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The Journal of Immunology, 2000, 164: 2937-2946.
Copyright © 2000 by The American Association of Immunologists

Induction of Rapid T Cell Activation, Division, and Recirculation by Intratracheal Injection of Dendritic Cells in a TCR Transgenic Model1

Bart N. Lambrecht2,*,{dagger}, Romain A. Pauwels{dagger} and Barbara Fazekas de St. Groth3,*

* Centenary Institute of Cancer Medicine and Cell Biology, Newtown, Sydney, Australia; and {dagger} Department of Respiratory Diseases, University Hospital, Ghent, Belgium


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Dendritic cells (DCs) are thought to be responsible for sensitization to inhaled Ag and induction of adaptive immunity in the lung. The characteristics of T cell activation in the lung were studied after transfer of Ag-pulsed bone marrow-derived DCs into the airways of naive mice. Cell division of Ag-specific T cells in vivo was followed in a carboxyfluorescein diacetate succinimidyl ester-labeled cohort of naive moth cytochrome c-reactive TCR transgenic T cells. Our adoptive transfer system was such that transferred DCs were the only cells expressing the MHC molecule required for presentation of cytochrome c to transgenic T cells. Ag-specific T cell activation and proliferation occurred rapidly in the draining lymph nodes of the lung, but not in nondraining lymph nodes or spleen. No bystander activation of non-Ag-specific T cells was induced. Division of Ag-specific T cells was accompanied by transient expression of CD69, while up-regulation of CD44 increased with each cell division. Divided cells had recirculated to nondraining lymph nodes and spleen by day 4 of the response. In vitro restimulation with specific Ag revealed that T cells were primed to proliferate more strongly and to produce higher amounts of cytokines per cell. These data are consistent with the notion that DCs in the lung are extremely efficient in selecting Ag-reactive T cells from a diverse repertoire. The response is initially localized in the mediastinal lymph nodes, but subsequently spreads systemically. This system should allow us to study the early events leading to sensitization to inhaled Ag.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The initiation of adaptive T cell immunity in the lung requires inhaled Ag to be taken up by professional APCs, processed into immunogenic peptides, and presented in the context of MHC molecules to naive T cells recirculating in the lymph nodes draining the lung (1, 2). An extensive network of Ag-presenting dendritic cells (DCs)4 lines the conducting airways and interstitium of the lung and is capable of presenting inhaled Ag to naive and memory T cells ex vivo (2, 3, 4, 5, 6). In the absence of inflammation or exogenous adjuvant, local T cell activation in the lung is limited by the immaturity of intraepithelial DCs and by the immunosuppressive effects of resident macrophages on T cell activation (2, 7). Furthermore, the function of DCs in capturing and presenting inhaled Ag is under strict microenvironmental control mediated via factors such as NO, TGF-ß, and PGE2 secreted by alveolar macrophages (7, 8). Inflammatory stimuli, such as those derived from microbial invasion, attract immunostimulatory monocytes and DCs from the circulation to the lung, and induce secretion of proinflammatory cytokines such as GM-CSF and TNF-{alpha} by resident epithelial cells, macrophages, and fibroblasts (9, 10, 11). Maturation and migration of airway DCs to the T cell area of the lymph nodes, analogous to the migration of Langerhans cells from the skin, are poorly understood, but are considered to be essential for mature DCs to interact with naive recirculating T cells (12, 13). It is likely that this initial interaction between DCs and T cells, leading to mutual activation, determines the quality of the subsequent immune response. Exogenous factors thought to influence the outcome of T cell priming, such as the timing, route, and dose of Ag administration, and the type of adjuvant, may act at the level of the DC-T cell interaction (14, 15, 16, 17, 18). Thus, understanding the induction and regulation of adaptive immunity by DCs should provide insights into the pathogenesis and treatment of infectious, neoplastic, and allergic diseases of the lung.

Despite a growing awareness of the central role of DCs in the induction and regulation of pulmonary immunity, few studies have directly addressed the outcome of T cell priming by airway DCs. To model the early events during T cell activation in response to inhaled Ag, we have injected Ag-pulsed myeloid DCs into the trachea of naive animals. The adoptive transfer of purified Ag-pulsed splenic DCs has been used by others to study T cell priming after intratracheal injection (19). Its usefulness in addressing the question of whether DCs alone are responsible for priming the naive immune system is limited by the possibility that Ag may be eluted from purified DCs and presented by more potent endogenous APCs (19, 20). Moreover, the study of naive T cell activation in vivo is technically demanding because of the low precursor frequency of Ag-specific T cells. We have made use of a model in which the adoptive transfer of a cohort of TCR transgenic (Tg) T cells to naive syngeneic hosts allows the fate of individual Ag-specific T cells to be followed after activation by intratracheal injection of Ag-pulsed DCs. Our experimental approach is unique in that expression of the MHC molecule required for presentation of Ag is restricted to the injected DCs, excluding cross-presentation by endogenous APCs. Using this model, the localization and kinetics of T cell activation, division, differentiation, and recirculation were determined in the primary immune response to an Ag delivered by professional APCs in the lung.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Mice

Tg mice were bred and maintained under specific pathogen-free conditions at the Centenary Institute animal facility. Approval for all animal experimentation was obtained from the Institutional Ethics Committee at the University of Sydney. TCR Tg mice specific for the COOH-terminal epitope of moth cytochrome c (MCC87–103) in the context of I-E{alpha}kßk, I-E{alpha}kßb, or I-E{alpha}dßb expressed the rearranged V{alpha}11 and Vß3 genes derived from the 5C.C7 T cell clone under the control of the endogenous TCR 3' ß enhancer, as previously described (21, 22). This Tg line, termed -D, expresses the transgene-encoded ß-chain on >95% of CD4+ T cells, and I-E-mediated thymic positive selection results in expression of the Tg {alpha}-chain (Tg{alpha}+, recognized by staining with the mAb RR8.1 anti-V{alpha}11 (23)) on 50–80% of peripheral CD4+ T cells, depending on the age and sex of the mouse. The remainder of CD4+ T cells express an endogenously rearranged {alpha}-chain, paired with the 5C.C7 ß-chain.

The 107-1 line (24), expressing I-E{alpha}d as a transgene under the control of the endogenous MHC class II promoter, and the 36-2 line (24), in which expression of the same I-E{alpha}d transgene is restricted to thymic epithelial cells, were originally the gift of D. Lo (Scripps Research Institute, La Jolla, CA). In these lines, the I-E{alpha}d transgene combines with the endogenous I-Eßb chain from the H-2b host (C57BL/6, I-E{alpha} negative) to form I-E{alpha}dßb, which mediates both positive selection of 5C.C7 TCR T cells in the thymus and presentation of MCC87–103 to the same T cells in the periphery. Donors of T cells for adoptive transfer into 107-1 or 36-2 recipients were F1 offspring of -D (C57BL/6 background) x 107-1 crosses, selected for expression of both transgenes (termed -D 107-1).

Antigens

MCC87–103 peptide, containing residues 87–103 derived from moth cytochrome c (KANERADLIAYLKQATK), was obtained from the Queensland Institute of Medical Research (Brisbane, QLD, Australia).

Isolation and adoptive transfer of TCR Tg T cells

Pooled peripheral lymph nodes (cervical, mediastinal, brachial, subscapular, inguinal, paraaortic, and popliteal) were harvested from -D 107-1 mice and cell suspensions prepared in tissue culture medium (TCM), as previously described (25). Red cell lysis, depletion of B cells and APCs, and labeling with carboxyfluorescein diacetate succinimidyl ester (CFSE) were performed as described (25). Viable cells were enumerated by eosin exclusion before transfer into unirradiated 107-1 or 36-2 recipients. On average, each recipient received 25 x 106 labeled cells i.v. via the lateral tail vein.

Isolation of APCs

DCs were grown from bone marrow precursors obtained from 107-1 donors using a modification of the Inaba protocol (26). In brief, bone marrow cells were resuspended in Tris-ammonium chloride lysis buffer solution and washed twice with TCM. Cells were incubated for 30 min on ice with a mixture of complement-fixing Abs: anti-B220 (RA3-3A1; Ref. 27), anti-CD4 (RL172.4; Ref. 28), anti-CD8 (3.168; Ref. 29), anti-MHC class II (M5/114; Ref. 30), and anti-Gr-1 (RB6-AC5; PharMingen, San Diego, CA). Prewarmed rabbit complement (C-SIX Labs, Mequon, WI) was added for 30 min at 37°C. Cells were washed twice and resuspended in DC culture medium (TCM supplemented with 2.5 ng/ml of murine rGM-CSF (Serotec, Kidlington, U.K.) and 0.5 µg/ml of N-{gamma}-monomethyl-L-arginine (Calbiochem, La Jolla, CA)) at a concentration of 0.6 x 106 cells/ml in 24-well tissue culture plates (Becton Dickinson, Mountain View, CA ). DCs were harvested after 7 days of culture by removing the weakly adherent clusters and replating the cells in plastic petri dishes for an additional 12 h. The nonadherent cell fraction was collected, washed twice in TCM, and enriched by density centrifugation (600 x g, 22°C, 20 min) over a metrizamide gradient (14.5% w/v in RPMI 1640, density 1.0745 g/ml; Sigma, St. Louis, MO). The purity of low density DCs was assessed by staining cells with M5/114-FITC (anti-MHC class II, prepared in house) and biotinylated mAb against I-E{alpha}d (14.4.4), B7-1(16-10A1), B7-2(GL-1), heat-stable Ag (J11D), CD44 (IM7), CD23 (B3B4), DEC-205 (NLDC-145), 33D1, and CD11c (N418), followed by avidin-Quantum Red (Sigma) and analyzing on a FACScan flow cytometer (Becton Dickinson). Abs were acquired from the American Type Culture Collection (Manassas, VA), prepared, and biotinylated in-house or obtained from PharMingen. Cytospins (Cytospin I, Shandon, U.K.) of ethanol-fixed cells were stained with M5/114, followed by secondary rabbit anti-rat HRP (Dako, Glostrup, Denmark), and signal was developed using diaminobenzidine substrate (DAB, SigmaFast; Sigma).

Immunization by intratracheal injection of peptide-pulsed DCs

After enrichment, DCs were pulsed for 1 h with 5 µM MCC87–103 dissolved in TCM. Control DCs were incubated in peptide-free TCM. After pulsing, cells were extensively washed to remove unbound peptide, resuspended at 12.5 x 106 cells/ml in PBS, and adoptively transferred into the trachea of 36-2 mice that had received a cohort of -D 107-1 T cells 2 days earlier (see above). For intratracheal injection, mice were anesthetized by i.p. injection of 2.5% avertin (Sigma), and a volume of 80 µl containing 106 DCs was injected under direct vision through the opening vocal cords using a 25 gauge metal catheter connected to the outlet of a micropipette. The control group of animals received an injection of 80 µl of PBS containing 100 µg MCC87–103 without any DCs.

For intranasal immunization, 100 µg MCC87–103 peptide in a final volume of 20 µl was administered to the nares under light ether anesthesia. Control animals received 20 µl PBS.

Flow cytometry

On days 2, 4, and 7 after the adoptive intratracheal transfer of DCs, mice were killed by CO2 asphyxiation, and the spleen and superficial cervical, deep cervical, parathymic, mediastinal, subscapular, brachial, inguinal, and mesenteric lymph nodes were removed and individual cell suspensions prepared as described (25). Cells were washed twice with PBS containing 5% FCS and 5 mM sodium azide (FACS wash). Aliquots of 106 cells were stained for five-color immunofluorescence in 96-well round-bottom microtiter plates (ICN, Costa Mesa, CA). All staining reactions were performed for 30 min on ice. As CFSE fluorescence of transferred T cells is detected in the FL-1 channel, no FITC-conjugated Abs were used. For the first staining combination, cells were incubated with RR8 supernatant (rat IgG1, anti-V{alpha}11), followed by secondary anti-rat Texas Red (Caltag, Burlingame, CA). After blocking with 1% normal rat serum for 10 min, a third layer of Abs consisting of anti-CD4-PE (PharMingen) and biotinylated anti-CD69 (PharMingen) was applied. The final layer consisted of streptavidin-allophycocyanin (av-APC; Molecular Probes, Eugene, OR). The second Ab combination consisted of anti-CD44 supernatant (IM7, rat IgG1), followed by anti-rat TR, blocking with rat serum, and application of RR8-bi and anti-CD4-PE, followed by av-APC. Propidium iodide (PI) was added to every staining combination for exclusion of dead cells, before analysis on a FACStarPlus (Becton Dickinson) equipped with a 488-nm argon-ion laser and a 610-nm dye laser. Cells were gated for lymphocytes on the basis of forward angle and side-scatter profiles, and 2–10 x 105 events were collected using Lysis II software (Becton Dickinson). Final analysis and graphical output were performed using FlowJo software (Treestar, Costa Mesa, CA).

Calculation of CFSE content and recruitment into cell division

The term "CFSE content" refers to the absolute amount of CFSE within the donor-derived cell population of interest, independent of the total number of cells derived from division. It therefore gives an estimate of the original number of CFSE-labeled donor cells from which the donor-derived, divided population has arisen, and can be used to calculate whether the number of cells at any site has been affected by recruitment, migration, or cell death, in addition to division.

Total CFSE content within adoptively transferred CD4+Tg{alpha}+ cells was calculated by dividing the number of cells in each cell division peak by 2i, where i = the cell division number of that particular peak. Thus, CFSE content is as follows: f = {Sigma}ni/2i, where ni = the number of cells in the ith division peak. No cells divided more than seven times, so that: f = n0 + n1/2 + n2/4 + n3/8 + n4/16 + n5/32 + n6/64 + n7/128.

The percentage of cells recruited into cell division was calculated by dividing the number of undivided cells by CFSE content, using the following formula: 100 x {1 - [n0/(n0 + n1/2 + n2/4 + n3/8 + n4/16 + n5/32 + n6/64 + n7/128)]}.

In vitro restimulation of T cells

On day 4 after intratracheal transfer of peptide-pulsed DCs, individual cell suspensions of draining and nondraining lymph nodes were prepared as above. For proliferation assays, six serial 2-fold dilutions, starting at 2 x 105 cells/well, were made in TCM in flat-bottom 96-well plates (Falcon; Becton Dickinson). To provide saturating numbers of APCs in each well, 105 irradiated (1500 R) syngeneic spleen cells were added per well. Cells in a final volume of 200 µl/well were stimulated by addition of 5 nM MCC87–103 peptide. After 66 h, cells were pulsed for 6 h by addition of 0.5 µCi/well of [3H]thymidine. Cells were harvested using an automated cell harvester, and [3H]thymidine incorporation was measured in a beta scintillation counter, as described (25).

For measurement of cytokine production, bulk cultures were set up in a final volume of 2 ml in 24-well plates, using 2 x 106 lymph node cells, 106 irradiated syngeneic spleen cells as APCs, and 10 µM MCC87–103. Supernatants were collected and frozen at 48 h for IL-4 and 72 h for IL-3 and IFN-{gamma} measurement.

Cytokine measurements

IFN-{gamma} was measured in a capture ELISA, as described (22). The limit of detection was 0.78 ng/ml. IL-3 was measured by [3H]thymidine incorporation of the IL-3-dependent cell line R6X, as described (22). One unit/milliliter was defined as the dilution that yielded 50% of maximal [3H]thymidine incorporation. The limit of detection was 0.03 U/ml. IL-4 was measured by [3H]thymidine incorporation of the IL-4-dependent cell line CT.4S in response to serial dilutions of culture supernatants (31). After incubation for 24 h, cells were pulsed and harvested as for proliferation assays. A standard curve was generated using serial dilutions of murine rIL-4; 1 U/ml was defined as the dilution that yielded 50% of maximal [3H]thymidine incorporation. The limit of detection was 0.02 U/ml.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Purification of DCs

DCs were generated in vitro from bone marrow progenitor cells, as described in Materials and Methods. More than 90% of cells had numerous surface extensions or veils and a typical indented or clover leaf-shaped nucleus (not shown). Staining for MHC class II revealed a pool of intracellular MHC molecules (in MHC class II compartments) in addition to surface expression (not shown) (32). On flow-cytometric analysis, cells were invariably positive for MHC class II (I-Ab and I-Eb/d), and expressed costimulatory molecules heat stable Ag, CD80, and CD86). All cells expressed the DC marker CD11c at high levels and the markers 33D1 and NLDC-145(DEC 205) at lower levels (data not shown). Thus, the cells had a phenotype typical of cultured DCs (26).

An MHC-restricted model for T cell immune responses induced by DCs

The adoptive transfer of Ag-laden splenic DCs into the trachea of a syngeneic host has been used to study T cell activation in the draining mediastinal lymph nodes of the rat (19, 33). However, this methodology carries the risk that Ag may be eluted from the transferred APCs and presented by endogenous APCs (19, 20). To avoid this technical difficulty, Ag presentation can be restricted to a subpopulation of injected cells on the basis of exclusive expression of an MHC allele required for presentation. Our experimental protocol (34, 35), designed to limit Ag presentation to the injected DCs, made use of two MHC Tg lines in which I-E{alpha}d was expressed on the H-2b background, allowing it to pair with endogenous I-Eßb to form a functional I-E molecule (24, 36) capable of presenting MCC87–103 peptide to naive Tg T cells expressing the 5C.C7 TCR. Mice from the 36-2 line, expressing I-E{alpha}d only in the thymus, sufficient for inducing tolerance to I-E in the T cell compartment, were used as adoptive hosts of purified responder Tg T cells and I-E-positive DCs. As the 36-2 host lacks expression of I-E on peripheral APCs, the injected DCs from the 107-1 donor, expressing I-E with a wild-type distribution, were the only APCs capable of presenting MCC87–103 to the transferred TCR Tg T cells (Fig. 1Go). Two days before injection of DCs, unirradiated 36-2 mice received a cohort of purified, CFSE-labeled -D 107-1 Tg T cells. In the absence of antigenic stimulation, undivided transferred T cells were traceable by detection of CFSE staining for many weeks after adoptive transfer, indicating that there was no rejection of donor T cells by the host (data not shown; 35).



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  		FIGURE 1. A MHC-restricted model in which transferred I-E+ DCs are the only APCs capable of presenting MCC peptide (MCC87–103) to adoptively transferred TCR Tg T cells. Mice from the 107-1 line (in which Tg I-E{alpha}d is expressed with a wild-type distribution on all APCs, paired with endogenous I-Eßb) served both as donors of I-E+ DCs for adoptive transfer and as parents of (-D TCR Tg x 107-1)F1 T cell donors. Mice from the 36-2 Tg host are tolerant of I-E+ cells as a result of thymic expression of the I-E{alpha}d transgene, but express no I-E on peripheral APCs. On day -2, 25 x 106 CFSE-labeled MCC-reactive (107-1 x -D) responder T cells were injected i.v. into 36-2 hosts. On day 0, recipients were immunized by intratracheal injection of 106 MCC87–103-pulsed or PBS-pulsed DCs. A control group of 36-2 mice received MCC87–103 in the absence of DCs. The response was followed in all accessible nodes and spleen on days 2, 4, and 7 after immunization.

 
T cell proliferation in draining lymph nodes after intratracheal injection of DCs

On day 0 of the experiment, 106 MCC-pulsed or unpulsed DCs were injected into the trachea of 36-2 mice. On days 2, 4, and 7, flow cytometry was used to track cell division of adoptively transferred CFSE-labeled CD4+ cells in all accessible lymph nodes and spleen. Fig. 2GoA shows the distribution of the T cell response 2 days after intratracheal injection of MCC-pulsed DCs. In the draining mediastinal lymph nodes, a subpopulation of CD4+ Tg{alpha}+ T cells (specific for MCC87–103) had already undergone two cell divisions within 2 days of transfer of MCC-pulsed DCs. No division was seen in nondraining lymph nodes, including the superficial cervical, deep cervical, mesenteric, and peripheral (pooled brachial, subscapular, inguinal, and paraaortic) groups. Fig. 2GoB illustrates the kinetics of T cell division in the mediastinal lymph nodes after transfer of MCC-pulsed (upper panels) or unpulsed DCs (middle panels). On day 4, up to six cell divisions could be visualized within the CD4+ Tg{alpha}+ population. By day 7, the total number of divided cells had decreased substantially due, at least in part, to recirculation (see below). Immunization with PBS-pulsed DCs failed to induce a response in CD4+ Tg{alpha}+ T cells (Fig. 2GoB, middle panels). Moreover, very few CFSE-labeled CD4+ Tg{alpha}- cells (i.e., those not specific for MCC87–103) underwent cell division in response to pulsed or unpulsed DCs (Fig. 2GoB, upper and middle panels). The presence of divided cells within the Tg{alpha}- gate on day 7 of the response (Fig. 2GoB, top right panel) was an artifact due to a poor stain for the Tg{alpha}-chain on that day (as indicated by the decrease in the intensity of fluorescence in the Tg{alpha} channel), so that divided Tg{alpha}+ cells overlapped the region in which Tg{alpha}- cells would normally be located. Comparison with a day 7 plot from an experiment with a clearer Tg{alpha} stain (e.g., Fig. 4GoB, top right panel) indicated that the small degree of spontaneous division within the CD4+ Tg{alpha}- population gave a CFSE profile distinct from that in Fig. 2GoB.



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  		FIGURE 2. Local response of CFSE-labeled T cells after intratracheal injection of I-E+ DCs. Lymph node T cells were gated for scatter characteristics, expression of CD4, and exclusion of PI. The cohort of adoptively transferred T cells can readily be identified by strong green fluorescence of CFSE (x-axis). About 50% of CFSE-positive CD4+ cells express the transgene-encoded TCR {alpha}-chain, recognized by the RR8 Ab (y-axis). Cell division in Tg{alpha}+ cells can be observed by sequential halving of CFSE fluorescent signal. A, CFSE division profile of T cells in the various lymph nodes sampled 2 days after immunization with 106 MCC-pulsed DCs. B, Division profiles in draining mediastinal lymph nodes of the lung, taken at 2, 4, and 7 days after intratracheal immunization with MCC-pulsed DCs (upper panels), PBS-pulsed DCs (middle panels), or free MCC peptide (lower panels).

 


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  		FIGURE 4. Local response of CFSE-labeled T cells after intranasal administration of free MCC87–103 peptide to 107-1 mice with a wild-type distribution of I-E. Two days before immunization, mice received a cohort of MCC-reactive TCR Tg T cells. Lymph node T cells were gated based on scatter characteristics, expression of CD4, and exclusion of PI. A, Distribution of the response 2 days after intranasal administration of 100 µg MCC87–103. B, Kinetics of the response in mediastinal lymph nodes 2, 4, and 7 days after administration of MCC87–103 (upper panels) or PBS (lower panels).

 
The 36-2 host has been shown to express Tg I-E only in the thymus, and thus to be incapable of presenting MCC87–103 peptide (24, 36). We also tested whether donor-derived APCs were capable of presenting peptide in the experiment described above by injecting free MCC87–103 into the lower airways. A minor degree of peptide-dependent proliferation of CD4+ Tg{alpha}+ T cells was seen on day 7 (Fig. 2GoB, lower panels). This is consistent with the previously documented response to donor-derived B cells (34), which in this particular experiment were minor contaminants of the T cell preparation. However, the response was far lower than that to the injected DCs, with no early proliferation on day 2 and recruitment of only a few cells into one to two divisions by day 7. Thus, this response was unlikely to have influenced that to peptide-pulsed DCs, which preceded it, and was of greater magnitude.

Fig. 3Go quantitatively summarizes the data derived from mediastinal nodes in the experiment described above. The total number of CD4+ Tg{alpha}+ cells was significantly higher in MCC-DC- than PBS-DC-immunized mice on all days of the response, with the peak at day 4 (Fig. 3GoA). Although this was in part due to cell division (Fig. 2GoB), specific recruitment of peptide-reactive cells to the mediastinal node was also apparent, as indicated by the increased CFSE content of the mediastinal node CD4+ Tg{alpha}+ population in the MCC-DC group compared with the PBS-DC controls on all days of the response (Fig. 3GoC). Calculation of the number of donor-derived cells recruited into cell division indicated that 23% of the cytochrome c-reactive CD4+ T cells had undergone cell division as a result of intratracheal immunization with peptide-pulsed DCs, giving rise to progeny that constituted 56% of the CD4+ Tg{alpha}+ cells at that site on day 4 (Fig. 3GoB). The total number and CFSE content of CD4+ Tg{alpha}- cells were not significantly different in the MCC-DC group compared with the PBS-DC controls (Fig. 3Go, A and C), confirming the specificity of the response to pulsed DCs.



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  		FIGURE 3. Quantitative summary of data from the experiment described in the legend to Fig. 2Go. A, The number of donor-derived CD4+ Tg{alpha}+ (left panel) or CD4+ Tg{alpha}- (right panel) T cells per 105 live mediastinal lymph node cells at various days of the response after immunization with 106 MCC-pulsed or PBS-pulsed DCs. Individual points represent the mean ± SEM from the group. B, The percentage of donor-derived CD4+ Tg{alpha}+ T cells in mediastinal lymph nodes that have divided at least once after immunization with MCC-pulsed DCs, PBS-pulsed DCs, or free MCC peptide. N.D., not determined. C, The CFSE content, calculated as described in Materials and Methods, within donor-derived CD4+ Tg{alpha}+ and CD4+ Tg{alpha}- populations in the mediastinal lymph nodes at various days of the response. Individual points represent the mean ± SEM from the group.

 
Proliferative response to intranasal peptide

Initiation of a pulmonary immune response by means of intratracheal injection of a bolus of purified bone marrow DCs may not reflect the in vivo situation in which Ag is presented by endogenous DCs. To compare Ag presentation by injected DCs with presentation by endogenous DCs, the response to purified MCC87–103 peptide was studied in mice with a wild-type distribution of I-E that had received a cohort of CFSE-labeled MCC-reactive T cells 2 days before intranasal instillation of 100 µg MCC87–103 peptide. As expected for a soluble Ag, the response to intranasal peptide was more widespread than that to pulsed DCs. Two days after intranasal administration of peptide, CD4+ Tg{alpha}+ T cells had already undergone three cell divisions in the lymph nodes draining the lungs and the nasal and oral cavities, including the mediastinal, parathymic, superficial, and deep cervical lymph nodes (Fig. 4GoA). A smaller degree of division was also seen in the mesenteric lymph nodes, probably due to swallowing of peptide. No division was seen in peripheral nondraining nodes (pooled brachial, subscapular, inguinal, and paraaortic). By day 4, some cells in the draining nodes had undergone seven divisions, and the majority had undergone at least four (Fig. 4GoB). These responses were again Ag specific, as the pattern of cell division observed in CD4+ Tg{alpha}- T cells was comparable with that seen in animals treated with PBS.

Fig. 5Go quantitatively summarizes the data from the mediastinal nodes, allowing comparison between the responses to peptide-pulsed DCs (Fig. 3Go) and free peptide presented by host DCs. In contrast to the increased number of CD4+ Tg{alpha}+ cells during the course of the response to peptide-pulsed DCs, intranasal peptide caused a generalized loss of CFSE+ cells within both CD4+ (both Tg{alpha}+ and Tg{alpha}-) and CD4- populations in all lymph nodes from mice in the peptide group, as indicated by a decrease in CFSE content (compare Figs. 5GoC and 3GoC). This loss suggested a nonspecific toxic effect of the peptide. As a result, the number of CD4+ Tg{alpha}+ cells in the mediastinal lymph nodes was the same in the peptide and PBS groups 4 days after immunization, despite the substantial amount of cell division in the peptide group (Fig. 5GoA). Comparison of the quantitative data in Fig. 5GoB with that in Fig. 3GoB indicated that the response to this dose of intranasal peptide stimulated earlier cell division and that a larger total proportion of peptide-specific cells had undergone division at each time point. Thus, 45% of Tg{alpha}+ cells had a CFSE division pattern consistent with at least one cell division by 2 days after immunization with MCC87–103, the figure rising to 82% by day 4. The proportion of mediastinal node CD4+ Tg{alpha}+ cells recruited into cell division by day 4 was 39%.



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  		FIGURE 5. Quantitative summary of data from the experiment described in the legend to Fig. 4Go. A, The number of donor-derived CD4+ Tg{alpha}+ (left panel) or CD4+ Tg{alpha}- (right panel) T cells per 105 live mediastinal lymph node cells at various days of the response after intranasal administration of 100 µg MCC peptide or PBS. Individual points represent the mean ± SEM from the group. B, The percentage of donor-derived CD4+ Tg{alpha}+ T cells in mediastinal lymph nodes that have divided at least once after intranasal immunization with MCC87–103 or PBS. C, CFSE content, calculated as described in Materials and Methods, within donor-derived CD4+ Tg{alpha}+ and CD4+ Tg{alpha}- populations in mediastinal lymph nodes. Individual points represent the mean ± SEM from the group.

 
Phenotype and recirculation pattern of T cells activated in draining lymph nodes

To analyze the pattern of T cell activation in draining and nondraining lymph nodes in response to injected DCs, five-color flow cytometry was used to relate cell division pattern (CFSE staining) in CD4+ Tg{alpha}+ T cells to expression of the lymphocyte activation markers CD69 (Fig. 6Go, A and B) and CD44 (Fig. 6GoC). CD69 can be up-regulated as early as 2 h after Ag encounter (our unpublished findings; 37). Ag-reactive cells demonstrated enhanced CD69 expression before undergoing cell division, as evident from the high level of expression by undivided cells on day 2 after injection of peptide-pulsed DCs (Fig. 6GoA, top left panel). At that time, CD69 was also expressed by all the divided CD4+ Tg{alpha}+ cells, the total representing 76% of the CFSE+ CD4+ Tg{alpha}+ cells in the mediastinal lymph nodes, compared with 1.4% of cells in animals immunized with unpulsed DCs. No significant up-regulation was seen in the inguinal lymph nodes (Fig. 6GoA, top right panel) or any of the other nondraining nodes (superficial and deep cervical, subscapular, brachial, parathymic, and mesenteric, not shown). By day 4 of the response, a significant number of divided cells no longer expressed CD69, total expression having decreased to 34.4% of donor-derived CD4+ Tg{alpha}+ cells (Fig. 6GoB). The pattern of CD69 expression as a function of cell division on days 2 (Fig. 6GoA), 4 (Fig. 6GoB), and 7 (not shown) indicated that expression was progressively down-regulated upon each cell division, dividing cells eventually losing expression altogether.



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  		FIGURE 6. Expression of the early activation marker CD69 by CD4+ Tg{alpha}+ T cells in relation to cell division number in draining and nondraining lymph nodes 2 days (A) and 4 days (B) after immunization with MCC-pulsed (upper row) or unpulsed (lower row) DCs. C, Expression of the memory marker CD44 by CD4+ Tg{alpha}+ T cells in draining and nondraining lymph nodes at day 4 of the same experiment. Cells were gated for PI- CD4+ Tg{alpha}+. Donor-derived Tg{alpha}+ T cells can be discriminated from host T cells expressing endogenously rearranged V{alpha}11 by CFSE fluorescence. Number of cell divisions is indicated.

 
In contrast, expression of the activation/memory marker CD44 (38) was increased with each cell division cycle, cells that had undergone the most divisions having the highest level of expression (Fig. 6GoC showing the expression of CD44 by CD4+ Tg{alpha}+ T cells at day 4 of the response). Again, this relationship was observed irrespective of the time after the initiation of the response (not shown).

By day 4, divided CFSE+ CD4+ Tg{alpha}+ T cells were present in nondraining nodes and spleen (Fig. 6Go, B and C, and not shown). The intensity of CFSE staining in these cells revealed that they had undergone a minimum of four cell divisions, suggesting that they had recirculated to nondraining nodes and spleen after dividing in the draining nodes. Interestingly, cells that had recirculated to distant sites such as the peripheral lymph nodes were uniformly negative for expression of CD69, while retaining high expression of CD44. Recirculating T cells generated by immunization of I-E-positive 107-1 mice with free MCC87–103 peptide (either intranasal, s.c., or i.v.) have the same phenotype (A. L. Smith and B. Fazekas de St. Groth, unpublished data).

In vitro restimulation of lymph node T cells

On day 4 after intratracheal injection of peptide-pulsed or unpulsed DCs, lymphocytes were purified from the draining mediastinal and pooled nondraining lymph nodes and restimulated in vitro with MCC87–103 peptide in the presence of irradiated spleen APC from I-E+ (107-1) donors (Fig. 7GoA). In mice primed with peptide-pulsed DCs, enhanced [3H]thymidine uptake was observed in lymphocytes from draining mediastinal lymph nodes compared with nondraining nodes. Enhanced proliferation was also evident in the absence of restimulation with exogenous MCC87–103 (not shown), suggesting that the lymphocyte preparation contained Ag derived from the peptide-pulsed DCs or that proliferation of peptide-specific T cells continued in vitro in the absence of additional peptide. The response of lymphocytes from the nondraining nodes of mice immunized with peptide-pulsed DCs was comparable with that of both draining and nondraining nodes of mice immunized with unpulsed DCs. The magnitude of this response is consistent with that of the primary in vitro response of naive, high affinity peptide-specific T cells (22).



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  		FIGURE 7. A, In vitro proliferation of mediastinal (filled symbols) and nondraining (open symbols) lymph node lymphocytes, taken 4 days after immunization with MCC-pulsed (squares) or unpulsed DCs (circles). Cultures of 2 x 105 T cells were restimulated with 5 nM MCC peptide in the presence of syngeneic I-E+ APCs (lacking in 36-2 hosts; see Materials and Methods). Proliferation was measured 72 h later by [3H]thymidine incorporation. B, Cytokine production during in vitro restimulation (at 10 mM peptide) of the cells described in A, taken 4 days after immunization with MCC-pulsed (filled bars) or unpulsed DCs (open bars). Supernatants were assayed after 48 h for IL-4 and 72 h for IL-3 and IFN-{gamma} using bioassay (IL-3, IL-4) and ELISA (IFN-{gamma}). In groups of peripheral lymph nodes, IFN-{gamma} and IL-4 levels were not detected (N.D.) above the limits of the assay.

 
As a further measure of lymphocyte priming by DCs, in vitro secretion of cytokines by lymphocytes from draining and nondraining nodes was determined (Fig. 7GoB). Production of IL-3 and IFN-{gamma} was specifically increased in the draining lymph nodes of animals immunized with peptide-pulsed DCs compared with unpulsed DCs. In nondraining nodes, there was an increase in the level of IL-3 (but not IFN-{gamma}) production in animals in the peptide-DC group compared with the control DC group. Overall, the level of IL-4 production was very low and no immunization- dependent increase was seen in this experiment in either draining or nondraining nodes.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The activation of naive T cells in vivo is difficult to detect because of the low frequency of high affinity Ag-specific T cells. We have previously generated TCR Tg mice in which a high proportion of CD4+ T cells respond to the model Ag MCC87–103 in the context of the MHC class II molecule I-E (21). Although unmanipulated TCR Tg animals make essentially normal high affinity T cell responses to immunogenic and tolerogenic Ags (Ref. 17 ; M. E. Wikstrom, A. L. Smith, and B. Fazekas de St. Groth, in preparation), adoptive transfer of a CFSE-labeled cohort of TCR Tg T cells into a normal recipient offers a number of experimental advantages. Thus, the cell division profile can be correlated with expression of activation markers and cytokines, and with in vivo distribution and recirculation of Ag-specific T cells to lymphoid and nonlymphoid sites (22, 35, 39). In addition, adoptive transfer also allows a distinction to be made between the effect of MHC-peptide recognition at two distinct stages of T cell development, namely during positive selection in the thymus and activation of mature T cells in the periphery. Thus, T cells can mature in a thymus expressing a high level of the MHC allele required for efficient positive selection, after which they may be transferred into a host in which only a subset of peripheral APCs expresses the required MHC (34).

The adoptive experimental model used in this study (Fig. 1Go) mimics the initiation of the adaptive immune response in the lung by DCs, the most relevant APCs of the pulmonary immune response (2, 5). Myeloid DCs were grown from bone marrow progenitors, pulsed with the model Ag MCC87–103 in vitro, and injected into the trachea of unirradiated, semiallogeneic mice, avoiding the problems of interpretation related to earlier semiallogeneic DC adoptive transfer models in which the host animals were immunodeficient and therefore had abnormal lymphoid microarchitecture (40). Our approach required a normal host tolerant of semiallogeneic donor DCs, such as the Tg 36-2 line, expressing the I-E molecule only in the thymus, but not on peripheral APCs (24, 36, 41). These animals are effectively tolerant of the I-E expressed by the cohort of MCC87–103-pulsed DCs.

Two days after intratracheal immunization with pulsed DCs, 76% of peptide-specific T cells in the mediastinal nodes had reacted to Ag presented by DCs, as evidenced by up-regulation of CD69 (Fig. 6GoA), although fewer than 4.2% had been recruited into cell division, their progeny comprising 9% of the total donor- derived CD4+ Tg{alpha}+ cells (Fig. 3GoB). Neither up-regulation of CD69 nor division was seen in CD4+ Tg{alpha}- cells (which are not specific for the MCC87–103-I-E complex) in immunized mice, nor in either CD4+ Tg{alpha}+ or CD4+ Tg{alpha}- cells in recipients of unpulsed DCs. This is an illustration of the extraordinary capacity of DCs to specifically select and activate Ag-reactive T cells from a diverse repertoire of T cells, without inducing bystander activation (1, 42). The rapidity of the T cell response in the mediastinal nodes is in agreement with published studies of the kinetics of DC migration into draining lymph nodes (13, 15). Others have shown that spleen DCs injected into the rat trachea reach the T cell area of mediastinal and parathymic lymph nodes within 24 h of transfer (33). In further studies, we have confirmed rapid migration of DCs injected into the trachea using green fluorescent protein-transfected mouse bone marrow-derived DCs (B. N. Lambrecht and R. A. Pauwels, manuscript in preparation). Migrated DCs could be detected in the mediastinal nodes 24 h after injection, but were no longer detectable in any lymphoid structure 48 h later. In the current study, we did not observe T cell division or CD69 expression in nondraining lymph nodes or spleen at day 2 of the response, providing further evidence of the highly localized nature of T cell priming by DCs (15, 43, 44).

By day 4 of the response, some Ag-specific T cells in the mediastinal lymph nodes had undergone as many as six cell divisions. Expression of the memory marker CD44 increased with increasing cell division number, whereas expression of CD69 decreased slightly until the fourth, and thereafter showed an abrupt drop to baseline level (Fig. 6Go). The early induction and subsequent decrease in CD69 expression upon activation of Ag-specific T cells have been described by us and others and occur irrespective of the outcome of T cell activation (tolerance or immunity) (22, 35, 45, 46, 47).

When lymphocytes from the mediastinal nodes were restimulated in vitro with MCC87–103, enhanced proliferation and cytokine production were observed in the group that received peptide-pulsed DCs (Fig. 7Go). The enhanced production of IFN-{gamma} in mediastinal lymph nodes suggests induction of an Ag-specific Th1 response. Very little production of IL-4 could be detected, and no significant difference was seen between the experimental groups, consistent with failure to prime Th2 responses in vivo. This Th1 bias is a consistent finding for in vivo priming of naive T cells expressing the 5C.C7 TCR (Ref. 22 ; B. Fazekas de St. Groth, unpublished data), which is known to be of high affinity for the MCC87–103-I-E complex. Preferential induction of Th1 responses by high affinity TCR-peptide-MHC interactions has been noted previously (48). Our recent data suggest that DCs induce predominantly Th2 responses in the airways when a low affinity TCR-peptide-MHC interaction is involved, such as that between OVA323–339-I-Ad and the DO11.10 TCR (manuscript in preparation). Thus, in our model, the bias toward Th1 or Th2 is primarily a function of the TCR affinity, rather than a difference between the responses of non-Tg and TCR Tg animals.

Between days 2 and 4, divided Ag-specific CD4+ T cells recirculated from the mediastinal lymph nodes to the nondraining nodes and spleen. Primary division in the nondraining nodes appeared very unlikely, because the cells detected in blood (not shown) and nondraining sites (Fig. 6Go) had all divided at least four times. The phenotype of recirculating cells was remarkable in that they expressed uniformly high levels of the memory marker CD44 while lacking expression of the activation marker CD69. This phenotype is consistent with that of differentiated effector T cells (38, 49). Absence of CD69 expression by recently divided, recirculating T cells is seen irrespective of the route of immunization (i.e., s.c., i.v., intratracheal) or the presence of adjuvant at the injection site (unpublished observations and this study). It is striking that thymocytes also down-regulate expression of CD69 before emigrating from the thymus (50), suggesting that the correlation between down-regulation of CD69 and cell recirculation after Ag recognition may be of functional importance.

The association between the cell division profile and regulated expression of activation/memory markers (Fig. 6Go) is reminiscent of the correlation between differentiation of both T and B lymphocytes and the cell cycle (51, 52, 53, 54). Thus, the probability of isotype switching to IgG1 (51), IgE (52), IgG2a, IgG2b, IgG3, and IgA (61) is a function of the number of cell divisions, irrespective of the time after stimulation. The production of both Th1 and Th2 cytokines is also a function of cell division rather than time (53, 54). Further in vivo studies will be required to establish whether expression of activation markers, migratory behavior, and pattern of cytokine synthesis are all a strict function of cell division number rather than time after stimulation, and whether this relationship is preserved after activation by various routes, doses, and formulations of Ag administration. It will also be interesting to study whether the program of T cell recirculation following the primary immune response can be modified by interference with the cell cycle, or epigenetic control of gene regulation (54).

The data described in this study strongly support the notion that naive, Ag-specific T cells can be primed in vivo by a single intratracheal injection of DCs. This conclusion is supported by evidence from a number of functional tests, including the use of CFSE to visualize T cell proliferation in situ and measurement of proliferation and cytokine production of T cells in vitro. Although contaminating donor-derived B cells appeared to have stimulated a minor degree of proliferation after direct intratracheal injection of free peptide, the response was both later and smaller than that to pulsed DCs, indicating that they would be very unlikely to serve as efficient presenting cells for peptide eluted from pulsed DCs. The use of a semiallogeneic adoptive transfer also excluded the possibility of elution of peptide to endogenous APCs. It was recently demonstrated that transfer of class II-restricted Ag from injected DCs to endogenous DCs can occur after s.c. injection of apoptotic cells, leading to efficient presentation of Ag by lymph node DCs (20). Our experimental model excludes transfer of Ag from injected to endogenous DCs, because the latter do not express the MHC molecule required for presentation. However, the model does not exclude transfer of already formed peptide-MHC complexes between injected and endogenous DCs, a phenomenon that has recently been demonstrated in vitro (55). As mentioned above, our as yet unpublished experiments have demonstrated direct migration of myeloid DCs expressing green fluorescent protein from the trachea to the mediastinal node (B. N. Lambrecht, manuscript in preparation) and of fluorescein-labeled myeloid DCs from the footpad to the popliteal node (35). However, there is evidence that peptide-MHC complexes from lymphoid DCs can be recognized in the draining node in the absence of viable, donor-derived lymphoid DCs (35), suggesting an alternative route of presentation for Ags associated with nonmigratory or nonviable APCs. It should also be noted that none of the above mechanisms are mutually exclusive, and since all serve to enhance the traffic of T cell antigenic epitopes from the periphery to the site of the primary immune response, they may all be involved in optimizing Ag presentation in vivo.

Previous investigators have induced T cell priming in response to instillation of DCs purified from the spleen (19). Although the functional status of splenic and bone marrow-derived DCs may differ from that of DCs isolated directly from the lung (56, 57), it is not clear whether any differences between lung-derived and spleen- or bone marrow-derived DCs would be retained in the face of conditioning by the microenvironment after intratracheal administration. Moreover, even the behavior of reinstilled ex vivo lung DCs may not provide an exact mimic of that of undisturbed resident lung DCs. For example, previous studies have demonstrated that splenic myeloid DCs home to the splenic T cell zones, rather than the marginal zone in which they are usually located (58), after purification and i.v. administration (59). Their behavior appears to be altered by the purification process itself, which, like administration of adjuvants such as LPS, stimulates maturation and migration (16).

A second difficulty in interpreting the response to injection of a bolus of DCs into the deeper airways is that this mode of administration may not reflect the in vivo situation in which Ag may be intercepted and presented by endogenous DCs in both the upper and lower airways. To examine this aspect, the response to purified MCC87–103 peptide administered intranasally was compared with that to prepulsed intratracheal DCs. The primary proliferative response to intranasal peptide in I-E+ (line 107-1) hosts was more widespread, involving the nodes draining the nose, trachea, lung, oral cavity, and to a lesser extent the gut, presumably as a result of both inhaling and swallowing the peptide. Indeed, it was recently shown that oral administration of MCC87–103 peptide leads to local activation of T cells in the mesenteric lymph nodes of Tg mice expressing the 5C.C7 ß-chain paired with an endogenous repertoire of {alpha}-chains (46). After intranasal peptide, multiply divided Ag-specific CD4+ CD69- CD44high cells were first seen in nondraining sites on day 4 (not shown), suggesting a very similar pattern of activation, division, and recirculation to that induced by intratracheal peptide-pulsed DCs (Figs. 2Go and 6Go). The primary response to intranasal peptide was somewhat faster and recruited a higher proportion of Ag-specific T cells into cell division, presumably because peptide was presented by a much larger number of host-derived MHC class II+ DCs migrating from the airways to the draining nodes (6, 13). However, the total number of CD4+ Tg{alpha}+ cells in the mediastinal lymph nodes at the peak of response was substantially lower than in MCC-DC-immunized mice, as a result of two independent factors. First, a nonspecific toxic effect of soluble peptide (Fig. 5GoC) was manifest as an Ag-nonspecific decrease in donor-derived T and B cell numbers after administration of peptide by the intranasal route. We have noted a similar decrease after i.v. administration of free peptide (A. L. Smith and B. Fazekas de St. Groth, unpublished data). The mechanism of this effect has not yet been established. Second, Ag-specific recruitment of T cells to the mediastinal nodes after MCC-DC, but not intranasal peptide immunization, was apparent (Fig. 3GoC). The lack of recruitment into the mediastinal nodes after intranasal peptide was surprising, but may have resulted from the wide distribution of peptide in the other nodes draining the nasal and oral cavities, which effectively immobilized the majority of CD4+ Tg{alpha}+ cells at the sites in which they first encountered Ag.

It should be noted that despite the initial similarities in the pattern of T cell activation via intranasal peptide and intratracheal instillation of peptide-pulsed DCs, the functional outcome of the two protocols may be fundamentally different. Whereas mucosal administration of free peptide in the absence of inflammatory stimuli is an intrinsically tolerogenic event (our unpublished results; Refs. 46 and 60), the outcome of intratracheal DC immunization is immunity (19). Administration of free peptide in the absence of inflammation may fail to activate DCs, leading to an abortive response. In addition, direct access of free peptide to the circulation, particularly after oral administration, may allow access to lymphoid DCs that have been postulated to be the primary initiators of deletional tolerance (17). A third possibility, for which we and others have preliminary evidence, is that the immune response in the microenvironment of the superficial cervical nodes draining the nose is of a different character to that in the mediastinal nodes, generating a regulatory population of cells that maintains tolerance by an active, Ag-dependent mechanism (unpublished results; 60).

In conclusion, we demonstrate that myeloid DCs are potent inducers of the adaptive T cell response after migration into the mediastinal nodes. The T cell response is initially compartmentalized to the draining nodes, but subsequently spreads systemically. The experimental system we describe in this work will allow us to further characterize the early events of the pulmonary immune response induced by DCs, and to determine the extent to which other stimuli such as adjuvants and cytokines can influence the outcome of the response. This information should help us to better understand situations in which an aberrant pulmonary immune response leads to allergic sensitization and asthma.


   Acknowledgments
 
We thank David Lo for provision of I-E Tg animals, Kate Scott for screening of Tg mice, and Karen Knight and the staff of the Centenary Institute animal house facility for excellent animal husbandry.


   Footnotes
 
1 B.N.L. was a recipient of a scholarship of the Fund for Scientific Research Vlaanderen (F.W.O.), and was supported by a travel grant from the Horlait Dapsens Foundation, Belgium. B.F.d.S.G. was supported by a Wellcome Trust Senior Research Fellowship. This work was supported by the National Health and Medical Research Council of Australia, the Wellcome Trust, and the Medical Foundation of the University of Sydney. Back

2 Current address: Department of Pulmonary Medicine, Erasmus University Rotterdam, Dr. Molewaterplein 50, 3015 GE Rotterdam, The Netherlands. E-mail address: Back

3 Address correspondence and reprint requests to Dr. Barbara Fazekas de St. Groth, Centenary Institute of Cancer Medicine and Cell Biology, Locked Bag No. 6, NSW 2042 Newtown, Australia. E-mail address: Back

4 Abbreviations used in this paper: DC, dendritic cell; CFSE, carboxyfluorescein diacetate succinimidyl ester; MCC, moth cytochrome c; PI, propidium iodide; TCM, tissue culture medium; Tg, transgenic. Back

Received for publication August 9, 1999. Accepted for publication January 4, 2000.


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 Introduction
 Materials and Methods
 Results
 Discussion
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