were secondarily stained with streptavidin-percp. Cells were stained for DC (CD11c (HL3), DEC205 (NLDC145; Cedarlane Laboratories, Toronto, Canada); lineage markers (CD8 (53-6.7) and CD11b (M1/70)); B cells (B220 (RA3-6B2)); granulocytes (Gr-1 (RB6-8C5)); macrophages (Mac-3 (M3/84)); MHC class II (I-Ab); intercellular adhesion marker (CD54 (3E2)); and costimulatory molecules (CD40 (3/23), CD80 (B7-1 (1G10)), CD86 (B7-2 (GL1)). Analysis of flow cytometry data was performed using FlowJo (Tree Star, Ashland, OR) software. Cytokine measurement and Ag uptake assays To assess cytokine production, we cultured immunomagnetic bead-purified liver or spleen DC in 100 l of complete medium with or without the addition of GM-CSF (20 ng/ml from J558 supernatant; gift of Ralph Steinman, Rockefeller University, New York, NY) on a 96-well flat-bottom Falcon tissue culture plate. LPS (1 g/ml; Sigma-Aldrich) or CpG ODN 1826 (CpG; 10 g/ml; Oligos, Etc., Wilsonville, OR) was added to some wells. After 36 h, cell culture supernatants were harvested and tested by cytometric bead array for IL-2, IL-4, IL-5, IL-6, IL-10, IL-12 (p70), IFN-, and TNF- (BD PharMingen) according to the manufacturer s protocols. Briefly, cell culture supernatant was mixed with equal amounts of cytokine capture beads and PE detection reagent for a 2-h incubation period, before washing and flow cytometric measurement of mean fluorescence. Production of IFN- by plasmacytoid DC was assessed by culturing FACS-sorted cells in complete medium with GM-CSF for 48 h. CpG was added to some wells at the beginning of the culture period. Supernatant IFN- content was measured by sandwich ELISA (Antigenix America, Huntington Station, NY) according to the manufacturer s protocol. Data presented are representative of three repetitions of this experiment with similar results. To measure in vitro Ag uptake, CD11c liver or spleen DC were incubated with 100 g of FITC-dextran or FITC-OVA in duplicate wells for varying periods at 37 C. At the end of each time period, the cells were washed with ice-cold PBS and stained with subtype DC markers before analysis by flow cytometry. T cell assays For MLR, DC were added in various numbers to allogeneic T lymphocytes (purified using Thy-1.2 (CD90.2) immunomagnetic microbeads; Miltenyi) in 96-well U-bottom plates and then pulsed with [ 3 H]thymidine (1 Ci/well) on day 3 for an additional 24 h. In vitro Agspecific T cell activation was assayed with an H-2K b -restricted CD8 T cell hybridoma specific for OVA (SIINFEKL peptide (OVA)) (13). DC were plated at various concentrations with OVA-restricted T cells and OVA (1 g/ml) in a 96-well plate for 3 days. T cell activation was determined by measuring supernatant IL-2 concentration with ELISA. Results Liver DC are less mature than spleen DC Because liver DC from normal mice are not well described, we first isolated them and analyzed their phenotype by flow cytometry. To estimate the relative size of liver and spleen DC, we determined the mean forward scatter for each population. The forward scatter of liver DC was 428 whereas that of spleen DC was 495, demonstrating that liver DC are smaller on average (Fig. 1A). We also found that normal CD11c liver DC had a markedly different surface marker profile from spleen DC isolated from the same cohort of mice. Liver DC expressed considerably lower levels of MHC class II, as well as the T cell costimulatory molecules CD40, CD80, and CD86, than spleen DC (Fig. 1B). Liver DC were not only more immature overall than spleen DC, but they were also more heterogeneous in their expression of the maturation markers than spleen DC, which had relatively uniform expression. Nevertheless, microscopic examination of Giemsa-stained liver and splenic DC demonstrated similar appearing cells with large, eccentric nuclei and some cytoplasmic vacuoles (not shown). However, more of the splenic DC had membranous ruffling which was consistent with their higher degree of maturation. Liver DC capture less Ag than spleen DC The differences in maturity between liver and spleen DC suggested that they would also have disparate function. We isolated hepatic and splenic CD11c cells and tested their ability to capture the protein OVA and the carbohydrate dextran in vitro, both of which are taken up primarily via macropinocytosis. Although splenic DC were more mature, we found that they captured more dextran and OVA protein as evidenced by higher mean fluorescence at all time points after incubation (Fig. 2). However, this observed difference in the rate and magnitude of uptake may not be physiologically relevant because nearly 100% of both liver and spleen DC showed evidence of having captured either Ag after 5 min of incubation (not shown). Liver DC have less in vitro T cell-stimulatory ability than spleen DC The hallmark of DC function is their unparalleled capacity to activate T cells. Therefore, we compared the differential stimulation of allogeneic T cells by liver and splenic DC in vitro. Not surprisingly, given their lower baseline maturity level, freshly isolated liver DC consistently induced approximately one-third as much allogeneic T cell proliferation as did splenic DC (Fig. 3A). Additionally, when 10 4 DC were mixed with 10 5 allogeneic T cells in MLR cultures, the supernatant of splenic DC contained more IFN- and IL-2, consistent with greater Th1 activation (Fig. 3B). Having established the lower allogeneic T cell stimulatory capacity of liver DC, we wanted to examine their Ag-specific T cell stimulation. We did this by culturing liver or spleen DC with an OVA-specific H-2K b -restricted T cell hybridoma in vitro in the presence of Ag. We measured the IL-2 content of supernatant from

4 The Journal of Immunology 1011 FIGURE 1. Freshly isolated murine liver DC are smaller and less mature than spleen DC. A, Forward scatter vs side scatter plots of CD11c liver and spleen DC demonstrate that bulk liver DC are smaller than spleen DC. B, Surface expression of maturation markers by bulk CD11c liver and spleen DC. Shaded areas represent surface expression of the indicated markers. Areas within solid lines show background isotype staining. Data are representative of eight separate experiments with similar results. this culture to assess T cell activation and found a pattern of activation similar to that of the MLR (Fig. 3C). Specifically, liver DC generated about one-third as much Ag-specific T cell stimulation as splenic DC. Liver and spleen DC have different cytokine profiles The cytokines secreted by DC are believed to have important autocrine and paracrine effects both in vivo and in vitro. We therefore wanted to determine whether the differences in T cell stimulation between liver and spleen DC were due to differences in cytokine secretion. We tested IFN-, TNF-, IL-2, IL-4, IL-5, IL-6, IL-10, and IL-12 levels in DC cultured for 36 h in RPMI 1640 with GM-CSF (Fig. 4). The exclusion of GM-CSF from the culture medium proportionately decreased the magnitude of each cytokine without altering the differential pattern of expression (not shown). Unstimulated liver DC made 10 times as much IL-6 and TNF- as did splenic DC. Both DC made minimal to no IL-10, IL-12, or IFN-. To simulate cytokine production under physiological conditions of bacterial infection, we added the Toll-like receptor ligands LPS or CpG to the DC cultures. Overall, the effects of LPS treatment were modest. The primary differences were that liver DC production of IL-6 was doubled and spleen DC secreted somewhat more IL-6, IL-10, and TNF-. In contrast, the effects of CpG treatment on cytokine production by both liver and spleen DC were profound. Both had massive increases in their IL-6, IFN-, and TNF- production. Interestingly, liver DC up-regulated their production of IL-12 nearly 100-fold, whereas splenic DC similarly increased secretion of IL-10. These data indicate that despite their lower basal ability to activate T cells, liver DC are capable of providing a potent activating signal upon stimulation. Supernatant IL-2, IL-4, or IL-5 was not detected in any of the groups tested. FIGURE 2. Liver DC are less efficient at in vitro Ag uptake than spleen DC. To compare the ability of liver DC to capture Ag via macropinocytosis, we incubated bead-purified CD11c cells from either organ at 37 C with FITCconjugated dextran or OVA for various periods before flow cytometric analysis. Bars, SDs of duplicates within the same assay. Results are representative of four separate experiments with similar results.

5 1012 LIVER DC SUBTYPES FIGURE 3. Liver DC achieve less T cell stimulation. Liver and spleen DC were isolated using immunomagnetic beads. A and B, To assess their allostimulatory capacity in vitro, various numbers of DC were cultured with BALB/c T cells for 3 days before the removal of a small amount of supernatant from the top wells for cytokine measurement and the addition of [ 3 H]thymidine for a final 24-h period. C, The Ag-specific activating ability of liver and spleen DC was tested by culturing the DC with OVA-specific T cell hybridoma cells for 3 days and measuring IL-2 production. Bars, SDs of triplicate wells from the same assay. Data are consistent among two to five repetitions of each assay., p 0.05 by a one-sided t test of combined data from two separate experiments. Liver DC are comprised of multiple subtypes with heterogeneous maturation The heterogeneity of phenotypic maturity, the reduced Ag uptake and T cell stimulation, and the differences in cytokine production suggested that there might be further distinctions between the subtypes of liver and spleen DC. We attempted to delineate DC subsets by analyzing the relative surface expression of the lymphoid marker CD8 and the myeloid molecule CD11b, which are the most wellestablished markers to define murine DC subtypes in the spleen and other lymphoid organs (14). The liver contained four distinct CD11c subtypes that were CD8 CD11b, CD8 low/ CD11b low/, CD8 CD11b, and CD8 CD11b (Fig. 5). Consistent with published results (14), we found two predominant populations of CD11c cells in the spleen: CD8 CD11b low/ and CD8 CD11b. In accord with our observation that bulk liver DC are immature in comparison with spleen DC (Fig. 1B), we found that, to varying degrees, all liver DC subtypes were less mature than either spleen DC subtype. The CD8 CD11b cell populations from both the liver and spleen were phenotypically quite similar in that they both had low expression of the endocytic receptor DEC205, but relatively high staining for maturation markers (Fig. 5). Additionally, the CD8 low/ CD11b low/ group of liver DC was similar to CD8 CD11b low/ spleen DC in that both populations had high DEC205 and moderate to high maturation marker staining even though they differed in the magnitude of CD8 expression. In the spleen, DEC205 is consistently coexpressed with CD8, whereas DEC205 is found only in a portion of CD8 liver DC, which is a pattern similar to that shown previously for lymph node DC (14, 15). The CD8 CD11b cells in the liver were mostly DEC205 and were somewhat less mature. The population of liver CD11c DC that has no measurable expression of CD8 or CD11b was also particularly immature, with low to moderate MHC class II and CD86 expression and no detectable CD40 or CD80. Immature plasmacytoid DC (pdc) have recently been identified in the spleen, lymph node and bone marrow and have been shown to have the capacity either to induce tolerance in the steady state or to produce IFN- upon CpG or viral stimulation (16). Therefore, we wanted to know whether these cells were also present in the liver. We found that a proportion of the CD8 CD11b, CD8 low/ CD11b low/, and CD8 CD11b liver DC, but not the CD8 CD11b liver DC, stained for the pdc marker B220 (Fig. 5). Although 19% of the cells were B220 CD11c intermediate pdc, a relatively small (5%) percentage of spleen DC had a similar plasmacytoid phenotype (Fig. 6). Liver pdc had higher expression of the granulocyte marker Ly-6G and were even more immature than spleen pdc in terms of MHC class II, CD40, CD80, and CD86 expression. Additionally, pdc from both organs differed in their expression of CD8 and CD11b. Liver pdc had very low CD8 expression and no detectable CD11b, in contrast with the moderate expression of both markers by splenic pdc. Additionally, as has been found in other organs, supernatant of pdc cultured in the presence of CpG and GM-CSF contained 1.4 ng/ ml/10 6 cells of IFN- after 48 h, whereas IFN- was undetectable in supernatant from the remaining CpG-stimulated liver DC subtypes or from liver pdc cultured in the absence of the CpG motif. Our finding that there are subtypes of liver DC that differ with respect to maturation suggested that there might be differences among the respective efficiencies of liver DC subtypes in capturing fluorescently conjugated Ags (Fig. 7). Indeed, there were relative differences among the various liver and spleen DC subtypes. CD8 CD11b liver DC had the highest uptake of dextran of all of the cells even though, as a whole, liver DC were less efficient (Fig. 2). All of the liver DC subtypes were less capable than both splenic DC subtypes at taking up OVA, but CD8 CD11b cells remained the most efficient liver DC (Fig. 7). Similarly, CD8 CD11b low/ spleen DC were better at taking up both Ags than were CD8 CD11b spleen DC.

6 The Journal of Immunology 1013 FIGURE 4. Liver and spleen DC in vitro cytokine secretion profile. Liver and spleen DC were isolated using immunomagnetic beads and cultured at a concentration of cells/ml in RPMI 1640 supplemented with GM-CSF (20 ng/ml) alone ( ), or GM-CSF plus CpG (10 g/ml; o), or LPS (1 g/ml; f). After 36 h, supernatant from each group was tested for the presence of IL-2, IL-4, IL-5, IL-6, IL-10, IL-12 (p70), IFN-, and TNF- by cytometric bead array. Supernatant IL-2, IL-4, or IL-5 levels were undetectable. Bars, SDs of duplicates within the same assay. Results for cytokines with detectable levels are shown. Similar results were obtained from three separate experiments. The T cell-immunostimulatory capacity of CD8 CD11b and CD8 CD11b liver DC is similar to that of spleen DC Once we had established the presence of additional subtypes of DC with distinct phenotypes and maturational states in the liver, as well as with distinct abilities to capture Ags, we wanted to determine whether they differed in their stimulation of T cells. Therefore, we performed flow cytometric cell sorting to separate out the four liver and two spleen DC subtypes described above and repeated the allogeneic and Ag-specific T cell-stimulatory assays with each group. Interestingly, we found that both CD8 CD11b and CD8 CD11b liver DC were as capable of triggering in vitro allogeneic T cell proliferation as were the same subtypes of spleen DC (Fig. 8A). In contrast, liver CD8 low/ CD11b low/ DC were less efficient at triggering MLR proliferation, whereas CD8 CD11b liver DC produced only minimal response. Th1 cytokine secretion followed a pattern similar to that of T cell proliferation in this assay (Fig. 8B). We found similar results in the Ag-specific T cell assay (Fig. 8C). Myeloid DC from both the liver and spleen had equally high immunostimulatory ability, whereas spleen CD8 CD11b low/ and liver CD8 CD11b and CD8 low/ CD11b low/ DC were somewhat less efficient in this regard. In this assay, CD8 CD11b liver DC were again only weakly immunostimulatory. Discussion DC are likely to play a critical role in defining the liver as a major regulator of peripheral tolerance. Defining the biology of liver DC may facilitate their therapeutic exploitation to activate or suppress immunity. Our current understanding of liver DC has been limited by their scarcity and difficulty of isolation. Initially, investigators used MHC class II staining detected by immunohistochemistry as a surrogate marker of liver DC (8). Using MHC class II to identify DC is problematic, given that MHC class II is also expressed by other more abundant cells such as liver sinusoidal endothelial cells. We used CD11c to identify DC, because it is currently the most widely used marker of murine DC. However, other cell types may express CD11c; therefore, its use is a limitation of this as well as

7 1014 LIVER DC SUBTYPES FIGURE 5. CD11c liver DC are composed of four subtypes based on CD8 and CD11b expression, whereas there are only two subtypes in the spleen. CD45 liver and spleen leukocytes were isolated using immunomagnetic beads before staining for four-color FACS. The combined CD8 and CD11b expression of CD11c cells was used to delineate the various subtypes with distinct profiles based on DEC205, maturation marker, and B220 expression. Areas within solid lines represent matched isotype staining. Data are representative of four separate experiments. other investigations. In the early studies, MHC class II-positive cells in the liver were found to have minimal expression of costimulatory molecules; however, we have recently reported that, using anti-cd11c immunohistochemistry after blocking nonspecific staining due to Fc receptors, there is an exceedingly small number of DC present in normal murine liver (4, 8). In the current FIGURE 6. Liver DC are composed of a large proportion of B220 pdc with low CD8 expression. Liver and spleen DC were prepared as in Fig. 5, and stained cells expressing high levels of B220 and intermediate CD11c were assessed for their expression of the shown surface molecules. Isotypes are shown as open histograms. Similar data were found from four repetitions.

8 The Journal of Immunology 1015 FIGURE 7. Liver DC subtypes have variable Ag uptake efficiencies. CD11c liver NPC were incubated with either FITC-conjugated dextran or OVA at 37 C for varying periods. Cells were then washed with ice-cold PBS and subsequently stained with anti-cd11b and anti-cd8 Abs to delineate DC subtypes. Bars, SDs of duplicates within a single experiment. Data represent two repetitions with consistent results. study, we show that liver DC actually express low to moderate levels of costimulatory molecules. Other indirect information regarding liver DC has been extrapolated from cultures of NPC grown in GM-CSF (3). These DC-like cells were immature, and their phenotype was skewed by the presence of GM-CSF, which we have shown to expand a uniformly myeloid population in vivo (4). Despite the belief that liver DC have little immunostimulatory capacity, we previously found that CD11c liver DC isolated from normal mice using a laborious process relying on density centrifugation as well as immunomagnetic bead sorting could generate significant T cell allostimulation (4). In the present study, we have compared in detail the phenotype and function of steady state FIGURE 8. Liver DC subtypes vary in their ability to activate T cells. CD45 immunomagnetic bead presorted liver and spleen leukocytes were FACS sorted into the six subtypes shown in Fig. 6. The allogeneic (A and B) and Ag-specific syngeneic T cell-activating (C) abilities of these cells were then compared. Bars, SDs of triplicate wells in the same assay. Data are representative of two separate experiments with similar results.

9 1016 LIVER DC SUBTYPES murine liver DC with that of spleen DC, which are the most well studied freshly isolated murine DC. Our phenotypic analysis of CD11c cells was based on the differential expression of CD8 and CD11b because of their established utility in defining lymphoid and myeloid DC subtypes in the spleen and lymph node (14, 15). We were able to delineate four distinct subtypes of liver DC which were, on the whole, less mature than spleen DC (Fig. 5). We found that the liver contained both myeloid (CD8 CD11b ) and lymphoid (CD8 CD11b ) DC populations similar to those in the spleen. In addition, there were CD8 low/ CD11b low/ cells that expressed DEC205 and resemble a subset of DEC205 DC found in murine lymph nodes (14, 15). There was also an abundance of immature plasmacytoid B220 DC (Fig. 6) like those that have previously been found in the thymus, spleen, lymph nodes, bone marrow, and liver (10, 16). Although the surface marker expression by liver pdc was homogeneous, it appeared that there may be subtypes of spleen pdc based on heterogeneous expression of Ly6-G as well as the maturation markers tested. It is widely accepted that myeloid DC function as immunostimulatory APCs. However, the in vivo function of other DC subtypes remains controversial especially with regard to their role in promoting either tolerance or immunity (11). Importantly, CD8 DC have recently been shown to play a key role in the maintenance of peripheral tolerance by cross-presentation of tissue-associated Ags (12). Additionally, targeting of Ags to the DEC205 receptor in vivo has been found to lead to Ag-specific tolerance induction in the steady state, and this effect is abrogated when DC are matured by an anti-cd40 mab (17, 18). Therefore, our identification of relatively immature CD8 CD11b and CD8 low/ CD11b low/ in the liver is particularly germane and may in part explain intrahepatic tolerance. Our phenotypic data differ considerably from that of a recent report by Lian et al. (10) who showed several subtypes of myeloid and pdc, but did not identify lymphoid DC displaying CD8 or DEC205. There are two differences between our methods that may in part explain the discrepant results. First, our method of isolation was different in that we routinely injected the portal vein with collagenase before removal of the liver to maximize the number of DC released. The other group omitted this step to minimize the release of autofluorescent cells. We eliminated the autofluorescent population with immunomagnetic bead presorting for CD45 cells. Another disparity is that we analyzed the phenotype and function of all CD11c cells and did not remove the cells that were NK1.1. We did not discard the CD11c NK1.1 cells in this study for several reasons. First, a developmental link between human NK cells and DC has previously been shown (19). Additionally, all CD11c cells in this study rapidly took up both OVA and dextran, consistent with DC function (Fig. 2). Furthermore, NK1.1 cells make up 40% of the liver CD11c population but represent only 10% of the total NK1.1 population. The majority of CD11c NK1.1 cells in the liver have surface expression of MHC class II and CD86 and comprise a proportion of the CD8 CD11b, CD8 CD11b, and the CD8 CD11b liver DC subtypes described in this study. Moreover, they are also found in the spleen, thymus, bone marrow, and mesenteric lymph nodes (not shown). Finally, the promiscuity of NK cell marker expression on liver DC is not limited to NK1.1, because DX5, which is also commonly used to identify NK cells, is found on 50% of CD11c NK1.1 liver DC (not shown). Further study of whether there is any significance of NK cell marker expression by liver DC is clearly indicated. To assess the level of liver DC function, we compared them to the more well-characterized spleen DC. Our finding that bulk CD11c liver DC were less mature and less efficient than spleen DC in activating T cells supports the commonly accepted notion that liver DC have generally weaker immunostimulatory function (Fig. 3) and thereby may be more tolerogenic. However, our phenotypic delineation of four liver DC subtypes raised the possibility that each may have distinct functions. Indeed, we found that CD8 CD11b and CD8 CD11b liver DC induced similar T cell stimulation as did the same subtypes of spleen DC (Fig. 8). However, these two subtypes account for only 20% of the total population of liver DC, whereas they make up the vast majority of spleen DC. Therefore, although classical myeloid and lymphoid liver DC may be quite immunostimulatory, their respective roles in activating immunity in vivo may be relatively minor because of their infrequency. In contrast, the other two more prevalent subtypes of liver DC, CD8 low/ CD11b low/ and CD8 CD11b DC, were poor T cell stimulators. This suggests that either these two populations may serve as precursors to the more immunostimulatory subtypes in the setting of immune activation or they may contribute to tolerance induction in the steady state. Although bulk liver DC were weakly immunostimulatory, we found them to be capable of producing inflammatory cytokines. Upon activation with CpG oligonucleotides, liver DC produced massive amounts of IL-12, consistent with a previous report (10). Surprisingly, however, the levels were 10 times higher than those of spleen DC, further supporting the dichotomy between their functions (Fig. 4). Previous studies have demonstrated that a combination of stimulants including GM-CSF, IFN-, CpG, and IL-4 is required to elicit optimal IL-12 secretion by spleen DC (20). We also found an inverse relationship between liver and spleen DC IL-10 production after CpG stimulation, because spleen DC produced far more of this cytokine. The absence of significant amounts of IL-10 in the liver DC cultures may have, in turn, facilitated the high production of IL-12 (21). Thus, the predominant subtypes of liver DC produce little T cell activation. In contrast, the small proportion of liver DC that are of the classical lymphoid or myeloid subtype induces as much allogeneic and Ag-specific T cell stimulation as do their splenic counterparts. The relative frequency of the various subtypes explains why liver DC overall are only weakly immunostimulatory under steady state conditions, although they are capable of producing inflammatory cytokines when stimulated. Our findings have important implications for the role of liver DC in peripheral tolerance and immunity. Acknowledgments We thank Jan Hendrikx, Vinod Sahi, and Patrick Anderson from the Memorial Sloan-Kettering Cancer Center Flow Cytometry Core Facility for their invaluable assistance. References 1. Thomson, A. W., P. J. O Connell, R. J. Steptoe, and L. 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