| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
* Institute of Experimental Immunology, Zurich, Switzerland;
Research Department, Kantonsspital St. Gallen, St. Gallen, Switzerland;
Medizinische Hochschule Hannover Institut für Immunologie, Hannover, Germany;
Department of Nephrology and Hypertension, University of Schleswig-Holstein, Campus Kiel, Kiel, Germany; and
¶ Max Delbrück Center for Molecular Medicine, Berlin, Germany
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
CCR7 and its ligands, CCL19 and CCL21, are crucial for the positioning of T cells and dendritic cells (DC)3 within T cell zones of secondary lymphoid organs (8, 9, 10). Naive T cells express high levels of CCR7, and its down-regulation is closely correlated with T cell activation (3, 11). Furthermore, it has been suggested that memory T cell populations can be functionally distinguished by the presence or the absence of CCR7. CCR7lowCD62Llow effector memory cells preferentially migrate to peripheral nonlymphoid organs (12, 13) and exhibit immediate effector function. It has been suggested that these cells may eventually differentiate to CCR7highCD62Lhigh central memory cells (14), which are retained within secondary lymphoid organs as Ag-experienced nonpolarized cells lacking immediate effector function.
Numerous studies support the idea that CCR7 and its chemokine ligands, CCL19 and CCL21, also have a fundamental impact on priming and maintenance of immune reactions by influencing T cell and DC migration (8, 9). We have recently shown that plt/plt mice lacking the CCR7 ligands CCL19 and CCL21-Ser mount rapid antiviral T and B cell responses and exhibit normal formation of memory CTL (15). In the present study we used CCR7-deficient mice to further dissect the role of this receptor in the distribution, migration, and function of antiviral effector and memory CTLs in lymphoid/central and nonlymphoid/peripheral organs.
Protective immune responses against the noncytopathic lymphocytic choriomeningitis virus (LCMV) largely depend on the induction of antiviral CTL, which destroy infected cells in a contact-dependent and perforin-mediated manner (16). We examined the role of CCR7 during the effector and memory phases of antiviral immune responses using LCMV as a model infection. Our results indicate that the lack of organized T cell zones in CCR7–/– mice limits the maximal expansion of antiviral CTL. Furthermore, the CCR7 deficiency crucially affects memory T cell distribution between lymphoid and nonlymphoid organs. The absence of CCR7, however, did not preclude viral clearance and generation of protective recall responses, suggesting a role for CCR7 in the efficient expansion of antiviral CTL and the homeostatic recirculation of memory T cells.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
All mice used in this study were kept at the Institut für Labortierkunde, University of Zurich (Zurich, Switzerland). CCR7–/– mice (8) were backcrossed onto the 129/Ola background for at least five generations. For adoptive transfer experiments, CCR7–/– mice were backcrossed to C57BL/6 for nine generations, then crossed to P14 (17) or H8 transgenic mice (18). Genotyping for CCR7 deficiency was performed as described previously (8). Experiments were performed with sex-matched CCR7–/– mice and control heterozygous littermates at the age of 8–12 wk. Heterozygous control mice express slightly less CCR7 on lymphocytes compared with wild-type mice; however, this does not to impinge on the ability of CCR7+/– mice to mount efficient anti-LCMV CTL responses (data not shown).
Antibodies
Anti-CD8-FITC, anti-CD8-PE, anti-IFN-
-FITC, anti-CD62L-FITC, anti-CD8-PerCP, anti-CCR5-PE, anti-CD43-PE, anti CD44-PE, anti-Thy1.2.-allophycocyanin, anti-V
2-TCR-PE, and streptavidin-allophycocyanin were obtained from BD Pharmingen (Basel, Switzerland). To assess CCR7 expression, lymphocytes were incubated for 1 h at 4°C with 80 µl of COS cell supernatant containing 1 µg/ml CCL19-Ig, generated as previously described (19) with minor modifications (20). Cells were washed and incubated for 30 min at 4°C with biotin-SP-conjugated goat anti-human IgG (Fc-specific; Jackson ImmunoResearch Laboratories, West Grove, PA), followed by streptavidin-allophycocyanin and other directly labeled Abs. When combined with tetramer stainings, CCL19-Ig labeling followed the initial incubation with the tetramer complexes. For blood samples, erythrocytes were lysed with FACS Lysing Solution (BD Pharmingen) before analysis. Cells were analyzed with a FACSCalibur flow cytometer using the CellQuest software (BD Biosciences, Mountain View, CA).
Viruses and peptides
LCMV, WE strain, originally obtained from Dr. F. Lehmann-Grube (Heinrich-Pette-Institut, Hamburg, Germany), was propagated on L929 cells at a low multiplicity of infection and titrated as previously described (21). Recombinant vaccinia virus (VV) expressing gp33 as a minigene (VV-gp33C) was provided by Dr. M. van den Broek (University of Zurich) and titrated as previously described (22). The LCMV-GP peptides KAVYNFATM (gp33) and FQPGNGQFI (np396) were purchased from Neosystem (Strasbourg, France).
Cytotoxic T cell response
Specific ex vivo cytotoxicity was determined in a standard 51Cr release assay as previously described (22). The supernatants of the cytotoxicity assay cultures were counted in a Cobra II gamma counter (Canberra Packard, Mississauga, Canada). The percentage of specific lysis was calculated as (experimental release – spontaneous release)/(total release – spontaneous release) x 100. Spontaneous release was always <20%.
Footpad swelling reaction
A delayed-type hypersensitivity (DTH) response was induced by injecting 50 PFU of LCMV-WE into the hind footpads. Footpad thickness was measured by a spring-loaded caliper. Increased footpad thickness was expressed as the percent swelling relative to the thickness before injection.
Isolation of liver, lung, and splenic white pulp lymphocytes
Perfused livers were smashed through a metal grid. Lymphocytes were purified by Ficoll (Biochrom, Berlin, Germany) gradient centrifugation (600 x g, 15 min). Lungs were minced with razor blades and incubated in balanced salt solution containing 1 mg/ml DNase (Fluka, Buchs, Switzerland) and 2 mg/ml collagenase I (Sigma-Aldrich, St. Louis, MO) at 37°C for 30 min. Cell aggregates were dispersed by passing the digest through an 18-gauge syringe, and lymphocytes were isolated by Ficoll gradient centrifugation. White pulps of the spleen were isolated by digestion with collagenase V and III as previously described (23).
Construction of tetrameric class I-peptide complexes and flow cytometry
MHC class I (H-2Db) monomers complexed with gp33 were produced as previously described (24) and tetramerized by addition of streptavidin-PE (Molecular Probes, Eugene, OR). At the indicated time points after immunization, animals were bled, and single cell suspensions were prepared from spleens and lymph nodes. Aliquots of 5 x 105 cells or three drops of blood were stained using 50 µl of a solution containing tetrameric class I-peptide complexes at 37°C for 10 min, followed by staining with anti-CD8-FITC (BD Pharmingen) at 4°C for 20 min. The cells were analyzed by flow cytometric gating on viable leukocytes. Absolute cell counts were determined by counting leukocytes in an improved Neubauer chamber and, for blood samples, using an automated Advia counter (Bayer, Germany) in the central hematology laboratory of University Hospital Zurich.
Intracellular cytokine staining
Spleens were removed at the indicated time points after infection with LCMV. Single cell suspensions of 1 x 106 splenocytes, lymph node cells, or liver or lung lymphocytes were incubated for 5 h at 37°C in 96-well, round-bottom plates in 200 µl of culture medium containing 25 U/ml IL-2 and 5 µg/ml brefeldin A (Sigma-Aldrich). Cells were stimulated with PMA (50 ng/ml) and ionomycin (500 ng/ml) as the positive control or were left untreated as the negative control. For analysis of peptide-specific responses, 106 cells were stimulated with 10–6 M gp33 peptide and then surface stained as described previously (15). The percentage of CD8+ T cells producing IFN- was determined using a FACSCalibur flow cytometer.
Adoptive transfer and DC homing
Bone marrow-derived DC were generated from H8 mice as previously described (25). Assessment of DC-induced CTL priming was performed as follows. MACS-sorted P14 or P14 x CCR7–/– CD8+ T cells (3 x 105) were adoptively transferred together with 2 x 105 H8-DC or H8-CCR7–/–-DC into B6PLThy1.1 recipients. Five days later, lymphocytes from spleens, mesenteric lymph node (MLN), and livers were enumerated and stained for Thy1.2, CD8, CD62L, and CD44. The impact of CCR7 deficiency on intrasplenic positioning of DC and CTL was assessed by staining DC with the live dye 5-(and-6)-(((4-chloromethyl)benzoyl)amino)tetramethyl-rhodamine (Molecular Probes, Eugene, OR) and staining TCR transgenic CD8 T cells with CFSE (Molecular Probes) before adoptive transfer. Because only 
10% of the i.v. administered DC reach the spleen (26), both DC and T cells were injected directly into the spleens of B6 recipients to reach a sufficient cell density for histomorphological evaluation. Five hours postinjection of 5 x 106 cells of both cell types, spleens were sectioned, acetone-fixed, counterstained with 4',6-diamino-2-phenylindoledihydrochloride (Chemicon International, Temecula, CA), and analyzed by fluorescence microscopy using a BX61 fluorescence microscope (Olympus, Volketswil, Switzerland). To assess the proliferation of CD8+ T cells, MACS-sorted P14 CD8 T cells or P14-CCR7–/– CD8 T cells were labeled with CFSE, and 5 x 106 cells were adoptively transferred into C57BL/6 or CCR7–/– mice, respectively, before infection with 200 PFU of LCMV. Two, 4, or 7 days later, lymphocytes from spleens, MLN, and blood were stained for CD8 and V2 TCR and analyzed for CFSE dilution by flow cytometry. The homing efficacy of CCR7-deficient vs wild-type DC after i.v. injection was assessed by labeling with 50 µCi of 51Cr (Amersham Biosciences, Arlington Heights, IL) for 45 min at 37°C. Cells were washed three times with balanced salt solution, and the labeling efficiency per 5 million DC was determined using a Cobra II gamma counter (PerkinElmer Canberra Packard, Foster City, CA). Five million 51Cr-labeled DC were injected i.v. per recipient mouse. At different time points, organs of perfused mice were collected and assayed for the content of radioactivity.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Differential expression of CCR7 and CD62L has been reported to characterize effector and memory T cells in both humans (12) and mice (7). We thus assessed the surface phenotype of virus-specific CTL in lymphoid and nonlymphoid compartments after infection of heterozygous CCR7+/– mice with LCMV (Fig. 1). Due to the very low numbers of LCMV-specific CTL in naive animals, CCR7 and CD62L expression was assessed on CD8+ T cells (Fig. 1, upper row). As expected, CD8+ T cells were uniformly found to be CCR7high, even in peripheral organs such as lung and liver. Furthermore, CD62L was highly expressed on CD8+ T cells in all organs tested, except in lungs, where 67% of the cells had down-regulated CD62L.
|
Reduced clonal burst of effector CTL in CCR7–/– mice
CCR7–/– mice show aberrantly formed lymphoid T cell zones, strongly reduced T cell numbers in lymph nodes, and homing defects of DC and naive T cells (8). To assess the functional significance of this phenotype for the generation of acute antiviral CTL responses, CCR7–/– and CCR7+/– mice were infected with 200 PFU of LCMV-WE, and the number and surface phenotype of LCMV-specific, tet-gp33-positive cells in the CD8+ T cell pool were determined in the indicated organs by flow cytometry (Fig. 2). On day 8 after infection, CCR7–/– mice displayed reduced relative (Fig. 2A) and absolute numbers of CD8+tet-gp33+ CTL in all organs tested (Fig. 2B). However, the distribution of effector CTL was similar in both CCR7–/– mice and CCR7+/– mice, indicating that the absence of CCR7 did not lead to an aberrant localization of effector CTL during the acute phase of infection. A time-course experiment revealed that CTL expansion in CCR7–/– mice peaked on day 12 in spleen, blood, and lymph nodes, but, nevertheless, was weaker than that in CCR7+/– mice (Fig. 2C). This was also mirrored by a lower footpad swelling reaction of CCR7–/– mice, a DTH reaction that integrates all parameters of a peripheral virus infection (Fig. 2D). The surface phenotypes (CD62LlowCD44highCCR5lowCD43high) of gp33-specific effector CTL from CCR7–/– and CCR7+/– mice were identical (Fig. 2E). These data suggest that the lack of CCR7 plays a role in determining the kinetics and the overall magnitude of antiviral effector CTL responses, but does not affect effector CTL localization or differentiation.
|
To further substantiate the finding that the differentiation of effector CTL is independent of CCR7, the cytolytic activity of LCMV-specific CTL at the single cell level was assessed using a standard chromium release assay. The direct cytolytic activity of CCR7–/– mice was comparable to that in CCR7+/– mice after normalization of the values for the numbers of tetramer-positive cells present in spleen and lymph node cell preparations (Fig. 3A). Moreover, IFN- production by individual gp33-specific CD8+ T cells in response to cognate peptide was not impaired by the lack of CCR7 when assessed on day 14 postinfection (Fig. 3B). CCR7+/– heterozygous and CCR7+/+ wild-type mice generated equivalent anti-LCMV CTL responses (data not shown), thus excluding an impact of the slight differences in CCR7 expression between wild-type and heterozygous mice. Unlike in CCR7+/– mice, LCMV was not eliminated from lymphoid and nonlymphoid organs of CCR7–/– mice on day 8 postinfection (Fig. 3C), probably due to the reduced numbers of effector CTL in these mice. As shown above, gp33-specific CTL continued to expand in CCR7–/– mice between days 8 and 12, eventually leading to virus clearance.
|
Next, we assessed the impact of CCR7-mediated positioning of DC and CTL within secondary lymphoid organs on the efficacy of CTL priming. To this end, we used an adoptive transfer approach. CCR7–/– mice were crossed with P14 transgenic mice encoding a TCR specific for LCMV-gp33 (17) or H8 transgenic mice (18), which ubiquitously express LCMV-gp33. All mice were on the C57BL/6 background to facilitate adoptive transfer of lymphocytes into C57BL/6 and the congenic B6PLThy1.1 strain. Tracing experiments with 51Cr-labeled DC revealed that the homing patterns of CCR7-deficient and wild-type DC after i.v. injection were comparable (Fig. 4A), so that differential expansion of effector CTL could not be attributed to differential DC numbers within secondary lymphoid organs. Priming of adoptively transferred gp33-specific P14 or P14 x CCR7–/– CD8+ T cells by H8 or H8 x CCR7–/– DC CTL was examined by flow cytometry using the Thy1.2 marker. Expansion of P14-CCR7–/– CTL by H8-CCR7–/– DC was reduced by 50% compared with transfer of both wild-type populations (Fig. 4B). Activation of CTL in both situations was comparable, as determined by the expression of the activation markers CD44 and CD62L (Fig. 4C). Surprisingly, the priming efficacy was significantly reduced when CCR7 expression on the transferred cell populations was disparate, i.e., CCR7-negative H8 DC failed to prime CCR7-positive P14 CD8 T cells and vice versa (Fig. 4, B and C). These results strongly suggest that productive cognate interaction between Ag-bearing DC and responding CTL requires homing of both cell populations to the same compartment, be it the T cell zone or the red pulp/marginal zone. Indeed, visualization of the intrasplenic positioning of CCR7-competent DC and CD8+ T cells revealed that DC and T cells homed mainly to the T cell zone (Fig. 4D), whereas CCR7-deficient DC and T cells migrated exclusively to the red pulp or the marginal zone (Fig. 4G). Importantly, DC and CD8+ T cells from CCR7-disparate donors were usually not able to establish productive contacts, because CCR7-positive cells homed to the T cell zone, whereas CCR7-negative cells were sequestered in the red pulp and the marginal zone (Fig. 4, E and F). Taken together, these data show that DC-CTL contact in the T cell zone is not an absolute requirement for efficient CTL priming. Nevertheless, a microenvironment that facilitates concentration of both cell populations and increases the likelihood of their cognate interaction appears to be important for maximal amplification of the immune response.
|
We next assessed, using adoptive transfer of CFSE-labeled P14 x CCR7–/– CD8+ T cells into CCR7–/– mice and of P14 CD8+ T cells into C57BL/6 mice, whether the reduced magnitude in the LCMV-induced CTL response in CCR7–/– mice (Fig. 2) is due to an impaired Ag-specific proliferation. On day 2 after infection with 200 PFU of LCMV-WE, neither CCR7-positive nor -negative P14 CD8 T cells had entered the proliferation phase (Fig. 5A). However, P14 T cells were present at higher frequencies in spleens and lymph nodes of C57BL/6 recipients, most likely reflecting the more efficient recruitment of CCR7-competent T cells to secondary lymphoid organs. On day 4 after infection, P14 CD8+ T cells had already undergone at least five rounds of cell divisions, in contrast to P14 x CCR7–/– cells that had not yet started the proliferation program (Fig. 5B). Nevertheless, on day 7 postinfection, both P14 and P14 x CCR7–/– cells had efficiently proliferated (Fig. 5C). These data support the idea that the altered kinetics of CTL expansion in CCR7-deficient mice after LCMV infection are due to a weaker recruitment of CD8+ T cells to secondary lymphoid organs and a delayed onset of Ag-specific proliferation.
|
We next set out to examine the role of CCR7 in the distribution of memory CTL after viral infection. For this purpose, we determined the absolute numbers of gp33-specific CTL in different lymphoid and nonlymphoid organs during the memory phase of the LCMV infection. Absolute numbers of gp33-specific memory CTL on day 80 postinfection were lower in all organs of CCR7–/– mice compared with CCR7+/– mice (Fig. 6, first data point). Comparison of these values with the expansion of CTL on day 40 postinfection (Fig. 2C) indicated that the decay of memory CTL in CCR7–/– mice did not differ from that observed in CCR7+/– mice. However, it is noteworthy that memory CTL were efficiently excluded from lymph nodes, indicating that the absence of CCR7 impacts mainly memory CTL distribution.
|
CCR7 expression does not correlate with the activity of antiviral memory CTL from lymphoid vs nonlymphoid organs
Memory CTL from peripheral nonlymphoid organs exert rapid effector function (28) and are thought to provide a mechanism for rapid pathogen containment. It has been suggested that the presence of such effector memory T cells in peripheral organs is associated with loss of CCR7 expression and gain of effector function (12). We therefore assessed the immediate IFN- production of CCR7-deficient or -competent memory CTL from different lymphoid and nonlymphoid organs on day 80 after infection with LCMV-WE after a short term in vitro stimulation with gp33. Both CCR7–/– and CCR7+/– gp33-specific memory CTL from all organs tested readily produced IFN- (Fig. 7A). These data indicate that memory CTL from both lymphoid and nonlymphoid organs can mount immediate effector function, and that this function is not dependent on CCR7 expression. Although the absolute numbers of gp33-specific memory CD8+ T cells producing IFN- after short term peptide stimulation was decreased in CCR7–/– mice compared with CCR7+/– mice (Fig. 7A), these numbers did not significantly differ from the values obtained for the respective groups by tetramer analysis (compare Fig. 6). When memory CTL from lymphoid vs peripheral organs were tested in a 5-h 51Cr release assay, both memory CTL from spleens and lungs of CCR7–/– mice and CCR7+/– controls showed efficient target lysis (Fig. 7B). Furthermore, the expression of the cell surface markers CD44, CD62L, CCR5, and the activation-associated isoform of CD43 (1B11) was similar on gp33-specific memory CTL from lymphoid and nonlymphoid organs of CCR7–/– mice and CCR7+/– (Fig. 7C). Taken together, these data further support our conclusion that CCR7 is important for the distribution of memory CTL in lymphoid and nonlymphoid organs, but that CCR7 expression does not correlate to their functional differentiation.
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
In vitro studies using polarized T cells or T cell lines and in vivo studies (3, 11) have shown a correlation between effector function and low CCR7 expression. In accordance with these findings, we found that CCR7 is highly expressed on naive CTL in lymphoid and nonlymphoid organs, and that virus-specific CTL in blood and peripheral organs of mice had down-regulated CCR7 during acute virus infection. However, it is interesting to note that a significant proportion of LCMV-specific CTL in spleen and lymph nodes had not lost CCR7 and CD62L expression during the effector phase. In contrast to previous studies describing antiviral effector CTL to be CCR7low in LCMV infections (11, 14), we have not exclusively used transgenic CD8+ T cells, but also followed the complete gp33-specific effector CTL population by tetramer analysis. This population is polyclonal (29) and likely to be composed of T cells with different TCR affinities and activation requirements. Therefore, the expression levels of CCR7 and CD62L may vary at a given time point of effector differentiation. Moreover, our data corroborate a recent study by Unsoeld et al. (30), suggesting that during the acute phase of an antiviral immune response both CCR7-positive and -negative CTL from secondary lymphoid organs can exert full effector function. It appears therefore that the CCR7low phenotype of CTL is associated with acute effector function in peripheral nonlymphoid organs, but not in secondary lymphoid organs.
It has been shown that CCR7 deficiency results in impaired T cell responses, such as the complete abolishment of DTH reactions against FITC or keyhole limpet hemocyanin after a short (4-day) priming period (8). Similarly, we have demonstrated reduced LCMV-mediated footpad swelling in CCR7–/– mice. DTH reactions depend strongly on DC-mediated Ag transport from the site of inoculation to secondary lymphoid organs (31). We consider it likely that severely impaired DTH reactions in CCR7–/– mice are the result of a nearly complete blockade of DC migration from skin to local lymph nodes, and that only limited amounts of DC are available for T cell priming (8, 32). However, the induction of CTL responses during LCMV infection is probably less dependent on DC-mediated Ag influx to secondary lymphoid organs, because this rapidly disseminating virus can spread independently of cell trafficking. Therefore, the reduced CTL expansion in CCR7–/– mice after LCMV infection is presumably accentuated by ectopic DC and CTL positioning in secondary lymphoid organs. The presented adoptive transfer experiments using CCR7-deficient DC and CTL support this interpretation because ectopic CTL priming in the red pulp and/or the marginal zone resulted in reduced priming efficacy.
CCR7-deficient mice and plt/plt mice, which lack the CCR7 ligand chemokines CCL19 and CCL21-Ser within secondary lymphoid organs (9), exhibit comparable morphology in lymph nodes and spleens. However, unlike CCR7–/– mice, plt/plt mice are able to mount nearly unimpaired DTH reactions (33) and antiviral CTL responses (15). The plt/plt mice express the leucine isoform of CCL21 (CCL21b) outside lymphoid tissues in lymphatic vessels. In addition, CCL21b is expressed constitutively in nonlymphoid organs (10, 34) and may therefore improve the recruitment of antiviral CCR7-positive effector CTL to peripheral tissues. Furthermore, a recent study reported that peripheral CCL21 caused lymph node congestion and augmented the initiation of T cell responses during inflammation (32). The expression of CCL21b in lymph vessels and peripheral tissues in plt/plt mice thus most likely contributes to the preserved antiviral CTL responses after LCMVinfection, as observed in our previous study (15), whereas DC and T cells in CCR7–/– mice remain unresponsive to the peripheral effect of CCL21b.
It is noteworthy that the alternative priming sites for antiviral CTL in the absence of CCR7 or its ligand chemokines, e.g., the marginal zone or the red pulp in the spleen, still facilitate efficient DC-CTL interaction. CTL priming outside the T cell zone resulted in appropriate differentiation of CTL, which were able to produce effector cytokines, exhibited significant cytolytic activity, and mediated antiviral protection in both CCR7–/– mice as well as plt/plt mice (this study and Ref. 15). Therefore, it is reasonable to assume that CCR7-mediated organization of the lymphoid T cell zone increases the likelihood of productive DC-T cell contacts and is thus necessary for the maximal amplification of virus-induced CTL responses.
During the memory phase of an anti-LCMV immune response, a significant proportion of specific CTL retained in peripheral organs had down-regulated both CCR7 and CD62L, whereas those found in lymph nodes and splenic white pulp exhibited mainly the central memory-like CD62LhighCCR7high phenotype. Our data are therefore compatible with the concept that CD62LlowCCR7low effector memory CTL reside primarily in the periphery, and that CD62LhighCCR7high central memory CTL localize mainly to secondary lymphoid organs. However, the observed functional characteristics of those cell subsets differ from those proposed in the original model of central vs effector memory (12). Antiviral CTL from both peripheral nonlymphoid and secondary lymphoid organs rapidly produced IFN- after specific restimulation and displayed significant cytolytic activity. This confirms previous studies showing that immediate effector function of memory T cells is not strictly associated with down-regulation of CCR7 (14, 30, 35, 36). In particular, our data are consistent with a recent study demonstrating significant ex vivo lytic activity of memory CTL from both spleen and liver (14). The somewhat longer incubation time for the direct CTL assay after infection with LCMV-Armstrong was probably necessary in that study, because the clonal burst size after LCMV-Armstrong infection is weaker than that after LCMV-WE infection (37). Regarding the functional activity of lymphoid vs nonlymphoid memory CTL, recent data suggest that the memory CTL pool of most nonlymphoid organs is mobile, and thus in constant exchange with the lymphoid memory pool (38, 39), and that both pools can exocytose IFN- and lytic granules to the same extent in response to specific Ag (14, 39). Therefore, a strict functional distinction of peripheral vs lymphoid memory CTL is not supported in several systems and readouts.
One study demonstrating stronger direct CTL activity from peripheral organs than from spleen (28) used vesicular stomatitis virus, a virus that does not rely on CTL for antiviral protection because it is cleared from the host within hours via neutralizing Abs (40). The maintenance of memory CTL activity in such an infection may rely on mechanisms differing from those that mediate the maintenance of memory CTL against persistent viruses such as LCMV.
Taken together, we have shown in this study that CCR7 plays a key role in the homeostatic recirculation of antiviral memory CTL. However, the strict differentiation between resting central vs active effector memory T cells based on their CCR7 expression probably does not reflect the complex in vivo situation as seen, for example, during a viral infection. Furthermore, our data underscore the importance of organized lymphoid structures for the generation of primary immune responses and delineate the role of CCR7-mediated lymphocyte and DC migration in the induction and maintenance of antiviral immunity.
|
Acknowledgments |
|---|
|
Footnotes |
|---|
1 Address correspondence and reprint requests to Dr. Tobias Junt, Institute of Experimental Immunology, Department of Pathology, University of Zurich, Schmelzbergstrasse 12, CH-8091 Zurich, Switzerland. E-mail address: tobias.junt{at}usz.ch or 
2 Current address: Research Department, Kantonsspital St. Gallen, 9007 St. Gallen, Switzerland. E-mail address: burkhard.ludewig{at}kssg.ch
3 Abbreviations used in this paper: DC, dendritic cell; DTH, delayed-type hypersensitivity; LCMV, lymphocytic choriomeningitis virus; MLN, mesenteric lymph node; VV, vaccinia virus.
Received for publication March 17, 2004. Accepted for publication September 30, 2004.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
8.1 chain transgenic mice. EMBO J. 8:719.[Medline]
This article has been cited by other articles:
|
|
 |
|
  T.-B. Kang, G.-S. Oh, E. Scandella, B. Bolinger, B. Ludewig, A. Kovalenko, and D. Wallach Mutation of a Self-Processing Site in Caspase-8 Compromises Its Apoptotic but Not Its Nonapoptotic Functions in Bacterial Artificial Chromosome-Transgenic Mice J. Immunol., August 15, 2008; 181(4): 2522 - 2532. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 G. Bouma, S. Burns, and A. J. Thrasher Impaired T-cell priming in vivo resulting from dysfunction of WASp-deficient dendritic cells Blood, December 15, 2007; 110(13): 4278 - 4284. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
N. Czeloth, A. Schippers, N. Wagner, W. Muller, B. Kuster, G. Bernhardt, and R. Forster Sphingosine-1 Phosphate Signaling Regulates Positioning of Dendritic Cells within the Spleen J. Immunol., November 1, 2007; 179(9): 5855 - 5863. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. Unsoeld, K. Mueller, U. Schleicher, C. Bogdan, J. Zwirner, D. Voehringer, and H. Pircher Abrogation of CCL21 chemokine function by transgenic over-expression impairs T cell immunity to local infections Int. Immunol., November 1, 2007; 19(11): 1281 - 1289. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. A. Schneider, J. G. Meingassner, M. Lipp, H. D. Moore, and A. Rot CCR7 is required for the in vivo function of CD4+ CD25+ regulatory T cells J. Exp. Med., April 16, 2007; 204(4): 735 - 745. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. Scandella, K. Fink, T. Junt, B. M. Senn, E. Lattmann, R. Forster, H. Hengartner, and B. Ludewig Dendritic Cell-Independent B Cell Activation During Acute Virus Infection: A Role for Early CCR7-Driven B-T Helper Cell Collaboration J. Immunol., February 1, 2007; 178(3): 1468 - 1476. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
U. E. Hopken, A. M. Wengner, C. Loddenkemper, H. Stein, M. M. Heimesaat, A. Rehm, and M. Lipp CCR7 deficiency causes ectopic lymphoid neogenesis and disturbed mucosal tissue integrity Blood, February 1, 2007; 109(3): 886 - 895. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Alfonso-Perez, S. Lopez-Giral, N. E. Quintana, J. Loscertales, P. Martin-Jimenez, and C. Munoz Anti-CCR7 monoclonal antibodies as a novel tool for the treatment of chronic lymphocyte leukemia J. Leukoc. Biol., June 1, 2006; 79(6): 1157 - 1165. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Junt, K. Fink, R. Forster, B. Senn, M. Lipp, M. Muramatsu, R. M. Zinkernagel, B. Ludewig, and H. Hengartner CXCR5-Dependent Seeding of Follicular Niches by B and Th Cells Augments Antiviral B Cell Responses J. Immunol., December 1, 2005; 175(11): 7109 - 7116. [Abstract] [Full Text] [PDF]
|
|
| ||||||||
and 
) and CCR7+/– (
and 
) mice was tested on 51Cr-labeled EL4 cells pulsed with gp33 or np396. E:T cell ratios were corrected for the number of tetramer-positive CD8+ cells (see Fig. 2
