The objective of the study was to characterize better the immunologic mechanisms underlying a previously developed animal model of chemical-induced asthma. BALB/c and severe combined immunodeficiency disease (SCID) mice received toluene diisocyanate (TDI) or vehicle on each ear on day 1 and/or day 7. On day 10, they were intranasally challenged with TDI or vehicle. Ventilatory function was monitored by whole body plethysmography for 40 min after challenge. Reactivity to methacholine was measured 23 h later: enhanced pause and actual resistance measurements. Pulmonary inflammation was assessed 1, 6, and 24 h after challenge by bronchoalveolar lavage (BAL). Tumor necrosis factor-α and macrophage inflammatory protein (MIP)-2 levels were measured in BAL. Immunological parameters included total IgE, IgG1, and IgG2a in serum, lymphocyte populations in auricular and cervical lymph nodes, and IL-4 and IFN-γ levels in supernatants of lymph node cells, cultured with or without concanavalin A. Ventilatory changes suggestive of airway obstruction and increased methacholine reactivity were observed in all TDI-sensitized and TDI intranasally instilled mice, except in SCID mice. A neutrophil influx, accompanied by an increase in MIP-2 levels, was found in BAL of all responding groups 6 and 24 h after intranasal challenge. In BALB/c mice an increased level of CD19+ B cells was found in the auricular lymph nodes. IL-4 and IFN-γ levels were increased in supernatants of concanavalin A-stimulated auricular lymph node cells from BALB/c mice completely treated with TDI. These results indicate that our model is dependent on the presence of lymphocytes, but it is not characterized by a preferential stimulation of Th1 or Th2 lymphocytes.
- occupational asthma
- severe combined immunodeficiency disease mice
- murine model
- T lymphocyte subtypes
although the clinical presentation of occupational asthma does not depend much on the causative agent, it is common to distinguish occupational asthma caused by sensitization to high-molecular-weight agents from that caused by sensitization to low-molecular-weight chemicals. The former agents consist of (glyco)proteins of biological origin, which may cause IgE-mediated allergy and asthma in a fashion similar to common inhalant allergens that cause atopic asthma. Sensitization to low-molecular-weight chemicals results from a response of the immune system to haptens conjugated with endogenous proteins. However, the exact pathways and mechanisms of sensitization to such chemicals and the pathogenesis of the subsequent respiratory reactions are much less well understood, as they seem to differ from those of the classic IgE-mediated asthma (16).
We recently developed (18, 19, 21) a mouse model of chemical-induced asthma using toluene diisocyanate (TDI). This model, which involves dermal applications of TDI followed by an intranasal challenge with a nonirritant amount of TDI, reproduces several important features of human asthma, including 1) an immediate ventilatory response, consistent with bronchoconstriction, shortly after the challenge, 2) nonspecific hyperresponsiveness to methacholine 1 day after the challenge, and 3) airway inflammation (admittedly with neutrophils rather than eosinophils) (19). Furthermore, we have validated (20) this model with trimellitic anhydride, a known respiratory sensitizer, and dinitrochlorobenzene, a known dermal sensitizer.
Here we present studies carried out to understand the nature and mechanisms of the immune reactions underlying TDI-induced asthma. First we verified whether the observed respiratory responses were dependent on lymphocytes by applying our protocol to mice with severe combined immunodeficiency disease (SCID). We also determined, in wild-type BALB/c mice having undergone one or two dermal sensitizations, the kinetics of pulmonary inflammation at various time points (1, 6, and 24 h) after intranasal challenge with TDI. Furthermore, we characterized lymphocyte subpopulations in auricular and cervical lymph nodes draining the sites of skin and nose application of TDI, respectively.
Our results indicate that our model of TDI-induced asthma is dependent on the presence of lymphocytes, but it is not characterized by a preferential stimulation of Th1 or Th2 lymphocytes.
MATERIALS AND METHODS
Toluene-2,4-diisocyanate (TDI) (98%; Fluka CAS 584-84-9) and acetyl-β-methylcholine (methacholine) were obtained from Sigma Aldrich (Bornem, Belgium). Acetone, diethyl ether, and formaldehyde were obtained from VWR (Leuven, Belgium). Pentobarbital sodium (Nembutal) was obtained from Sanofi Santé animale (CEVA, Brussels, Belgium). The vehicle (AOO) used to dissolve TDI consisted of a mixture of 2 volumes of acetone and 3 volumes of olive oil (“extra virgin,” Carbonell). Concentrations of TDI are given as percentage (vol/vol) in AOO.
Male BALB/c mice (∼20 g, 6 wk old) were obtained from Harlan. The mice were housed in a conventional animal house with 12:12-h dark-light cycles. They received lightly acidified water and pelleted food (Trouw Nutrition, Ghent, Belgium) ad libitum. Specific pathogen-free male BALB/c-derived SCID mice (5–7 wk old) were obtained from the REGA institute (Katholieke Universiteit Leuven). They were housed in sterilized cages with filter tops and fed sterilized food and water ad libitum. All experimental procedures were approved by the local Ethical Committee for Animal Experiments.
Groups of Animals and Treatment Protocol
All mice received dermal applications of 20 μl of vehicle or 20 μl of 0.3% TDI on each ear on days 1 and 7. On day 10 they received, under light anesthesia with diethyl ether, an intranasal instillation of 10 μl of vehicle or 10 μl of TDI (0.1%) in each nostril. In all experiments, treatment with TDI is indicated as 1 and treatment with vehicle is indicated as 0. Thus, the 1/1/1 group consists of mice that received dermal applications of TDI (days 1 and 7) and an intranasal instillation of TDI (day 10), and the 0/0/0 control group consists of mice that received the AOO vehicle on all occasions. Other groups were 0/0/1, 1/0/1, and 0/1/1. The mice were killed by an overdose (90 mg/kg ip) of pentobarbital sodium at 1, 6, or 24 h after intranasal instillation.
Measurements of Ventilatory Function: Early Ventilatory Response and Airway Hyperreactivity
On day 10, just before the intranasal instillation, the ventilatory function of each mouse was recorded, under resting conditions, for 5 min in a whole body plethysmograph (EMKA Technologies, Paris, France). Immediately after the intranasal instillation, mice were placed again in the whole body plethysmograph (zero time point) and their ventilatory parameters were measured for 40 min. Every 30 s, the enhanced pause (Penh), which is a composite index indicative of airways obstruction (5), was calculated. The area under the curve (AUC) of Penh against time between 0 and 40 min was calculated for each individual mouse, and this figure was used for statistical analyses.
On day 11, i.e., 23 h after intranasal instillation, reactivity to methacholine was assessed in the whole body plethysmograph according to the procedure of Hamelmann et al. (5). Briefly, Penh was calculated for each mouse under resting conditions (baseline) and after inhalation, over 1 min, of nebulized methacholine (successively 0, 10, 25, 50, and 100 mg/ml). An average Penh over 30 s was calculated over 3 min (6 measurements), and the mean of these six values was used for each concentration. For each mouse, Penh was plotted against methacholine concentration (from 0 to 100 mg/ml) and the AUC was calculated.
In additional experiments carried out to validate the findings obtained by whole body plethysmography, airway responses were measured with a FlexiVent system (FlexiVent, SCIREQ, Montreal, QC, Canada). On day 11, i.e., 23 h after intranasal instillation, measurement of airway resistance (R) to methacholine was assessed. Mice were anesthetized with an intraperitoneal injection of pentobarbital sodium (70 mg/kg). The trachea was exposed and tracheostomized, and an 18-gauge metal needle was inserted. Mice were connected to a computer-controlled small-animal ventilator. The mice were quasi-sinusoidally ventilated with a tidal volume of 10 ml/kg at a frequency of 150 breaths/min and a positive end-expiratory pressure of 2 cmH2O to achieve a mean lung volume close to that during spontaneous breathing. After measurement of a baseline, each mouse was challenged with methacholine aerosol, generated with an in-line nebulizer and administered directly through the ventilator for 5 s, with increasing concentrations (0, 0.625, 1.25, 2.5, 5, and 10 mg/ml). R was measured with a “snapshot” protocol each 20 s for 2 min. The mean of these six values was used for each methacholine concentration. For each mouse, R was plotted against methacholine concentration (from 0 to 10 mg/ml) and the AUC was calculated.
Bronchoalveolar lavage (BAL) was performed 1, 6, or 24 h after intranasal instillation (in SCID mice only after 6 and 24 h). The lungs were lavaged in situ three times with 0.7 ml of sterile saline (0.9% NaCl) at room temperature, and the recovered fluid was pooled. Cells were counted with a Bürker hemocytometer (total cells), and the BAL fluid was centrifuged (1,500 g, 10 min). The supernatant was frozen (−80°C) until further analysis. For differential cell counts, 250 μl (100,000 cells/ml) of the resuspended cells was spun (1,400 g, 6 min) (Cytospin 3, Shandon, TechGen, Zellik, Belgium) onto microscope slides, air dried, and stained (Diff-Quik method). For each sample, 200 cells were counted for the number of macrophages, eosinophils, neutrophils, and lymphocytes.
Levels of tumor necrosis factor (TNF)-α (Biosource, Nivelles, Belgium) and macrophage inflammatory protein (MIP)-2 (R&D Systems, Abingdon, UK) were measured in undiluted BAL fluid by a sandwich enzyme-linked immunosorbent assay (ELISA), according to the manufacturer's instructions. Lower limits of detection were 3.0 and 1.5 pg/ml for TNF-α and MIP-2, respectively.
Lymph Node Cell Collection
Retroauricular and superficial cervical lymph nodes were obtained from BALB/c mice 24 h after intranasal instillation. The lymph nodes from 4 or 5 mice were pooled and kept on ice in RPMI-1640 (Invitrogen, Merelbeke, Belgium). Cell suspensions were obtained by pressing the lymph nodes through a cell strainer (100 μm) (BD Biosciences) and rinsing with 10 ml of tissue culture medium (RPMI-1640). Cells were counted with a Bürker hemocytometer. Lymphocytes were washed three times and suspended in complete tissue culture medium (RPMI-1640 supplemented with 10% heat-inactivated fetal bovine serum, 10 mg/ml streptomycin, 100 IU/ml penicillin, 1 mM sodium pyruvate, and nonessential amino acids) at concentrations of 107 cells/ml.
Five hundred thousand cells were stained with anti-CD3 phycoerythrin [(PE)]-, anti-CD4 fluorescein isothiocyanate (FITC)-, anti-CD25 [peridinin chlorophyll protein (PerCP)]-, or anti-CD19 (PE)-labeled antibodies (BD Biosciences, Erembodegem, Belgium), according to standard procedures, with control samples being labeled with isotype-matched control antibodies (BD Biosciences). Flow cytometry (CellQuest, BD Biosciences) was performed with at least 104 cells.
Cells were also seeded into 48-well culture plates at a density of 2 × 106/ml and incubated for 18 h without or with concanavalin A (ConA; 2.5 μg/ml) (Sigma, Bornem, Belgium). Cells were then centrifuged, and supernatants were stored at −80°C. Concentrations of interleukin (IL)-4 and interferon (IFN)-γ were measured in undiluted supernatants by standard ELISA technique, according to the manufacturer's instructions (Biosource). Lower limits of detection were 5 and 1 pg/ml for IL-4 and IFN-γ, respectively.
Total Serum IgE, IgG1, and IgG2a
At 1, 6, or 24 h after intranasal instillation, blood was drawn (before BAL) from the retroorbital plexus and pooled for three mice. Serum samples were stored at −80°C for further analysis. The OptEIA Mouse IgE and IgG2a set from Pharmingen (BD Biosciences) was used to measure total serum IgE (diluted 1/70) and total serum IgG2a (diluted 1/10,000). Measurements were performed according to the manufacturer's instructions. For measurement of total serum IgG1 (diluted 1/100,000), we coated our plates with purified anti-mouse IgG1, a standard was created with purified mouse IgG1, and further measurements were performed according to the manufacturer's instructions (BD Biosciences) with the use of biotinylated anti-mouse IgG1 and avidin-horseradish peroxidase conjugate.
All data were analyzed with the nonparametric Kruskal-Wallis test (GraphPad Prism 3.01). A level of P < 0.05 was considered significant.
Nonparametric Spearman correlation test and linear curve fitting were performed with SPSS 9.0 to describe correlations between “early-phase” ventilatory response and neutrophil influx or MIP-2 levels in BAL, 24 h after intranasal challenge, in the animals from the groups exhibiting a response (1/1/1, 1/0/1, and 0/1/1).
Figure 1 shows no significant changes in ventilatory responses following the intranasal instillation of TDI in TDI-naive animals (0/0/1 groups) either in wild-type BALB/c mice or in SCID mice. In the groups of wild-type BALB/c mice that had previously received dermal applications of TDI—once on day 1 (1/0/1), once on day 7 (0/1/1), or twice (days 1 and 7, 1/1/1)—there were significant and large increases in Penh after instillation of TDI, with average increases in AUC by 8–10 times compared with the 0/0/0 and 0/0/1 groups. These changes were not found in similarly treated SCID mice.
Figure 2 shows the airway responsiveness to methacholine on day 11 as measured by whole body plethysmography. The BALB/c mice that had received TDI on the skin (0/1/1, 1/0/1, and 1/1/1) and were then challenged with TDI showed an increased methacholine responsiveness compared with the 0/0/1 and 0/0/0 control groups. The mean AUC of the 1/0/1 and 0/1/1 groups was increased threefold compared with the 0/0/0 group. The 1/1/1 group showed a fourfold increase compared with the 0/0/0 group and a threefold increase compared with the 0/0/1 control group. SCID mice treated with either TDI or vehicle showed no increased airway responsiveness to methacholine compared with the control groups.
Figure 3 shows the airway responsiveness to methacholine on day 11 as measured by FlexiVent, by which a true value of R is obtained. Dermally TDI-treated mice (0/1/1, 1/0/1, and 1/1/1) showed an augmentation of airway resistance after exposure to increasing concentration of methacholine compared with the 0/0/0 control group. Only the completely treated TDI group (1/1/1) showed a significant 1.5-fold increase in AUC compared with both the 0/0/1 and 0/0/0 control groups. The 0/1/1 and 1/0/1 groups showed a significant 1.3- and 1.4-fold increase compared with the 0/0/0 control group, respectively.
Figure 4A shows the total cell counts (left) and neutrophil percentages (right) 1, 6, and 24 h after intranasal instillation in wild-type BALB/c mice, and Fig. 4B shows the same indexes in SCID mice 6 and 24 h after intranasal instillation. One hour after intranasal instillation, the groups did not differ from each other in terms of total cell counts. Six hours after intranasal instillation, total cell numbers were significantly different from the 0/0/0 and 0/0/1 groups in the 0/1/1 group. Twenty-four hours after intranasal instillation, all these groups (0/1/1, 1/0/1, and 1/1/1) showed significant increases (∼3-fold) in total cell numbers in BAL. In the SCID mice there were no differences in total cell counts 6 or 24 h after intranasal instillation.
In the 0/0/0 control animals, the average proportions of neutrophils in BAL were 1.8%, 1.0%, and 1.6% at 1, 6, and 24 h, respectively. At no time were there significant changes in cell type proportions in BAL after the intranasal instillation of TDI in TDI-naive animals (0/0/1 group) (neutrophils: 1.4%, 1.6%, 1.2%). At 1 h after intranasal instillation of TDI, significantly increased percentages of neutrophils were found in the 1/0/1 and 1/1/1 groups (4% and 6%, respectively) compared with the concurrent 0/0/0 and 0/0/1 control groups. At 6 h, all three groups pretreated dermally with TDI (0/1/1, 1/0/1, and 1/1/1) showed increased percentages of neutrophils (8%, 16%, and 16%, respectively). Twenty-four hours after intranasal challenge, the neutrophil influx was further increased to 20% in the 1/1/1 group, 36% in the 1/0/1 group, and 29% in the 0/1/1 group. In SCID mice, small, nonsignificant increases in BAL neutrophils were found in 1/1/1 and 0/1/1 groups. In all groups, mainly macrophages and neutrophils were present in BAL, with only one or two eosinophils and lymphocytes being found per cytospin.
Because ventilatory function data and BAL data were obtained from the same animals, it was possible to verify whether early ventilatory changes correlated with subsequent inflammatory changes in the lungs within individual animals. Thus there was a significant correlation between values of Penh (expressed as AUC) and proportions of BAL neutrophils (expressed in %) at 24 h, when mice from 0/1/1, 1/0/1, and 1/1/1 groups were considered (Spearman correlation coefficient of 0.74, P < 0.01; n = 18).
Figure 5 shows MIP-2 concentrations in BAL fluid 1, 6, and 24 h after intranasal instillation. Six hours after intranasal challenge, significantly increased levels of BAL MIP-2 were found in the 0/1/1 group compared with the 0/0/0 control group. Twenty-four hours after intranasal instillation, a significant increase was observed in all airway-responding groups (0/1/1, 1/0/1, and 1/1/1) compared with the 0/0/0 control group. The MIP-2 concentration in BAL 24 h after intranasal instillation of the 1/0/1 group was also statistically different from that of the 0/0/1 control group. MIP-2 concentrations significantly correlated with the percentage of neutrophils in BAL (Spearman correlation coefficient 0.3, P < 0.05; n = 18) in the 1/1/1, 1/0/1, and 0/1/1 groups. TNF-α levels in BAL fluid showed considerable variability in all groups and at all time points; no significant differences were found between any of the groups (data not shown).
Lymph Node Cells
Table 1 shows results of flow cytometric analyses of lymphocyte subpopulations in auricular and cervical lymph nodes removed 24 h after intranasal instillation of TDI or vehicle. An increase in the proportion of CD19+ lymphocytes (B cells) was apparent in the dermally TDI-treated groups (1/1/1, 1/0/1, and 0/1/1) compared with the 0/0/0 and 0/0/1 groups, but this increase was only statistically significant (compared to the 0/0/0 control group) for the 0/1/1 and the 1/1/1 groups. The proportions of other lymphocyte subpopulations (CD3+CD4+ and CD3+CD4+CD25+) did not differ significantly between groups. There were no differences in CD3+CD8+ lymphocytes, and the expression of CD25+ on CD3+CD8+ T cells was barely detectable (data not shown). In the cervical lymph nodes no significant differences in lymphocyte subpopulations between the control and treatment groups was found.
In vitro cytokine release.
Figure 6, A and B, show the levels of IL-4 and IFN-γ, respectively, in supernatants of auricular (left) and cervical (right) lymph node cells cultured for 18 h without and with ConA stimulation. The spontaneous release of IL-4 and IFN-γ was low and not statistically different between the groups. We therefore calculated an average spontaneous release and indicated this as a dotted line in Fig. 6. In ConA-stimulated auricular lymphocytes, levels of both IL-4 and IFN-γ were significantly increased in the supernatants from the 1/1/1 group compared with those from the 0/0/1 and 0/0/0 control groups. No differences were found in the lymphocytes of the cervical lymph nodes.
Serum Total IgE, IgG1, and IgG2a
Figure 7 shows total serum IgE, IgG1, and IgG2a. We pooled the values of the different time points (1, 6, and 24 h) because no differences were found between these three time points. Total IgE levels were significantly higher than in controls (0/0/0 and 0/0/1) only in mice from the 1/1/1 group (Fig. 7A). All dermally TDI-treated mice showed a significantly increased level of total serum IgG1 compared with the 0/0/1 and 0/0/0 control mice (Fig. 7B). There were no differences in total serum IgG2a levels among all groups (Fig. 7C).
The present study confirms that both the ventilatory responses and the pulmonary inflammation observed in our model of chemical-induced asthma (19, 21) depend on the presence of lymphocytes, since these responses did not occur in SCID mice. Furthermore, we show that the influx of neutrophils in the lungs starts a few hours after challenge and that activation of various types of lymphocytes takes place in this model of chemical-induced asthma.
The individual values in Figs. 1 and 2 reveal substantial variations in the magnitude of the responses within the responsive groups. This variability is not simply due to experimental variation in the measurement of Penh; it reflects authentic and consistent individual variations in biological responses. Correlation analysis demonstrated that, within the responding groups, the mice exhibiting the largest increases in Penh also tended to have most neutrophils in BAL. Consequently, the absence of responses in SCID mice, compared with immunocompetent BALB/c mice, is reliable even though fewer SCID mice were studied. A few SCID mice of the dermally treated groups did exhibit small alterations in Penh and increases in BAL neutrophils, but these (admittedly unexplained) changes were of low magnitude. The lymphocyte dependence of the ventilatory and inflammatory responses further supports the idea that they are based on sensitization to TDI and not on a “toxic” response. No other studies of chemical-induced asthma have been performed in SCID mice. However, others have shown the lymphocyte dependence of TDI-induced airway hyperreactivity development with athymic mice, but they did not study early responses to allergen challenge and the protocols used in these experiments differed considerably from our model (11, 17).
Whole body plethysomography has been criticized as being inadequate for assessing airway mechanics (8, 12). Although an increase in Penh does not necessarily reflect bronchial obstruction correctly, there is no doubt (because our design included all necessary control groups) that early ventilatory responses and increases in methacholine responsiveness occurred in TDI-sensitized and then intranasally instilled mice. Furthermore, we validated the measurements of bronchial responsiveness obtained by whole body plethysmograph by more reliable measurements of airway resistance.
In the three responsive groups (0/1/1, 1/0/1, and 1/1/1), an increase in BAL neutrophils appeared 1 h after instillation of TDI, with further increases at 6 and 24 h (Fig. 4). MIP-2, a chemoattractant for neutrophils in mice (1), also increased (Fig. 5). However, MIP-2 was only measurably increased when the influx of neutrophils had been already manifested. This is presumably due to the lower measurement variability for neutrophils than for MIP-2. Concentrations of TNF-α in BAL were not changed at all. However, this does not completely rule out that the irritant properties of TDI play a role in its effects as a potent asthmogen.
The absence of lymphocytes in BAL is puzzling, particularly in view of our SCID experiment, which indicates that lymphocytes are necessary in our model. This needs more research. The influx of neutrophils rather than eosinophils is not contradictory with allergic asthma. Neutrophilic inflammation has also been observed in TDI asthmatics (13). The nature of the pulmonary inflammation in asthma is presumably heavily dependent on the time course of the disease and the pattern of the exposure.
Although the involvement of Th2 cytokines and IgE is well accepted in atopic asthma, examples are known in which asthma exists in the absence of detectable levels of IgE (2, 14, 22). Haptens do not necessarily need intracellular processing and protein binding to induce an antigenic response, as some chemicals can cross-link major histocompatibility complex II molecules of antigen-presenting cells and T cell receptor (3, 4).
We found an increased percentage of B cells in the draining lymph nodes and significantly increased total serum IgE and IgG1 in completely TDI-treated mice (1/1/1), thus supporting the notion that Th2 stimulation had occurred. However, the absence of significant changes in the other two groups (0/1/1 and 1/0/1), in which the respiratory responses were equally pronounced, suggests that the increase in total IgE is not critical in our model.
Both IL-4 and IFN-γ were increased in lymphocytes obtained from the auricular nodes of the completely TDI-treated group (1/1/1) (Fig. 6). These experiments do not provide definitive answers to the question of the nature of the lymphocyte activation in TDI-induced asthma, but they suggest that both Th1 and Th2 lymphocytes are increased, at least at the time of the analysis. This was also observed in other mouse models of isocyanate asthma (6, 9, 10). Both the Th and the Tc lymphocytes appear to be important for the ventilatory, inflammatory, and immunologic responses in mouse models of TDI-induced asthma, since these responses are almost gone in CD4- and CD8-knockout mice (9). We did not find any differences in the cervical lymph nodes after a single intranasal instillation. However, multiple intranasal challenges yield the same cytokine profile (mixed Th1-Th2) in the cervical lymph nodes as in the auricular lymph nodes (unpublished data).
A limitation of our experiments is that only one time point was studied. Although the sampling of lymph nodes was done 1 day after nasal instillation of TDI in all groups, the interval between the lymphocyte analysis and the last dermal application of TDI varied between groups (9 days for group 1/0/1 and 3 days for groups 0/1/1 and 1/1/1). Another limitation is that we did not characterize the mediastinal lymph nodes draining the lungs, because they did not contain enough cells to perform further characterization, at least after a single intranasal instillation.
The occurrence of respiratory responses in the 0/1/1 group is interesting and also intriguing. We did not anticipate that this “control” group would show any reaction, because we believed that the interval between the first dermal application (on day 7) and the nasal instillation (on day 10) would be too short for an immune response to be mounted. However, the respiratory responses in the 0/1/1 group proved to be qualitatively similar to those found in the 1/0/1 and 1/1/1 groups, in which the delay between sensitization (day 1) and challenge (day 10) was long enough, based on conventional wisdom (7). Our findings with the 0/1/1 group suggest that the time required for mounting an allergic response to a chemical may be as short as 72 h after sensitization. This may be a particular feature of chemical sensitizers, as opposed to protein antigens. Thus as shown previously, a single dermal application of 2,4-dinitrofluorobenzene can induce contact dermatitis if reapplied after 4 days (15).
In conclusion, our studies confirm that respiratory allergic responses to a chemical sensitizer such as TDI may occur after dermal sensitization. The complete mechanisms of these reactions remain to be elucidated, but the present findings indicate that the reactions are lymphocyte dependent, but without fitting a simplistic paradigm of Th2 stimulation or IgE-mediated mechanisms.
We thank the Belgian Federal Science Policy Office (PS/01/43) and Fund for Scientific Research Flanders (FWO) (project 7.0024.00) for funding this research.
↵* M. Tarkowski and J. A. J. Vanoirbeek contributed equally to this work.
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- Copyright © 2007 the American Physiological Society