Intestinal Dendritic Cell Subsets: Differential Effects of Systemic TLR4 Stimulation on Migratory Fate and Activation In Vivo
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免疫学杂志 2005年第3期
Abstract
Dendritic cells (DC) present peripheral Ags to T cells in lymph nodes, but also influence their differentiation (tolerance/immunity, Th1/Th2). To investigate how peripheral conditions affect DC properties and might subsequently regulate T cell differentiation, we examined the effects of a potent DC-activating, TLR-4-mediated stimulus, LPS, on rat intestinal and hepatic DC in vivo. Steady-state rat intestinal and hepatic lymph DC are E2 integrinhigh (CD103) and include two subsets, signal regulatory protein (SIRP)hi/low, probably representing murine CD8–/+ DC. Steady-state lamina propria DC are immature; surface MHC class IIlow, but steady-state lymph DC are semimature, MHC class IIhigh, but CD80/86low. Intravenous LPS induced rapid lamina propria DC emigration and increased lymph DC traffic without altering SIRPhigh/SIRPlow proportions. CD80/86 expression on lymph or mesenteric node DC was not up-regulated after i.v. LPS. In contrast, i.v. LPS stimulated marked CD80/86 up-regulation on splenic DC. CD80/86 expression on intestinal lymph DC, however, was increased after in vitro culture with TNF- or GM-CSF, but not with up to 5 μg/ml LPS. Steady-state SIRPlow DC localized to T cell areas of mesenteric nodes, spleen, and Peyer’s patch, whereas SIRPhigh DC were excluded from these areas. Intravenous LPS stimulated rapid and abundant SIRPhigh DC accumulation in T cell areas of mesenteric nodes and spleen. In striking contrast, i.v. LPS had no effect on DC numbers or distribution in Peyer’s patches. Our results suggest that any explanation of switching between tolerance and immunity as well as involving changes in DC activation status must also take into account differential migration of DC subsets.
Introduction
Intestinal immune responses toward orally acquired Ags must be tightly regulated. Despite demonstrating a profound hyporesponsiveness to dietary and commensal bacterial Ags, the gut mounts protective responses toward pathogens. It is likely that conditions within the local gut microenvironment play a key role in determining the course of the immune response. A better understanding of the regulation of intestinal immune responses is crucial for developing treatment of inflammatory bowel disease and food allergies and for the design of effective intestinal vaccines.
Naive T cells, particularly CD4+ cells, are not thought to be precommitted to any particular differentiation pathway before their activation, but after activation, may differentiate along one of several distinct pathways. Exogenous stimuli (such as the properties of an Ag itself or the effects that pathogens/Ags exert on their local microenvironment) may influence their differentiation fate. Antigenic encounters primarily occur in the periphery; however, T cells are activated in draining lymph nodes (LN).4 To select those patterns of gene expression needed for appropriate differentiation, LN T cells need to be reactive to environmental conditions in the periphery. A central and only partially understood question is how information about these conditions is relayed from the periphery to the LN.
Dendritic cells (DC) acquire Ags in peripheral tissues and transport them to LN, where they present processed peptides to T cells and probably native Ags to B cells (1, 2). DC in general are highly sensitive to environmental cues, and local microenvironmental conditions affect DC phenotype and function (3, 4, 5). DC are therefore prime candidates for the transduction of information from the periphery to LN. Despite some current dogma, much evidence shows that DC are continually migrating in afferent lymph from tissues such as intestine, skin, and liver in the absence of overt inflammatory stimulation (6, 7). DC can exist in different states of maturation/activation that are reflected in the fate of the CD4+ T cells they activate. Under steady-state conditions, DC in LN have been shown to express low levels of costimulatory molecules and can induce tolerogenic or hyporesponsive T cells in vivo (8, 9). It is an attractive hypothesis that perturbations of the steady-state (e.g., inflammation) modulate the information delivered by the DC to the T cell, leading to an active, rather than a tolerogenic, immune response. Although this concept is supported by many studies that have investigated environmental effects on DC in vitro, whether similar effects occur in vivo is largely unknown. Studying environmental effects on DC in vivo is problematic, because the majority of techniques for DC isolation are themselves likely to induce phenotypic and functional changes in DC independent of true physiological or pathological stimuli in the local microenvironment. Of the various in vivo DC populations that can be studied, the DC most likely to be informative in relation to information transduction from the periphery are those migrating in afferent lymph. We have used the approach of mesenteric (MLNX) or celiac (CoeLNX) lymphadenectomy (10, 11, 12), followed by thoracic duct cannulation, to isolate DC migrating from the periphery. By collecting and manipulating these cells at 4°C, we can preserve the patterns of gene expression within migrating DC in vivo (6, 7, 10, 13, 14, 15 ; reviewed in Ref.16). This model thus provides the opportunity to directly identify changes in the properties of migrating DC that accompany perturbations in peripheral tissues and that may relate to differential activation of naive T cells.
Using this model we have shown previously that two phenotypically and functionally distinct subsets of DC migrate in rat intestinal lymph (both E2 integrinhigh), which can be distinguished because one subset coexpresses CD4 and high levels of signal regulatory protein (SIRP) (6). The same two subsets have been identified in rat spleen (17, 18), but how these phenotypically similar DC relate functionally is not yet fully clear. In both lymph and spleen, SIRPhigh DC are more potent than SIRPlow DC in the activation of allogeneic CD4+ and CD8+ T cells, naive Ag-specific CD4+ T cells in vivo, and sensitized Ag-specific CD4+ T cells in vitro (6, 17, 18). In addition, SIRPlow lymph DC selectively transport remnants of apoptotic intestinal epithelial cells to the T cell areas (TCAs) of mesenteric LNs (MLN) (7). This selective uptake of apoptotic material is a function shared by murine CD8+ DC and may indicate that the SIRPlow DC is the rat homologue involved in the induction of peripheral tolerance (19, 20, 21).
We have described in detail the properties of DC migrating in lymph from the intestine under steady-state conditions (6, 7, 10, 13, 14, 15, 22) and have also shown that i.v. LPS administration caused a substantial increase in the output of DC in intestinal lymph and that this effect can be partially blocked by an anti-TNF- Ab (23). In this study we have examined the effects of i.v. LPS on the properties of intestinally derived DC subsets, with reference to DC in MLN, liver-draining lymph, and spleen for comparison to gain insight into how such changes might affect the quality of the immune response. Our results show that i.v. LPS induces major changes in DC migration from the gut in the absence of significant changes to DC surface phenotype. We also show that proinflammatory stimulation is associated with the appearance of the previously excluded SIRPhigh DC subset in TCAs of MLN and spleen. We suggest that these alterations in migration may relate to the regulation of the switch between tolerance and active immunity.
Materials and Methods
Rats and surgical procedures
Rats (PVG (RT1c)) were bred and maintained under specific pathogen-free conditions in the Sir William Dunn School of Pathology (Oxford, U.K.) and were males between 12 and 16 wk of age. All procedures were approved by and conducted in accordance with Home Office guidelines. MLNX and thoracic duct cannulation were performed as previously described (10). CoeLNX was conducted as described by Matsuno et al. (24). Cannulations always involved multiple rats to ensure sufficient cell numbers for experimentation.
Expansion of DC numbers in vivo using Flt3 ligand
To boost DC numbers for in vitro studies, rats were injected s.c. daily for 8 days with 15 μg of recombinant human Fms-like receptor tyrosine kinase 3 ligand (Flt3L) in 200 μl of sterile PBS. Flt3L was provided by Immunex.
Reagents
All cell culture was performed in RPMI 1640 medium supplemented with 10% FCS, 50 μM 2-ME, 100 μg/ml penicillin, 50 U/ml streptomycin, and 100 μg/ml L-glutamine (all from Invitrogen Life Technologies). Recombinant rat TNF- and recombinant rat GM-CSF were purchased from R&D Systems and were used at a final concentration of 50 ng/ml for in vitro cell stimulation. Salmonella typhimurium LPS and 7-aminoactinomycin D (7AAD) were purchased from Sigma-Aldrich. FITC-, Cy3-, and Cy5-conjugated tyramide used for immunofluorescence were obtained from NEN. Normal rat, mouse, and donkey sera were obtained from Serotec. OX62 MACS beads were purchased from Miltenyi Biotec. Goat anti-mouse IgG Dynabeads were obtained from Dynal Biotech U.K. Cy5 for Ab conjugation was purchased from Amersham Biosciences.
Antibodies
Monoclonal Abs. mAbs were either neat tissue culture supernatants (donated by M. Puklavec, Sir William Dunn School of Pathology) or commercially available purified IgG. All mAbs were raised in mice unless otherwise stated. Unconjugated mAbs were anti-human factor I, IgG1 isotype control (OX21), anti-human HLA class I, IgG2a isotype control (W6/32), anti-MHC class II (OX6), anti-CD8 (OX8), anti-Ig -chain (OX12), anti-RT-1Ac MHC class I (OX27), anti-CD4 (OX38), anti-SIRP (OX41), anti-CD11b (OX42), and anti-E2 integrin (OX62). Anti-CD11c (8A2), anti-sialoadhesin (ED3), and anti-TCR (R73) mAbs were purchased from Serotec. Anti-CD80 (3H5) and anti-CD86 (24F) were provided by Dr. H. Yagita (Juntendo University School of Medicine, Tokyo, Japan).
Conjugated mAbs. FITC- and PerCP-conjugated anti-MHC class II (OX6), FITC-conjugated anti-CD4 (OX38), and FITC-conjugated hamster anti-mouse/rat CD40 (HM40-3) were obtained from BD Pharmingen. PE-conjugated anti-E2 integrin (OX62) was purchased from Serotec. PE-conjugated CD80 (3H5) and CD86 (24F) mAbs were obtained from Biocarta (Oxford, U.K.). FITC- and Cy5-conjugated OX62 and OX41 and HRP-conjugated OX41 mAbs were made in-house using a peroxidase labeling kit according to the manufacturer’s instructions (Roche).
Secondary Abs. FITC-, Cy3-, Cy5-, and HRP-conjugated donkey anti-mouse IgG Abs were obtained from Jackson ImmunoResearch Laboratories. Texas Red-conjugated goat anti-mouse IgG/IgM/IgA Ab was purchased from Southern Biotechnology Associates. PE-conjugated goat anti-mouse IgG was obtained from Sigma-Aldrich.
Primary cell preparations
Intestinal and hepatic pseudoafferent lymph DC. Thoracic duct lymph collected from lymphadenectomized rats was passed through a sterile 70-μm pore size cell strainer (BD Biosciences), and RBC were lysed using ammonium chloride solution. Lymph DC were typically enriched within total lymph cells by density gradient centrifugation. Total lymph cells were resuspended in PBS/2% FCS at a concentration of 2 x 107 cells/ml, and 108 cells were overlaid onto 2 ml of Nycoprep 1.068 medium (Nycomed). After centrifugation at 2000 rpm for 20 min at room temperature, the collected interface cells typically consisted of 15–30% DC. In some experiments DC were enriched to 20–40% by depletion of T and B cells. Total lymph cells were labeled using OX12 (anti-Ig) and OX52 (pan T cell) mAbs, followed by goat anti-mouse IgG Dynabeads. Labeled cells were then magnetically removed. Highly purified lymph DC were obtained using OX62-MACS beads and positive selection using AUTOMACS according to the manufacturer’s instructions.
Spleen, MLN, and cervical LN (CLN) DC. Spleens, MLN, and CLN were teased apart using watchmaker’s forceps into prewarmed RPMI 1640 medium containing 0.5 mg/ml collagenase D and 0.5 mg/ml DNase (Roche). Tissues were digested for 45 min at 37°C with periodic agitation. Undigested stromal material was removed by passage over a 70-μm pore size filter before RBC lysis using ammonium chloride solution. DC were enriched within splenocytes or MLN cells by magnetic selection using OX62-MACS beads.
MLR
Lymph was collected from MLNX PVG (RT1c) rats over 16 h after i.v. injection of PBS or 50 μg of LPS, and DC were enriched by density gradient centrifugation. Lymph DC were irradiated and then titrated against 2 x 105 responder CLN cells from DA (RT1a) rats. Cultures were performed in quadruplicate at 37°C for 3 days, after which, T cell proliferation was measured by thymidine incorporation and expressed as cpm. Background proliferation counts of responder cells alone were subtracted from proliferation in cultures containing DC.
Flow cytometry
All Ab and serum dilutions were made in PBA (PBS/1% BSA/10 mM sodium azide). All Ab incubation steps were conducted for 30 min over ice, and excess unbound Ab was removed by three washes in PBA. Cells were first blocked with 10% normal rat serum to block potential FcR binding. Abs were neat tissue culture supernatants or purified IgG diluted to 1–10 μg/ml. Relevant isotype-matched control Abs were included. Unconjugated primary Abs were detected using FITC- or PE-conjugated anti-mouse Abs. Where appropriate, 10 μg/ml 7AAD was added during the final minute of immunolabeling to label dead cells. Cells were fixed using 4% paraformaldehyde in PBS. Fluorescence was recorded using a FACSCalibur (BD Biosciences), and data were analyzed using CellQuest software (BD Biosciences), excluding dead cells.
Tissue immunofluorescence
Fresh tissues were frozen in Tissue-Tek OCT compound (Sakura Finetek Europe) over isopentane on dry ice. Sections were cut at 5 μm, dried, and stored at –20°C until use. After thawing, sections were fixed for 10 min in cold 100% ethanol and rinsed in PBS. Nonspecific protein binding was minimized by blocking for 30 min at room temperature with PBS/3% BSA before immunolabeling. Abs were neat tissue culture supernatants or purified IgG, typically diluted to 1–10 μg/ml in PBS/3% BSA. Appropriate isotype-matched negative control Abs were included in all immunostaining procedures to ensure specificity of staining. Ab incubations were conducted in a moist chamber at room temperature for 2 h or overnight at 4°C. Excess unbound Ab was washed off using three changes of PBS. In multicolor immunolabeling procedures, cross-reaction between anti-mouse secondary detection Ab and subsequently added directly conjugated mouse mAbs was prevented by blocking with 10% normal mouse serum for 20 min at room temperature. Sections were mounted in water-soluble Vectashield (Vector Laboratories), and coverslips were secured using clear nail varnish. Sections were stored in the dark to minimize photobleaching.
HRP/tyramide amplification of immunofluorescence
Unconjugated mAbs giving a weak signal when detected using a standard fluorophore-conjugated secondary Ab were detected using HRP-conjugated secondary mAb and were developed using fluorophore-conjugated tyramide. Endogenous tissue peroxidase was first quenched within fixed tissue sections for 30 min at 37°C using 0.7 U/ml glucose oxidase enzyme (Sigma-Aldrich) in prewarmed 0.1 M phosphate buffer containing 25 mM D-glucose and 1 mM sodium azide. Sections were then rinsed in PBS, preblocked, and immunolabeled as described above. Tyramide development of HRP was conducted for 15–20 min at room temperature according to the manufacturer’s instructions (NEN).
Image acquisition
Tissue immunofluorescence was recorded using an MRC 1024 laser scanning confocal microscope (Bio-Rad) or an Axioplan epifluorescence microscope (Carl Zeiss). For the confocal microscope, digital images were collected using COMOS software (Bio-Rad); for the epifluorescence microscope, images were obtained using a Spot camera and Spot Advanced software (Diagnostic Instruments). Single-label fluorescence sections were observed alongside multilabeled sections to ensure no bleed-through of signal from one channel to another. Digital images were processed and overlaid using Photoshop software (Adobe Systems). Figures show representative images of many sections studied using tissues derived from a number of different rats.
Results
In situ distribution of DC in the steady-state small intestine
DC were identified in cryosections of rat ileum by immunolabeling for E2 integrin, CD11c, or MHC class II Ags. In Peyer’s patches (PP), large, irregular E2 integrin+ cells were detected in the subepithelial dome region underlying the follicle-associated epithelium and in the interfollicular TCAs between B cell follicles (Fig. 1a). These cells were confirmed to be DC by double-labeling for CD11c and MHC class II (Fig. 1, c and e). Large, irregular E2 integrin+ and CD11c+ cells were also distributed throughout the lamina propria (Fig. 1b). These cells intensely coexpressed intracellular MHC class II, but were Ig negative, indicating that the MHC class II was unlikely to have been derived from the ingestion of apoptotic B cells (data not shown). In the lamina propria, E2 integrin was also expressed by MHC class II– intraepithelial lymphocytes as previously described (25).
FIGURE 1. Detection of DC within the steady-state rat ileum. Cryosections of ileum immunolabeled for DC-associated Ags in PP and lamina propria (LP). a and b, E2 integrin immunolabeling (green) of the subepithelial dome region (SED), interfollicular regions (IFR), and B cell follicles (B) of PP and LP (magnification: a, x100; b, x200). c–e, Double labeling for MHC class II (green) and CD11c (red) in the subepithelial dome (c, x200), interfollicular region of PP (e, x400), and lamina propria (d, x630). f and h, Double labeling for E2 integrin (red) and MHC class II (green) in lamina propria and intestinal epithelial cell (IEC) layer (f, x630). g, i, and j, Double labeling for MHC class II (green) and CD86 (red) in subepithelial dome of PP (g, x400), interfollicular region of PP (i, x400) and lamina propria (j, x200).
The cellular distribution of MHC class II differed markedly between PP and lamina propria. In lamina propria, although some MHC class II was surface expressed, the bulk appeared granular and intracellular (characteristic of retention within intracytoplasmic vesicles; Fig. 1, d, f, and h). In contrast, DC of the subepithelial dome and interfollicular regions of PP showed surface expression of MHC class II that colocalized well with E2 integrin or CD11c expression (Fig. 1, c and e). Most large, irregular cells in the subepithelial dome and interfollicular regions of PP (Fig. 1, g and i, respectively) coexpressed MHC class II and CD86, but lamina propria DC expressing granular, cytoplasmic MHC class II were CD86– (Fig. 1j). That lamina propria DC expressed intracellular MHC class II and lacked CD86 expression suggests that they are more developmentally immature than PP DC.
Intravenous LPS stimulates DC migration from the lamina propria
We have previously shown that i.v. LPS stimulates an increased traffic of DC in lymph draining the small intestine (23). To relate this to events occurring in the intestine, cryosections of ileum from rats that had been injected i.v. with either 50 μg of LPS or PBS 6, 12, 24, 48, or 72 h previously were labeled with OX62 (anti-E2 integrin) mAb. Compared with tissue from normal rats, PBS administration had no effect on the relative frequency and distribution of lamina propria E2 integrin+ cells (represented as steady-state in Fig. 2a). Intravenous LPS caused a time-dependent loss of DC from the lamina propria, detectable by 6 h (not shown), and by 12 h, virtually no DC were present in the lamina propria (Fig. 2c). By 24 h, small numbers of DC were present (Fig. 2e), and steady-state numbers were reached by 72 h (Fig. 2g). CD11c immunolabeling gave similar results (Fig. 2, b, d, f, and h). Similar results were obtained using an anti-MHC class II mAb (data not shown). E2 integrin+/CD11c– intraepithelial lymphocytes were detectable at all time points studied after PBS or LPS administration. Thus, i.v. LPS administration leads to a significant, but only transient, loss of DC from lamina propria.
FIGURE 2. Intravenous LPS administration leads to a loss of DC immunolabeling within the lamina propria of the small intestine. Cryosections of ileum from rats i.v. injected with PBS or 50 μg of LPS and killed at various times. Sections were immunolabeled for E2 integrin (a, c, e, and g) or CD11c (b, d, f, and h). Intestinal epithelial cells (IEC) and lamina propria (LP) are indicated. All panels show x630 magnification.
Intravenous LPS stimulates increased migration of DC in lymph draining both intestine and liver
We have previously shown that i.v. LPS administration is associated with an increased traffic of DC in lymph draining the intestine of MLNX rats (23). To determine whether the increase in output of DC into lymph was true of other organs, we investigated the effects of i.v. LPS on the frequency and phenotype of DC trafficking in pseudoafferent lymph from CoeLNX rats. The steady-state output of DC into liver-draining lymph was less (85%) than that into intestinal lymph. There was a selective increase in DC traffic from both organs, peaking between 6 and 12 h after i.v. LPS (Fig. 3a). Maximum output from the intestine increased by 10- to 20-fold, whereas output from the liver increased by 60- to 70-fold. In both cases, the increase in DC output from the periphery was transient, and DC traffic had returned to steady-state levels by 24 h. A subsequent injection of LPS given 24 h after the first, resulted in only a minor transient increase in lymph DC output, consistent with the lamina propria being depleted of DC (Fig. 3b).
FIGURE 3. Increased DC frequency in intestinal and hepatic lymph after i.v. LPS injection. Thoracic duct lymph from MLNX and CoeLNX rats collected 0–6, 6–12, and 12–22 h under steady-state conditions or after i.v administration of 50 μg of LPS. a, DC output after i.v. LPS administration, expressed as the fold increase relative to steady-state output over the same time. Absolute DC numbers trafficking in lymph during each collection period were determined by multiplying the percent proportion of DC (identified by E2 integrin expression by flow cytometry) by the total number of lymph cells. Results shown are representative of one of three similar experiments. b, Total number of intestinal lymph DC collected at intervals over 48 h. LPS (50 μg) was injected at 0 and 24 h, and the output of DC during each interval was determined. Similar results were obtained in two independent experiments. c, Four-color flow cytometric analysis of DC draining the intestine (MLNX) and liver (CoeLNX) using mAbs to E2 integrin, MHC class II, SIRP, and CD4. The four lower dot plots show lymph DC subset proportions before (control) and 6–12 h after LPS (+LPS) administration using the gates shown in two upper dot plots. d, Output of CD4low/SIRPlow and CD4high/SIRPhigh DC subsets draining the intestine (MLNX) and liver (CoeLNX) after i.v. LPS administration, expressed as the fold increase relative to steady-state output over the same time. Similar results were obtained in two independent experiments.
We have previously shown that intestinal pseudoafferent lymph contains two populations of DC, distinguishable by their expression of CD4 and SIRP (6). DC were identified by their high E2 integrin and MHC class II expression, and the proportions of CD4high/SIRPhigh and CD4low/SIRPlow DC subsets contained within steady-state intestinal and hepatic pseudoafferent intestinal lymph were determined. DC were detectable in nonenriched MLNX and CoeLNX lymph, representing 0.3% of total lymph cells (Fig. 3c, upper panels). CD4high/SIRPhigh lymph DC represented the major subset of DC trafficking in both intestinal and hepatic pseudoafferent lymph under steady-state conditions, accounting for between 60 and 70% of total DC (Fig. 3c, lower panels). The number and relative proportions of CD4high/SIRPhigh and CD4low/SIRPlow DC remained constant over a period of at least 44 h after cannulation in control rats, showing that surgery did not stimulate differential changes in DC subset migration from the gut or liver (data not shown).
To determine whether LPS administration altered the relative proportions of migrating DC subsets, rats were injected i.v. with 50 μg of LPS, and hepatic or intestinal pseudoafferent lymph was collected at intervals over 22 h. In gut-draining lymph, the output of both subsets increased transiently in parallel, showing that LPS challenge did not stimulate a preferential increase in one particular subset. In liver-draining lymph, there was a more pronounced increase in the output of CD4high/SIRPhigh DC compared with CD4low/SIRPlow DC (Fig. 3d). Thus, i.v. LPS administration resulted in a transient increase in lymph DC traffic draining both intestine and liver. SIRPhigh/low DC proportions were unchanged in lymph draining the intestine after LPS, whereas SIRPhigh DC proportions increased in liver-draining lymph.
Intravenous LPS administration increases DC frequency in mesenteric nodes
Given that i.v. LPS administration led to a transient loss of DC immunolabeling in the lamina propria and increased DC traffic in lymph draining the intestine, we predicted that this would be reflected in an increased migration of DC into the draining MLN. Sections of MLN from rats injected i.v. 0, 6, 12, or 24 h earlier with PBS or 50 μg of LPS were double-labeled for MHC class II and TCR. PBS administration had no effect on DC distribution compared with tissue from noninjected rats at all time points studied (data not shown). After LPS administration, DC frequency in the TCA of MLN was slightly increased by 6 h (data not shown), and by 12 h, it was several-fold increased compared with that in sections from rats injected i.v. 12 h earlier with PBS (compare Fig. 4, a and b). By 24 h, DC frequency was reduced compared with that at 12 h, but remained elevated compared with steady-state levels (data not shown). Similar staining patterns were obtained using anti-E2 integrin and anti-CD11c mAbs (data not shown). The increase in DC numbers in the MLN correlates directly with the loss of DC from the lamina propria 12 h after LPS administration and with the peak output of DC from gut into lymph.
FIGURE 4. Intravenous LPS induces an accumulation of DC in the draining MLN. a and b, Cryosections of MLN from rats injected i.v. 12 h earlier with PBS or 50 μg of LPS were immunolabeled for TCR (red) and MHC class II (green), TCAs (magnification, x200). c, MLN DC enriched by OX62-MACS beads immunolabeled for E2 integrin and MHC class II. The absolute number of DC recovered at each time point was expressed as a proportion of the starting MLN cell number. Values represent the mean and SD of three independent experiments for each time point. Data were analyzed using unpaired one-tailed Student’s t test.
To quantitate changes in DC numbers in MLN, nodes taken at intervals after LPS administration were digested with collagenase, and DC were enriched by positive selection using OX62-MACS beads. DC were identified by flow cytometry by coexpression of E2 integrin and MHC class II. Absolute numbers of DC were expressed as a percentage of the total MLN cell numbers (Fig. 4c). Absolute DC numbers were increased above steady-state by 6 h, and a 4-fold increase in MLN DC proportions was observed 12 h after LPS administration (p = 0.024), consistent with our immunohistochemical data. Significantly increased numbers of MLN DC 12 h after i.v. LPS administration is consistent with the loss of lamina propria DC and the peak output of DC in lymph draining the intestine.
Effects of i.v. LPS on the anatomical distribution of DC subsets in secondary lymphoid organs
DC subsets were localized within secondary lymphoid tissue by immunolabeling for DC-associated markers together with the OX41 mAb. In addition to delineating rat DC subsets, the SIRP Ag detected by the OX41 mAb has been described on a subpopulation of macrophages (26, 27). We therefore paid particular attention to ensure that our immunolabeling procedure would allow us to distinguish between these cell types. Although SIRPlow DC were detectable by flow cytometry, they were not detectable by immunohistochemistry and thus appear SIRP–. We retain the term SIRPlow for consistency.
MLN. In cryosections from normal rats, double labeling for E2 integrin and SIRP identified both SIRPhigh and SIRPlow DC subsets in the interfollicular traffic areas of the cortex (Fig. 5a), consistent with the migration of both subsets in intestinal lymph and their entry into MLN via the afferent lymphatics. A few SIRPhigh and SIRPlow DC were also found in the subfollicular area of the superficial cortex, just beneath the lymph follicle. In contrast, DC in TCAs were almost exclusively SIRPlow, and SIRPhigh DC were rarely seen (Fig. 5b). The exclusion of SIRPhigh DC from the MLN TCA was confirmed by triple immunolabeling for MHC class II, TCR, and SIRP. MHC class II+/SIRPhigh and MHC class II+/SIRPlow DC were found in the cortical interfollicular traffic areas (Fig. 5c), whereas frequent SIRPlow DC, but few SIRPhigh DC, were detectable within the TCA (Fig. 5d). Thus, SIRPhigh DC are excluded from the steady-state MLN TCA.
FIGURE 5. Intravenous LPS administration is associated with the appearance of SIRPhigh DC in the MLN TCA. a and b, Cryosections of MLN from control rats were immunolabeled for E2 integrin (green) and SIRP (red) to localize DC subsets in the MLN cortex (a) and TCA (b); SIRPlow DC (shown in green) and SIRPhigh DC (shown in yellow). IFTA, Cortical interfollicular traffic areas; B, B cell follicles; T, T cell area. c and d, Immunolabeling for TCR (red), MHC class II (green), and SIRP (blue) on cryosections of control MLN. MHC class II+/SIRPlow DC (shown as large, irregular green cells) and MHC class II+/SIRPhigh DC (light blue). e and f, MLN cryosections from rats injected 12 h earlier with PBS (e) or 50 μg of LPS (f) immunolabeled for CD11c (red), MHC class II (green), and SIRP (blue) to localize DC subsets in the MLN TCA. CD11c+/MHC class II+/SIRPlow DC are shown in yellow, and CD11c+/MHC class II+/SIRPhigh DC are shown in white. Magnification for all panels, x200.
To determine whether the DC subset distribution in MLN changed after i.v LPS administration, cryosections from rats injected 6, 12, or 24 h earlier with 50 μg of LPS or PBS were labeled for MHC class II, CD11c, and SIRP. Tissue from normal rats or rats injected with PBS at all of the time points studied showed the presence of SIRPlow DC, but few SIRPhigh TCA DC (Fig. 5e). Six hours after administration of LPS, SIRPhigh DC were detected in the MLN (data not shown), and by 12 h significant numbers were present (Fig. 5f). The increase in SIRPhigh DC in the TCA 12 h after LPS challenge was accompanied by an increase in SIRPlow DC. Both subsets were still detectable in the TCA 24 h after LPS administration (data not shown).
PP. Cryosections from normal rats immunolabeled for E2 integrin and SIRP identified both SIRPlow and SIRPhigh DC in the subepithelial dome region underlying the follicle-associated epithelium (Fig. 6a), although there were relatively fewer SIRPhigh DC. SIRP+/E2 integrin– cells were also found at the subepithelial dome and in the interfollicular region, and these may be macrophages. As with the MLN, SIRP was undetectable on essentially all DC in the TCA of the interfollicular region of PP (Fig. 6b). However, in complete contrast to the results for MLN, i.v. LPS had no overt effect on either the relative number or the distribution of DC in PP. Both SIRPlow and SIRPhigh DC remained detectable in the subepithelial dome region at all time points studied after LPS (Fig. 6c), and SIRPhigh DC were rarely found in the PP TCA (Fig. 6d). Although SIRP-expressing cells were detectable in normal lamina propria using OX41 mAb, these cells did not coexpress E2 integrin or MHC class II. SIRPhigh DC were not detectable in the lamina propria after i.v. LPS administration at any of the time points studied (data not shown).
FIGURE 6. Intravenous LPS administration fails to modulate the relative frequency and distribution of rat PP DC subsets. Cryosections of PP from rats injected i.v. 12 h earlier with PBS (a and b) or 50 μg of LPS (c and d) immunolabeled for E2 integrin (green) and SIRP (red). SIRPlow DC are shown in green, and SIRPhigh DC are shown in yellow. SED, subepithelial dome region; B, B cell follicle; IFR, interfollicular regions. Magnification: a–c, x200; d, x400.
Spleen. Cryosections of rat spleen labeled using OX41 mAb showed that SIRPhigh cells were abundant within the steady-state spleen, particularly in the red pulp (Fig. 7a). These cells are probably macrophages, because they failed to express CD11c or E2 integrin (data not shown). SIRPhigh cells were also detected in the outer periarteriolar lymphoid sheath, where they formed a tight cuff around the TCR+ TCAs (Fig. 7b). These SIRPhigh cells expressed MHC class II and CD11c (Fig. 7c), but did not coexpress sialoadhesin (data not shown), indicating that they were DC rather than macrophages. Although CD11c+, MHC class II+/SIRPlow DC were found in the splenic TCA under steady-state conditions, SIRPhigh DC were rarely found. The cuff of SIRPhigh DC was not continuous around B cell follicles.
FIGURE 7. SIRPhigh DC are excluded from the TCAs of the steady-state rat spleen. a–c, Cryosections of spleen from control rats: single stained for SIRP (a), double labeled for SIRP (green) and TCR (red; b), or triple labeled for MHC class II (green), CD11c (red), and SIRP (blue; c). CD11c+/MHC class II+/SIRPlow DCs are shown in yellow, and CD11c+/MHC class II+/SIRPhigh are shown in white. d and e, Cryosections from rats injected i.v. with PBS (d) or 50 μg of LPS (e) 6 h earlier immunolabeled for MHC class II (green), TCR (red), and SIRP (blue). MHC class II+/SIRPhigh DC are shown in light blue, and MHC class II+/SIRPlow DC are shown in green. RP, red pulp; MZ, marginal zone; B, B cell follicles; T, TCAs. Magnification: a–c, x200; d and e, x400.
Intravenous PBS administration had no effect on the distribution of SIRPlow and SIRPhigh DC subsets in splenic white pulp compared with tissue from noninjected rats at all time points studied (data not shown). In contrast, i.v. LPS administration led to a loss of MHC class II+/SIRPhigh DC immunolabeling on the outskirts of the TCA and the appearance of MHC class II+/SIRPhigh DC within the TCA. This effect was observed by 6 h after LPS administration (compare Fig. 7, d and e). Interestingly, the frequency of SIRPhigh DC surrounding TCAs under steady-state conditions was comparable to the frequency of SIRPhigh DC present in the TCA after LPS challenge, suggesting a direct migration of DC from the outer periarteriolar lymphoid sheath to TCA. The overall frequency of SIRPlow TCA DC was increased by 12 h after LPS injection, correlating with a loss of SIRPlow DC from the red pulp (data not shown). At 12 and 24 h after i.v. LPS, SIRPhigh DC remained detectable within the TCA (data not shown). These data show that i.v. LPS administration is associated with the appearance of the previously excluded SIRPhigh DC subset in the TCA of MLN and spleen, but not PP.
Effects of i.v. LPS administration on the surface phenotype of lymph, MLN, and splenic DC
Intestinal pseudoafferent lymph DC were collected under steady-state conditions or during the peak output after i.v. LPS administration, and their surface phenotype was analyzed by flow cytometry. Surprisingly, LPS had little effect on the expression of E2 integrin, CD11c, MHC class I, CD40, and CD11b by DC trafficking in lymph draining the intestine. MHC class II expression was reduced, and CD80 and CD86 levels were increased by only 1.5- to 2-fold. Similar results were obtained for hepatic lymph DC collected after i.v. LPS administration (data not shown). Steady-state and LPS-induced, intestinally derived lymph DC showed a similar potency in stimulating the proliferation of allogeneic T cells, consistent with their comparable expressions of molecules known to be important for T cell activation (Fig. 8a).
FIGURE 8. Surface phenotype comparison of steady-state and LPS-induced intestinal lymph DC: comparison with hepatic lymph DC, MLN DC, and spleen DC. a, Comparison of the allostimulatory potential of intestinally derived lymph DC from PBS- or LPS-injected rats. Equal numbers of irradiated, density enriched, intestinally derived lymph DC from PBS or LPS-injected PVG (RT1c) rats were cultured with 2 x 105 total responder cervical LN cells from DA (RT1a) rats and cultured for 3 days. T cell proliferation was measured by thymidine incorporation and expressed as cpm after subtracting the background proliferation of responder cells alone. Error bars represent the SD of the mean of quadruplicate cultures. Data shown are from one of three similar experiments. b, Intestinal and hepatic lymph was collected at intervals over 22 h under steady-state conditions or after i.v. LPS administration. DC were enriched by depleting T and B cells and were immunolabeled for E2 integrin, SIRP, CD4, and either CD80 or CD86. Similar results were obtained in two independent experiments. c, Enriched DC immunolabeled for E2 integrin, MHC class II, and CD80, CD86, or CD40. Intestinal and hepatic lymph DC were collected between 6 and 12 h after i.v PBS or LPS injection. MLN and spleen DC were isolated 12 h after i.v. PBS or LPS injection. Filled histograms, DC from PBS-injected rats; solid line, DC from LPS-injected rats. Data are representative of at least three independent experiments.
The increases in surface costimulatory molecule expression by LPS-induced lymph DC were not restricted to one subset in particular. Both CD4low/SIRPlow and CD4high/SIRPhigh intestinal and hepatic lymph DC showed transiently increased surface expression of CD80 and CD86, with DC collected after the peak output showing similar surface expression levels as steady-state DC (Fig. 8b).
We wanted to compare the effect of i.v. LPS administration on intestinal and hepatic lymph DC with that on DC isolated from secondary lymphoid tissue. Lymph DC were collected under steady-state conditions or during their peak output after i.v. LPS administration. MLN and splenic DC were isolated 12 h after i.v. LPS or PBS administration by collagenase digestion, density separation, and MACS enrichment using OX62 beads. DC (identified as E2 integrin+/MHC class II+ cells) were additionally labeled for costimulatory molecules and examined by flow cytometry (Fig. 8c). Interestingly, intestinally derived lymph DC and MLN DC expressed comparable levels of CD80 and CD86, showing that mere entry of DC into the MLN from lymph did not induce phenotypic maturation, and only minor increases in CD80 and CD86 expression were observed on MLN DC 12 h after i.v. LPS, consistent with the minimal effect of i.v. LPS on intestinal lymph DC phenotype. In contrast, splenic DC showed substantial up-regulation of surface costimulatory molecule expression, with 5-fold increases in both CD80 and CD86 and a 2- to 3-fold increase in CD40 expression 12 h after LPS administration.
These data suggest that the environmental conditions created after i.v. LPS administration are insufficient to stimulate substantial up-regulation of molecules critical for T cell activation on DC derived from gut and liver. In contrast, splenic DC were phenotypically matured in response to i.v. LPS administration. Thus, DC in different tissue sites respond differently to the same stimulus.
Modulation of lymph DC surface phenotype during in vitro culture with GM-CSF and TNF-, but not LPS
To ensure that DC derived from the gut were not refractory to stimulation in general, in vitro experiments were conducted to examine the ability of known inducers of DC maturation to stimulate intestinally derived lymph DC. To collect sufficient DC, rats were pretreated with human Flt-3 ligand, which induces a 5- to 10-fold increase in intestinal lymph DC output without causing detectable changes in phenotype or function (U. Yrlid and G. G. MacPherson, manuscript in preparation).
Purified intestinal lymph DC from Flt-3 ligand-treated rats were cultured in vitro for 12 or 24 h in culture medium alone or in medium supplemented with 50 ng/ml recombinant rat GM-CSF, 50 ng/ml recombinant rat TNF-, or doses of LPS up to 5 μg/ml. The DC surface phenotype was subsequently assessed by flow cytometry (Fig. 9). Compared with freshly isolated lymph DC, only minor increases (<2-fold) in surface expression of MHC class I, MHC class II, and CD40 molecules were seen after culture in medium alone, and the presence of GM-CSF, TNF-, or LPS had no additive effect (not shown). A 24-h culture in medium alone failed to stimulate changes in surface CD80 and CD86 expression; however, the presence of 50 ng/ml GM-CSF or TNF- stimulated a profound increase in surface expression of these costimulatory molecules (up to a log-fold increase in CD86 with GM-CSF after 24 h). Surprisingly, the presence of up to 5 μg/ml LPS during culture had no effect on CD80/CD86 expression. Therefore, intestinally derived lymph DC can be phenotypically matured, but the nature of the stimulus is critical.
FIGURE 9. In vitro modulation of intestinally derived lymph DC surface phenotype by GM-CSF and TNF-, but not by LPS. Intestinal lymph DC from steady-state Flt3L-pretreated rats were enriched to >90% purity by density gradient centrifugation and cultured at 37°C in 5% CO2 for 12 and 24 h at 106/ml in medium alone or in medium supplemented with 50 ng/ml GM-CSF, 50 ng/ml TNF-, or up to 5 μg/ml LPS. Cells were then labeled for anti-E2 integrin and CD80/CD86 and were analyzed by flow cytometry. Histograms compare surface expression levels of CD80 and CD86 on freshly isolated lymph DC (filled histograms), 12-h cultured lymph DC (dashed line), and 24-h cultured lymph DC (solid line). Data are representative of one of three similar experiments.
Discussion
Intestinal Ags may induce tolerance or immunity, probably depending on the presence or the absence of proinflammatory conditions (9, 28). DC migratory behavior and sensitivity to environmental cues suggest that they play a central role in the tolerance/immunity decision. How DC respond to such stimuli in vivo, however, is poorly understood. Study of peripheral DC is difficult due to their low frequency, and their isolation induces changes in gene expression that may mimic those occurring in inflammatory conditions. In contrast, by collecting DC actively trafficking in lymph we can follow the complete migratory path of DC from peripheral tissues to draining LN. These DC are near physiological and are likely to be transducing information that will determine the outcome of naive T cell activation.
We have shown previously that i.v. LPS increases DC traffic in intestinal lymph in a TNF--dependent manner (23). In the current study we have examined the effects of i.v. LPS on the surface phenotype, migration, and tissue distribution of DC from lamina propria and PP, intestinal lymph, and MLN and compared the effects on intestinal DC with those on DC in spleen and exiting the liver.
Steady-state lamina propria contains many DC, mostly immature, in that MHC class II is largely intracellular. Intravenous LPS caused an almost complete loss of lamina propria DC by 12 h. That this loss represents migration rather than down-regulation of DC-associated Ags is suggested by the concomitant increase in DC migrating in intestinal lymph. By 24 h, a few DC are present, but steady-state DC numbers were not restored until 72 h, closely reflecting normal intestinal DC turnover (10). Proinflammatory stimuli have similar effects in other tissues; intratracheal bacteria, viruses, and LPS stimulate increased DC migration from rat respiratory tract epithelium (3, 29, 30) and murine epidermis, heart, and kidney (31). Intravenous LPS also empties the liver of DC (U. Yrlid and G. G. MacPherson, unpublished observations). In contrast to the effects of luminal stimuli on DC in the respiratory tract, LPS injected intraintestinally has no detectable effect on intestinal DC migration (G. G. MacPherson and C. Jenkins, unpublished observations)
In contrast to the immaturity of lamina propria DC, DC migrating in lymph are uniformly surface MHC class IIhigh (6). These DC left the intestine only moments before their collection on ice; thus, lamina propria DC must synchronously up-regulate surface MHC class II expression and undergo the changes required for intestinal exit. Lymph DC collected after LPS-stimulated migration are also surface MHC class IIhigh, suggesting that redistribution of MHC class II to the surface is an integral part of the phenotypic changes accompanying migration. That hepatic lymph DC behave identically shows that these changes are not dependent on intestinal commensal bacteria.
Lymph DC are surface MHC class IIhigh, but express little CD80 or CD86, showing that expression of MHC class II and costimulatory molecules need not be regulated coherently in vivo. Lymph DC are therefore similar to the semimature DC described by Lutz for in vitro bone marrow-derived DC (32, 33, 34). This phenotype is not restricted to lymph DC, because freshly isolated MLN DC showed a similar phenotype.
Intestinal lymph contains at least two distinct DC populations (6), and LPS administration stimulates increased migration of both populations. Steady-state DC outputs into hepatic and intestinal lymph are similar, but LPS stimulates a much greater increase in DC output into hepatic lymph. This could represent a larger population of DC in the liver or differences in the level of LPS/proinflammatory cytokines reaching and stimulating DC migration from the two tissues (35).
LPS and TNF- are potent in vitro stimuli for DC maturation/activation (36, 37, 38, 39). Because we have shown that LPS causes TNF--dependent changes in DC migration, we were surprised to find only minor increases in CD80 and CD86 expression on lymph or MLN DC at any time after LPS administration. This does not represent a general hyporesponsiveness of intestinal DC; lymph DC cultured with TNF- or GM-CSF showed marked up-regulation of surface CD80 and CD86, whereas high doses of LPS had no effect. Others have shown that the phenotype of gut DC can be modulated by proinflammatory cytokines. PP DC cultured in vitro with TNF- showed phenotypic maturation and enhanced T cell stimulatory capacity (40), and DC isolated from the GALT of Flt3L-treated mice after i.p. IL-1 administration expressed increased surface CD80 and CD86 (41). That DC showed enhanced costimulatory molecule expression after culture in vitro with TNF-, but not when isolated ex vivo after LPS administration might be caused by levels of TNF- entering the gut after i.v. LPS being much lower than the doses we used for in vitro culture. Furthermore, after i.v. LPS administration, migration, but not phenotypic maturation, of lymph DC is observed, and this may reflect dissociation of signaling pathways in DC or differences in the levels of proinflammatory cytokines required. Others have suggested that DC migration and phenotypic maturation may be independently regulated events (42).
In contrast to lymph DC, i.v. LPS caused a marked increase in costimulatory molecule expression on spleen DC. These differential effects could relate to DC’s peripheral microenvironments. The intestine is continually exposed to LPS from commensal organisms, and this may induce a refractory state in intestinal DC, whereas splenic DC would be unaffected. However, liver lymph DC show properties similar to those of intestinal DC, although we have been unable to detect significant levels of LPS in portal venous blood (U. Yrlid and G. G. MacPherson, unpublished observations).
Intravenous LPS induces major changes in the distribution of DC subsets in secondary lymphoid organs. This study is the first to describe such changes in rat DC subsets under proinflammatory conditions. In the steady-state, DC are differentially distributed in PP, MLN, and spleen. CD4high/SIRPhigh DC are found in the interfollicular traffic areas of the MLN cortex and in a narrow layer underlying the follicles. In spleen, they localize to the outer periarteriolar lymphoid sheath, where marginal metallophils are present. CD4low/SIRPlow DC, however, are found in large numbers throughout TCAs as classical interdigitating DC. It is unlikely that CD4low/SIRPlow DC derive from CD4high/SIRPhigh DC that have down-regulated expression of these molecules upon entry into the TCAs because 1) down-regulation does not occur in culture; and 2) after LPS administration, SIRPhigh DC are seen in TCAs. It remains to be determined why SIRPhigh DC are excluded from the TCAs of lymphoid tissues and what changes are associated with their entry after LPS administration.
The appearance of SIRPhigh DC in TCAs of MLN and spleen following i.v. LPS administration is most likely due to alterations in their migration, rather than to up-regulation of the SIRP molecule by SIRPlow DC already in the TCA. SIRPlow and SIRPhigh DC represent distinct subsets; both are detectable in normal intestine, and we have shown previously that both subsets appear in intestinal lymph with similar kinetics, indicating that one is not the precursor of the other (6). If proinflammatory stimuli were directly stimulating SIRP expression by SIRPlow DC already in TCAs, then we might expect the frequency of SIRPlow DC to decrease. In fact, an increased frequency of SIRPlow DC was observed in MLN and splenic TCAs after LPS administration. Furthermore, similar frequencies of SIRPhigh DC are found surrounding splenic TCAs before LPS and within TCAs after LPS administration. This implies a direct migration of SIRPhigh DC from one site to the other. We cannot induce SIRP expression on SIRPlow lymph DC in vitro by proinflammatory stimuli; however, these experiments are hampered by the poor survival of SIRPlow DC in culture (6).
The steady-state segregation of SIRPlow and SIRPhigh DC in rat secondary lymphoid tissues and migration of SIRPhigh DC into TCAs after LPS stimulation are analogous to the behavior of DC subsets in other species. Bovine lymph DC are subdivided by expression of a SIRP homologue, MyD-1. MyD-1+ cells localize to the cortex of skin-draining LN and splenic red pulp and marginal zone, but were found in TCAs after infection (43). Furthermore, our observations fit closely with those of murine CD8–/+ subsets. In murine PP and spleen, CD8+ DC are constitutively in TCAs, whereas CD8– DC are excluded. After microbial challenge, however (LPS, soluble toxoplasma Ag, or bacteria), CD8– DC migrate into TCAs (44, 45, 46, 47). We have other functional evidence to suggest that SIRPlow and SIRPhigh DC are the rat equivalents of murine CD8+ and CD8– DC, respectively (U. Yrlid and G. G. MacPherson, manuscript in preparation).
In complete contrast to the effects in MLN and spleen, i.v. LPS administration had no detectable effect on the number or distribution of DC subsets in rat PP. This lack of response is probably not due to an inherent lack of mobility of PP DC, because others have reported modulation of PP DC migration in response to microbial stimulation in rodents (47, 48, 49). The differences may relate to DC maturity, because, in contrast to lamina propria DC, most PP DC are mature (surface MHC class IIhigh). Maturity could determine sensitivity to TNF- or other mediators released in response to LPS. Alternatively, because we do not know the source of the TNF-, it could be that PP DC are shielded from it. The differences may have physiological significance, in that PP DC are more likely to contact commensal bacterial LPS under nonpathological conditions (via M cell transport) than are lamina propria DC, and it would be important that PP DC not have proinflammatory responses to commensal bacteria.
Current hypotheses suggest that whether Ag induces active immunity or tolerance depends on DC activation status. Our results suggest an alternative or complementary mechanism. We suggest that under steady-state conditions, SIRPlow DC are the primary DC that interact directly with T cells in secondary lymphoid organs and that these DC may be specialized to induce tolerance. This is supported by the findings that SIRPlow DC in rats and cattle are weaker stimulators of CD4+ and CD8+ T cells, that SIRPlow DC in the rat intestine acquire and transport remnants of apoptotic epithelial cells to the TCAs of MLN under steady-state conditions, and that SIRPlow (but not SIRPhigh DC) are constitutively found in the TCAs of rat secondary lymphoid tissue. That SIRPhigh DC are more potent T cell stimulators, but are normally excluded from contact with T cells, suggests that they may be involved in the generation of active T cell responses and are able to override the activity of SIRPlow DC when they migrate into TCAs under proinflammatory conditions. The functional differences between the subsets cannot be explained by differential expression of MHC and costimulatory molecules (6, 50), and the molecules conferring these differences have not been identified as yet.
Acknowledgments
We thank Simon Milling and Andrew Lucas for their critical review of the manuscript.
Footnotes
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 E.L.T. was supported by a Medical Research Council Research Studentship and the E. P. Abraham Trust. U.Y. is supported by a Wellcome Trust Traveling Research Fellowship and the E.P. Abraham Trust.
2 Current address: Edward Jenner Institute for Vaccine Research, Compton, Berkshire RG20 7NN, U.K.
3 Address correspondence and reprint requests to Dr. G. Gordon MacPherson, Sir William Dunn School of Pathology, South Parks Road, Oxford, U.K. OX1 3RE. E-mail address: gordon.macpherson@path.ox.ac.uk
4 Abbreviations used in this paper: LN, lymph node; 7AAD, 7-aminoactinomycin; DC, dendritic cell; CLN, cervical LN; CoeLNX, coeliac lymphadenectomy; Flt3L, Fms-like receptor tyrosine kinase 3 ligand; MLN, mesenteric LN; MLNX, mesenteric lymphadenectomy; PP, Peyer’s patch; SIRP, signal regulatory protein ; TCA, T cell area.
Received for publication June 1, 2004. Accepted for publication November 9, 2004.
References
Banchereau, J., F. Briere, C. Caux, J. Davoust, S. Lebecque, Y. J. Liu, B. Pulendran, K. Palucka. 2000. Immunobiology of dendritic cells. Annu. Rev. Immunol. 18:767.
MacPherson, G. G., N. Kushnir, M. Wykes. 1999. Dendritic cells, B cells and the regulation of antibody synthesis. Immunol. Rev. 172:325.
McWilliam, A. S., S. Napoli, A. M. Marsh, F. L. Pemper, D. J. Nelson, C. L. Pimm, P. A. Stumbles, T. N. Wells, P. G. Holt. 1996. Dendritic cells are recruited into the airway epithelium during the inflammatory response to a broad spectrum of stimuli. J. Exp. Med. 184:2429.
Stumbles, P. A., D. H. Strickland, C. L. Pimm, S. F. Proksch, A. M. Marsh, A. S. McWilliam, A. Bosco, I. Tobagus, J. A. Thomas, S. Napoli, et al 2001. Regulation of dendritic cell recruitment into resting and inflamed airway epithelium: use of alternative chemokine receptors as a function of inducing stimulus. J. Immunol. 167:228.
Randolph, G. J., K. Inaba, D. F. Robbiani, R. M. Steinman, W. A. Muller. 1999. Differentiation of phagocytic monocytes into lymph node dendritic cells in vivo. Immunity 11:753
Liu, L., M. Zhang, C. Jenkins, G. G. MacPherson. 1998. Dendritic cell heterogeneity in vivo: two functionally different dendritic cell populations in rat intestinal lymph can be distinguished by CD4 expression. J. Immunol. 161:1146.
Huang, F. P., N. Platt, M. Wykes, J. R. Major, T. J. Powell, C. D. Jenkins, G. G. MacPherson. 2000. A discrete subpopulation of dendritic cells transports apoptotic intestinal epithelial cells to T cell areas of mesenteric lymph nodes. J. Exp. Med. 191:435.
Hawiger, D., K. Inaba, Y. Dorsett, M. Guo, K. Mahnke, M. Rivera, J. V. Ravetch, R. M. Steinman, M. C. Nussenzweig. 2001. Dendritic cells induce peripheral T cell unresponsiveness under steady state conditions in vivo. J. Exp. Med. 194:769.
Steinman, R. M., D. Hawiger, K. Liu, L. Bonifaz, D. Bonnyay, K. Mahnke, T. Iyoda, J. Ravetch, M. Dhodapkar, K. Inaba, et al 2003. Dendritic cell function in vivo during the steady state: a role in peripheral tolerance. Ann. NY Acad. Sci. 987:15
Pugh, C. W., G. G. MacPherson, H. W. Steer. 1983. Characterization of nonlymphoid cells derived from rat peripheral lymph. J. Exp. Med. 157:1758.
Matsuno, K., T. Ezaki, S. Kudo, Y. Uehara. 1996. A life stage of particle-laden rat dendritic cells in vivo: their terminal division, active phagocytosis, and translocation from the liver to the draining lymph. J. Exp. Med. 183:1865.
Kudo, S., K. Matsuno, T. Ezaki, M. Ogawa. 1997. A novel migration pathway for rat dendritic cells from the blood: hepatic sinusoids-lymph translocation. J. Exp. Med. 185:777.
MacPherson, G. G., S. Fossum, B. Harrison. 1989. Properties of lymph-borne (veiled) dendritic cells in culture. II. Expression of the IL-2 receptor: role of GM-CSF. Immunology 68:108.
MacPherson, G. G.. 1989. Properties of lymph-borne (veiled) dendritic cells in culture. I. Modulation of phenotype, survival and function: partial dependence on GM-CSF. Immunology 68:102.
Liu, L. M., G. G. MacPherson. 1991. Lymph-borne (veiled) dendritic cells can acquire and present intestinally administered antigens. Immunology 73:281.
Turnbull, E., G. MacPherson. 2001. Immunobiology of dendritic cells in the rat. Immunol. Rev. 184:58
Trinite, B., C. Voisine, H. Yagita, R. Josien. 2000. A subset of cytolytic dendritic cells in rat. J. Immunol. 165:4202.
Voisine, C., F. X. Hubert, B. Trinite, M. Heslan, R. Josien. 2002. Two phenotypically distinct subsets of spleen dendritic cells in rats exhibit different cytokine production and T cell stimulatory activity. J. Immunol. 169:2284.
Ferguson, T. A., J. Herndon, B. Elzey, T. S. Griffith, S. Schoenberger, D. R. Green. 2002. Uptake of apoptotic antigen-coupled cells by lymphoid dendritic cells and cross-priming of CD8+ T cells produce active immune unresponsiveness. J. Immunol. 168:5589.
Iyoda, T., S. Shimoyama, K. Liu, Y. Omatsu, Y. Akiyama, Y. Maeda, K. Takahara, R. M. Steinman, K. Inaba. 2002. The CD8+ dendritic cell subset selectively endocytoses dying cells in culture and in vivo. J. Exp. Med. 195:1289.
Liu, K., T. Iyoda, M. Saternus, Y. Kimura, K. Inaba, R. M. Steinman. 2002. Immune tolerance after delivery of dying cells to dendritic cells in situ. J. Exp. Med. 196:1091.
Liu, L. M., G. G. MacPherson. 1995. Antigen processing: cultured lymph-borne dendritic cells can process and present native protein antigens. Immunology 84:241
MacPherson, G. G., C. D. Jenkins, M. J. Stein, C. Edwards. 1995. Endotoxin-mediated dendritic cell release from the intestine: characterization of released dendritic cells and TNF dependence. J. Immunol. 154:1317.
Matsuno, K., S. Kudo, T. Ezaki, K. Miyakawa. 1995. Isolation of dendritic cells in the rat liver lymph. Transplantation 60:765.
Brenan, M., M. Puklavec. 1992. The MRC OX-62 antigen: a useful marker in the purification of rat veiled cells with the biochemical properties of an integrin. J. Exp. Med. 175:1457.
Robinson, A. P., T. M. White, D. W. Mason. 1986. Macrophage heterogeneity in the rat as delineated by two monolonal antibodies MRC OX-41 and MRC OX-42 the latter recognizing completement receptor type 3. Immunology 57:239.
Adams, S., L. J. van der Laan, E. V. Wilson, C. Renardel de Lavalette, E. A. Dopp, C. D. Dijkstra, D. L. Simmons, T. K. van den Berg. 1998. Signal-regulatory protein is selectively expressed by myeloid and neuronal cells. J. Immunol. 161:1853.
Steinman, R. M., M. C. Nussenzweig. 2002. Avoiding horror autotoxicus: the importance of dendritic cells in peripheral T cell tolerance. Proc. Natl. Acad. Sci. USA 99:351.
McWilliam, A. S., D. Nelson, J. A. Thomas, P. G. Holt. 1994. Rapid dendritic cell recruitment is a hallmark of the acute inflammatory response at mucosal surfaces. J. Exp. Med. 179:1331.
McWilliam, A. S., A. M. Marsh, P. G. Holt. 1997. Inflammatory infiltration of the upper airway epithelium during Sendai virus infection: involvement of epithelial dendritic cells. J. Virol. 71:226.
Roake, J. A., A. S. Rao, P. J. Morris, C. P. Larsen, D. F. Hankins, J. M. Austyn. 1995. Dendritic cell loss from nonlymphoid tissues after systemic administration of lipopolysaccharide, tumor necrosis factor, and interleukin 1. J. Exp. Med. 181:2237.(Emma L. Turnbull, Ulf Yrl)
Dendritic cells (DC) present peripheral Ags to T cells in lymph nodes, but also influence their differentiation (tolerance/immunity, Th1/Th2). To investigate how peripheral conditions affect DC properties and might subsequently regulate T cell differentiation, we examined the effects of a potent DC-activating, TLR-4-mediated stimulus, LPS, on rat intestinal and hepatic DC in vivo. Steady-state rat intestinal and hepatic lymph DC are E2 integrinhigh (CD103) and include two subsets, signal regulatory protein (SIRP)hi/low, probably representing murine CD8–/+ DC. Steady-state lamina propria DC are immature; surface MHC class IIlow, but steady-state lymph DC are semimature, MHC class IIhigh, but CD80/86low. Intravenous LPS induced rapid lamina propria DC emigration and increased lymph DC traffic without altering SIRPhigh/SIRPlow proportions. CD80/86 expression on lymph or mesenteric node DC was not up-regulated after i.v. LPS. In contrast, i.v. LPS stimulated marked CD80/86 up-regulation on splenic DC. CD80/86 expression on intestinal lymph DC, however, was increased after in vitro culture with TNF- or GM-CSF, but not with up to 5 μg/ml LPS. Steady-state SIRPlow DC localized to T cell areas of mesenteric nodes, spleen, and Peyer’s patch, whereas SIRPhigh DC were excluded from these areas. Intravenous LPS stimulated rapid and abundant SIRPhigh DC accumulation in T cell areas of mesenteric nodes and spleen. In striking contrast, i.v. LPS had no effect on DC numbers or distribution in Peyer’s patches. Our results suggest that any explanation of switching between tolerance and immunity as well as involving changes in DC activation status must also take into account differential migration of DC subsets.
Introduction
Intestinal immune responses toward orally acquired Ags must be tightly regulated. Despite demonstrating a profound hyporesponsiveness to dietary and commensal bacterial Ags, the gut mounts protective responses toward pathogens. It is likely that conditions within the local gut microenvironment play a key role in determining the course of the immune response. A better understanding of the regulation of intestinal immune responses is crucial for developing treatment of inflammatory bowel disease and food allergies and for the design of effective intestinal vaccines.
Naive T cells, particularly CD4+ cells, are not thought to be precommitted to any particular differentiation pathway before their activation, but after activation, may differentiate along one of several distinct pathways. Exogenous stimuli (such as the properties of an Ag itself or the effects that pathogens/Ags exert on their local microenvironment) may influence their differentiation fate. Antigenic encounters primarily occur in the periphery; however, T cells are activated in draining lymph nodes (LN).4 To select those patterns of gene expression needed for appropriate differentiation, LN T cells need to be reactive to environmental conditions in the periphery. A central and only partially understood question is how information about these conditions is relayed from the periphery to the LN.
Dendritic cells (DC) acquire Ags in peripheral tissues and transport them to LN, where they present processed peptides to T cells and probably native Ags to B cells (1, 2). DC in general are highly sensitive to environmental cues, and local microenvironmental conditions affect DC phenotype and function (3, 4, 5). DC are therefore prime candidates for the transduction of information from the periphery to LN. Despite some current dogma, much evidence shows that DC are continually migrating in afferent lymph from tissues such as intestine, skin, and liver in the absence of overt inflammatory stimulation (6, 7). DC can exist in different states of maturation/activation that are reflected in the fate of the CD4+ T cells they activate. Under steady-state conditions, DC in LN have been shown to express low levels of costimulatory molecules and can induce tolerogenic or hyporesponsive T cells in vivo (8, 9). It is an attractive hypothesis that perturbations of the steady-state (e.g., inflammation) modulate the information delivered by the DC to the T cell, leading to an active, rather than a tolerogenic, immune response. Although this concept is supported by many studies that have investigated environmental effects on DC in vitro, whether similar effects occur in vivo is largely unknown. Studying environmental effects on DC in vivo is problematic, because the majority of techniques for DC isolation are themselves likely to induce phenotypic and functional changes in DC independent of true physiological or pathological stimuli in the local microenvironment. Of the various in vivo DC populations that can be studied, the DC most likely to be informative in relation to information transduction from the periphery are those migrating in afferent lymph. We have used the approach of mesenteric (MLNX) or celiac (CoeLNX) lymphadenectomy (10, 11, 12), followed by thoracic duct cannulation, to isolate DC migrating from the periphery. By collecting and manipulating these cells at 4°C, we can preserve the patterns of gene expression within migrating DC in vivo (6, 7, 10, 13, 14, 15 ; reviewed in Ref.16). This model thus provides the opportunity to directly identify changes in the properties of migrating DC that accompany perturbations in peripheral tissues and that may relate to differential activation of naive T cells.
Using this model we have shown previously that two phenotypically and functionally distinct subsets of DC migrate in rat intestinal lymph (both E2 integrinhigh), which can be distinguished because one subset coexpresses CD4 and high levels of signal regulatory protein (SIRP) (6). The same two subsets have been identified in rat spleen (17, 18), but how these phenotypically similar DC relate functionally is not yet fully clear. In both lymph and spleen, SIRPhigh DC are more potent than SIRPlow DC in the activation of allogeneic CD4+ and CD8+ T cells, naive Ag-specific CD4+ T cells in vivo, and sensitized Ag-specific CD4+ T cells in vitro (6, 17, 18). In addition, SIRPlow lymph DC selectively transport remnants of apoptotic intestinal epithelial cells to the T cell areas (TCAs) of mesenteric LNs (MLN) (7). This selective uptake of apoptotic material is a function shared by murine CD8+ DC and may indicate that the SIRPlow DC is the rat homologue involved in the induction of peripheral tolerance (19, 20, 21).
We have described in detail the properties of DC migrating in lymph from the intestine under steady-state conditions (6, 7, 10, 13, 14, 15, 22) and have also shown that i.v. LPS administration caused a substantial increase in the output of DC in intestinal lymph and that this effect can be partially blocked by an anti-TNF- Ab (23). In this study we have examined the effects of i.v. LPS on the properties of intestinally derived DC subsets, with reference to DC in MLN, liver-draining lymph, and spleen for comparison to gain insight into how such changes might affect the quality of the immune response. Our results show that i.v. LPS induces major changes in DC migration from the gut in the absence of significant changes to DC surface phenotype. We also show that proinflammatory stimulation is associated with the appearance of the previously excluded SIRPhigh DC subset in TCAs of MLN and spleen. We suggest that these alterations in migration may relate to the regulation of the switch between tolerance and active immunity.
Materials and Methods
Rats and surgical procedures
Rats (PVG (RT1c)) were bred and maintained under specific pathogen-free conditions in the Sir William Dunn School of Pathology (Oxford, U.K.) and were males between 12 and 16 wk of age. All procedures were approved by and conducted in accordance with Home Office guidelines. MLNX and thoracic duct cannulation were performed as previously described (10). CoeLNX was conducted as described by Matsuno et al. (24). Cannulations always involved multiple rats to ensure sufficient cell numbers for experimentation.
Expansion of DC numbers in vivo using Flt3 ligand
To boost DC numbers for in vitro studies, rats were injected s.c. daily for 8 days with 15 μg of recombinant human Fms-like receptor tyrosine kinase 3 ligand (Flt3L) in 200 μl of sterile PBS. Flt3L was provided by Immunex.
Reagents
All cell culture was performed in RPMI 1640 medium supplemented with 10% FCS, 50 μM 2-ME, 100 μg/ml penicillin, 50 U/ml streptomycin, and 100 μg/ml L-glutamine (all from Invitrogen Life Technologies). Recombinant rat TNF- and recombinant rat GM-CSF were purchased from R&D Systems and were used at a final concentration of 50 ng/ml for in vitro cell stimulation. Salmonella typhimurium LPS and 7-aminoactinomycin D (7AAD) were purchased from Sigma-Aldrich. FITC-, Cy3-, and Cy5-conjugated tyramide used for immunofluorescence were obtained from NEN. Normal rat, mouse, and donkey sera were obtained from Serotec. OX62 MACS beads were purchased from Miltenyi Biotec. Goat anti-mouse IgG Dynabeads were obtained from Dynal Biotech U.K. Cy5 for Ab conjugation was purchased from Amersham Biosciences.
Antibodies
Monoclonal Abs. mAbs were either neat tissue culture supernatants (donated by M. Puklavec, Sir William Dunn School of Pathology) or commercially available purified IgG. All mAbs were raised in mice unless otherwise stated. Unconjugated mAbs were anti-human factor I, IgG1 isotype control (OX21), anti-human HLA class I, IgG2a isotype control (W6/32), anti-MHC class II (OX6), anti-CD8 (OX8), anti-Ig -chain (OX12), anti-RT-1Ac MHC class I (OX27), anti-CD4 (OX38), anti-SIRP (OX41), anti-CD11b (OX42), and anti-E2 integrin (OX62). Anti-CD11c (8A2), anti-sialoadhesin (ED3), and anti-TCR (R73) mAbs were purchased from Serotec. Anti-CD80 (3H5) and anti-CD86 (24F) were provided by Dr. H. Yagita (Juntendo University School of Medicine, Tokyo, Japan).
Conjugated mAbs. FITC- and PerCP-conjugated anti-MHC class II (OX6), FITC-conjugated anti-CD4 (OX38), and FITC-conjugated hamster anti-mouse/rat CD40 (HM40-3) were obtained from BD Pharmingen. PE-conjugated anti-E2 integrin (OX62) was purchased from Serotec. PE-conjugated CD80 (3H5) and CD86 (24F) mAbs were obtained from Biocarta (Oxford, U.K.). FITC- and Cy5-conjugated OX62 and OX41 and HRP-conjugated OX41 mAbs were made in-house using a peroxidase labeling kit according to the manufacturer’s instructions (Roche).
Secondary Abs. FITC-, Cy3-, Cy5-, and HRP-conjugated donkey anti-mouse IgG Abs were obtained from Jackson ImmunoResearch Laboratories. Texas Red-conjugated goat anti-mouse IgG/IgM/IgA Ab was purchased from Southern Biotechnology Associates. PE-conjugated goat anti-mouse IgG was obtained from Sigma-Aldrich.
Primary cell preparations
Intestinal and hepatic pseudoafferent lymph DC. Thoracic duct lymph collected from lymphadenectomized rats was passed through a sterile 70-μm pore size cell strainer (BD Biosciences), and RBC were lysed using ammonium chloride solution. Lymph DC were typically enriched within total lymph cells by density gradient centrifugation. Total lymph cells were resuspended in PBS/2% FCS at a concentration of 2 x 107 cells/ml, and 108 cells were overlaid onto 2 ml of Nycoprep 1.068 medium (Nycomed). After centrifugation at 2000 rpm for 20 min at room temperature, the collected interface cells typically consisted of 15–30% DC. In some experiments DC were enriched to 20–40% by depletion of T and B cells. Total lymph cells were labeled using OX12 (anti-Ig) and OX52 (pan T cell) mAbs, followed by goat anti-mouse IgG Dynabeads. Labeled cells were then magnetically removed. Highly purified lymph DC were obtained using OX62-MACS beads and positive selection using AUTOMACS according to the manufacturer’s instructions.
Spleen, MLN, and cervical LN (CLN) DC. Spleens, MLN, and CLN were teased apart using watchmaker’s forceps into prewarmed RPMI 1640 medium containing 0.5 mg/ml collagenase D and 0.5 mg/ml DNase (Roche). Tissues were digested for 45 min at 37°C with periodic agitation. Undigested stromal material was removed by passage over a 70-μm pore size filter before RBC lysis using ammonium chloride solution. DC were enriched within splenocytes or MLN cells by magnetic selection using OX62-MACS beads.
MLR
Lymph was collected from MLNX PVG (RT1c) rats over 16 h after i.v. injection of PBS or 50 μg of LPS, and DC were enriched by density gradient centrifugation. Lymph DC were irradiated and then titrated against 2 x 105 responder CLN cells from DA (RT1a) rats. Cultures were performed in quadruplicate at 37°C for 3 days, after which, T cell proliferation was measured by thymidine incorporation and expressed as cpm. Background proliferation counts of responder cells alone were subtracted from proliferation in cultures containing DC.
Flow cytometry
All Ab and serum dilutions were made in PBA (PBS/1% BSA/10 mM sodium azide). All Ab incubation steps were conducted for 30 min over ice, and excess unbound Ab was removed by three washes in PBA. Cells were first blocked with 10% normal rat serum to block potential FcR binding. Abs were neat tissue culture supernatants or purified IgG diluted to 1–10 μg/ml. Relevant isotype-matched control Abs were included. Unconjugated primary Abs were detected using FITC- or PE-conjugated anti-mouse Abs. Where appropriate, 10 μg/ml 7AAD was added during the final minute of immunolabeling to label dead cells. Cells were fixed using 4% paraformaldehyde in PBS. Fluorescence was recorded using a FACSCalibur (BD Biosciences), and data were analyzed using CellQuest software (BD Biosciences), excluding dead cells.
Tissue immunofluorescence
Fresh tissues were frozen in Tissue-Tek OCT compound (Sakura Finetek Europe) over isopentane on dry ice. Sections were cut at 5 μm, dried, and stored at –20°C until use. After thawing, sections were fixed for 10 min in cold 100% ethanol and rinsed in PBS. Nonspecific protein binding was minimized by blocking for 30 min at room temperature with PBS/3% BSA before immunolabeling. Abs were neat tissue culture supernatants or purified IgG, typically diluted to 1–10 μg/ml in PBS/3% BSA. Appropriate isotype-matched negative control Abs were included in all immunostaining procedures to ensure specificity of staining. Ab incubations were conducted in a moist chamber at room temperature for 2 h or overnight at 4°C. Excess unbound Ab was washed off using three changes of PBS. In multicolor immunolabeling procedures, cross-reaction between anti-mouse secondary detection Ab and subsequently added directly conjugated mouse mAbs was prevented by blocking with 10% normal mouse serum for 20 min at room temperature. Sections were mounted in water-soluble Vectashield (Vector Laboratories), and coverslips were secured using clear nail varnish. Sections were stored in the dark to minimize photobleaching.
HRP/tyramide amplification of immunofluorescence
Unconjugated mAbs giving a weak signal when detected using a standard fluorophore-conjugated secondary Ab were detected using HRP-conjugated secondary mAb and were developed using fluorophore-conjugated tyramide. Endogenous tissue peroxidase was first quenched within fixed tissue sections for 30 min at 37°C using 0.7 U/ml glucose oxidase enzyme (Sigma-Aldrich) in prewarmed 0.1 M phosphate buffer containing 25 mM D-glucose and 1 mM sodium azide. Sections were then rinsed in PBS, preblocked, and immunolabeled as described above. Tyramide development of HRP was conducted for 15–20 min at room temperature according to the manufacturer’s instructions (NEN).
Image acquisition
Tissue immunofluorescence was recorded using an MRC 1024 laser scanning confocal microscope (Bio-Rad) or an Axioplan epifluorescence microscope (Carl Zeiss). For the confocal microscope, digital images were collected using COMOS software (Bio-Rad); for the epifluorescence microscope, images were obtained using a Spot camera and Spot Advanced software (Diagnostic Instruments). Single-label fluorescence sections were observed alongside multilabeled sections to ensure no bleed-through of signal from one channel to another. Digital images were processed and overlaid using Photoshop software (Adobe Systems). Figures show representative images of many sections studied using tissues derived from a number of different rats.
Results
In situ distribution of DC in the steady-state small intestine
DC were identified in cryosections of rat ileum by immunolabeling for E2 integrin, CD11c, or MHC class II Ags. In Peyer’s patches (PP), large, irregular E2 integrin+ cells were detected in the subepithelial dome region underlying the follicle-associated epithelium and in the interfollicular TCAs between B cell follicles (Fig. 1a). These cells were confirmed to be DC by double-labeling for CD11c and MHC class II (Fig. 1, c and e). Large, irregular E2 integrin+ and CD11c+ cells were also distributed throughout the lamina propria (Fig. 1b). These cells intensely coexpressed intracellular MHC class II, but were Ig negative, indicating that the MHC class II was unlikely to have been derived from the ingestion of apoptotic B cells (data not shown). In the lamina propria, E2 integrin was also expressed by MHC class II– intraepithelial lymphocytes as previously described (25).
FIGURE 1. Detection of DC within the steady-state rat ileum. Cryosections of ileum immunolabeled for DC-associated Ags in PP and lamina propria (LP). a and b, E2 integrin immunolabeling (green) of the subepithelial dome region (SED), interfollicular regions (IFR), and B cell follicles (B) of PP and LP (magnification: a, x100; b, x200). c–e, Double labeling for MHC class II (green) and CD11c (red) in the subepithelial dome (c, x200), interfollicular region of PP (e, x400), and lamina propria (d, x630). f and h, Double labeling for E2 integrin (red) and MHC class II (green) in lamina propria and intestinal epithelial cell (IEC) layer (f, x630). g, i, and j, Double labeling for MHC class II (green) and CD86 (red) in subepithelial dome of PP (g, x400), interfollicular region of PP (i, x400) and lamina propria (j, x200).
The cellular distribution of MHC class II differed markedly between PP and lamina propria. In lamina propria, although some MHC class II was surface expressed, the bulk appeared granular and intracellular (characteristic of retention within intracytoplasmic vesicles; Fig. 1, d, f, and h). In contrast, DC of the subepithelial dome and interfollicular regions of PP showed surface expression of MHC class II that colocalized well with E2 integrin or CD11c expression (Fig. 1, c and e). Most large, irregular cells in the subepithelial dome and interfollicular regions of PP (Fig. 1, g and i, respectively) coexpressed MHC class II and CD86, but lamina propria DC expressing granular, cytoplasmic MHC class II were CD86– (Fig. 1j). That lamina propria DC expressed intracellular MHC class II and lacked CD86 expression suggests that they are more developmentally immature than PP DC.
Intravenous LPS stimulates DC migration from the lamina propria
We have previously shown that i.v. LPS stimulates an increased traffic of DC in lymph draining the small intestine (23). To relate this to events occurring in the intestine, cryosections of ileum from rats that had been injected i.v. with either 50 μg of LPS or PBS 6, 12, 24, 48, or 72 h previously were labeled with OX62 (anti-E2 integrin) mAb. Compared with tissue from normal rats, PBS administration had no effect on the relative frequency and distribution of lamina propria E2 integrin+ cells (represented as steady-state in Fig. 2a). Intravenous LPS caused a time-dependent loss of DC from the lamina propria, detectable by 6 h (not shown), and by 12 h, virtually no DC were present in the lamina propria (Fig. 2c). By 24 h, small numbers of DC were present (Fig. 2e), and steady-state numbers were reached by 72 h (Fig. 2g). CD11c immunolabeling gave similar results (Fig. 2, b, d, f, and h). Similar results were obtained using an anti-MHC class II mAb (data not shown). E2 integrin+/CD11c– intraepithelial lymphocytes were detectable at all time points studied after PBS or LPS administration. Thus, i.v. LPS administration leads to a significant, but only transient, loss of DC from lamina propria.
FIGURE 2. Intravenous LPS administration leads to a loss of DC immunolabeling within the lamina propria of the small intestine. Cryosections of ileum from rats i.v. injected with PBS or 50 μg of LPS and killed at various times. Sections were immunolabeled for E2 integrin (a, c, e, and g) or CD11c (b, d, f, and h). Intestinal epithelial cells (IEC) and lamina propria (LP) are indicated. All panels show x630 magnification.
Intravenous LPS stimulates increased migration of DC in lymph draining both intestine and liver
We have previously shown that i.v. LPS administration is associated with an increased traffic of DC in lymph draining the intestine of MLNX rats (23). To determine whether the increase in output of DC into lymph was true of other organs, we investigated the effects of i.v. LPS on the frequency and phenotype of DC trafficking in pseudoafferent lymph from CoeLNX rats. The steady-state output of DC into liver-draining lymph was less (85%) than that into intestinal lymph. There was a selective increase in DC traffic from both organs, peaking between 6 and 12 h after i.v. LPS (Fig. 3a). Maximum output from the intestine increased by 10- to 20-fold, whereas output from the liver increased by 60- to 70-fold. In both cases, the increase in DC output from the periphery was transient, and DC traffic had returned to steady-state levels by 24 h. A subsequent injection of LPS given 24 h after the first, resulted in only a minor transient increase in lymph DC output, consistent with the lamina propria being depleted of DC (Fig. 3b).
FIGURE 3. Increased DC frequency in intestinal and hepatic lymph after i.v. LPS injection. Thoracic duct lymph from MLNX and CoeLNX rats collected 0–6, 6–12, and 12–22 h under steady-state conditions or after i.v administration of 50 μg of LPS. a, DC output after i.v. LPS administration, expressed as the fold increase relative to steady-state output over the same time. Absolute DC numbers trafficking in lymph during each collection period were determined by multiplying the percent proportion of DC (identified by E2 integrin expression by flow cytometry) by the total number of lymph cells. Results shown are representative of one of three similar experiments. b, Total number of intestinal lymph DC collected at intervals over 48 h. LPS (50 μg) was injected at 0 and 24 h, and the output of DC during each interval was determined. Similar results were obtained in two independent experiments. c, Four-color flow cytometric analysis of DC draining the intestine (MLNX) and liver (CoeLNX) using mAbs to E2 integrin, MHC class II, SIRP, and CD4. The four lower dot plots show lymph DC subset proportions before (control) and 6–12 h after LPS (+LPS) administration using the gates shown in two upper dot plots. d, Output of CD4low/SIRPlow and CD4high/SIRPhigh DC subsets draining the intestine (MLNX) and liver (CoeLNX) after i.v. LPS administration, expressed as the fold increase relative to steady-state output over the same time. Similar results were obtained in two independent experiments.
We have previously shown that intestinal pseudoafferent lymph contains two populations of DC, distinguishable by their expression of CD4 and SIRP (6). DC were identified by their high E2 integrin and MHC class II expression, and the proportions of CD4high/SIRPhigh and CD4low/SIRPlow DC subsets contained within steady-state intestinal and hepatic pseudoafferent intestinal lymph were determined. DC were detectable in nonenriched MLNX and CoeLNX lymph, representing 0.3% of total lymph cells (Fig. 3c, upper panels). CD4high/SIRPhigh lymph DC represented the major subset of DC trafficking in both intestinal and hepatic pseudoafferent lymph under steady-state conditions, accounting for between 60 and 70% of total DC (Fig. 3c, lower panels). The number and relative proportions of CD4high/SIRPhigh and CD4low/SIRPlow DC remained constant over a period of at least 44 h after cannulation in control rats, showing that surgery did not stimulate differential changes in DC subset migration from the gut or liver (data not shown).
To determine whether LPS administration altered the relative proportions of migrating DC subsets, rats were injected i.v. with 50 μg of LPS, and hepatic or intestinal pseudoafferent lymph was collected at intervals over 22 h. In gut-draining lymph, the output of both subsets increased transiently in parallel, showing that LPS challenge did not stimulate a preferential increase in one particular subset. In liver-draining lymph, there was a more pronounced increase in the output of CD4high/SIRPhigh DC compared with CD4low/SIRPlow DC (Fig. 3d). Thus, i.v. LPS administration resulted in a transient increase in lymph DC traffic draining both intestine and liver. SIRPhigh/low DC proportions were unchanged in lymph draining the intestine after LPS, whereas SIRPhigh DC proportions increased in liver-draining lymph.
Intravenous LPS administration increases DC frequency in mesenteric nodes
Given that i.v. LPS administration led to a transient loss of DC immunolabeling in the lamina propria and increased DC traffic in lymph draining the intestine, we predicted that this would be reflected in an increased migration of DC into the draining MLN. Sections of MLN from rats injected i.v. 0, 6, 12, or 24 h earlier with PBS or 50 μg of LPS were double-labeled for MHC class II and TCR. PBS administration had no effect on DC distribution compared with tissue from noninjected rats at all time points studied (data not shown). After LPS administration, DC frequency in the TCA of MLN was slightly increased by 6 h (data not shown), and by 12 h, it was several-fold increased compared with that in sections from rats injected i.v. 12 h earlier with PBS (compare Fig. 4, a and b). By 24 h, DC frequency was reduced compared with that at 12 h, but remained elevated compared with steady-state levels (data not shown). Similar staining patterns were obtained using anti-E2 integrin and anti-CD11c mAbs (data not shown). The increase in DC numbers in the MLN correlates directly with the loss of DC from the lamina propria 12 h after LPS administration and with the peak output of DC from gut into lymph.
FIGURE 4. Intravenous LPS induces an accumulation of DC in the draining MLN. a and b, Cryosections of MLN from rats injected i.v. 12 h earlier with PBS or 50 μg of LPS were immunolabeled for TCR (red) and MHC class II (green), TCAs (magnification, x200). c, MLN DC enriched by OX62-MACS beads immunolabeled for E2 integrin and MHC class II. The absolute number of DC recovered at each time point was expressed as a proportion of the starting MLN cell number. Values represent the mean and SD of three independent experiments for each time point. Data were analyzed using unpaired one-tailed Student’s t test.
To quantitate changes in DC numbers in MLN, nodes taken at intervals after LPS administration were digested with collagenase, and DC were enriched by positive selection using OX62-MACS beads. DC were identified by flow cytometry by coexpression of E2 integrin and MHC class II. Absolute numbers of DC were expressed as a percentage of the total MLN cell numbers (Fig. 4c). Absolute DC numbers were increased above steady-state by 6 h, and a 4-fold increase in MLN DC proportions was observed 12 h after LPS administration (p = 0.024), consistent with our immunohistochemical data. Significantly increased numbers of MLN DC 12 h after i.v. LPS administration is consistent with the loss of lamina propria DC and the peak output of DC in lymph draining the intestine.
Effects of i.v. LPS on the anatomical distribution of DC subsets in secondary lymphoid organs
DC subsets were localized within secondary lymphoid tissue by immunolabeling for DC-associated markers together with the OX41 mAb. In addition to delineating rat DC subsets, the SIRP Ag detected by the OX41 mAb has been described on a subpopulation of macrophages (26, 27). We therefore paid particular attention to ensure that our immunolabeling procedure would allow us to distinguish between these cell types. Although SIRPlow DC were detectable by flow cytometry, they were not detectable by immunohistochemistry and thus appear SIRP–. We retain the term SIRPlow for consistency.
MLN. In cryosections from normal rats, double labeling for E2 integrin and SIRP identified both SIRPhigh and SIRPlow DC subsets in the interfollicular traffic areas of the cortex (Fig. 5a), consistent with the migration of both subsets in intestinal lymph and their entry into MLN via the afferent lymphatics. A few SIRPhigh and SIRPlow DC were also found in the subfollicular area of the superficial cortex, just beneath the lymph follicle. In contrast, DC in TCAs were almost exclusively SIRPlow, and SIRPhigh DC were rarely seen (Fig. 5b). The exclusion of SIRPhigh DC from the MLN TCA was confirmed by triple immunolabeling for MHC class II, TCR, and SIRP. MHC class II+/SIRPhigh and MHC class II+/SIRPlow DC were found in the cortical interfollicular traffic areas (Fig. 5c), whereas frequent SIRPlow DC, but few SIRPhigh DC, were detectable within the TCA (Fig. 5d). Thus, SIRPhigh DC are excluded from the steady-state MLN TCA.
FIGURE 5. Intravenous LPS administration is associated with the appearance of SIRPhigh DC in the MLN TCA. a and b, Cryosections of MLN from control rats were immunolabeled for E2 integrin (green) and SIRP (red) to localize DC subsets in the MLN cortex (a) and TCA (b); SIRPlow DC (shown in green) and SIRPhigh DC (shown in yellow). IFTA, Cortical interfollicular traffic areas; B, B cell follicles; T, T cell area. c and d, Immunolabeling for TCR (red), MHC class II (green), and SIRP (blue) on cryosections of control MLN. MHC class II+/SIRPlow DC (shown as large, irregular green cells) and MHC class II+/SIRPhigh DC (light blue). e and f, MLN cryosections from rats injected 12 h earlier with PBS (e) or 50 μg of LPS (f) immunolabeled for CD11c (red), MHC class II (green), and SIRP (blue) to localize DC subsets in the MLN TCA. CD11c+/MHC class II+/SIRPlow DC are shown in yellow, and CD11c+/MHC class II+/SIRPhigh DC are shown in white. Magnification for all panels, x200.
To determine whether the DC subset distribution in MLN changed after i.v LPS administration, cryosections from rats injected 6, 12, or 24 h earlier with 50 μg of LPS or PBS were labeled for MHC class II, CD11c, and SIRP. Tissue from normal rats or rats injected with PBS at all of the time points studied showed the presence of SIRPlow DC, but few SIRPhigh TCA DC (Fig. 5e). Six hours after administration of LPS, SIRPhigh DC were detected in the MLN (data not shown), and by 12 h significant numbers were present (Fig. 5f). The increase in SIRPhigh DC in the TCA 12 h after LPS challenge was accompanied by an increase in SIRPlow DC. Both subsets were still detectable in the TCA 24 h after LPS administration (data not shown).
PP. Cryosections from normal rats immunolabeled for E2 integrin and SIRP identified both SIRPlow and SIRPhigh DC in the subepithelial dome region underlying the follicle-associated epithelium (Fig. 6a), although there were relatively fewer SIRPhigh DC. SIRP+/E2 integrin– cells were also found at the subepithelial dome and in the interfollicular region, and these may be macrophages. As with the MLN, SIRP was undetectable on essentially all DC in the TCA of the interfollicular region of PP (Fig. 6b). However, in complete contrast to the results for MLN, i.v. LPS had no overt effect on either the relative number or the distribution of DC in PP. Both SIRPlow and SIRPhigh DC remained detectable in the subepithelial dome region at all time points studied after LPS (Fig. 6c), and SIRPhigh DC were rarely found in the PP TCA (Fig. 6d). Although SIRP-expressing cells were detectable in normal lamina propria using OX41 mAb, these cells did not coexpress E2 integrin or MHC class II. SIRPhigh DC were not detectable in the lamina propria after i.v. LPS administration at any of the time points studied (data not shown).
FIGURE 6. Intravenous LPS administration fails to modulate the relative frequency and distribution of rat PP DC subsets. Cryosections of PP from rats injected i.v. 12 h earlier with PBS (a and b) or 50 μg of LPS (c and d) immunolabeled for E2 integrin (green) and SIRP (red). SIRPlow DC are shown in green, and SIRPhigh DC are shown in yellow. SED, subepithelial dome region; B, B cell follicle; IFR, interfollicular regions. Magnification: a–c, x200; d, x400.
Spleen. Cryosections of rat spleen labeled using OX41 mAb showed that SIRPhigh cells were abundant within the steady-state spleen, particularly in the red pulp (Fig. 7a). These cells are probably macrophages, because they failed to express CD11c or E2 integrin (data not shown). SIRPhigh cells were also detected in the outer periarteriolar lymphoid sheath, where they formed a tight cuff around the TCR+ TCAs (Fig. 7b). These SIRPhigh cells expressed MHC class II and CD11c (Fig. 7c), but did not coexpress sialoadhesin (data not shown), indicating that they were DC rather than macrophages. Although CD11c+, MHC class II+/SIRPlow DC were found in the splenic TCA under steady-state conditions, SIRPhigh DC were rarely found. The cuff of SIRPhigh DC was not continuous around B cell follicles.
FIGURE 7. SIRPhigh DC are excluded from the TCAs of the steady-state rat spleen. a–c, Cryosections of spleen from control rats: single stained for SIRP (a), double labeled for SIRP (green) and TCR (red; b), or triple labeled for MHC class II (green), CD11c (red), and SIRP (blue; c). CD11c+/MHC class II+/SIRPlow DCs are shown in yellow, and CD11c+/MHC class II+/SIRPhigh are shown in white. d and e, Cryosections from rats injected i.v. with PBS (d) or 50 μg of LPS (e) 6 h earlier immunolabeled for MHC class II (green), TCR (red), and SIRP (blue). MHC class II+/SIRPhigh DC are shown in light blue, and MHC class II+/SIRPlow DC are shown in green. RP, red pulp; MZ, marginal zone; B, B cell follicles; T, TCAs. Magnification: a–c, x200; d and e, x400.
Intravenous PBS administration had no effect on the distribution of SIRPlow and SIRPhigh DC subsets in splenic white pulp compared with tissue from noninjected rats at all time points studied (data not shown). In contrast, i.v. LPS administration led to a loss of MHC class II+/SIRPhigh DC immunolabeling on the outskirts of the TCA and the appearance of MHC class II+/SIRPhigh DC within the TCA. This effect was observed by 6 h after LPS administration (compare Fig. 7, d and e). Interestingly, the frequency of SIRPhigh DC surrounding TCAs under steady-state conditions was comparable to the frequency of SIRPhigh DC present in the TCA after LPS challenge, suggesting a direct migration of DC from the outer periarteriolar lymphoid sheath to TCA. The overall frequency of SIRPlow TCA DC was increased by 12 h after LPS injection, correlating with a loss of SIRPlow DC from the red pulp (data not shown). At 12 and 24 h after i.v. LPS, SIRPhigh DC remained detectable within the TCA (data not shown). These data show that i.v. LPS administration is associated with the appearance of the previously excluded SIRPhigh DC subset in the TCA of MLN and spleen, but not PP.
Effects of i.v. LPS administration on the surface phenotype of lymph, MLN, and splenic DC
Intestinal pseudoafferent lymph DC were collected under steady-state conditions or during the peak output after i.v. LPS administration, and their surface phenotype was analyzed by flow cytometry. Surprisingly, LPS had little effect on the expression of E2 integrin, CD11c, MHC class I, CD40, and CD11b by DC trafficking in lymph draining the intestine. MHC class II expression was reduced, and CD80 and CD86 levels were increased by only 1.5- to 2-fold. Similar results were obtained for hepatic lymph DC collected after i.v. LPS administration (data not shown). Steady-state and LPS-induced, intestinally derived lymph DC showed a similar potency in stimulating the proliferation of allogeneic T cells, consistent with their comparable expressions of molecules known to be important for T cell activation (Fig. 8a).
FIGURE 8. Surface phenotype comparison of steady-state and LPS-induced intestinal lymph DC: comparison with hepatic lymph DC, MLN DC, and spleen DC. a, Comparison of the allostimulatory potential of intestinally derived lymph DC from PBS- or LPS-injected rats. Equal numbers of irradiated, density enriched, intestinally derived lymph DC from PBS or LPS-injected PVG (RT1c) rats were cultured with 2 x 105 total responder cervical LN cells from DA (RT1a) rats and cultured for 3 days. T cell proliferation was measured by thymidine incorporation and expressed as cpm after subtracting the background proliferation of responder cells alone. Error bars represent the SD of the mean of quadruplicate cultures. Data shown are from one of three similar experiments. b, Intestinal and hepatic lymph was collected at intervals over 22 h under steady-state conditions or after i.v. LPS administration. DC were enriched by depleting T and B cells and were immunolabeled for E2 integrin, SIRP, CD4, and either CD80 or CD86. Similar results were obtained in two independent experiments. c, Enriched DC immunolabeled for E2 integrin, MHC class II, and CD80, CD86, or CD40. Intestinal and hepatic lymph DC were collected between 6 and 12 h after i.v PBS or LPS injection. MLN and spleen DC were isolated 12 h after i.v. PBS or LPS injection. Filled histograms, DC from PBS-injected rats; solid line, DC from LPS-injected rats. Data are representative of at least three independent experiments.
The increases in surface costimulatory molecule expression by LPS-induced lymph DC were not restricted to one subset in particular. Both CD4low/SIRPlow and CD4high/SIRPhigh intestinal and hepatic lymph DC showed transiently increased surface expression of CD80 and CD86, with DC collected after the peak output showing similar surface expression levels as steady-state DC (Fig. 8b).
We wanted to compare the effect of i.v. LPS administration on intestinal and hepatic lymph DC with that on DC isolated from secondary lymphoid tissue. Lymph DC were collected under steady-state conditions or during their peak output after i.v. LPS administration. MLN and splenic DC were isolated 12 h after i.v. LPS or PBS administration by collagenase digestion, density separation, and MACS enrichment using OX62 beads. DC (identified as E2 integrin+/MHC class II+ cells) were additionally labeled for costimulatory molecules and examined by flow cytometry (Fig. 8c). Interestingly, intestinally derived lymph DC and MLN DC expressed comparable levels of CD80 and CD86, showing that mere entry of DC into the MLN from lymph did not induce phenotypic maturation, and only minor increases in CD80 and CD86 expression were observed on MLN DC 12 h after i.v. LPS, consistent with the minimal effect of i.v. LPS on intestinal lymph DC phenotype. In contrast, splenic DC showed substantial up-regulation of surface costimulatory molecule expression, with 5-fold increases in both CD80 and CD86 and a 2- to 3-fold increase in CD40 expression 12 h after LPS administration.
These data suggest that the environmental conditions created after i.v. LPS administration are insufficient to stimulate substantial up-regulation of molecules critical for T cell activation on DC derived from gut and liver. In contrast, splenic DC were phenotypically matured in response to i.v. LPS administration. Thus, DC in different tissue sites respond differently to the same stimulus.
Modulation of lymph DC surface phenotype during in vitro culture with GM-CSF and TNF-, but not LPS
To ensure that DC derived from the gut were not refractory to stimulation in general, in vitro experiments were conducted to examine the ability of known inducers of DC maturation to stimulate intestinally derived lymph DC. To collect sufficient DC, rats were pretreated with human Flt-3 ligand, which induces a 5- to 10-fold increase in intestinal lymph DC output without causing detectable changes in phenotype or function (U. Yrlid and G. G. MacPherson, manuscript in preparation).
Purified intestinal lymph DC from Flt-3 ligand-treated rats were cultured in vitro for 12 or 24 h in culture medium alone or in medium supplemented with 50 ng/ml recombinant rat GM-CSF, 50 ng/ml recombinant rat TNF-, or doses of LPS up to 5 μg/ml. The DC surface phenotype was subsequently assessed by flow cytometry (Fig. 9). Compared with freshly isolated lymph DC, only minor increases (<2-fold) in surface expression of MHC class I, MHC class II, and CD40 molecules were seen after culture in medium alone, and the presence of GM-CSF, TNF-, or LPS had no additive effect (not shown). A 24-h culture in medium alone failed to stimulate changes in surface CD80 and CD86 expression; however, the presence of 50 ng/ml GM-CSF or TNF- stimulated a profound increase in surface expression of these costimulatory molecules (up to a log-fold increase in CD86 with GM-CSF after 24 h). Surprisingly, the presence of up to 5 μg/ml LPS during culture had no effect on CD80/CD86 expression. Therefore, intestinally derived lymph DC can be phenotypically matured, but the nature of the stimulus is critical.
FIGURE 9. In vitro modulation of intestinally derived lymph DC surface phenotype by GM-CSF and TNF-, but not by LPS. Intestinal lymph DC from steady-state Flt3L-pretreated rats were enriched to >90% purity by density gradient centrifugation and cultured at 37°C in 5% CO2 for 12 and 24 h at 106/ml in medium alone or in medium supplemented with 50 ng/ml GM-CSF, 50 ng/ml TNF-, or up to 5 μg/ml LPS. Cells were then labeled for anti-E2 integrin and CD80/CD86 and were analyzed by flow cytometry. Histograms compare surface expression levels of CD80 and CD86 on freshly isolated lymph DC (filled histograms), 12-h cultured lymph DC (dashed line), and 24-h cultured lymph DC (solid line). Data are representative of one of three similar experiments.
Discussion
Intestinal Ags may induce tolerance or immunity, probably depending on the presence or the absence of proinflammatory conditions (9, 28). DC migratory behavior and sensitivity to environmental cues suggest that they play a central role in the tolerance/immunity decision. How DC respond to such stimuli in vivo, however, is poorly understood. Study of peripheral DC is difficult due to their low frequency, and their isolation induces changes in gene expression that may mimic those occurring in inflammatory conditions. In contrast, by collecting DC actively trafficking in lymph we can follow the complete migratory path of DC from peripheral tissues to draining LN. These DC are near physiological and are likely to be transducing information that will determine the outcome of naive T cell activation.
We have shown previously that i.v. LPS increases DC traffic in intestinal lymph in a TNF--dependent manner (23). In the current study we have examined the effects of i.v. LPS on the surface phenotype, migration, and tissue distribution of DC from lamina propria and PP, intestinal lymph, and MLN and compared the effects on intestinal DC with those on DC in spleen and exiting the liver.
Steady-state lamina propria contains many DC, mostly immature, in that MHC class II is largely intracellular. Intravenous LPS caused an almost complete loss of lamina propria DC by 12 h. That this loss represents migration rather than down-regulation of DC-associated Ags is suggested by the concomitant increase in DC migrating in intestinal lymph. By 24 h, a few DC are present, but steady-state DC numbers were not restored until 72 h, closely reflecting normal intestinal DC turnover (10). Proinflammatory stimuli have similar effects in other tissues; intratracheal bacteria, viruses, and LPS stimulate increased DC migration from rat respiratory tract epithelium (3, 29, 30) and murine epidermis, heart, and kidney (31). Intravenous LPS also empties the liver of DC (U. Yrlid and G. G. MacPherson, unpublished observations). In contrast to the effects of luminal stimuli on DC in the respiratory tract, LPS injected intraintestinally has no detectable effect on intestinal DC migration (G. G. MacPherson and C. Jenkins, unpublished observations)
In contrast to the immaturity of lamina propria DC, DC migrating in lymph are uniformly surface MHC class IIhigh (6). These DC left the intestine only moments before their collection on ice; thus, lamina propria DC must synchronously up-regulate surface MHC class II expression and undergo the changes required for intestinal exit. Lymph DC collected after LPS-stimulated migration are also surface MHC class IIhigh, suggesting that redistribution of MHC class II to the surface is an integral part of the phenotypic changes accompanying migration. That hepatic lymph DC behave identically shows that these changes are not dependent on intestinal commensal bacteria.
Lymph DC are surface MHC class IIhigh, but express little CD80 or CD86, showing that expression of MHC class II and costimulatory molecules need not be regulated coherently in vivo. Lymph DC are therefore similar to the semimature DC described by Lutz for in vitro bone marrow-derived DC (32, 33, 34). This phenotype is not restricted to lymph DC, because freshly isolated MLN DC showed a similar phenotype.
Intestinal lymph contains at least two distinct DC populations (6), and LPS administration stimulates increased migration of both populations. Steady-state DC outputs into hepatic and intestinal lymph are similar, but LPS stimulates a much greater increase in DC output into hepatic lymph. This could represent a larger population of DC in the liver or differences in the level of LPS/proinflammatory cytokines reaching and stimulating DC migration from the two tissues (35).
LPS and TNF- are potent in vitro stimuli for DC maturation/activation (36, 37, 38, 39). Because we have shown that LPS causes TNF--dependent changes in DC migration, we were surprised to find only minor increases in CD80 and CD86 expression on lymph or MLN DC at any time after LPS administration. This does not represent a general hyporesponsiveness of intestinal DC; lymph DC cultured with TNF- or GM-CSF showed marked up-regulation of surface CD80 and CD86, whereas high doses of LPS had no effect. Others have shown that the phenotype of gut DC can be modulated by proinflammatory cytokines. PP DC cultured in vitro with TNF- showed phenotypic maturation and enhanced T cell stimulatory capacity (40), and DC isolated from the GALT of Flt3L-treated mice after i.p. IL-1 administration expressed increased surface CD80 and CD86 (41). That DC showed enhanced costimulatory molecule expression after culture in vitro with TNF-, but not when isolated ex vivo after LPS administration might be caused by levels of TNF- entering the gut after i.v. LPS being much lower than the doses we used for in vitro culture. Furthermore, after i.v. LPS administration, migration, but not phenotypic maturation, of lymph DC is observed, and this may reflect dissociation of signaling pathways in DC or differences in the levels of proinflammatory cytokines required. Others have suggested that DC migration and phenotypic maturation may be independently regulated events (42).
In contrast to lymph DC, i.v. LPS caused a marked increase in costimulatory molecule expression on spleen DC. These differential effects could relate to DC’s peripheral microenvironments. The intestine is continually exposed to LPS from commensal organisms, and this may induce a refractory state in intestinal DC, whereas splenic DC would be unaffected. However, liver lymph DC show properties similar to those of intestinal DC, although we have been unable to detect significant levels of LPS in portal venous blood (U. Yrlid and G. G. MacPherson, unpublished observations).
Intravenous LPS induces major changes in the distribution of DC subsets in secondary lymphoid organs. This study is the first to describe such changes in rat DC subsets under proinflammatory conditions. In the steady-state, DC are differentially distributed in PP, MLN, and spleen. CD4high/SIRPhigh DC are found in the interfollicular traffic areas of the MLN cortex and in a narrow layer underlying the follicles. In spleen, they localize to the outer periarteriolar lymphoid sheath, where marginal metallophils are present. CD4low/SIRPlow DC, however, are found in large numbers throughout TCAs as classical interdigitating DC. It is unlikely that CD4low/SIRPlow DC derive from CD4high/SIRPhigh DC that have down-regulated expression of these molecules upon entry into the TCAs because 1) down-regulation does not occur in culture; and 2) after LPS administration, SIRPhigh DC are seen in TCAs. It remains to be determined why SIRPhigh DC are excluded from the TCAs of lymphoid tissues and what changes are associated with their entry after LPS administration.
The appearance of SIRPhigh DC in TCAs of MLN and spleen following i.v. LPS administration is most likely due to alterations in their migration, rather than to up-regulation of the SIRP molecule by SIRPlow DC already in the TCA. SIRPlow and SIRPhigh DC represent distinct subsets; both are detectable in normal intestine, and we have shown previously that both subsets appear in intestinal lymph with similar kinetics, indicating that one is not the precursor of the other (6). If proinflammatory stimuli were directly stimulating SIRP expression by SIRPlow DC already in TCAs, then we might expect the frequency of SIRPlow DC to decrease. In fact, an increased frequency of SIRPlow DC was observed in MLN and splenic TCAs after LPS administration. Furthermore, similar frequencies of SIRPhigh DC are found surrounding splenic TCAs before LPS and within TCAs after LPS administration. This implies a direct migration of SIRPhigh DC from one site to the other. We cannot induce SIRP expression on SIRPlow lymph DC in vitro by proinflammatory stimuli; however, these experiments are hampered by the poor survival of SIRPlow DC in culture (6).
The steady-state segregation of SIRPlow and SIRPhigh DC in rat secondary lymphoid tissues and migration of SIRPhigh DC into TCAs after LPS stimulation are analogous to the behavior of DC subsets in other species. Bovine lymph DC are subdivided by expression of a SIRP homologue, MyD-1. MyD-1+ cells localize to the cortex of skin-draining LN and splenic red pulp and marginal zone, but were found in TCAs after infection (43). Furthermore, our observations fit closely with those of murine CD8–/+ subsets. In murine PP and spleen, CD8+ DC are constitutively in TCAs, whereas CD8– DC are excluded. After microbial challenge, however (LPS, soluble toxoplasma Ag, or bacteria), CD8– DC migrate into TCAs (44, 45, 46, 47). We have other functional evidence to suggest that SIRPlow and SIRPhigh DC are the rat equivalents of murine CD8+ and CD8– DC, respectively (U. Yrlid and G. G. MacPherson, manuscript in preparation).
In complete contrast to the effects in MLN and spleen, i.v. LPS administration had no detectable effect on the number or distribution of DC subsets in rat PP. This lack of response is probably not due to an inherent lack of mobility of PP DC, because others have reported modulation of PP DC migration in response to microbial stimulation in rodents (47, 48, 49). The differences may relate to DC maturity, because, in contrast to lamina propria DC, most PP DC are mature (surface MHC class IIhigh). Maturity could determine sensitivity to TNF- or other mediators released in response to LPS. Alternatively, because we do not know the source of the TNF-, it could be that PP DC are shielded from it. The differences may have physiological significance, in that PP DC are more likely to contact commensal bacterial LPS under nonpathological conditions (via M cell transport) than are lamina propria DC, and it would be important that PP DC not have proinflammatory responses to commensal bacteria.
Current hypotheses suggest that whether Ag induces active immunity or tolerance depends on DC activation status. Our results suggest an alternative or complementary mechanism. We suggest that under steady-state conditions, SIRPlow DC are the primary DC that interact directly with T cells in secondary lymphoid organs and that these DC may be specialized to induce tolerance. This is supported by the findings that SIRPlow DC in rats and cattle are weaker stimulators of CD4+ and CD8+ T cells, that SIRPlow DC in the rat intestine acquire and transport remnants of apoptotic epithelial cells to the TCAs of MLN under steady-state conditions, and that SIRPlow (but not SIRPhigh DC) are constitutively found in the TCAs of rat secondary lymphoid tissue. That SIRPhigh DC are more potent T cell stimulators, but are normally excluded from contact with T cells, suggests that they may be involved in the generation of active T cell responses and are able to override the activity of SIRPlow DC when they migrate into TCAs under proinflammatory conditions. The functional differences between the subsets cannot be explained by differential expression of MHC and costimulatory molecules (6, 50), and the molecules conferring these differences have not been identified as yet.
Acknowledgments
We thank Simon Milling and Andrew Lucas for their critical review of the manuscript.
Footnotes
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 E.L.T. was supported by a Medical Research Council Research Studentship and the E. P. Abraham Trust. U.Y. is supported by a Wellcome Trust Traveling Research Fellowship and the E.P. Abraham Trust.
2 Current address: Edward Jenner Institute for Vaccine Research, Compton, Berkshire RG20 7NN, U.K.
3 Address correspondence and reprint requests to Dr. G. Gordon MacPherson, Sir William Dunn School of Pathology, South Parks Road, Oxford, U.K. OX1 3RE. E-mail address: gordon.macpherson@path.ox.ac.uk
4 Abbreviations used in this paper: LN, lymph node; 7AAD, 7-aminoactinomycin; DC, dendritic cell; CLN, cervical LN; CoeLNX, coeliac lymphadenectomy; Flt3L, Fms-like receptor tyrosine kinase 3 ligand; MLN, mesenteric LN; MLNX, mesenteric lymphadenectomy; PP, Peyer’s patch; SIRP, signal regulatory protein ; TCA, T cell area.
Received for publication June 1, 2004. Accepted for publication November 9, 2004.
References
Banchereau, J., F. Briere, C. Caux, J. Davoust, S. Lebecque, Y. J. Liu, B. Pulendran, K. Palucka. 2000. Immunobiology of dendritic cells. Annu. Rev. Immunol. 18:767.
MacPherson, G. G., N. Kushnir, M. Wykes. 1999. Dendritic cells, B cells and the regulation of antibody synthesis. Immunol. Rev. 172:325.
McWilliam, A. S., S. Napoli, A. M. Marsh, F. L. Pemper, D. J. Nelson, C. L. Pimm, P. A. Stumbles, T. N. Wells, P. G. Holt. 1996. Dendritic cells are recruited into the airway epithelium during the inflammatory response to a broad spectrum of stimuli. J. Exp. Med. 184:2429.
Stumbles, P. A., D. H. Strickland, C. L. Pimm, S. F. Proksch, A. M. Marsh, A. S. McWilliam, A. Bosco, I. Tobagus, J. A. Thomas, S. Napoli, et al 2001. Regulation of dendritic cell recruitment into resting and inflamed airway epithelium: use of alternative chemokine receptors as a function of inducing stimulus. J. Immunol. 167:228.
Randolph, G. J., K. Inaba, D. F. Robbiani, R. M. Steinman, W. A. Muller. 1999. Differentiation of phagocytic monocytes into lymph node dendritic cells in vivo. Immunity 11:753
Liu, L., M. Zhang, C. Jenkins, G. G. MacPherson. 1998. Dendritic cell heterogeneity in vivo: two functionally different dendritic cell populations in rat intestinal lymph can be distinguished by CD4 expression. J. Immunol. 161:1146.
Huang, F. P., N. Platt, M. Wykes, J. R. Major, T. J. Powell, C. D. Jenkins, G. G. MacPherson. 2000. A discrete subpopulation of dendritic cells transports apoptotic intestinal epithelial cells to T cell areas of mesenteric lymph nodes. J. Exp. Med. 191:435.
Hawiger, D., K. Inaba, Y. Dorsett, M. Guo, K. Mahnke, M. Rivera, J. V. Ravetch, R. M. Steinman, M. C. Nussenzweig. 2001. Dendritic cells induce peripheral T cell unresponsiveness under steady state conditions in vivo. J. Exp. Med. 194:769.
Steinman, R. M., D. Hawiger, K. Liu, L. Bonifaz, D. Bonnyay, K. Mahnke, T. Iyoda, J. Ravetch, M. Dhodapkar, K. Inaba, et al 2003. Dendritic cell function in vivo during the steady state: a role in peripheral tolerance. Ann. NY Acad. Sci. 987:15
Pugh, C. W., G. G. MacPherson, H. W. Steer. 1983. Characterization of nonlymphoid cells derived from rat peripheral lymph. J. Exp. Med. 157:1758.
Matsuno, K., T. Ezaki, S. Kudo, Y. Uehara. 1996. A life stage of particle-laden rat dendritic cells in vivo: their terminal division, active phagocytosis, and translocation from the liver to the draining lymph. J. Exp. Med. 183:1865.
Kudo, S., K. Matsuno, T. Ezaki, M. Ogawa. 1997. A novel migration pathway for rat dendritic cells from the blood: hepatic sinusoids-lymph translocation. J. Exp. Med. 185:777.
MacPherson, G. G., S. Fossum, B. Harrison. 1989. Properties of lymph-borne (veiled) dendritic cells in culture. II. Expression of the IL-2 receptor: role of GM-CSF. Immunology 68:108.
MacPherson, G. G.. 1989. Properties of lymph-borne (veiled) dendritic cells in culture. I. Modulation of phenotype, survival and function: partial dependence on GM-CSF. Immunology 68:102.
Liu, L. M., G. G. MacPherson. 1991. Lymph-borne (veiled) dendritic cells can acquire and present intestinally administered antigens. Immunology 73:281.
Turnbull, E., G. MacPherson. 2001. Immunobiology of dendritic cells in the rat. Immunol. Rev. 184:58
Trinite, B., C. Voisine, H. Yagita, R. Josien. 2000. A subset of cytolytic dendritic cells in rat. J. Immunol. 165:4202.
Voisine, C., F. X. Hubert, B. Trinite, M. Heslan, R. Josien. 2002. Two phenotypically distinct subsets of spleen dendritic cells in rats exhibit different cytokine production and T cell stimulatory activity. J. Immunol. 169:2284.
Ferguson, T. A., J. Herndon, B. Elzey, T. S. Griffith, S. Schoenberger, D. R. Green. 2002. Uptake of apoptotic antigen-coupled cells by lymphoid dendritic cells and cross-priming of CD8+ T cells produce active immune unresponsiveness. J. Immunol. 168:5589.
Iyoda, T., S. Shimoyama, K. Liu, Y. Omatsu, Y. Akiyama, Y. Maeda, K. Takahara, R. M. Steinman, K. Inaba. 2002. The CD8+ dendritic cell subset selectively endocytoses dying cells in culture and in vivo. J. Exp. Med. 195:1289.
Liu, K., T. Iyoda, M. Saternus, Y. Kimura, K. Inaba, R. M. Steinman. 2002. Immune tolerance after delivery of dying cells to dendritic cells in situ. J. Exp. Med. 196:1091.
Liu, L. M., G. G. MacPherson. 1995. Antigen processing: cultured lymph-borne dendritic cells can process and present native protein antigens. Immunology 84:241
MacPherson, G. G., C. D. Jenkins, M. J. Stein, C. Edwards. 1995. Endotoxin-mediated dendritic cell release from the intestine: characterization of released dendritic cells and TNF dependence. J. Immunol. 154:1317.
Matsuno, K., S. Kudo, T. Ezaki, K. Miyakawa. 1995. Isolation of dendritic cells in the rat liver lymph. Transplantation 60:765.
Brenan, M., M. Puklavec. 1992. The MRC OX-62 antigen: a useful marker in the purification of rat veiled cells with the biochemical properties of an integrin. J. Exp. Med. 175:1457.
Robinson, A. P., T. M. White, D. W. Mason. 1986. Macrophage heterogeneity in the rat as delineated by two monolonal antibodies MRC OX-41 and MRC OX-42 the latter recognizing completement receptor type 3. Immunology 57:239.
Adams, S., L. J. van der Laan, E. V. Wilson, C. Renardel de Lavalette, E. A. Dopp, C. D. Dijkstra, D. L. Simmons, T. K. van den Berg. 1998. Signal-regulatory protein is selectively expressed by myeloid and neuronal cells. J. Immunol. 161:1853.
Steinman, R. M., M. C. Nussenzweig. 2002. Avoiding horror autotoxicus: the importance of dendritic cells in peripheral T cell tolerance. Proc. Natl. Acad. Sci. USA 99:351.
McWilliam, A. S., D. Nelson, J. A. Thomas, P. G. Holt. 1994. Rapid dendritic cell recruitment is a hallmark of the acute inflammatory response at mucosal surfaces. J. Exp. Med. 179:1331.
McWilliam, A. S., A. M. Marsh, P. G. Holt. 1997. Inflammatory infiltration of the upper airway epithelium during Sendai virus infection: involvement of epithelial dendritic cells. J. Virol. 71:226.
Roake, J. A., A. S. Rao, P. J. Morris, C. P. Larsen, D. F. Hankins, J. M. Austyn. 1995. Dendritic cell loss from nonlymphoid tissues after systemic administration of lipopolysaccharide, tumor necrosis factor, and interleukin 1. J. Exp. Med. 181:2237.(Emma L. Turnbull, Ulf Yrl)