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Division of Pulmonary and Critical Care Medicine, and
Division of Rheumatology and Immunology, University of Virginia Health System, Charlottesville, VA 22908;
Section of Molecular Genetics and Microbiology and Institute for Cellular and Molecular Biology, University of Texas, Austin, TX 78712; and
Center for Comparative Medicine, University of Virginia Health System, Charlottesville, VA 22908
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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There is increasing evidence that proinflammatory chemokines and their receptors also have a profound effect on pulmonary allergic responses (14, 15). Among the chemokines detected in airways of patients with allergic asthma is monocyte chemotactic protein (MCP)-1 (16). Various roles have been attributed to MCP-1 in promoting allergic responses, including recruitment of basophils (17) and clumping of mast cells (18), IgE-independent induction of basophil (17) and mast cell (18) histamine release, chemotaxis of T cells (19), and enhancement of polarization of naive T cells to IL-4-producing Th2 cells (20, 21). Further evidence that MCP-1 may play a role in allergic asthma can be found in studies that observe that MCP-1 immunoneutralization resulted in reduction in AHR, pulmonary inflammation, and production of lymphocyte-derived inflammatory mediators in allergen-challenged animals (22, 23, 24). Because CCR2 is the major receptor for MCP-1, it may also play a role in allergic bronchial hyperreactivity and inflammation.
Mice deficient in CCR2 exhibit a strong Th1 to Th2 switch in their immune responses to infectious challenge, as characterized by large increases in production of IL-4, IL-5, and anti-pathogen-specific Abs, including IgE (25, 26). Considering this bias toward Th2 immunity in CCR2-deficient mice, we wanted to determine how these animals would respond in a noninfectious, Ag-driven model of allergic asthma. Consistent with the results from the infection models, CCR2-deficient mice sensitized to OVA and then rechallenged with aerosolized Ag showed a strong pulmonary Th2 response relative to the response of wild-type (wt) mice. The response included high levels of IL-5, Ag-specific Ig, and MCP-1, as well as profound bronchial eosinophilia, increased AHR, and Goblet cell hyperplasia. Taken together, these results provide evidence for an important role for CCR2 signaling pathways in Th2 immune responses. Defects in these pathways contribute to an enhanced Th2 phenotype, which in this case leads to the exacerbation of clinical features in this model of pulmonary allergy.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The experimental CCR2-deficient mice CCR2-/- on a mixed 129/Ola and C57BL/6 background (27) and wt mice on a mixed 129 x C57BL/6 background were bred in the Animal Resource Facility at the University of Virginia (Charlottesville, VA). Inbred C57BL/6 and 129/SvJ mice were purchased from The Jackson Laboratory (Bar Harbor, Maine). All animal procedures were approved by the Animal Research Committee at the University of Virginia, and all procedures conform to the Guide for the Care and Use of Laboratory Animals by the Institute of Laboratory Animal Resources. For lung BAL and fixation of lung tissue, deep anesthesia was induced by i.p. injection of ketamine (80 mg/kg) and xylazine (8 mg/kg) followed by euthanasia by exsanguination.
OVA sensitization and challenge
On days 0 and 7, mice received an i.p. injection of 25 µg of chicken egg OVA (Sigma, St. Louis, MO) mixed with 2 mg of alum (Pierce, Rockford, IL) in 100 µl of 0.9% NaCl (normal saline). Beginning 14 days after this initial sensitization, animals were challenged every other day for 16 days (8 treatments in total) with either inhalation of 1% OVA in normal saline or inhalation of normal saline alone. For each 20-min challenge, animals were placed in a 1.3-liter plastic chamber connected to the aerosol output from a Ultra-Neb99 nebulizer (DeVilbiss, Somerset, PA).
Measurement of AHR
AHR was measured as we have described previously (28) by using the enhanced expiratory pause (Penh) in plethysmographic box pressure during expiration (Buxco Electronics, Sharon, CT). Free-moving conscious mice were evaluated in closed plethysmographic chambers through which continuous airflow refreshed the air supply. Penh was calculated by computer analysis of plethysmographic (box) pressure curves generated by each inhalation (negative pressure) and exhalation (positive pressure wave) with BioSystem XA software, version 1.5 (Buxco Electronics, Sharon, CT). Expiratory pressure was measured against time. Total exhalation time (Te) was the time from initiation of exhalation until the box pressure became zero. With an arbitrarily defined percentage of 65%, the computer divided the expiratory pressure-time curve into an area 65% of which represented the time from onset of expiration to the time-point representing 65% of the area. The time under this area was the relaxation time (Tr). The subsequent 35% of area of the pressure-time curve at the latter part of expiration was the pause time (Tp), such that Te = Tr + Tr. Penh was calculated by the computer by multiplying the ratio of Tp/Tr by the ratio of peak expiratory box pressure (PEP) to peak inspiratory box pressure (PIP) by the following equation: Penh = (Tp/Tr) x (PEP/PIP). Penh measurements have been validated by Hamelmann et al. (29) in regard to identification of AHR in OVA-sensitized and -challenged mice, compared with OVA-challenged nonsensitized mice. The heightened increase in Penh with methacholine challenge in OVA-sensitized/-challenged mice was accompanied by parallel enhancement in lung airway resistance (RL) responses to methacholine, with a high degree of correlation between Penh and RL. Moreover, these investigators found no effect of breathing pattern or respiratory rate on Penh. The pressure-time curve areas used to calculate Tr and Tp by these investigators were 64% and 36%, respectively, which are nearly identical with 65% and 35% used in the present experiments.
AHR was evaluated within 24 h of the last day of aerosol challenge (day 29). Mice were placed in separate plethysmographic chambers and allowed to acclimatize for at least 15 min before analysis. Control measurements were obtained over a 5-min control period. Afterward, increasing concentrations of methacholine in saline of 6.2, 12.5, 25, and 50 mg/ml were nebulized into the chambers for 3 min. At the end of each 3-min methacholine aerosolization, Penh measurements were taken for two consecutive 5-min periods. This was followed by challenge with the next dose of methacholine. In the analysis of Penh responses, it was found that there were no differences in Penh measurements in the first 5-min period after each methacholine challenge. However, statistically significant differences were encountered in Penh in the second 5-min period after methacholine. Therefore, the data represent measurements obtained in the second 5-min period after each methacholine challenge.
BAL (right lung) and lung fix-inflation (left lung)
After deep anesthesia with ketamine/xylazine, a 22-gauge Teflon catheter was inserted into the proximal trachea and secured with 5–0 silk suture, and the animal was ventilated (Model 687; Harvard Apparatus, South Natick, MA) at a rate of 100 and an inflation pressure of 16 cm H2O. Through a midline celiotomy, animals were euthanized by exsanguination through withdrawal of blood from the inferior vena cava. The blood was placed in tubes containing ethylenediamine tetraacetate for total and differential leukocyte counts and measurement of Ig levels and MCP-1.
A median sternotomy incision was performed and the left hilum was cross-clamped, as evidenced by cessation of left lung inflation by the ventilator. The right lung was lavaged through the tracheal cannula with 10 ml of 0.9% NaCl warmed to 37°C. The left hilar cross-clamp was removed, and both lungs were fix-inflated with 10% neutral buffered formalin or 2.5% glutaraldehyde in 0.1 M sodium phosphate buffer to an inflation pressure of 25 cm H2O.
Identification and quantitation of leukocytes
Total leukocyte counts in peripheral blood and BAL fluid were measured with a hemacytometer. The percentages of leukocyte subsets were determined by microscopic examination of blood smears and cytocentrifuged BAL leukocytes stained in Diff-Quik (Dade Diagnostics, Aguada, Puerto Rico).
Lung histology
After fixation, lung tissue was embedded in paraffin, and sections were mounted onto glass slides by the Research Histology Core Laboratory of the Center for Research in Reproduction at the University of Virginia. Deparaffinized sections of lung tissue were stained with hematoxylin and eosin for quantitation of inflammatory infiltrates and with periodic acid/Schiff (PAS) for quantitation of bronchial mucous gland hypertrophy. Videomicroscopic images of lung sections were analyzed with ImagePro software (Media Cybernetics, Silver Spring, MD) calibrated with a micrometer slide. In PAS-stained lung sections used for bronchial mucous gland hyperplasia, the mucosal circumference of an axial section of a bronchus was measured, which allowed conversion of total mucosal PAS-positive cells to the number of PAS-positive cells per 100 µm of mucosal perimeter. The area of inflammatory cuffing around pulmonary arterioles and bronchioles also was measured by using ImagePro software.
Ag stimulation of T cells
Mice were given a single footpad and s.c. injection of OVA/alum
in concentrations identical with the sensitization dose used in the
airway hyperreactivity studies. Twelve days later, the mice were
euthanized and regional draining lymph nodes and spleen were harvested.
Tissues from two mice of each genotype were combined. The lymph node
cells were dispersed in RPMI 1640 containing 100 U/ml of
collagenase D (Boehringer Mannheim, Indianapolis, IN). The spleen was
injected with RPMI 1640 containing collagenase D, 400 U/ml, and teased
apart for dispersion of splenic cells. Cells from the lymph nodes and
spleen were filtered through 100-µm cell strainers (Falcon; Becton
Dickinson, Franklin Lakes, NJ) into separate tubes, and lymphocytes
were separated by density centrifugation with sodium
diatrizoate/polysucrose (1.083 g/ml; Sigma). Mononuclear cells
(interface) were aspirated and resuspended in RPMI 1640 containing
penicillin/streptomycin and 50 µM 2-ME. After irradiation (3000 rad)
of the splenic mononuclear cells, lymphocytes (0.5 x
106 cells/well) and spleen cells containing APCs
(2 x 106 cells/well) were cocultured in
individual wells of a 24-well culture plate in the presence of
increasing concentrations of OVA (0, 10, 50, or 250 µg/ml). All cell
isolation procedures were conducted at 4°C. The plates were placed in
a 37°C incubator for 72 h in 5% CO2, and
the supernatant from each well was aspirated and frozen at -70°C
until assays for IL-4, IL-5, and IFN-
were performed by ELISA, as
described previously (28). Cytokine production is
expressed as the fold increase in cytokine production (OD) from
unstimulated cells (RPMI 1640) to cells incubated with maximum Ag (OVA,
250 µg/ml), and in ng/ml determined from a standard curve.
Measurement of serum Igs
Serum Igs were measured by ELISA, as described previously (28). Microtiter wells were coated with OVA, and diluted serum samples and standards were placed in each well. The plates were incubated at room temperature for 2 h and then washed with PBS. HRP-labeled Ab to mouse IgG1 or IgG2a, alkaline phosphatase-labeled anti-IgG2b, or biotinylated Ab to mouse IgE were added to individual wells and the plates were incubated for 2 h at room temperature. To measure IgE, avidin-labeled HRP was added to the appropriate wells after washing unbound Ab. After washing, each well was incubated with appropriate substrate and read in an automated microtiter plate at either 490 or 405 nm. The difference between the OD of the positive Ig standard and the negative control was arbitrarily assumed to represent 1000 U. The ODs of the unknown samples then were converted to units per sample by dividing the difference of the unknown OD and background by the difference between the positive control and background and multiplying the quotient by 1000.
CCR2-/- characterization
Tail DNA was isolated from nine of the
CCR2-/- mutants and six of the wt
CCR2+/+ mice for genotypic confirmation by PCR.
Two PCR primers for murine CCR2 (Ref. 30 ; GenBank U47035)
were synthesized for sense position 55 (5'-AATATGTTACCTCAGTTCATCCAC)
and antisense position 249 (5'-ACCAAAGATGAATACCAGGGA), which flank the
BamHI insertion site of the 1.8-kb pgk-neocassette that
disrupted the CCR2 gene (27). A third probe was
synthesized to the 529 sense position of neomycin (GenBank AF080389).
Preliminary experiments revealed that two separate PCRs were necessary
to genotype the animals. PCR with the two CCR2 primers yielded the
expected DNA product of 
200 bp, confirming the intact CCR2 gene in
the CCR2+/+ mice. PCR of DNA with the CCR2 sense
probe and the neomycin sense probe yielded a DNA product of around 1000
bp, identifying the presence of the neomycin gene and, hence, the
CCR2-/- mutant.
Assessment of mononuclear phagocyte chemotaxis to MCP-1 also was done to assess the disruption of CCR2. Peritoneal exudative cells were elicited in wt and CCR2-/- mutant mice by peritoneal lavage of ketamine/xylazine anesthetized animals at 72 h after i.p. injection of 2.5 ml of 0.2% sodium caseinate (31). The harvested peritoneal exudative leukocytes were washed, quantified, and resuspended in RPMI 1640 containing 1 mg/ml BSA (fraction V; Sigma) at a concentration of 2 x 106 cells/ml. Leukocyte chemotaxis was measured as described (32) through an acellular Poretics 5-µm non-PVP polycarbonate membrane (Osmonics, Livermore, CA) with a 48-well chemotaxis chamber (Neuro Probe, Gaithersburg, MD). In duplicate, medium alone (RPMI 1640/BSA) or with recombinant murine MCP-1 (R&D Systems, Minneapolis, MN) were loaded into the lower chambers, and the upper wells then were filled with 50 µl of RPMI 1640 containing 2 x 106 of unseparated leukocytes/ml. After incubation in 5% CO2 at 37°C for 90 min, the membrane was scraped to remove nonmigratory leukocytes. The cells were fixed in methanol, stained with Diff-Quik, and the mononuclear cells were counted within the width of a high power field (total magnification x1000) across the horizontal diameter of the well. In peritoneal leukocytes from CCR2+/+ mice, a dose-dependent response in macrophage chemotaxis was observed in response to mMCP-1 (0.1, 1, 10, and 100 nM; data not shown). In contrast, very little chemotaxis occurred to MCP-1 by elicited peritoneal leukocytes from CCR2-/- mice, supporting evidence for the CCR2-/- null mutation.
ELISA for MCP-1
Plasma MCP-1 levels were measured from blood collected from each mouse at the time of euthanasia in tubes containing potassium ethylenediamine tetraacetate with a commercially available ELISA for murine MCP-1 (R&D Systems).
RNase protection assay
Lungs were removed from OVA- or saline-challenged CCR2+/+ and CCR2-/- mice sacrificed under anesthesia, and total RNA extraction was performed according to the method of Chomczynski and Sacchi (33). Total RNA was further purified by using RNeasy minispin columns (Qiagen, Valencia, CA). RNase protection assay was performed with a commercially available assay (BD PharMingen, San Diego, CA) according to the manufacturer’s instructions. Riboprobes labeled with 32P were synthesized by in vitro transcription. For each protocol, 10 µg of whole lung mRNA pooled from two mice (5 µg per mouse) was incubated at 56°C for 16–20 h with the labeled riboprobe template set. Unbound riboprobes not hybridizing to mRNA were removed by adding RNase followed by phenol/chloroform-isoamyl alcohol extraction. Protected riboprobes binding to lung RNA then were electrophoresed over an acrylamide/bis-acrylamide gel (19:1). After gel transfer to filter paper, each gel was dried and placed against Kodak X-OMAT AR film in cassettes at -80°C for development of autoradiographs.
Statistical analysis of data
All data were analyzed by the SAS general linear models procedure (SAS Institute, Cary, NC; Ref. 34). This procedure is similar to ANOVA but is preferred when there are unequal numbers of observations between treatment groups. When an overall treatment effect was identified at the p < 0.05 level, specific differences between treatment groups were determined by Duncan’s multiple range test (35). For each variable, factorial analysis of the general linear models procedure was used to analyze for effects of genotype (CCR2+/+ vs CCR2-/-), treatment (OVA vs saline), and interaction between genotype and treatment in the mixed-background CCR2+/+ and CCR2-/- mice. The factorial analysis procedure also was used in all CCR2+/+ groups to evaluate for effect of genetic background (mixed background, inbred C57BL/6, inbred 129/Sv), either alone or through interaction with treatment (OVA). In the text and figures, data are expressed as the mean ± SEM.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In the CCR2+/+ and
CCR2-/- mice in the mixed-background, control
Penh levels were comparable among all groups (not shown). As depicted
in Fig. 1
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Penh (Penh after
methacholine dose - Penh control) rose significantly
(p < 0.05) with increasing methacholine doses
in both genotypes, regardless of whether challenged with OVA or saline.
At the highest methacholine dose (50 mg/ml), increased airway
hyperreactivity was observed in the OVA-challenged
CCR2-/- mutants, with Penh significantly
higher (p < 0.05) in the OVA-challenged
CCR2-/- mutants than Penh observed in
OVA-challenged CCR2+/+ mice. All methacholine
Penh responses for the four protocols depicted in Fig. 1 were analyzed
for the effects of methacholine dose, OVA-treatment, and CCR2 status. A
highly significant effect (p < 0.05) of
methacholine dose effect was present. Although there were not
significant effects of CCR2 or OVA treatment alone, there was
interaction (p < 0.05) between methacholine
dose, OVA-challenge, and genotype (CCR2+/+ vs
CCR2-/-).
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Penh between OVA- and saline-challenged
CCR2+/+ mice in C57BL/6 and 129/Sv backgrounds
(data not shown). Fig. 2 depicts the
Penh responses to the highest methacholine dose in all groups
studied. Multivariate analysis of the Penh responses to methacholine
for the CCR2+/+ mice in mixed and inbred
backgrounds identified a significant methacholine dose effect
(p < 0.05), and there was significant
interaction (p < 0.05) between methacholine
dose and genetic background. However, there was neither interaction
between genetic background (mixed, C57BL/6, or 129/Sv) and OVA
challenge, or significant interaction between methacholine dose,
genetic background, and OVA challenge on AHR responses. This suggests
that the enhanced AHR in the OVA-challenged
CCR2-/- mutants is unrelated to genetic
background effects.
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In the mixed background, an increase in total leukocytes and
percentage of eosinophils and a striking increase in absolute numbers
of eosinophils were observed in the OVA-challenged
CCR2-/- mutants compared with comparably
challenged CCR2+/+ mice (Figs. 3 and 4).
For total BAL leukocyte numbers, percentage of eosinophils, and
absolute number of eosinophils, factorial analysis in the mixed
background (left), revealed significant effects
(p < 0.05) of OVA challenge and CCR2 status
with interaction (p < 0.05) between the two
effects. In the CCR2+/+ inbred mice, although
absolute BAL leukocytes did not increase in either genotype, OVA
challenge led to an increase in both the percentage of eosinophils and
total number of eosinophils in 129/Sv mice. Factorial analysis across
the three genotypes of CCR2+/+ mice (mixed
background and inbred C57BL/6 and 129/Sv strains) revealed significant
(p < 0.05) OVA and genetic background effects
on the percentage of eosinophils in BAL, with evidence for interaction
(p < 0.05) between these effects. These
observations suggest that increase in percentage of eosinophils in BAL
leukocytes in the CCR2 mutants may be influenced by genetic background.
However, in regard to absolute numbers of BAL eosinophils, although
there was evidence of an OVA effect in the
CCR2+/+ mice, (p <
0.05), there was no evidence of genetic background effect or any
interaction between OVA and genetic background. Thus, enhancement in
BAL eosinophilia in OVA-challenged CCR2-/-
mutants is unrelated to genetic background of the mutants.
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B), including profound
cellular infiltrates that cuffed small bronchioles and pulmonary
arterioles. Lung histological changes in OVA-challenged
CCR2+/+ mice were barely detectable (Fig. 5A), with evidence of slight accumulation of inflammatory
cells around the small pulmonary arterioles. There was absence of
histological changes in the saline-challenged mice of either genotype
in the mixed background (not shown).
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, left). The increase in
peribronchiolar inflammatory cuffing in the OVA-treated mutants was not
related to the size of bronchioles used for morphometry, as the
diameters of bronchioles were comparable at 279 ± 25 and 244
± 19 µm, respectively. Factorial analysis revealed significant
(p < 0.05) effects of OVA treatment and CCR2
in peribronchial cuffing. Moreover, there was interaction
(p < 0.05) between OVA and CCR2 effects.
Although a small degree of peribronchiolar cuffing was observed in the
inbred CCR2+/+ strains challenged with OVA, the
changes were not significantly different from inbred controls
challenged with saline (Fig. 6, right). Factorial analysis
of all CCR2+/+ mice revealed a significant
OVA-treatment effect (p < 0.05), but no
genetic background effect was evident on peribronchial inflammatory
cuffing. Thus, enhanced peribronchial cuffing of inflammatory cells in
the OVA-challenged CCR2-/- mutant is likely
related to the CCR2 deletion rather than genetic background
effects.
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, left). Increased
perivascular inflammatory infiltrates in the OVA-challenged mutants was
not related to the size of arterioles studied, with comparable
pulmonary arteriolar diameters of 87 ± 9 and 71 ± 8 µm,
respectively. In the mixed background, there was evidence on factorial
analysis of significant (p < 0.05) OVA and
CCR2 effects on vascular cuffing. Moreover, there was interaction
(p < 0.05) between OVA and CCR2 effects.
Although OVA challenge did not lead to an increase in perivascular
cuffing in CCR2+/+ C57BL/6 inbred mice, OVA
treatment was associated with an increase in perivascular infiltrates
in CCR2+/+ 129/Sv mice compared with respective
saline-treated control group (Fig. 7, right). Factorial
analysis of all CCR2+/+ groups, although not
revealing a background effect, identified a significant
(p < 0.05) OVA effect, with significant
interaction (p < 0.05) between OVA and genetic
background. These observations suggest that inflammatory cuffing around
pulmonary arterioles in the CCR2-/- mice may be
related to differences in genetic background between the
mixed-background CCR2+/+ and
CCR2-/- mice.
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and 9) compared with OVA-challenged
CCR2+/+ mice. In the mixed background, there were
increased (p < 0.05) numbers of PAS-positive
mucous-producing cells in the bronchial mucosa of OVA-challenged
mutants compared with OVA-challenged CCR2+/+ mice
(Fig. 9, left). Factorial analysis revealed significant
effects of both OVA (p < 0.05) and CCR2 status
(p < 0.05) on the number of PAS-positive
cells, and there was evidence of interaction (p
< 0.05) between these effects. There also was an increase in Goblet
cell hyperplasia with OVA challenge in both C57BL/6 and 129/Sv
CCR2+/+ inbred mice (Fig. 9, right)
compared with respective saline-challenged controls. Factorial analysis
of all CCR2+/+ mice revealed significant effects
for OVA and genetic background in the number of Goblet cells, with
interaction between the two effects. This suggests that enhancement in
Goblet cell hyperplasia in the OVA-challenged
CCR2-/- mutants could be influenced by genetic
background effects.
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OVA-specific serum IgG1 levels were elevated in the OVA-challenged
mutants above all other groups (Fig. 10). OVA-specific IgE also was
significantly elevated in OVA-challenged
CCR2-/- mutants above all other groups (Fig. 10). In addition, circulating IgG1 was elevated in the
saline-challenged OVA-sensitized CCR2-/- null
mutants over the saline-challenged OVA-sensitized
CCR2+/+ mice. Factorial analysis identified
significant (p < 0.05) effects of OVA
treatment and CCR2 status on both circulating
IgG1 and IgE levels. However, there was no
interaction identified between the two effects. IgG2a or IgG2b levels
were unchanged with OVA challenge in CCR2-/-
and CCR2+/+ mice (data not shown).
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There was a striking 8-fold elevation (p
< 0.05) in serum MCP-1 levels (Fig. 11) in the OVA-sensitized
CCR2-/- mutants over OVA-sensitized
CCR2+/+ mice regardless of whether the challenge
was saline or OVA. OVA challenge had no effect on circulating MCP-1
levels in either genotype. Factorial analysis identified a significant
(p < 0.05) CCR2 effect but no OVA-treatment
effect on serum MCP-1 levels.
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There was evidence for differences in lung expression of chemokine
receptors in CCR2-/- vs
CCR2+/+ mice (Fig. 12). This was not attributable to
nonspecific up-regulation because lung expression of L32 and GADPH was
comparable in all groups. In saline-challenged
CCR2-/- mutants (lane 4)
there was decreased expression of CCR1, CCR3, and CCR5 compared
with saline-challenged CCR2+/+ mice
(lane 2). However, despite decreased basal
expression, OVA challenge led to enhanced expression of CCR3 in
CCR2-/- mutants (lane 5),
compared with comparably treated CCR2+/+ mice
(lane 3). Moreover, there was up-regulation in CCR4
(faint band) in OVA-challenged CCR2-/- mutants,
whereas CCR4 was undetectable in all other groups. In both
CCR2-/- mutants and
CCR2+/+ mice, OVA challenge resulted in an
increase in lung expression of CCR1, CCR3, and CCR5 compared with
respective saline-challenged controls for each genotype. Interestingly,
although substantially less than in the CCR2+/+
mice, there was evidence of mild hybridization of the
32P-labeled CCR2 riboprobe to lung RNA from the
CCR2-/- mutants (Fig. 12, lanes 3
and 4). This is probably attributable to the fact that the
216-nt CCR2 riboprobe from BD PharMingen is complementary to a CCR2
sequence quite downstream from the BamHI neocassette
insertion site at position 209. In these and subsequent experiments in
which whole lung RNA is used for evaluation of expression of CCR and
cytokines, increase in mRNA could occur through emigration of
inflammatory cells into the lung as well as occurring through an
increased expression on cells normally residing within the lung
compartments.
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). In
addition, a faint band representing IL-5 mRNA was detected in
OVA-challenged CCR2-/- mutant, but like IL-13,
IL-5 expression was undetectable in any other group. IL-4 expression
was not evident in CCR2-/- mutants or either of
the CCR2+/+ groups. IFN- expression was
faintly detected in all groups, and expression appeared to be
comparable in all groups.
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Ag (OVA) stimulation of cells from regional lymph nodes of
sensitized CCR2-/- mice (mixed background)
resulted in enhanced (p < 0.05) IL-5
production compared with lymph node cells from
CCR2+/+ mice (Fig. 14). However, there were no differences
in IL-4 or IFN- production.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The results in these experiments differ from a recent report that CCR2-deficient mice had attenuated rather than enhanced airway reactivity with Ag sensitization and challenge with cockroach Ag (24), but agree with a recent report of enhanced AHR, increased lung expression of IL-5 and IL-13, and increased serum IgE in CCR2-/- mutants challenged with intratracheal Asperigillus conidia compared with comparably challenged wt (CCR2+/+) mice (36). In regard to the study (24) reporting attenuation of responses to cockroach Ag challenge in CCR2-/- mutants, the study and our experiments had many differences in methods, including Ag, adjuvant, route and schedule of Ag administration, and measurement of AHR. No histological data was presented in the cockroach-challenged mutants. Therefore, reasons for differences between the studies are not clear. These experiments also differ from observations of diminished Th2-mediated lung granuloma formation in CCR2-/- mice induced by Schistosoma egg Ag (37). However, it appears likely that diminished granuloma size was related to diminished migration of macrophages to the site of the Ag, a phenomenon observed by ourselves and others with diminished migration of macrophages to the peritoneal cavity after i.p. casein or thioglycollate administration (27, 32, 38).
Possible explanations for enhanced AHR and lung inflammation in the
Ag-sensitized CCR2 mutants in the present study include changes in
lymphocyte or macrophage migration to the lung, enhanced release of
histamine, and enhanced polarization of CD4+
lymphocytes to a Th2 phenotype, with increased release of Th2
cytokines. Although MCP-1 may increase histamine release by mast cells
and basophils (17, 18), it is unlikely that enhanced
histamine alone can lead to the lung and airway inflammation observed
in the CCR2-/- mutants. As mentioned above,
CCR2-/- mutants have diminished macrophage
migration to sites of inflammatory stimulation, and it is possible that
diminished migration of macrophages to the lungs of CCR2 mutants may
have led to enhanced airway inflammation and responsiveness in the
OVA-sensitized mutants. CCR2-/- mutants also
have deficient migration of Langerhans cells, dendritic cells in the
skin, to regional lymph nodes (39). This suggests that
dendritic cell migration to the lungs or regional lymph nodes may also
be impaired in CCR2-/- mutants. Moreover,
CCR2-/- mutant mice are deficient in CD8
subset of dendritic cells in the spleen (39). Because
these cells stimulate Th1 differentiation of T cells, a deficiency of
this subset of dendritic cells in the lungs or regional lymph nodes
might result in diminished Th1 activity, with decrease in IFN-
production. Reduction in IFN- production could enhance Th2 activity,
because previous investigators have observed that IFN-, a Th1
cytokine, may attenuate Th2 responses (40, 41). It also is
possible that differences in T cell localization to the lungs were
responsible for enhanced airway responsiveness. Although Th1 and Th2
cells both express CCR2, CCR2 is more abundantly expressed on Th1
cells, whereas CCR4 and CCR3 are prevalent on Th2 cells
(42). These data suggest that in regard to T cell
localization to the lung, Th1 localization would be more impaired in
CCR2-/- mice than Th2 localization. However,
insufficient localization of Th1 cells is a plausible explanation for
enhanced reactivity in the CCR2-/- mutants, in
view of the above-mentioned IFN- suppression of Th2 responses
(40, 41). In support of the notion that IFN- activity
may influence Th2 activity, chemokine profiles of children with atopic
asthma reveal correlation of disease with low IFN- activity
(43). These observations raise the question of whether
enhanced airway responses and inflammation in the CCR2 mutants is
directly related to factors that induce Th2 polarization or indirectly
related to diminished Th1 activity with reduced levels of IFN-.
Assessment of whole-lung cytokine expression reveals greater
differences in expression of Th2 cytokines (IL-5, IL-13) than in
IFN- expression. However, reliance on whole-lung expression can be
misleading, because localized attenuation in expression of IFN- may
still have contributed to enhancement in Th2 activity. Future studies
in which localized IFN- expression is assessed are required to
assess this possibility. Irrespective of the mechanism, there is
mounting evidence of enhanced Th2 cytokine activity in CCR2 mutants. We
observed enhanced release of IL-5 and absence of IFN- production
after Ag stimulation of lymphocytes from regional lymph nodes of
sensitized CCR2-/- mice and up-regulation in
lung expression of IL-5 and IL-13 in OVA-challenged CCR2 mutants. If
the exaggerated airway responses in the OVA-challenged CCR2 mutant are
related to increase in Th2 activity, it is unclear why the
up-regulation in Th2 cytokines includes IL-5 and IL-13 but not IL-4.
This up-regulated cytokine array is not unique and has been reported
recently in Asperigillus-challenged
CCR2-/- mutants, with increase in lung IL-5 and
IL-13 compared with wt mice, whereas changes in IL-4 between
CCR2-/- and wt mice were comparable
(36). Despite absence of IL-4 up-regulation in these two
experiments, compared with intact mice, there was enhancement in AHR
and lung inflammation, possibly attributable to the actions of IL-5 and
of IL-13, which jointly share the IL-4 receptor. IL-13 itself is a
potent mediator of allergic airway responses (44),
possibly through stimulation of production of monocyte-derived
chemokine (45, 46). This Th2 cytokine profile evident in
Asperigillus- or OVA-challenged CCR2 mutants appears to be
stimulus specific, as other investigators have reported that
Cryptococcus neoformans (25) and
Leishmania major (39) increase IL-4 production
in CCR2-/- cells. Lastly, there is indirect
evidence of IL-4 activity in the OVA-challenged CCR2-/-
mutants, with Ig subclass switching.
The 8-fold increase in circulating MCP-1 in CCR2-/- mice compared with wt mice has been suggested previously in lung homogenates or BAL of mice challenged with A. conidia (36) or C. neoformans (25). However, in these previous studies, discordance in MCP-1 levels between the CCR2-/- mutant and wt mice did not occur until after infection with Cryptococcus or sensitization to Asperigillus.
In view of the increase in circulating MCP-1 in the CCR2-/- mutants, the question could be raised as to its involvement in AHR and inflammation. Certainly, MCP-1 immunoneutralization results in attenuation of allergic AHR (22, 23, 24), and MCP-1-deficient mice have impaired Th2 responses (20). However, because CCR2 is the putative receptor for MCP-1 on hematopoietic cells (30, 47), these observations seem to be at odds with observations of enhanced Th2 responses in CCR2-/- mutants. The answer may possibly be related to elevation of circulating MCP-1 in the CCR2-/- mutants by the hypothesis that MCP-1 mediates AHR and lung inflammation through a non-CCR2 mechanism. Evidence is conflicting about MCP-1 agonist activity for other CCR on leukocytes besides CCR2. MCP-1 induces chemotaxis of mouse lymphoma cells transfected with CCR3 (48) and stimulates calcium flux in CCR4 mRNA-injected Xenopus oocytes (49). However, mice deficient in CCR4 have allergic airway responses that are comparable to intact mice, making CCR4 less likely as a mediator of heightened airway responses in the present study (50). There is growing evidence that MCP-1 is able to stimulate nonhemapoietic cells through other chemokine receptors, such as MCP-1-induced expression of tissue factor by smooth muscle cells that are devoid of CCR2 (51). Recently, an alternate receptor for MCP-1, CCR11, has been characterized in lung, liver, and intestine, but this receptor is not expressed on hematopoietic cells, and its relationship to the present observations remains unclear (52). For the MCP-1 mediation through a non-CCR2 hypothesis to be plausible, the issue of comparable elevation of plasma MCP-1 in saline-challenged as well as OVA-challenged mutants must be resolved. If MCP-1 is a mediator of exaggerated responses, why are responses not also exaggerated in sensitized saline-challenged mice as well? One potential answer can be found in observations by Karpus et al. (21) that primary Ag exposure of T cells in the presence of MCP-1 leads to enhanced Th2 phenotype, evidenced by IL-4 release, when the primed T cells are restimulated with Ag in the absence of further MCP-1 exposure. If this mechanism were present in CCR2-/- mice, elevation of MCP-1 might influence T cell differentiation into Th2 phenotype, which would not be evident until the animals were challenged with Ag. However, the absence of an increase in IL-4 in the OVA-challenged CCR2-/- mutants leaves open to question whether MCP-1 played such a role in these studies. Lastly, it must also be considered that the enhanced Th2 responses in the CCR2-/- mice could be mediated by abrogation of CCR2 stimulation by other cytokines or up-regulation of other CCR2-active chemokines including MCP-2 (47) MCP-3 (48), MCP-4 (48), and mouse MCP-5 (53). Thus, possible explanations for the results of the present study should include MCP-1 stimulation of non-CCR2 receptors, prevention of non-MCP-1 CC chemokine interactions with CCR2, or up-regulation of other CCR2-active chemokines. The latter possibility is supported by observations of increase in lung expression of MCP-3 and MCP-5 in CCR2-/- mutants at 4 and 8 days after challenge with the Th2 stimulus Schistosoma egg Ag (37) Subsequent experiments are required to assess these possibilities.
Although we observed enhanced airway hyperreactivity in OVA-challenged CCR2-/- mutants over saline-challenged mice of the same genotype, we did not see augmentation in AHR in OVA-challenged CCR2+/+ mice in mixed-background, inbred C57BL/6 or 129/Sv strains, compared with their respective saline-challenged controls. This may in part have been related to the C57BL/6 background, because it has been observed previously that OVA treatment augmented methacholine-induced AHR compared with saline-treated mice in BALB/c, but not in C57BL/6 or B6D2F1 mice (54). It is unclear why OVA challenge did not enhance AHR in 129/Sv.
In summary, our findings confirm enhanced Th2 responses in mice deficient in CCR2. Moreover, this study is the first to report enhanced airway responses to methacholine, airway eosinophilia, and peribronchial and perivascular cuffing in CCR2 mutants compared with wt mice. Ag stimulation of isolated lymphocytes identified an increased production of IL-5 in OVA-sensitized CCR2-/- mutants, which differed from responses in OVA-sensitized CCR2+/+ controls. Circulating MCP-1 levels were elevated 8-fold in OVA-sensitized CCR2 mutants above OVA-sensitized wt mice, regardless of whether they were challenged with OVA or saline. These studies suggest that CCR2 has an immunomodulatory role in Th2 polarization, but further studies are necessary to determine mechanisms responsible for enhancement in allergic airway responses.
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2 Address correspondence and reprint requests to Dr. C. Edward Rose, Jr., Department of Internal Medicine, P.O. Box 800546, University of Virginia Health System, Charlottesville, VA 22908.
3 Abbreviations used in this paper: AHR, airway hyperreactivity; BAL, bronchoalveolar lavage fluid; MCP, monocyte chemotactic protein; wt, wild type; Penh, enhanced pause; PAS, periodic acid/Schiff.
Received for publication July 20, 2000. Accepted for publication February 15, 2001.
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