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Clinical significance of monocyte heterogeneity.
Publication date (Electronic):
2015
Journal:
Clinical and translational medicine
Publisher:
Springer Science and Business Media LLC
Keywords:
Atherosclerosis,
Autoimmune Disease,
CD14,
CD16,
Cardiovascular,
Human,
Ly6C,
Macrophage,
Monocyte,
Mouse
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Abstract
Monocytes are primitive hematopoietic cells that primarily arise from the bone marrow, circulate in the peripheral blood and give rise to differentiated macrophages. Over the past two decades, considerable attention to monocyte diversity and macrophage polarization has provided contextual clues into the role of myelomonocytic derivatives in human disease. Until recently, human monocytes were subdivided based on expression of the surface marker CD16. "Classical" monocytes express surface markers denoted as CD14(++)CD16(-) and account for greater than 70% of total monocyte count, while "non-classical" monocytes express the CD16 antigen with low CD14 expression (CD14(+)CD16(++)). However, recognition of an intermediate population identified as CD14(++)CD16(+) supports the new paradigm that monocytes are a true heterogeneous population and careful identification of specific subpopulations is necessary for understanding monocyte function in human disease. Comparative studies of monocytes in mice have yielded more dichotomous results based on expression of the Ly6C antigen. In this review, we will discuss the use of monocyte subpopulations as biomarkers of human disease and summarize correlative studies in mice that may yield significant insight into the contribution of each subset to disease pathogenesis.
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Most cited references82
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The three human monocyte subsets: implications for health and disease.
Kok Loon Wong, Wei Hseun Yeap, June Tai … (2012)
Human blood monocytes are heterogeneous and conventionally subdivided into two subsets based on CD16 expression. Recently, the official nomenclature subdivides monocytes into three subsets, the additional subset arising from the segregation of the CD16+ monocytes into two based on relative expression of CD14. Recent whole genome analysis reveal that specialized functions and phenotypes can be attributed to these newly defined monocyte subsets. In this review, we discuss these recent results, and also the description and utility of this new segregation in several disease conditions. We also discuss alternative markers for segregating the monocyte subsets, for example using Tie-2 and slan, which do not necessarily follow the official method of segregating monocyte subsets based on relative CD14 and CD16 expressions.
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Inflammation switches the differentiation program of Ly6Chi monocytes from antiinflammatory macrophages to inflammatory dendritic cells in the colon
Aymeric Rivollier, Jianping He, Abhisake Kole … (2012)
Immune responses in the intestine need to be tightly regulated, particularly in the colon where the ability to combat pathogenic microbes must be balanced with the need to prevent exuberant inflammatory responses to the large pool of diverse commensal microorganisms. How this occurs is not yet clear; however, there is increasing evidence that specialized gut-resident DCs and macrophages (MPs), in addition to providing innate defense against both pathogenic and commensal microorganisms, are directly involved in driving regulatory responses. These include the differentiation, expansion, and maintenance of regulatory T cell (T reg cell) populations and the induction of IgA against commensal bacteria (Manicassamy and Pulendran, 2009). However, there is less consensus regarding the definition and functions of any one of several specialized MP and DC populations, particularly because the inconsistent use of an ever increasing number of surface markers has resulted in new levels of complexity. We and others originally defined DCs in the intestine based on the traditional lymphoid organ DC markers CD11c, CD8α, and CD11b (Iwasaki and Kelsall, 1999, 2001; Chirdo et al., 2005). More recently, a new organizational scheme of intestinal DC and MP populations has been proposed based on their expression of the nonoverlapping markers CD103 (αE integrin) and CX3CR1 (fractalkine receptor; Schulz et al., 2009). CD103+ DCs were identified in the small intestine (SI) lamina propria (LP), mesenteric LNs (MLNs), and Peyer’s patches of mice (Johansson-Lindbom et al., 2005) and recently in human colonic tissues (Jaensson et al., 2008). CD103+ DCs have a high capacity to produce the vitamin A metabolite retinoic acid and TGF-β (Coombes et al., 2007), which gives them enhanced ability to drive the expression of the gut-homing molecules α4β7 and CCR9 on naive T cells during priming (Iwata et al., 2004; Jaensson et al., 2008). They also have been shown to induce the de novo differentiation of foxp3+ T reg cells (Coombes et al., 2007; Sun et al., 2007) and contribute to IgA+ B cell differentiation from naive precursors in vitro (Mora et al., 2006). In the SI LP, CD103+ DCs are predominantly CD11b+CD8α− (Coombes et al., 2007; Denning et al., 2007b; Sun et al., 2007), appear to be the main, if not the only, DC population that constitutively migrates to the MLNs via a CCR7-dependent mechanism, and are critical for the induction of oral tolerance, as well as for the suppression of colitis development by T reg cells (Annacker et al., 2005; Worbs et al., 2006; Bogunovic et al., 2009; Schulz et al., 2009). CD103+ DCs do not express CX3CR1 and, whether they are CD11b+ or CD11b−, expand in response to Flt3-L and GM-CSF and are generated by adoptively transferred pre-DCs (Bogunovic et al., 2009; Varol et al., 2009). In contrast, CX3CR1+ cells do not express CD103 and were initially described as DCs in the terminal ileum, extending dendrites between epithelial cells (intestinal epithelial cells [IECs]) to actively take up bacteria and soluble antigens from the intestinal lumen. These cells were thus proposed to play a key role in the capture and transport of intestinal antigens to the MLNs (Rescigno et al., 2001; Niess et al., 2005; Vallon-Eberhard et al., 2006; Hapfelmeier et al., 2008; Bogunovic et al., 2009; Schulz et al., 2009). However, the identity of the CD11c+CX3CR1+ cells as bona fide DCs has been challenged by studies in the SI LP that illustrated their MP-like vacuolar system (Bogunovic et al., 2009) and their poor ability to drive naive T cell stimulation and differentiation (Bogunovic et al., 2009; Schulz et al., 2009), as well as their inability to migrate to draining MLNs (Schulz et al., 2009). In addition, CX3CR1+ cells have a different ontogeny in the intestine, relative to CD103+ DCs, and appear to be derived from Gr-1hi monocytes (MOs) in an M-CSF–dependent manner (Bogunovic et al., 2009; Varol et al., 2009). Both pro- and antiinflammatory properties have been documented for CX3CR1+ cells in the LP. CX3CR1+ cells seem to play a critical role in the oral tolerance induction process, by expanding foxp3+ T reg cells in the SI LP, subsequent to their priming in the MLNs (Hadis et al., 2011) but also support inflammatory immune responses under normal conditions and in both innate and CD45RBhi CD4+ T cell transfer colitis models (Varol et al., 2009; Niess and Adler, 2010). Therefore, whether LP CX3CR1+ cells are MPs or DCs and whether these cells play pro- or antiinflammatory functions require further clarification. One key point to consider when addressing this question is the fact that LP CX3CR1+ cells represent a heterogeneous group of cells expressing low to high levels of CX3CR1 (Schulz et al., 2009; Varol et al., 2009). Independently of CX3CR1 expression, several intestinal MP populations have also been characterized. In the human, two CD68+ MP populations coexist in the intestine of patients with inflammatory bowel disease: a steady-state CD14− inflammatory anergic population that neither expresses innate response receptors (Rogler et al., 1998; Smith et al., 2001; Smythies et al., 2005) nor produces proinflammatory cytokines in response to an array of inflammatory stimuli but retains avid phagocytic and bactericidal activity (Smythies et al., 2005) and an additional CD14+ proinflammatory population that accumulates in patients with Crohn’s disease and contributes to the pathogenesis of this disease (Rugtveit et al., 1997; Kamada et al., 2008). In the mouse SI, CD11c−F4/80+CD11b+ MPs express IL-10, induce the differentiation of foxp3+ T reg cells in the presence of exogenous TGF-β, and suppress the differentiation of Th17 by CD11c+CD11b+ DCs in vitro (Denning et al., 2007b). In the mouse colon, MPs, defined as CD11c−/loF4/80+CD11b+ cells, seem to represent a heterogeneous pool of cells, and discrepancies in the literature regarding their pro- or antiinflammatory properties in the steady-state and during colitis remain to be resolved (Platt et al., 2010; Takada et al., 2010). Therefore, the identification and characterization of the colonic LP (cLP) MP pool is only beginning to be unveiled in the mouse, and the pro- or antiinflammatory nature of these cells is still unclear. To clarify the discrepancies in the definition, function, and origin of mucosal DCs and MPs, we undertook experiments in mice to extensively phenotypically and functionally characterize DC and MP populations in the colon during steady-state conditions and during inflammation using the CD45RBhi T cell transfer model of colitis. Using a variety of criteria, including extensive phenotypic analyses and gene expression arrays, our data indicate that F4/80hiCX3CR1hi cells are MPs that constitute the majority of the steady-state cLP mononuclear phagocytes (MNPs). These MPs can be divided into CD11c+ and CD11c− populations, both of which are poor at presenting antigens to T cells, and produce large amounts of IL-10 and other antiinflammatory cytokines. In contrast, DCs express low to intermediate levels of F4/80 and CX3CR1, are CD11c+, and comprise at least three separate populations: CD103+CX3CR1−CD11b− DCs, CD103+CX3CR1−CD11b+ DCs, but also CD103−CX3CR1intCD11b+ DCs. CD103+CX3CR1−CD11b− DCs are unique in their ability to efficiently cross-present antigens to CD8+ T cells, whereas both CD103+CX3CR1−CD11b+ DCs and CD103−CX3CR1intCD11b+ DCs are more efficient at presenting antigens to CD4+ T cells. More importantly, we identified Ly6Chi MOs as precursors for the F4/80hiCX3CR1hiCD11c+ MP population exclusively. IL-10 constitutively produced by these cells and by CD11c− MPs actively suppresses IL-12p35 and IL-23p19 production from cLP MNPs in noninflammatory conditions in vivo. In contrast, during colitis MOs differentiate into the CD103−CX3CR1intCD11b+ DC population, which massively infiltrates the cLP and plays proinflammatory functions by producing IL-12, IL-23, iNOS, and TNF and driving the differentiation of IFN-γ–producing T cells. These experiments are the first to definitively identify CX3CR1hiCD11c+ as MPs and not DCs and to demonstrate the dual capacity of Ly6Chi MOs to differentiate into regulatory or inflammatory populations depending on the local environmental conditions. RESULTS The steady-state cLP contains two major subsets of MPs To characterize the phenotype of cLP MP and DC populations, we used a combination of CD45, MHC-II, lineage markers (lin: CD3, CD19, TCR-β, TCR-γδ), CD11c, and F4/80. After pregating on CD45+ cells to exclude any contaminating IECs, total cLP MNPs, gated as MHC-IIhilin− cells, could be subgrouped into four populations based on CD11c and F4/80 expression (Fig. 1, A and C). Two classical CD11c−F4/80hi MP and CD11chiF4/80− DC populations represented 39.02 ± 6.19% and 3.51 ± 1.24% of total MNPs, respectively. Interestingly, the cLP also contained a large population of cells coexpressing high levels of CD11c and F4/80 (30.83 ± 5.71%). Finally, a population of CD11c+ cells with intermediate levels of F4/80 expression (10.01 ± 2.59%) could also be identified (Fig. 1, A and C). For better clarity, the four described cLP MP/DC populations will be hereafter referred to as subsets 1, 2, 3, and 4 as illustrated in Fig. 1 A. Electron microscopy revealed that both subsets 1 and 2 exhibited classical morphological and ultrastructural characteristics of MPs such as numerous phagocytic vacuoles, eccentric nuclei, and primary and secondary lysosomal granules (Fig. 1 D). In contrast, subset 4 was of smaller size and had more typical DC morphology, with less cytoplasm, cell surface dendritic projections, and lack of large intracytoplasmic vesicles. Finally, subset 3 exhibited characteristics more similar to DCs than MPs, as is shown in Fig. 1 D. Further analysis of CX3CR1, CD103, and CD11b expression on these four subsets revealed that MP subsets 1 and 2 are homogeneously CX3CR1hiCD11b+CD103−, whereas DC subset 4 is homogeneously CX3CR1−CD11b−CD103+. In contrast, subset 3 expresses low to intermediate levels of CX3CR1 and contains at least two separate populations of CX3CR1−CD11b+CD103+ (CD103+ subset 3) and CX3CR1intCD11b+CD103− (CD103− subset 3) cells (Fig. 1 B). A more extensive flow cytometry phenotypic analysis revealed that subsets 1 and 2 both expressed the classical MP marker CD14 but also differed in their expression of Mac-3, as well as the transferrin receptor CD71, the M-CSF receptor (CD115), and the T cell co-stimulatory molecule CD70 (Fig. 1 E; Atarashi et al., 2008). The DC subset 4 did not express the MP markers CD14, Mac-3, CD70, CD115, or CD71. Subset 3 exhibited a phenotype very similar to subset 4 but seemed to express some low levels of CD14. DNA microarray analysis further supported the MP identity of subsets 1 and 2, which expressed much higher levels of F4/80, CX3CR1, CD14, CD68, mannose receptor (Mrc1), MP scavenger receptor (Msr1), lysozyme (Lyz), Fc-γ receptors (Fcgr1, Fcgr2b, Fcgr3, and Fcgr4), complement receptors (C5ar1 and C3ar1), and cathepsins (Cts), compared with subsets 3 and 4 (Fig. 1 F). In addition however, using stringent statistical analysis, 21 genes were differentially expressed between subset 1 and 2, supporting the fact that these subsets are distinct (Fig. 1 G). For example, the scavenger receptor CD163 and the coagulation factor XIII A1 polypeptide (FXIIIa1) were 12.2 ± 2.96– and 10.89 ± 3.18–fold higher in subset 1 versus 2. Furthermore, in vivo BrdU incorporation experiments were also consistent with subsets 1 and 2 being MPs, as they both exhibited much slower turnover rates, compared with DC subsets 3 and 4 (Fig. S1). Figure 1. Definition and characterization of four DC/MP subsets in the steady-state colon. (A) Representative FACS analysis showing the gating strategy to define colonic MP and DC subsets. After pregating on CD45+MHC-IIhilin− cells, cLP cells were subdivided into four different populations (MP subsets 1 and 2 and DC subsets 3 and 4) based on their CD11c and F4/80 expression. (B) CX3CR1/CD103 and CD11b/CD103 expression profiles on the four MP and DC populations defined in A. Plots are representative of >20 experiments with 3–10 pooled mouse colons. (C) Quantification of the four cLP MP/DC populations (defined in A), expressed as percentages of total MHC-IIhilin− cells. Data are mean ± SEM of three independent experiments with at least three pooled mouse colons. (D) Electron micrographs of cLP subsets 1–4 FACS sorted from 10 pooled mouse colons. (E) Surface phenotype of subsets 1–4. Specific markers (black lines) and isotype controls (gray-filled areas) are shown. Plots are representative of at least three independent experiments with 3–10 pooled colons each. (F) Microarray analyses of the genes differentially expressed in subsets 1–4. (H) Microarray analyses of the genes differentially expressed in subset 1 versus 2. Each replicate represents data obtained for one subset FACS sorted from 10 pooled colons. We next investigated the functional properties of colonic MPs and DCs. First, their capacity to internalize fluorescent microbeads was assessed, as a measure of their phagocytic activity. A higher percentage of subsets 1 and 2 phagocytosed beads, and these cells were more likely to contain multiple beads compared with subsets 3 and 4 (Fig. 2, A and B). Colonic subsets 1 and 2 also exhibited very low T cell priming capacities, similar to splenic MPs, and lower than subsets 3 and 4 or splenic DCs in both allogeneic (Fig. 2 C) and antigen-specific T cell proliferation assays using OT-I and OT-II TCR transgenic T cells (Fig. 2, D and E). Of note, subset 4 was less efficient at OT-II T cell priming but more efficient at OT-I T cell priming, compared with subset 3 or splenic DCs, suggesting its superior ability to cross-present antigens. Surprisingly, no striking difference in the expression of MHC-II or of the co-stimulatory molecules CD80, CD86, PD-L1, or PD-L2 could be detected between the different subsets (Fig. S2 A). However, subsets 3 and 4 were uniquely able to strongly up-regulate CCR7 expression 24 h after in vitro stimulation with LPS, suggesting their potential selective ability to migrate to the MLNs to prime T cells (Fig. S2 B). Figure 2. Functional characterization and localization of cLP MP/DC subsets and T cell priming capacities of subsets 1–4. (A and B) Phagocytic capacities of subsets 1–4. cLP cells from C57BL/6 mice were incubated in the presence of fluorescent microbeads for 45 min and analyzed for their bead content. (A) Percentage of each subset containing at least one bead. (B) Percentage of each subset containing one bead, two beads, three beads, four beads, or more after pregating on total bead+ cells. Results are mean ± SD and are representative of two independent experiments with three mice each. (C) Allo-stimulatory capacities of C57BL/6 cLP subsets 1–4 and control splenic DCs and MPs co-cultured with CD4+ T cells from the spleen of BALB/c at the indicated ratios. (D and E) Proliferation of OT-II (D) and OT-I (E) cells after co-culture with graded doses of OVA protein–pulsed cLP subsets 1–4 and control splenic (SPL) DCs and MPs. Proliferation was determined by thymidine incorporation assay after 4 d (C and D) or 3 d (E) of culture. Results are expressed as mean ± SD of cpm triplicates and are representative of at least two independent experiments using pooled cells from 10 mice per experiment. Unpaired Student’s t tests were performed to compare subset 2 versus 3 (blue statistics) and subset 2 versus 4 (green). The comparison of subset 2 versus 1 never reached statistical significance. *, P 30% of cells of this subset had turned over (taken up BrdU) in untreated WT mice (Fig. S1). Consistent with their subset 2 surface phenotype, FACS-sorted cLP MO-derived (CD45.1+) cells expressed high levels of IL-10 and TNF but low levels of other proinflammatory cytokine mRNA (Fig. 5 G). Together, these data indicate that Ly6Chi MOs selectively give rise to the cLP MP subset 2 and not subset 1 in the noninflamed colon. MOs generate an inflammatory CD103− DC subset 3 after adoptive transfer into RAG−/− colitic mice We next used the adoptive transfer model of colitis (Powrie et al., 1993) to investigate the changes in cLP MP/DC populations during chronic intestinal inflammation and to determine which cLP populations were derived from MOs under these conditions. RAG−/− mice adoptively transferred with naive CD45RBhiCD4+ T cells developed severe intestinal inflammation within 7–8 wk after transfer, as indicated by the four- to fivefold increase in the total number of hematopoietic cells (CD45+) and MHC-IIhi MPs/DCs in the colon (Fig. 6, A and B), as well as by histological scoring (not depicted). In contrast, control RAG−/− mice that received a mixture of CD45RBhi+lo T cells (containing T reg cells) did not develop intestinal inflammation during the same time course (Fig. 6 B; Read et al., 2000). A selective increase in the percentage of subset 3 was observed in the inflamed cLP of CD45RBhi- versus CD45RBhi+lo-transferred mice, whereas the percentages of subsets 1, 2, and 4 were decreased (Fig. 6, C and D). In colitic mice, subset 3 outnumbered subset 2 by two- to threefold. Interestingly, this accumulating subset 3 from CD45RBhi colitic mice was almost exclusively CD103− compared with control CD45RBhi+lo mice (Fig. 6, D and E), suggesting that most cells in subset 3 either down-regulated CD103 expression during colitis or were derived from newly recruited precursors that differentiated into a CD103− subset 3 in the colitic environment. In support of the latter possibility, the CD103− subset 3 expressed E-cadherin, a marker previously shown to be expressed on inflammatory DCs recruited to the colon of colitic mice (Fig. S5 A; Siddiqui et al., 2010), as well as the monocytic lineage marker CX3CR1 (Fig. S5 B). Moreover, Ly6Chi MOs adoptively transferred into colitic mice rapidly migrated to the inflamed colon (Fig. 6 F) and largely developed into cells that were phenotypically similar to the endogenous CD103− subset 3 that massively infiltrated the cLP (Fig. 6 G). Additionally, a larger proportion of the MO-derived cells in the colitis environment corresponded to subset 3 and possibly 4, compared with what was observed in the adoptive transfer experiments in noninflamed mice (20–25% vs. 80% of which correspond to subset 2; Fig. 1 C) restores their IL-12 production to levels comparable with splenic CD11c+ DCs (Monteleone et al., 2008). Second, mice with a myeloid-specific deletion of the transcription factor STAT3, both downstream of the IL-10 receptor and upstream of IL-10 production, develop a chronic enterocolitis (Takeda et al., 1999). Furthermore, colitis development in IL-10−/− mice results from the absence of suppression of MyD88-dependent commensal-induced inflammation by IL-10 (Rakoff-Nahoum et al., 2006). We provide evidence that commensal bacteria contribute to the production of IL-10 by cLP subsets 1 and 2, but other mechanisms are also involved, and none require MyD88 signaling. The functions of CD11c+ and CD11c− MPs may be largely redundant, as each shares phenotypic and functional characteristics, including the constitutive production of IL-10 in the cLP, and largely similar baseline gene expression. However, our data also suggest that these two MP populations have unique origins and biological functions. Thus, although both MP subsets 1 and 2 have a slow turnover rate compared with LP DC populations, only the CD11c+ MP subset 2 is readily derived from MOs after cell depletion. No MO-derived subset 1 could be detected in our MO transfer experiments at day 8 or 15, a time point when the endogenous population of subset 1 was already fully reconstituted (not depicted). It is therefore unlikely that MOs are precursors of MP subset 1 or that subset 2 is an intermediate differentiation state between MOs and subset 1 under these noninflammatory conditions. Finally, in our hands, Ly6Chi MOs also appeared to give rise to at least some CD103+ cells (falling into subsets 3 and 4) during colitis (Fig. 6 F). Although this finding is inconsistent with current data indicating a pre-DC origin of CD103+ DCs, to our knowledge, the origin of CD103+ DCs has not yet been evaluated in the presence of ongoing inflammation, nor has CD103 expression on MO-derived CX3CR1+ cells. This finding is even more intriguing, considering that CD103+ DCs have recently been shown to lose their tolerogenic properties during colitis (Laffont et al., 2010). Interestingly in the lungs, Jakubzick et al. (2008) documented that Ly6Chi MOs preferentially repopulate CD103+ DCs, whereas Ly6Clo MOs repopulate CD11bhi DCs, further supporting the concept that the DC/MP differentiation potential of MOs may vary depending on the organ and environment to which these precursors home. These results are consistent with two recent articles demonstrating that MOs are precursors of MHC-II+CD11c+CD11b+CD103−CX3CR1+ cells in the SI LP and cLP in steady-state conditions, whereas CD103+ DCs were mostly derived from pre-DCs (Bogunovic et al., 2009; Varol et al., 2009). However, these studies did not analyze the origin of CD11c− cell populations and so did not address the differential origin of intestinal MP populations. In addition, our DNA microarray analyses further support the possibility that subsets 1 and 2 represent distinct MP populations. cLP subset 1 expresses higher levels of CD163, FXIIIa1, and Lyve1, which have been previously associated with MPs involved in wound healing or tissue repair (Philippidis et al., 2004; Schledzewski et al., 2006; Quatresooz et al., 2008). In comparison, subset 2 expresses higher levels of DC-specific transmembrane protein (DC-STAMP), which was recently shown to be involved in regulating DC functions and preventing autoimmunity development (Sawatani et al., 2008). Further work is necessary to understand the respective roles and origins of subsets 1 and 2. In summary, our data illustrate a dual role of MOs in gut MP/DCs homeostasis. MOs may contribute to the maintenance of colon homeostasis in the steady-state by generating the cLP CX3CR1hi MP subset 2, which produces regulatory cytokines, including IL-10, but may also contribute to colitis pathogenesis by generating a CX3CR1intCD103− DC subset 3 that exhibits proinflammatory properties in the inflamed colon. The identification of the colonic factors controlling the differentiation of MOs into pro- or antiinflammatory MNPs and determining the mechanisms regulating IL-10 production by subsets 1 and 2 will be important steps in understanding intestinal pathophysiology and could open promising therapeutic options for patients with inflammatory bowel diseases. MATERIALS AND METHODS Mice C57BL/6 and C57BL/6 SJL mice were obtained from the National Cancer Institute and used at 6–12 wk of age. C57BL/6 GF animals were bred at the National Institute of Allergy and Infectious Diseases colony from Taconic before transfer to the GF facility at the National Institutes of Health and screened weekly for viral, bacterial, and fungal contamination. Control SPF, C57BL/6 Rag2−/− , C57BL/6 Rag1−/− OT-I, and OT-II transgenic mice were also purchased from Taconic. IL-10–GFP transcriptional reporter mice (Vert-X mice) were obtained from C. Karp (Cincinnati Children’s Hospital Medical Center, Cincinnati, OH). CX3CR1GFP/GFP, B6.129P2-IL10tm1Cgn/J, and CD11cDTR mice were purchased from The Jackson Laboratory. CX3CR1+/GFP mice were obtained by crossing CX3CR1GFP/GFP with WT C57BL/6 mice. All mice were maintained at an American Association for the Accreditation of Laboratory Animal Care–accredited animal facility at the National Institute of Allergy and Infectious Diseases and housed in accordance with the procedures outlined in the Guide for the Care and Use of Laboratory Animals under an animal study proposal approved by the National Institute of Allergy and Infectious Diseases Animal Care and Use Committee. Experimental animal models T cell transfer colitis. Rag2−/− mice were injected i.p. with 3 × 105 CD4+CD45RBhi T cells (colitic CD45RBhi mice) or 1.5 × 105 CD4+CD45RBhi + 1.5 × 105 CD4+CD45RBlo T cells (noncolitic CD45RBhi+lo mice) isolated from the spleens of WT mice. Mice were housed with nonsterile bedding and given nonsterile food and nonacidified water. Mice from the colitic group developed significant intestinal inflammation 7–8 wk after transfer. CD11cDTR bone marrow chimeric mice and CD11c+ cell depletion. C57BL/6 mice were irradiated (900 rads) and transferred i.p. with 3 × 106 total bone marrow cells isolated from CD11cDTR mice. CD11cDTR → C57BL/6 bone marrow chimeric mice received drinking water containing antibiotics (0.13 mg/ml Trimethoprim and 0.67 mg/ml Sulfamethoxazole) for 6 wk. For CD11c+ cell depletion, CD11cDTR → C57BL/6 bone marrow chimeric mice were injected i.p. with DTx (8 ng/g body weight; Sigma-Aldrich) as indicated. Isolation and FACS analysis of colon and spleen phagocyte subsets. After removing extraintestinal fat tissue and blood vessels, colons were flushed of their luminal content with HBSS, opened longitudinally, cut into 2-cm pieces, and incubated with HBSS containing 0.015% DTT (15 min, 37°C in a shaking water bath). Epithelial cells and mucus were removed by extensive washes in HBSS, 5% FCS, and 25 mM Hepes before colons were cut into smaller 2-mm2 pieces and digested in complete Iscove’s media containing 167 µg/ml Liberase TL and 30 µg/ml DNase I (Roche) for 60 min at 37°C in a shaking water bath. The digested cell suspension was then passed through 100- and 40-µm cell strainers and resuspended in 1.077 g/cm3 iso-osmotic metrizamide medium (NycoPrep; Accurate Chemical & Scientific Corp.). After centrifugation at 1,600 g for 15 min at room temperature, the low-density fraction was collected. For FACS analysis or sorting of colon samples, cells were stained with Fc block/anti-CD16/32 (2.4G2) and antibodies to CD11c (HL-3), MHC-II (AF6-120.1), F4/80 mAb (BM8), CD11b (M1/70), and CD103 (M290), as well as antibodies to B cell and T cell lineages (referred hereafter as lin): CD19 (RA3-6B2), CD3 (145-2C11), TCR-β (H57-597), and TCR-γδ (eBioGL3), all from eBioscience. When FACS analysis was performed directly on the digested colonic cell suspension (without NycoPrep gradient preenrichment), cells were pregated on hematopoietic cells using α-CD45.2 (104) antibody. Live/dead cells were identified using the Live/Dead Fixable violet dead cell stain kit (Invitrogen), according to the manufacturer’s instructions. For the FACS-sorting of spleen MPs and DCs, lineage (CD3/CD19/TCR-β/TCR-γδ)+ cells were first excluded. MPs were then sorted as CD11c−F4/80+ cells, whereas DCs were sorted as CD11c+F4/80− to int cells. Other antibodies used in this paper include CD45.1 (A20), CD14 (Sa2-8), Mac3 (M3/84), CD70 (FR70), CD71 (C2), CD115 (AFS98), CCR6 (clone 140706; R&D Systems), CCR7 (4B12), CD25 (7D4), CD40 (3/23), CD69 (H1.2F3), CD80 (16-10A1), CD86 (GL-1), PD-L1 (MI-H5), PD-L2 (122), Ly6C (AL-21), c-kit (2B8), Flt3 (A2F10), E-cadherin (36), and CX3CR1 (rabbit polyclonal; Abcam), as well as corresponding isotype controls, all purchased from BD or eBioscience, unless otherwise specified. Cells were analyzed on an LSR II flow cytometer (BD) or sorted using a FACSVantage or FACSAria machine (BD). Isolation and adoptive transfer of bone marrow MOs. 3 × 106 Fms-like tyrosine kinase 3 ligand (FLT3-L)–expressing B16 cells were injected s.c. in C57BL/6 SJL donor mice 12–14 d before MO isolation to increase their total bone marrow MO numbers. Bone marrow was flushed from the tibias and femurs of donor mice. After red blood cell lysis using ACK lysis buffer, CD115+ cells were MACS enriched using an α-CD115 biotinylated antibody (eBioscience) followed by antibiotin microbeads (Miltenyi Biotech) according to the manufacturer’s instructions. CD115-enriched cells were then stained for Ly6C (AL21), CD11b (M1/70), streptavidin-PE, and a mixture of lin antibodies (CD3 [145-2C11], CD4 [RM4-5], CD8 [53-6.7], CD19 [RA3-6B2], NK1.1 [PK136], CD11c [N418], and c-kit [2B8]). MOs were sorted based on lin−CD115+Ly6ChiCD11bhi. MO purity after sorting was consistently >98%. Immunofluorescence staining of colonic tissue sections. Colon tissue was frozen in OCT medium (Sakura). 8-µm cryosections were fixed in acetone (30 s, −20°C) and stained with hamster anti–mouse CD11c (N418; eBioscience) and rat anti–mouse F4/80-biotin (MCA 497B; AbD Serotec) or rat anti–mouse CD11b-biotin (M1/70; eBioscience) antibodies. CD11c staining was detected using goat anti–hamster horseradish peroxidase (Jackson ImmunoResearch Laboratories, Inc.) followed by Alexa Fluor 594–tyramide (Invitrogen). After two 15-min quenching sessions in Peroxidase Blocking Reagent (Dako), F4/80 or CD11b staining was revealed by streptavidin horseradish peroxidase and Alexa Fluor 488–tyramide (Invitrogen). Slides were counterstained with Hoechst nuclear stain (Sigma-Aldrich), mounted with Fluoromount-G medium (SouthernBiotech), and visualized using a microscope (Axiovert 200M; Carl Zeiss) and IPLab software (BioVision Technologies). Transmission electron microscopy. cLP DC/MP subsets were FACS sorted from C57BL/6 mice and resuspended in 50 µl PBS. Cell pellets from centrifuged samples were fixed overnight at 4°C with 2.5% glutaraldehyde/4% paraformaldehyde in 0.1 M sodium cacodylate buffer, pH 7.2, and then postfixed 30 min with 0.5% osmium tetroxide/0.8% potassium ferricyanide, 1 h with 1% tannic acid and overnight with 1% uranyl acetate at 4°C. Samples were dehydrated with a graded ethanol series and embedded in Spurr’s resin. Thin sections were cut with an RMC MT-7000 ultramicrotome (Ventana) and stained with 1% uranyl acetate and Reynold’s lead citrate before viewing at 80 kV on a transmission electron microscope (H-7500; Hitachi). Digital images were acquired with a Hammamatsu XR-100 bottom mount charge-coupled device system (Advanced Microscopy Techniques). Bead phagocytosis assay. 3 × 106 total cLP cells were incubated for 45 min at 37°C in the presence of 75 × 106 Fluoresbrite Plain Yellow Gold 1-µm Microspheres (Polysciences Inc.) and then washed five times in cold PBS. Cells were then stained for I-Ab, CD11c, F4/80, CD11b, CD103 and CD19, CD3, TCR-β, and TCR-γδ, and the number of fluorescent microspheres in each cLP DC/MP subset was determined by flow cytometry. Co-culture experiments Thymidine incorporation assays. In mixed lymphocyte reactions, cLP and splenic DC/MP subsets were FACS sorted from C57BL/6 mice and co-cultured for 5 d with MACS-enriched CD4+ T cells from BALB/c mouse spleens. In antigen-specific assays, FACS-sorted cLP and splenic DC/MP subsets were incubated for 2 h at 37°C in the presence of 0.5 mg/ml endotoxin-free OVA protein (EndoGrade OVA; Hyglos) and washed twice in complete RPMI before co-culture with OT-I (3 d) or OT-II cells (5 d), MACS enriched from LNs and spleen of RAG1 OT-I or RAG1 OT-II transgenic mice. In all assays, 105 T cells were cultured in round-bottom 96-well plates with the indicated ratios of APCs. 1 µCi [3H]thymidine was added in each well for the last 18 h of culture, and thymidine incorporation was measured on a Microbeta Trilux Luminescence Counter (PerkinElmer). T cell differentiation assays. FACS-sorted cLP or splenic DC/MP were incubated at 105 cells/ml for 2 h at 37°C in the presence of 0.5 mg/ml endotoxin-free OVA protein (EndoGrade OVA) and washed twice in complete RPMI before co-culture at the indicated ratios with 105 OT-II cells (5 d), MACS enriched from the LNs and spleen of RAG1 OT-II transgenic mice. All cultures were split and complemented with fresh media after 72 h. On day 5, cells were restimulated for 5 h with 50 ng/ml phorbol myristate acetate and 500 ng/ml ionomycin in the presence or not of GolgiStop (BD). Cells were then surface stained with APC-Cy7–conjugated α-CD4 mAb (RM4-5) and Live/Dead Fixable cell stain kit, followed by FITC-conjugated α-foxp3 (FJK-16s), PE-conjugated α–IL-17A (eBio17B7), and APC-conjugated IFN-γ (XMG1.2) intracellular mAbs (all from eBioscience) in accordance with the manufacturer’s protocol. After the 5-h restimulation period, culture supernatants were also harvested for analysis of their cytokine content. Gene expression analysis. cLP DC/MP subsets were FACS sorted from WT C57BL/6 mice, colitic CD45RBhi mice, or control CD45RBhi+lo mice, as indicated (Figs. 1 A and 6 C), and total RNA was isolated using TRIZOL reagent (Invitrogen) according to the manufacturer’s instructions. RNA quantity and quality were assessed using the 2100 Bioanalyzer (Agilent Technologies). RNA was reverse transcribed into cDNA (SuperScript III; Invitrogen), and real-time RT-PCR was performed on a 7900 HT instrument (Applied Biosystems; FAM-labeled primers were obtained from Applied Biosystems and are listed in Table S1). Gene expression was analyzed using GAPDH as an endogenous control. In the MO-adoptive transfer experiments, gene expression analysis was performed using the TaqMan PreAmp Cells-to-Ct kit (Applied Biosystems; Denning et al., 2007a) for cDNA synthesis from a low number of cells. cDNA was then subject to a 14-cycle, target-specific preamplification step before being analyzed by regular real-time RT-PCR on a 7900 HT instrument using a custom-designed TaqMan Express Plate of 45 target genes and 3 housekeeping endogenous control genes (PN 4391526; Applied Biosystems). The main findings were confirmed by regular RT-PCR on non-preamplified cDNA. Microarray analysis. Total RNA was extracted from biological replicates of freshly isolated, FACS-sorted cLP subsets 1–4. cDNA was prepared, labeled, and hybridized to the commercial GeneChip, mouse gene 1.0 ST containing the mouse genome for 16 h, according to standard manufacturer protocols (Affymetrix). Hybridized chips were stained and washed and were scanned using the GeneChip 3000 7Gplus scanner (Affymetrix). GeneChip Operating Software (GCOS version 1.4; Affymetrix) was used for the initial analysis of the microarray data to convert the image files to cell intensity files. All cell intensity files, representing individual biological replicates, were normalized using the RMA-sketch within Expression Console (EC version 1.1; Affymetrix) to produce the chip files and perform the quality assessment. A pivot table constructed from the individual chip files was created including calls and signal intensities for each probe set. The pivot table was then imported into GeneSpring GX 7.3 (Agilent Technologies), where hierarchical clustering using mean linkage with Pearson correlation similarity measure was used to produce the dendrogram indicating biological replicate and condition grouping. The pivot table was also imported into Genomics Suite software (Partek Inc.) to produce a principal components analysis plot as a second measure for the grouping of biological replicates. Analysis of variance was run from this dataset to produce a false discovery rate report producing multiple test-corrected p-values for each comparison of interest. The replicates of all test conditions and controls were combined, and quality filters based on combined calls, signal intensities, and fold change were used to evaluate individual probe set comparisons. Present and marginal calls were treated as the same, whereas absent calls were negatively weighted and eliminated from quality summations. Ratios of test/control values and associated analysis of variance p-values of all individual probe sets passing the above filters were summarized in Excel from GCOS, EC, GeneSpring, SAM, and Partek software. The resulting probe sets were input into Ingenuity Pathway Analysis (Ingenuity Systems) for a core analysis summarizing the genes into networks, functions, and pathways. The integrality of the microarray data is available from GEO DataSets under accession no. GSE27859. Analysis of DC/MP and T cell cytokine production For MP/DC cultures, culture supernatants were harvested after 24 h of culture in medium alone or LPS from Escherichia coli (Sigma-Aldrich). Cytokines concentrations were analyzed using the SearchLight multiplex cytokine immunoassays (Aushon Biosystems). In some cases, the amount of IL-10 was determined using a conventional double-sandwich ELISA (DuoSet mouse IL-10; R&D systems) according to the manufacturer’s instructions. IFN-γ and IL-17A concentration in T cell culture supernatants (after 5 h of restimulation) was determined using conventional double-sandwich ELISA (DuoSet mouse). BrdU incorporation experiments WT C57BL/6 mice were pulse-injected i.p. with 0.8 mg BrdU (Sigma-Aldrich) and then continuously given 0.8 mg/ml BrdU in drinking water supplemented with 1% sucrose. cLP cell suspensions were prepared at different time points and analyzed for BrdU incorporation with the FITC–anti-BrdU Flow kit (BD). Statistical analysis An unpaired Student’s t test was performed using Prism 4 software (GraphPad Software) in all cases, except for the microarray analysis (see above). Where appropriate, mean ± SD is represented on graphs. *, P < 0.05; **, P < 0.01; ***, P < 0.001. Online supplemental material Fig. S1 shows that subsets 1 and 2 have a much slower turnover than subsets 3 and 4. Fig. S2 shows that cLP subsets 3 and 4 but not 1 and 2 express CCR7 after in vitro activation. Fig. S3 shows that F4/80+ cells are excluded from the organized lymphoid structures of the cLP. Fig. S4 shows that DTx treatment efficiently depletes cLP subsets 1–4 without inducing significant inflammation in the colon of CD11cDTR → C57BL/6 chimeric mice. Fig. S5 shows that the MP subsets 1 and 2 and also the CD103− DC subset 3 express E-cadherin and CX3CR1 during colitis. Table S1 lists references of the Applied Biosystems FAM-labeled primers used in this study. Table S2 shows classification of colonic MPs and DCs according to their surface marker expression and proposed functions. Online supplemental material is available at http://www.jem.org/cgi/content/full/jem.20101387/DC1.
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- Abstract: found
- Article: not found
CD14, a receptor for complexes of lipopolysaccharide (LPS) and LPS binding protein.
R Ulevitch, J Mathison, Graham A Wright … (1990)
Leukocytes respond to lipopolysaccharide (LPS) at nanogram per milliliter concentrations with secretion of cytokines such as tumor necrosis factor-alpha (TNF-alpha). Excess secretion of TNF-alpha causes endotoxic shock, an often fatal complication of infection. LPS in the bloodstream rapidly binds to the serum protein, lipopolysaccharide binding protein (LBP), and cellular responses to physiological levels of LPS are dependent on LBP. CD14, a differentiation antigen of monocytes, was found to bind complexes of LPS and LBP, and blockade of CD14 with monoclonal antibodies prevented synthesis of TNF-alpha by whole blood incubated with LPS. Thus, LPS may induce responses by interacting with a soluble binding protein in serum that then binds the cell surface protein CD14.
Author and article information
Journal
Journal ID (iso-abbrev): Clin Transl Med
Title:
Clinical and translational medicine
Publisher:
Springer Science and Business Media LLC
ISSN:
2001-1326
ISSN
(Print):
2001-1326
Publication date
(Electronic):
2015
Volume: 4
Affiliations
[1
]
Department of Pediatrics and Neonatal-Perinatal Medicine, Georgia Regents University,
Augusta, Georgia ; Vascular Biology Center, Georgia Regents University, Augusta, Georgia
; Medical College of Georgia at Georgia Regents University, 1120 15th St, BIW-6033,
Augusta, GA 30912 USA.
[2
]
Herman B. Wells Center for Pediatric Research, Georgia Regents University, Augusta,
Georgia ; Department of Pediatrics and Neonatal-Perinatal Medicine, Indiana University
School of Medicine, Indianapolis, Indiana USA ; Department of Biochemistry and Molecular
Biology, Indiana University School of Medicine, 699 Riley Hospital Drive, RR208, Indianapolis,
IN 46202 USA.
Article
Publisher Item ID: 40DOI: 10.1186/s40169-014-0040-3
PMC ID: 4384980
PubMed ID: 25852821
SO-VID: e0b3ec60-a8b4-411c-8e93-036cc6fbc105
History
Data availability:
Keywords: Atherosclerosis, Autoimmune Disease, CD14, CD16, Cardiovascular, Human, Ly6C, Macrophage, Monocyte, Mouse