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Address correspondence to Stephen W. Chensue, M.D., Ph.D., Department of Pathology and Laboratory Medicine, VA Ann Arbor Healthcare System, 2215 Fuller Rd., Ann Arbor, MI 48105.
Department of Pathology and Laboratory Medicine, VA Ann Arbor Healthcare System, Ann Arbor, MichiganDepartment of Pathology, University of Michigan Health System, Ann Arbor, Michigan
Cigarette smoke (CS)–induced lung injury involves innate immune responses. The activation of innate effector cells is thought to require cross talk with dendritic cells (DCs) and macrophages, but the mediators of interaction are unknown. One candidate, CC chemokine receptor 4 (CCR4), is expressed by innate and adaptive effector cells, and its ligands are produced by DCs and macrophages. Using flow cytometry and confocal microscopy, we defined innate responses of lung myeloid DCs, macrophages, and conventional natural killer (NK) cells in mice exposed to CS over 4 days and examined the contribution of CCR4 using CCR4 knockout (CCR4−/−) mice. CS affected populations differently, causing an increase in F4/80+ macrophages, a reduction in parenchymal CD11c+CD11b+CD103− DCs, but no effect on mucosal CD11c+CD11b−CD103+ DCs. CS also induced a population of primed/activated CD69+ NK cells and bronchoepithelial expression of the stress-related NKG2D receptor–activating protein, retinoic acid early transcript 1. CS-exposed CCR4−/− mice were similar to controls regarding effects on DCs and macrophages but displayed substantially impaired NK priming/activation and reduced expression of transcripts for interferon gamma, CXCL10, and retinoic acid early transcript 1. Quantitative confocal microscopy revealed that lungs of CS-exposed CCR4−/− mice had significantly reduced contacts of NK cells with CD11c+ cells. These findings demonstrate that acute CS exposure elicits NK cell responses and suggest that CCR4 promotes NK cell priming/activation by mediating contacts with sentinel cells in the lung.
In recent years, the relationship between cigarette smoke (CS) and immunity has been subject to extensive investigation. Tobacco abuse can be viewed as a model of repeated lung injury with superimposed toxic and pharmacologic effects that elicit and modify pulmonary immune responses. Various studies suggest that CS-related chronic inflammatory conditions, such as chronic obstructive pulmonary disease, involve innate and adaptive immune responses, but much controversy remains as to how chronic lung injury is established and sustained.
Innate immunity in the lung is mediated by multiple elements, including the mucociliary system, epithelial-derived defensins, phagocytic leukocytes, dendritic cells (DCs), and lymphoid populations, such as conventional natural killer (NK) cells, NK T cells, and γ/δ T cells. Initiation of innate immune responses involves cell receptors that recognize microbial- or damage-associated molecular patterns. In particular, sentinel cells, such as DCs and macrophages, are pivotal not only in innate recognition but also in regulating immune responses through interactions with effector cells, such as NK cells.
Conventional NK cells, traditionally considered innate responders, represent an important component of the pulmonary immune response, mounting rapid and potent responses to infection, injury, and neoplasms. However, NK cells are now known to participate as innate and memory effectors possibly contributing to chronic inflammation.
but the mechanisms of NK cell maturation, priming, and activation are not fully understood.
In a model of pulmonary mycobacterial infection, we recently demonstrated that CC chemokine receptor 4 (CCR4) and its ligands contributed to early innate resistance to infection, which was related to NK cell activation.
Moreover, DCs and macrophages are known sources of CCR4 ligands. The priming and activation of NK cells is thought to involve cross talk with sentinel cells, such as DCs.
Based on these findings, we postulated that CCR4 might contribute to the CS-elicited priming/activation of NK cells by promoting contacts between effector and sentinel cells in the lung.
Using flow cytometric and confocal stereologic approaches, we carefully defined antigen-presenting cell and NK cell responses after acute CS exposure and then tested the effect of CCR4 gene knockout. The present findings support a model in which CCR4 promotes contacts between sentinel and CCR4+ effector cells, providing a means for rapid organ-based effector priming/activation that in the setting of CS exposure could contribute to chronic lung injury.
Materials and Methods
Mice
Eight- to 12-week-old male and female C57BL/6 mice were obtained from The Jackson Laboratory (Bar Harbor, ME). Mice lacking the CCR4 gene (CCR4−/−) were from Tularik Inc (South San Francisco, CA) and were generated as previously described and bred onto a C57BL/6 background.
Knockout status was confirmed by RT-PCR analysis using gene-specific primers and probes. Mice were maintained under specific pathogen-free conditions and were provided with food and water ad libitum in a University Committee on Use and Care of Animals–approved facility. All the studies were approved by the University of Michigan Committee on Use and Care of Animals.
CS Exposure
Smoke from standardized 3R4F research cigarettes (University of Kentucky, Lexington, KY) with the filters removed was generated by a TE-2 cigarette smoking machine (Teague Enterprises, Woodland, CA). This device is set up to provide a mixture of mainstream and sidestream smoke. Animals were exposed on 4 consecutive days for 1 hour per day in a 54-L glass and Plexiglas whole-body exposure chamber with an electric fan for chamber mixing in standard mouse caging units with wire cage tops, with water available ad libitum. For a gravimetric measure of total suspended particulate matter, high-retention glass fiber filters (Pall Corp., East Hills, NY) were weighed before exposure and were placed in line at the exhaust port for the duration of the exposure. Filters were weighed with correction for room humidity, and the means ± SD concentration of particulates collected during a 1-hour exposure was 10.87 ± 2.17 mg. Control animals were housed in an identical chamber exposed to room air with no smoke. Animals were sacrificed on day 5 for analysis.
Lung and Draining Lymph Node Collection and Processing
Smoke- and air-exposed lungs were perfused with 10 mL of cold RPMI 1640 medium (Sigma-Aldrich, St. Louis, MO), excised, and collected in 20 mL of cold RPMI 1640 medium. Lung lobes were dissected and distributed for concurrent studies as follows: right lung for flow cytometry, left lower lobe for confocal microscopy, and left upper lobe for transcript expression analysis. For flow cytometry, lung lobes were blended individually in a Waring blender on low speed for 30 seconds, and entire lung homogenate was centrifuged. Lungs were dispersed by incubation at 37°C for 10 minutes in 3 mL of RPMI medium containing 5% (v/v) fetal bovine serum (Atlanta Biologicals, Norcross, GA) and 150 U/mL of type IV collagenase (Sigma-Aldrich). Digested lungs were resuspended in 10 mL of RPMI 1640 medium and were sieved through a 40-μm cell strainer (BD Biosciences, San Jose, CA) and subsequently washed three times by centrifugation before staining. Draining lymph nodes and spleens were collected, mechanically dispersed by teasing, filtered, and washed by centrifugation before flow cytometric analysis.
Flow Cytometry
Cells were suspended in 2 mL of Dulbecco's PBS (DPBS) (Sigma-Aldrich) staining buffer with 2% (v/v) fetal bovine serum for staining. Cells were first incubated with 10 μg/mL of TruStain fcX anti-CD16/32 block (BioLegend, San Diego, CA) for 10 minutes at 4°C. Cell samples were then equally divided, and 100-μL antibody (Ab) cocktails were added using 0.5 μL of Ab per 100 μL of staining buffer for either the lymphoid marker panel [CD103-PE, B220-PE Texas Red (BD Pharmingen, San Jose, CA), CD4-Pacific Blue, CD8-APC-Cy7, CD3e-PerCP-Cy5.5, CD44-AF700, NK1.1-AF647, CD62L-AF488, CD25-PE-Cy7, CD69-PE-Cy5 (BioLegend)] or the DC marker panel [CD103-PE, B220-PE Texas Red (BD Pharmingen), CD4-Pacific Blue, CD8-APC-Cy7, F4/80-PerCP-Cy5.5, CD45.2 PE-Cy7, CD11c-PE-Cy5, CD11b-AF488 (BioLegend)]. For CCR4 detection, samples were stained with anti-mouse CCR4(CD194)–PE (BioLegend), with parallel control tubes treated with phosphatidylethanolamine-labeled isotype Ab. Preparations were incubated for 30 minutes at 4°C and were washed with 2 mL of staining buffer. Cells were then sieved through a 40-μm preseparation filter (Miltenyi Biotec Inc., Auburn, CA) before flow analysis.
A FACScan LSRII 12-color flow cytometer (BD Biosciences, San Diego, CA) and FlowJo software version 7.5.5 (Tree Star Inc., Ashland, OR) were used for data acquisition and analysis. Cells were suspended to 1 × 106/mL, and then a cocktail of fluorochrome-labeled antimurine surface marker Abs for either lymphoid or DC analyses was added as described previously herein. Target populations were gated from a minimum of a 100,000- to 1 million-cell interrogation by forward and 90° light scatter analysis, and then fluorescence intensity was gauged and compared with controls stained with isotype-matched control IgG. Individual fluorophores were detected directly in the appropriate channels.
Gene Expression Analysis
Semiquantitative RT-PCR was used to assess targeted transcript expression. After dissection, left upper lung lobes were immediately submersed in 7 mL of RNAlater (Life Technologies, Foster City, CA) and were frozen at −40°C for later isolation. Total RNA from individual lung lobes was isolated using the RNeasy mini kit (Qiagen Inc., Germantown, MD) according to the manufacturer's instructions. RNA samples were reverse transcribed using the ImProm-II reverse transcription system (Promega Corp., Madison, WI) according to the manufacturer's protocol. The cDNA samples were analyzed for gene expression levels of RAET-1, IL-10, IL-15, IL-18, CCL2, CCL4, CCL17, CCL20, CL22, CXCL9, CXCL10, CXCL12, and IFNγ using commercially available minor groove binder–labeled primer/probe sets (Life Technologies). Rodent glyceraldehyde-3-phosphate dehydrogenase was used as the internal control. The 7500 real-time PCR system (Life Technologies) was used for detection using the preprogrammed thermoprofile for amplification.
Confocal Microscopy
Excised lung lobes were postinflated with OCT (Sakura Finetek USA Inc., Torrance, CA) diluted 1:1 in PBS and were rapid frozen in OCT-filled mounting molds on dry ice. Frozen microtome sections (20 μm thick) were mounted on adhesive slides and fixed in ice-cold acetone. Sections were rehydrated in DPBS for 3 minutes and then were blocked with 10 μg/mL of TruStain fcX anti-CD16/32 (BioLegend) in DPBS for 10 minutes at room temperature. Slides were decanted, and primary Abs were added at a 1:100 dilution (v/v) in DPBS. Mouse specific Abs included rabbit anti-(CCL22) (Abcam, Cambridge, MA), anti–CD11c-AF488, anti–CD11c-PE, anti–MHC(IA)-Brilliant Violet 421 (BioLegend), and goat anti-NKp46/NCR1 (R&D Systems, Minneapolis, MN) or anti–RAET-1 (pan-specific)-PE (R&D Systems) and were incubated for 1 hour in the dark at room temperature. Anti-goat AF555 or anti-rabbit–fluorescein isothiocyanate secondary Abs (Life Technologies) were used at 1:200 (v/v) in DPBS, and slides were incubated for 30 minutes in the dark at room temperature. Slides were washed three times for 3 minutes each time with DPBS, and then ProLong gold mounting fluid (Life Technologies) was used for mounting coverslips. For nuclear staining, standard DAPI solution was used.
Confocal analysis was performed using a spinning disk confocal microscope (Olympus America Inc., Center Valley, PA) with a digital CCD camera (Hamamatsu Photonics, Hamamatsu, Japan) for image capture and an arc lamp illumination source providing excitation wavelengths of 350 to 700 nm and three-color emission analyses. The acquired digital images were processed and analyzed using Stereo Investigator software version 9 (MBF Bioscience, Williston, VT) with the capacity to perform single-layer or stacked image analysis for stereologic image reconstruction and automated point counting of defined fluorescent foci. Fluorescent images of the different emission colors were overlaid to detect foci of coexpression.
For lung parenchymal analysis, a 400 × 400-μm image area was captured at ×200 magnification, with at least 10 fields sampled per mouse. After counting, values were normalized to number per square millimeter of parenchyma. For airway analysis, images of bronchi with a means ± SD diameter of 158 ± 53 μm were captured at ×200 magnification, and the lengths of the airways were measured. DCs were defined as large cells with cell processes and coexpression of high-threshold CD11c+ and MHCII (IA) fluorescence (Supplemental Figure S1). Numbers of bronchiole-associated MCHII+CD11c+ or NK cells (NKp46+) were counted and were normalized to number per millimeter length of mucosa. Portions of staining cytoplasm were not counted. For NK cell contact analysis, the total number of close contacts between NK cells and large MCHII+CD11c+ and large MCHII−CD11c+ cells was counted and normalized to total identified NK cells. Captured images were subjected to blinded evaluation (S.W.C., V.R.S., B.M.) for quantitative morphometric analysis.
Statistical Analysis
Student’s t-test was used for direct comparisons with a parallel control group. One-way analysis of variance with Tukey post hoc pairwise testing was used for multigroup analyses. P < 0.05 indicates significance.
Results
Short-Term CS Exposure Elicits Alterations in Lung DC and Macrophage Populations
For this assessment of the innate response to a 4-day CS exposure protocol, we characterized effects on DC and macrophage populations. The lung harbors two major populations of myeloid DCs (mDCs) with distinct phenotypes: CD11c+CD11b+CD103−MHChi and CD11c+CD11b−CD103+MHChi.
CD103 is the designation for the membrane integrin molecule αE, which is expressed as a heterodimer with β7. The αE/β7 integrin dimer binds to the epithelial cell adhesion protein E-cadherin. Hence, CD103 is associated with mucosal epithelial interactions and is thought to allow CD103+ DC intercalation with gut or lung epithelium. Using a panel of inclusion and exclusion marker Abs, these populations can be clearly identified by multicolor flow cytometry. Furthermore, consistent with the study by Sung et al,
confocal microscopy demonstrated that cells with morphologic and phenotypic characteristic of CD103+ DCs were largely restricted to bronchi and bronchioles, whereas CD11c+CD103−MHChi cells were mostly located in the parenchymal interstitium (Supplemental Figure S1).
Using these approaches, we examined the effect of 4-day CS exposure on mDC subpopulations and on lung macrophage populations. CS substantially decreased the overall proportion of CD11c+MHCIIhi in gated CD11c+ large mononuclear cells (Figure 1A). Analysis of mDCs showed that there was an absolute decrease in the number of CD11b+CD103− DCs but no effect on CD11b−CD103+ DCs (Figure 1, B and C). In contrast, CD11c+F4/80+MHCIIlo macrophages increased by twofold (Figure 1D). This observation was supported by confocal microscopic analysis, which showed a decrease in the density of CD11c+MHCIIhi cells in the parenchyma but no effect on mucosa-associated populations (Figure 1, E and F).
Figure 1Effect of acute CS exposure on lung mDCs and macrophages. Lung lobes of C57BL/6 CS-exposed and air-exposed control mice were dissected and distributed for parallel flow cytometric and confocal microscopic analyses as described in Materials and Methods. A: Percentages of gated CD11c+MHCIIhi mDCs were reduced in CS-exposed mice. B: Representative flow cytometric plots of CD11b+CD103− and CD11b−CD103+ DCs show skewed reduction of CD11b+ DCs in CS-exposed mice. C: Numbers of gated CD11b+CD103− and CD11b−CD103+ DCs show an absolute reduction in CD103− DCs. D: Numbers of gated F4/80+ macrophages show an absolute increase in macrophages in CS-exposed mice. E: Representative confocal images of lung parenchyma show reduced density of cells with coexpression of CD11c and MHCII in CS-exposed mice. Arrows indicate cells with coexpression in merged images. Original magnification, ×200. F: Quantitative confocal analysis of parenchyma- and mucosa-associated CD11c+MCHII+ cells. Black bars, smoke; gray bars, air. Bars are means ± SD; five mice per group. ∗P < 0.05.
These studies demonstrated that short-term CS exposure reduced lung parenchymal but not mucosal DCs. It was suspected that this was due mainly to emigration rather than down-regulation of MHCII or in situ death because flow cytometric analysis of draining lymph nodes from CS-exposed mice showed significant increases in CD103− mDC numbers compared with controls (Supplemental Figure S2). However, DC tracking studies are required for further confirmation.
Short-Term CS Exposure Elicits NK Cell Activation
Conventional NK cells are a major immune effector population in the lung, representing approximately 10% of lymphoid cells.
Differential activation of killer cells in the circulation and the lung: a study of current smoking status and chronic obstructive pulmonary disease (COPD).
reported that long-term (>8 weeks) CS exposure induced NK cell priming with enhanced activation on stimulation with viral-related pathogen-associated molecules or ILs (IL-12/IL-18). In view of this report, we examined lung NK cells (identified as CD45+NK1.1+CD3−B220−) after 4-day CS exposure and monitored endogenous NK priming/activation by expression of the CD69 marker. Flow cytometric analysis revealed a trend toward increased numbers of lung NK cells in CS-exposed compared with air-exposed mice with significantly increased proportions of CD69+NK cells (Figure 2). We also performed confocal analysis to locate and quantify NK cells using NKp46, which provides a good marker of conventional NK cells because it is expressed by only a minute fraction of NK T cells. In addition, we assessed the expression of retinoic acid early transcript 1 protein (RAET-1), a stress-induced protein ligand for the NKG2D-activating receptor expressed by NK and CD8 T cells that promote cytotoxicity. NK cells were detected in parenchymal and mucosal locations of CS-exposed mice (Figure 3, A and B). In addition, RAET-1 was notably increased in the airway epithelium of CS-exposed mice compared with air-exposed controls. The quantitative confocal analysis mirrored the flow cytometric findings, indicating a trend toward more NK cells in the lung parenchyma with a statistically significant increase in mucosa-associated NK cells (Figure 3G). Transcript analysis for RAET-1 was also performed, and this supported the immunofluorescent findings, showing a fourfold increase in transcript levels in CS-exposed mice (Figure 3H).
Figure 2Acute CS exposure elicits an NK cell response with enhanced activation. Multicolor flow cytometric analysis was performed on individually dispersed right lower lung lobes of C57BL/6 CS-exposed and air-exposed control mice as described in Materials and Methods. A: Absolute numbers of gated conventional NK cells. Inset: Representative dot plots. B: Expression of CD69 activation marker in all gated NK cells. Black bars, smoke; gray bars, air. Bars are means ± SD; five mice per group. ∗P < 0.05.
Figure 3Confocal microscopic localization of NK cells and RAET-1 expression in lungs of CS-exposed and air-exposed control mice. Lungs of C57BL/6 CS-exposed and control mice were snap frozen, cryosectioned, and subjected to immunofluorescent staining with confocal analysis for the indicated markers. A: Airway of CS-exposed lung showing mucosa-associated NKp46+ cells (arrows). B: Lobular parenchyma of CS-exposed lung showing NKp46+ cells (arrows). C: RAET-1 expression in airway mucosal epithelium. D–F: Parallel regions and magnifications of air-exposed control lungs. G: Quantitative confocal analysis of NKp46 positive cells in parenchymal and mucosal locations. NK cells were rarely observed in association with airways of air-exposed controls. H: RAET-1 transcript levels in CS-exposed and control lungs. Original magnification: ×200 (A, B, D, E); ×100 (C and F). Black bars, smoke; gray bars, air. Bars are means ± SD. ∗P < 0.05.
These results demonstrated that in addition to effects on mDCs and macrophages, short-term CS exposure induced a stress response in bronchoepithelium and elicited a population of primed/activated NK cells.
CCR4 Is Required for Optimal CS-Elicited NK Cell Activation
We previously reported that CCR4 was required for NK cell activation during the innate-phase response to Mycobacterium bovis infection.
We further postulated that CCR4 and its ligands might also participate in the CS-elicited NK cell response. CCR4 is among a group of chemokine receptors expressed by subpopulations of CD4+ memory T cells and NK cells.
To assess CCR4 cellular distribution, we stained lung and splenic lymphoid cells from unchallenged C57BL/6 mice for CCR4. Lung conventional NK cells harbored the greatest population of CCR4-expressing cells (Figure 4A). These cells were also largely killer cell lectin-like receptor G1 (KLRG1) positive, consistent with a mature phenotype (data not shown). CCR4 was also detected in a significant proportion of splenic NK cells, but this was less than among lung NK cells, suggesting that CCR4+ NK cells were concentrating in the peripheral organ. In accord with previous reports, CCR4+ cells were detected among CD44+CD4+ memory but not CD44−CD4+ naive T cells. The net percentage expression is shown in Figure 4B. Note that CCR4 was also not significantly expressed by CD8+ T-cell populations (data not shown).
Figure 4Subpopulations of conventional NK cells express CCR4. Lungs and spleens of unchallenged C57BL/6 mice were subjected to multicolor flow cytometric analysis to assess CCR4 expression in lymphoid populations. A: Representative flow cytometry plots; anti-CCR4 staining of gated lung and spleen CD3-NK1.1+ NK cells, lung and spleen CD44hiCD3+ CD4+ memory phenotype T cells, and splenic naive CD3+CD44lo CD4+ T cells and the corresponding isotype control Ab staining plots of the same populations. B: Percentage of CCR4+ cells among CD44hiCD4+ memory T cells, CD44loCD4+ naive T cells, and CD44hiNK1+ conventional NK cells. Bars are means ± SD; derived from three individual mice. Analysis of variance with Tukey pairwise postanalysis showed the greatest proportion of CCR4+ cells among gated lung NK cells compared with all other gated populations. ∗P < 0.05.
Reportedly, the ligands for CCR4, CCL17, and CCL22 are constitutively expressed by DCs and macrophages in many organs, including the lungs, suggesting a homeostatic role for this receptor.
When we measured transcript levels for CCL17 and CCL22 in lungs of CS- and air-exposed control mice, we observed no induction above constitutive levels (Figure 5). In addition, consistent with previous studies of lung CD11b+ and CD103+ DCs,
confocal staining of lungs for CCL22 revealed constitutive protein expression in large CD11c+ cells in the parenchymal interstitium and airway mucosa (Supplemental Figure S3).
Figure 5CCR4 ligand transcripts are constitutively expressed in lungs of C57BL/6 mice. Transcript levels for the CCR4 agonists CCL22 and CCL17 were measured in lungs of CS- and air-exposed control mice. Black bars, smoke; gray bars, air. Bars are means ± SD; four to five separate mice.
Having established a potential role for CCR4 and its ligands in the lung, we next compared the 4-day CS-elicited innate response in CCR4+/+ and CCR4−/− mice. There was no significant influence of CCR4 deficiency on F4/80+ macrophage and DC populations in CS-exposed mice (Figure 6A), indicating that this receptor was not required for lung positioning or migration of these cells. There were also no differences in the levels of MHCII antigen expression between wild-type and knockout mice (Figure 6A). However, as shown in Figure 6B, CCR4 deficiency did affect the NK cell response. CCR4+/+ and CCR4−/− CS-exposed mice had increased numbers of lung NK cells compared with air-exposed controls, which also had comparable baseline resident NK cell populations (data not shown), but CCR4−/− mice showed significantly impaired endogenous NK cell priming/activation as assessed by CD69 expression.
Figure 6Effect of CCR4 gene knockout on the innate pulmonary response to acute CS exposure. A: Numbers of macrophage (MP) and mDC populations in lungs of CS-exposed CCR4+/+ and CCR4−/− mice. Inset: Relative MCHII expression of these populations as mean fluorescent index (MFI). B: NK cell activation response in lungs of CS-exposed CCR4+/+ and CCR4−/− mice. C: Lung cytokine, chemokine, and RAET-1 transcript expression in lungs of CS-exposed CCR4+/+ and CCR4−/− mice. Transcript levels were determined by semiquantitative RT-PCR. Arrows indicate where transcript levels in CCR4−/− mice significantly deviated from CS-exposed controls. ND, no transcripts detected. D: Quantitative confocal analysis of NK cell localization in lobular parenchyma and airway mucosa of CS-exposed CCR4+/+ and CCR4−/− mice. Bars are means ± SD; four mice per group. ∗P < 0.05.
It was possible that the present findings were due to altered development of organ-homing KLRG1–positive NK cells in CCR4−/− mice. However, differences could not be attributed to this because wild-type (CCR4+/+) and knockout (CCR4−/−) mice had comparable mature cytolytic organ–homing CD27−KLGR1+ and noncytolytic lymphoid tissue–homing CD27+KLRG1− populations in the blood (Supplemental Figure S4). Hence, CCR4 was likely participating in organ-based NK cell migration or activation events.
To assess global functional changes in lungs, we measured transcript expression for a selection of cytokines and chemokines, normalizing transcript induction as fold change of CS-exposed compared with air-exposed control mice. CCR4−/− mice displayed reduced transcript induction of IFN-γ and the IFN-γ–induced chemokine CXCL10 (Figure 6C). As noted previously herein, there were no significant changes in CCR4 ligand transcripts compared with constitutive levels. There were also no differences in the transcript levels for the NK priming- and maturation-related molecules IL-15 and IL-18. However, it was noted that knockout mice showed increased expression of IL-10 transcripts and a notable reduction in transcripts for the NKG2D receptor–activating ligand RAET-1. The CCR4 transcript analysis confirmed knockout status.
Quantitative confocal analysis was performed to determine differences in NK cell localization. There were no differences between CCR4+/+ and CCR4−/− mice among parenchymal NKp46+ populations, but CCR4−/− mice showed a significant reduction in mucosa-associated NKp46+ cells (Figure 6D), suggesting that CCR4 was required for optimum airway localization of NK cells.
CCR4 Permits Optimal NK Cell Contacts with Lung CD11c+ and CD11+MCHII+ Cells
The activation of mature NK cells involves a series of stages, one of which is NK cell priming. The latter is required to reverse the effect of inhibitory signals that maintain NK cells in a quiescent state. Priming involves cell-cell interaction of NK cells with mDCs or macrophages that bear the cell membrane cytokine IL-15.
IL-15 signaling triggers NK cells to express CD69, but they do not necessarily attain effector functions, such as IFN-γ production or cytotoxicity, until additional activating signals, such as NKG2D ligands, are encountered.
Based on the present findings, we postulated that CCR4 might contribute to NK cell priming events in the lung by promoting interactions with sentinel cells. Using quantitative confocal microscopy, we measured the number of NK (NKp46+) cells in tight contact with large CD11c+ and CD11c+MHCII+ cells in CS-exposed CCR4+/+ and CCR4−/− mice. Respective examples of NK cell contacts with CD11c+ and CD11c+MHCII+ cells in CS-exposed wild-type mice are shown (Figure 7, A and B). Such interactions were observed at parenchymal and peribronchiolar locations. Moreover, tight synapse-like contacts were demonstrable as yellow overlap zones along membranes (Figure 7B). Contacts were sometimes multiple and were more commonly observed in parenchymal locations, which may reflect the fact that a greater area of lobular parenchyma can be observed in tissue sections. In CS-exposed CCR4−/− mice, these contacts were less frequent (Figure 7, C and D). Quantitation revealed an approximately 60% reduction in contacts in lungs of CCR4−/− mice (Figure 7E).
Figure 7CCR4 knockout impairs NK cell contacts with CD11c+ in lungs of CS-exposed mice. A and B: Representative images of NKp46+ cells tightly contacting CD11c+ and CD11c+MHCII+ cells in lungs of CS-exposed wild-type (CCR4+/+) mice. Inset: Tight synapse-like contacts are seen as yellow overlap zones along membranes (arrow). C and D: Representative images of NKp46+ cells with impaired contacts with CD11c+ and CD11c+MHCII+ cells in lungs of CS-exposed CCR4−/− mice. Original magnification, ×200. E: Quantitative confocal analysis of NK cell contacts expressed as a percentage of NKp46+ cells making contact with large CD11c and CD11c+MHCII+ cells. Bars are means ± SD; four mice per group. ∗P < 0.05.
The movement and positioning of effector cells is thought to be mediated by chemokines, but the precise functions of the various members of this family of molecules must be defined before designing therapeutic interventions. We recently reported that CCR4 was required for optimum NK cell activation during innate stage elimination of mycobacteria.
Originally, CCR4 was purported to be restricted to adaptive Th2 CD4+ memory T cells, but subsequent studies demonstrated expression by a variety of effector populations, such as regulatory T cells, Th1, Th17, and NK cells.
The chemokine ligands for CCR4 are produced by macrophages and mDCs under homeostatic and challenge conditions, suggesting a role for these chemokines in regulating effector cell function through interactions with sentinel cells. To elucidate the potential role of CCR4 in CS-elicited innate responses in the lung, we first characterized mDC, macrophage, and NK cell responses in mouse lungs after 4-day CS exposure, representing a period before adaptive responses are mounted. NK cell responses are usually transient, being supplanted by adaptive responses, but they are critical in that they can dictate the nature of subsequent supplanting response. Furthermore, cross talk between DCs and NK cells is thought to determine the mutual maturation and activation state of these early responders.
However, the location and factors mediating DC–NK cell interactions have yet to be fully determined.
Unlike previous studies examining the role of CS effects on DC populations, we developed a flow cytometric strategy to separately examine CD103−CD11bhi and CD103+CD11blo mDC subpopulations, which represent predominantly parenchyma- and mucosa-associated sentinel DCs, respectively (Supplemental Figure S1). Short-term CS exposure affected these populations differently, causing at least a transient reduction in the parenchymal CD11bhiDC population but with no effect on the CD103+ mucosa-associated DCs. These results differ from those of Botelho et al,
who reported an accumulation of mDCs after 4-day CS exposure. However, that group used twice the exposure dose of the present study, hence differences may be due to dose-related effects on mDC kinetics. The present study also separately examined mDC subpopulations and performed corroborative direct in situ localization analysis, which may have added sensitivity.
The observation of differential effects of CS exposure on parenchymal and mucosal DCs is novel and suggests differences in emigration, survival, or replenishment of the populations. After organ challenge or injury, DCs normally emigrate to draining lymphoid tissue and are replenished from blood-borne precursors. Emigration was suggested by the analysis showing skewed appearance of CD103−CD11bhi mDCs in draining lymph nodes (Supplemental Figure S2). However, note that we cannot rule out NK cell–DC interactions that result in DC killing as a cause of the reduction in mDC populations.
In this hypothetical model, known as DC editing, NK cells directly lyze immature mDCs, resulting in the survival of mature DCs that initiate more vigorous immune responses. Differential editing of CD103−CD11bhi and CD103+CD11blo mDCs has not been defined, but these populations are developmentally and functionally different. Specifically, CD103+ DCs i) express high levels of class II MHC antigens, ii) have the capacity to present apoptotic cell antigens, iii) efficiently activate cytolytic T cells, and iv) are dependent on Batf3 gene expression for development.
Having a highly mature phenotype, CD103+ mDCs may be more resistant to editing compared with CD103−CD11bhi mDCs, resulting in a selective reduction of the latter. Alternatively, the present findings in CS-exposed mice may simply reflect different rates of parenchymal and mucosal mDC replenishment. Detailed trafficking and survival studies are needed to distinguish among these possibilities.
The finding of robust macrophage recruitment was not unexpected because it is a consistent observation in CS-exposed animals and humans.
In accord with these reports, the present confocal analysis demonstrated constitutive CCL22 expression in CD11c+ cells located in lung lobular interstitium and mucosa that were consistent with DCs and possibly macrophages (Supplemental Figure S3).
As well as effects on mDC and macrophage populations, short-term CS exposure elicited a population of endogenously activated CD69+ NK cells. Confocal microscopy of lungs located NK cells in the lobular interstitium of CS-exposed and control mice, consistent with the presence of a resident NK cell population.
In contrast to controls, CS elicited a significant increase in bronchiolar airway–associated NK cells. Moreover, short-term CS exposure also induced the NKG2D receptor–activating protein RAET-1 in bronchiolar epithelium similar to that reported in models of long-term CS exposure.
NKG2D receptors are expressed by NK and CD8+ T cells and promote cytotoxic function when ligated. In a recent report, NK cells from CS-exposed mice displayed enhanced killing of NKG2D ligand–expressing cells, and NKG2D receptor–deficient mice had reduced lung injury in a model of viral exacerbation of CS exposure.
These studies implicate cytotoxic activation of NK cells by CS-induced NKG2D stress proteins as the mechanism of lung injury. The present observations are consistent with these reports and further suggest a role for CCR4 and its ligands in this process.
CCR4 did not influence the distribution of lung mDCs or macrophages after short-term CS exposure. However, we detected a significant impairment of NK cell activation in CCR4−/− mice. The latter was associated with impaired induction of IFN-γ transcripts in CS-exposed lungs, which may reflect reduced production by NK cells as we previously demonstrated in lungs of mycobacteria-challenged CCR4−/− mice.
There was a corresponding enhancement of IL-10 transcripts, implying a shift to a tempered response, and this inverse relationship of IFN-γ and IL-10 is consistent with the commonly reported cross-regulation of these cytokines. Unexpectedly, lungs of CCR4−/− mice showed reduced RAET-1 induction, suggesting that factors other than direct CS-induced injury were eliciting this stress protein. One possibility is that CCR4 knockout affected additional signals required to amplify RAET-1 expression. For example, type I IFN-α promotes NKG2D ligand expression in cultured cells, increasing their sensitivity to NK cell–mediated lysis.
Plasmacytoid DCs are thought to be major sources of IFN-α. We did not examine plasmacytoid DCs in this study, but CCR4−/− mice displayed a reduction in transcripts for the chemokine CXCLI0, which is chemotactic for mouse NK cells and plasmacytoid DCs.
Teleologically, coordinate DC-mediated amplification of NKG2D ligand expression would promote more rapid cytotoxic elimination of damaged or virally infected epithelial cells, but further studies would be needed to test this hypothesis.
NK cell cross talk with CD11c+ mDCs has been extensively discussed in the literature, but the in vivo locations and physiologic framework for those interactions have been unclear. We show for the first time that such interactions can take place in the lung and seem to depend in large part on CCR4-mediated chemotaxis or adherence. The CCR4 ligand CCL22 was reported to be chemotactic for human NK cells soon after its discovery.
We extend those findings by demonstrating the presence of a significant population of CCR4+ NK cells in mouse lungs. Also, in accord with previous reports, we detected CCR4 expression in CD44+CD4+ memory phenotype T cells. NK and memory T cells express CD44, a hyaluronan receptor expressed by tissue-homing effector cells, which is also a co-stimulatory molecule for NK cells.
Macrophage-derived chemokine (MDC/CCL22) and CCR4 are involved in the formation of T lymphocyte-dendritic cell clusters in human inflamed skin and secondary lymphoid tissue.
we surmised that CCR4 may mediate DC contacts with conventional NK cells. DCs and macrophages are potential sources of IL-12, IL-15, and IL-18, cytokines known to influence NK cell expansion, maturation, and priming. Thus, compromised cross talk signaling due to reduced contacts in CS-exposed CCR4−/− mice would explain the observed impairment of NK cell activation.
We have now demonstrated CCR4 dependence of innate-stage NK cell activation in experimental infection and CS exposure models. We propose that during homeostasis, CCR4 promotes intraorgan contacts of tissue-homing CCR4+ NK cells with mDCs or macrophages. Under homeostatic conditions, this results in no effector response due to lack of costimulation. In contrast, organ infection/injury increases mDC expression of costimulatory molecules and cytokines, allowing for rapid first-contact effector cell priming/activation. Primed NK cells would then respond to nonhomeostatic inflammatory chemokines, such as CXCL10, reportedly produced by insulted airway epithelium
and lyze target cells on contact with appropriate ligands.
In conclusion, short-term CS exposure elicits a rapid innate activation of NK cells and induction of RAET-1 NKG2D ligand in the lung, which are significantly dependent on CCR4 expression. We further demonstrate a role for CCR4 in mediating in vivo contacts between NK and CD11c+MHCII+ cells, implicating CCR4 in effector cell–DC cross talk events in the lung. Further studies are warranted to test the therapeutic targeting of CCR4 on the immunopathology of long-term CS exposure.
Flow cytometric and confocal microscopic characterization of CD11b+CD103− and CD11b−CD103+ mDCs in the mouse lung. A: Identification of mDC subpopulations using multicolor flow cytometric analysis. CD11c+ cells were gated from CD45+ large mononuclear cells based on side and forward light scatter. The MHCII bright cells were selected and then were analyzed for CD11b and CD103 expression. Two distinct populations were discerned. B: Confocal microscopic localization of CD11c+MHCII+CD103- and CD11c+MHCII+CD103+ in mouse lungs. Large CD11c+MHCII+ cells were located in parenchymal and airway locations, but CD103 expression was primarily limited to CD11c+MHCII+ cells associated with bronchiolar airways. C: Confocal microscopic appearance of bronchiolar airway and lobular parenchymal CD11c+MHCII+ cells morphologically consistent with mDCs. Arrows indicate bronchiolar cells with processes extending beneath and into epithelium and lobular DCs with interstitial dendritic processes. Asterisks indicate the lumenal side of the airway. MHCII expression was often polarized to the antilumenal side of the cells. Original magnification: ×100 (B); ×200 (C).
Flow cytometric quantification of mDCs in draining mediastinal lymph nodes of CS-exposed and air-exposed control mice. Draining mediastinal lymph nodes were collected, dispersed, counted, and subjected to flow cytometric analysis to discern CD103− and CD103+ mDC subpopulations. Bars are means ± SD; five to seven mice per group. ∗P < 0.05.
Confocal immunofluorescence detection of constitutive CCL22 expression co-localizing with lung CD11c+ cells in airways and lobular parenchyma of naive C57BL/6 mouse lungs. Arrows indicate cells with CD11c and CCL22 co-localization. Original magnification, ×200.
CCR4 knockout mice display normal distributions of major blood NK cell populations. Tail vein anticoagulated blood samples of naive, unchallenged CCR4+/+ and CCR4−/− mice were subjected to multicolor flow cytometric analysis to assess peripheral tissue–homing KLRG1+ and secondary lymphoid tissue–homing CD27+ NK cell populations. Gated NK1.1+ cells negative for T- and B-cell markers were analyzed. Fluorophore-labeled Abs and isotype controls were obtained from BioLegend and eBioscience Inc. (both in San Diego, CA). Representative flow cytometric plots are shown. CCR4−/− generated normal profiles of conventional NK cell populations.
References
Hogg J.C.
Timens W.
The pathology of chronic obstructive pulmonary disease.
Differential activation of killer cells in the circulation and the lung: a study of current smoking status and chronic obstructive pulmonary disease (COPD).
Macrophage-derived chemokine (MDC/CCL22) and CCR4 are involved in the formation of T lymphocyte-dendritic cell clusters in human inflamed skin and secondary lymphoid tissue.
Supported by NIH National Institute of Allergy and Infectious Diseases grant A143460 (S.W.C.); in part by the Department of Veterans Affairs (S.W.C.); and by Flight Attendant Medical Research Institute grant CIA-103071 (P.M.).
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