The impact of splenectomy on antiviral T cell memory in mice

Sonja Mrusek, Simone Vallbracht and Stephan Ehl

Center for Pediatrics and Adolescent Medicine, University of Freiburg, Mathildenstrasse 1, 79106 Freiburg, Germany

Correspondence to: S. Ehl; E-mail: ehl{at}kikli.ukl.uni-freiburg.de


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
The contribution of the spleen to protective antiviral T cell memory was studied using the mouse model of infection with respiratory syncytial virus (RSV). Virus-specific CD8+ memory T cells were induced by local (intranasal or intracutaneous) or systemic (intravenous) immunization using RSV or vaccinia virus-recombinants expressing an RSV protein. After all three routes of immunization, the spleen was clearly identified as the main anatomic compartment harbouring virus-specific memory T cells. Surprisingly, however, splenectomy performed 30 days after immunization did not impair the efficacy of the memory T cell response to a subsequent RSV challenge infection. Irrespective of the route of priming, splenectomy did not influence the number or the functional activity of virus-specific memory T cells recruited to the lung following RSV challenge. More importantly, splenectomy did not impair pulmonary virus control by antiviral memory T cells in vivo. These findings were confirmed under experimental conditions where no neutralizing antibodies were induced by the priming infection. Thus, although most memory CD8+ T cells localize to the spleen after viral infections, this important lymphoid organ is dispensable for efficient recall responses. These findings have implications for the immunocompetence of splenectomized patients.

Keywords: lung, memory, spleen, T cells, viral infection


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
The spleen is a highly organized secondary lymphoid organ that provides an optimal structural framework for the generation of efficient immune responses. In particular, immunological tasks of the spleen include capture and presentation of antigen and retainment of APC, T cells and B cells to allow the required close cellular interactions. Absence of the spleen causes an increased susceptibility to systemic infections with encapsulated bacteria, which has been ascribed to lack of the ‘filter’ function of the spleen. Moreover, recent data indicate that the spleen is also critical for the generation and maintenance of B1a B cells (1) [CD27+IgM+ memory B cells in humans (2)] which mediate TI-2 antibody responses to polysaccharide components of bacterial cell walls.

The role of the spleen in antiviral immunity has been less well studied. No particular susceptibility to viral infections has been reported in asplenic patients. In spleenless mice lacking the orphan homeobox gene 11 (Hox11–/–), normal effector T cell responses to systemic infection with lymphocytic choriomeningitis virus (LCMV), vaccinia virus (VV) or vesicular stomatitis virus were detected in pooled lymph nodes (3). Moreover, compared to wild-type mice, there was no difference in T cell-mediated elimination of these viruses. While these data indicated that the spleen is dispensable for the generation of antiviral effector T cell responses, they did not answer the question whether the spleen plays a role in the maintenance of efficient antiviral T cell memory. This is of clinical relevance, because most viral infections (and antiviral immunizations) are encountered early in life and impairment of antiviral T cell memory after splenectomy could potentially impair immunocompetence to viral recall infections.

Why should the spleen play a role in antiviral T cell memory? Even after localized pulmonary viral infections, there is significant T cell expansion in systemic lymphoid tissues (4) and the largest number of memory T cells can be found in the spleen (5). Although several recent studies have shown that a significant number of functional antigen-specific CD8+ T cells also localized to non-lymphoid tissue during the memory phase of viral or intracellular bacterial infections (6,7), these cells could not divide or migrate back to the lymph node (6)—a process essential for the generation of an effective memory T cell response (8,9).

In this study, we induced antiviral CD8+ memory T cells using two different viruses by various routes of viral immunization and found that the majority of these cells localized to the spleen in the memory phase of infection. Surprisingly, however, splenectomy had no impact on the ability to rapidly recruit large numbers of functional memory T cells to the lung and to efficiently control a local pulmonary challenge infection.


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Mice
Specific-pathogen-free BALB/c mice were obtained from Charles River (Sulzburg, Germany) and used at 6–12 weeks of age. µMT mice were originally produced by Kitamura et al. (10). A ten-generation backcross to BALB/c was provided by Manfred Kopf (Molekulare Biomedizin, ETH Zürich, Switzerland). Mice were kept in a venti-rack (BioZone, Kent, UK). All animal experiments were performed in accordance with the local animal care commission.

Viruses
The RSV A2 strain was obtained from Peter Openshaw (Imperial College, London, UK), grown on HEp-2 cells and kept in liquid nitrogen. Mice were infected intranasally under methoxyflurane anesthesia with 2 x 106 pfu of RSV in 80 µl of RPMI. Recombinant vaccinia virus expressing the M2 protein of RSV (rVV M2) was a kind gift from Gail Wertz (University of Alabama, Birmingham, AL) and grown on BSC-40 cells. rVV expressing the nucleoprotein of LCMV (rVV NP) was obtained from Rolf Zinkernagel (University of Zürich, Switzerland). Mice were infected intravenously with 1 x 105 or 2 x 106 pfu in a volume of 200 µl or immunized intracutaneously (i.c.) by inoculating 10 µl of virus suspension (2 x 106 pfu) on shaved skin at the base of the tail by fine needle scratches (scarification). RSV titers were determined in homogenized lung samples on HEp-2 cells. Briefly, lung homogenates were titrated and transferred to a 96-well flat-bottom plate confluent with HEp-2 cells. After 24 h incubation, cells were fixed with methanol and incubated with goat anti-RSV biotin followed by streptavidin–HRP (Biogenesis, Poole, UK). Infected cells were detected using 3-amino-9-ethylcarbazole substrate and enumerated by light microscopy.

Splenectomy
Mice were anesthesized using ketamine and xylazine. After shaving the left side of the body, a 0.5 cm vertical incision was made in the skin. The peritoneum was opened, the spleen was carefully mobilized using forceps, and vessels and ligaments were heat cauterized using a heated needle. The peritoneal cavity was then closed with a silk suture and the skin was stapled.

Lymphocyte isolation
Single cell suspensions were prepared from spleen and pooled lymph nodes draining either the respiratory tract (cervical and mediastinal nodes, designated RTLN) or the lower extremities and the gastrointestinal tract (inguinal and mesenteric nodes, designated peripheral lymph nodes PLN). Lung (L) and liver (LIV) lymphocytes were prepared by mincing whole organ tissue through a 100 µm cell strainer (Falcon 2360) using a syringe plunger followed by gradient centrifugation over Ficoll (Ficoll-PaqueTM Plus, Amersham Biosciences) before analysis. Bone marrow cells (BM) were isolated by flushing both femurs and tibiae with 10 ml of PBS. Bronchoalveolar lavage (BAL) was performed as described previously (8).

Flow cytometry
Surface staining was performed for 30 min at 4°C using the following antibodies: CD3 (clone 145-2C11), CD8 (clone 53-6.7), CD62L (clone MEL-14 ) (BD Pharmingen, San Diego, CA). Staining with MHC Kd p82-90 tetramers (kindly provided by the NIAID tetramer facility) was performed prior to staining with other surface antibodies for 30 min at room temperature. For intracellular cytokine staining, cells were cultured for 3 h in V-bottom 96-well plates at 1–2 x 105 cells/well in a volume of 0.2 ml of RPMI 10% FCS supplemented with 1 µl/ml monensin (Golgistop, BD Pharmingen) and with p82 at a concentration of 0.1 µg/ml. Cells were harvested, washed, surface stained and then subjected to intracellular cytokine staining using the cytofix/cytoperm kit according to the manufacturer's instructions (BD Pharmingen). Cells were either stained with antibody against interferon-{gamma} (clone XMG1.2) or with an isotype control antibody (clone R3-43) (both from BD Pharmingen) and analyzed on a FACScan cytometer using Cellquest software. Staining for CCR7 was performed by incubating the cells with COS cell supernatant containing ~1 mg/ml CCL19-Ig [kindly provided by Heike Unsoeld (11)] at 4°C for 60 min followed by biotinylated polyclonal anti-human Fc{gamma} antibodies (Dianova, Hamburg, Germany) and allophycocyanin–streptavidin (BD PharMingen, San Diego, CA).

Cytotoxicity assays
CTL assays were performed under ‘Mini-Killer’ conditions as described previously (12). Briefly, effector cells were plated in 2-fold dilutions starting with 5 x 104 cells per well in a volume of 50 µl in a 96-well V-bottom plate (Greiner Labortechnik, Solingen, Germany). P815 cells were used as target cells and 2 x 103 cells were added in a volume of 50 µl per well for an initial effector:target ratio of 25:1. Target cells were labelled with 51Cr and with 10–6 Molar RSV p82-90 for 2 h at 37°C. The plates were centrifuged and incubated for 5 h at 37°C. Spontaneous 51Cr release was <20% in all assays.


    Results
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Rationale of the experimental system
The characteristics of a T cell memory response are likely to depend on the route of priming immunization, which influences both the numbers of T cells induced as well as the distribution of T cells between different anatomic compartments. These factors contribute to the extent and the kinetics of a recall response and may thus critically influence its biological efficacy. To analyze the role of the spleen in antiviral T cell memory in a more general fashion, we therefore immunized mice using two different viruses either by systemic (i.v.) or by local (i.n., i.c.) routes. Priming infection with RSV i.n. mimics natural infection or immunization with a live vaccine, and i.v. or i.c. infection with recombinant vaccinia virus expressing the RSV M2 protein mimics systemic or local vaccination. Mice were splenectomized 4 weeks after priming, rested for 3 weeks and then challenged with RSV intranasally. Recruitment and function of memory T cells and the kinetics of T cell-mediated virus control was compared to non-splenectomized mice. The experimental model of RSV infection was chosen, because control of challenge infection by memory T cells in this model not only requires the presence of sufficient and functional T cells, but is also dependent on their migration to the pulmonary site of challenge infection.

The spleen is the major anatomic compartment harbouring antiviral CD8+ memory T cells after local and systemic virus infections
Groups of BALB/c mice were immunized either with 2 x 106 pfu RSV i.n. or with 2 x 106 pfu rVV M2 containing the immunodominant CTL epitope p82-90 (p82) of RSV i.v. or by cutaneous scarification (i.c.). Mice infected with rVV expressing the nucleoprotein of lymphocytic choriomeningitis virus (rVV NP) were used as a specificity control. Mice were sacrificed 50 days after infection, lymphocytes were isolated from various tissues and the percentage and the absolute number of CD8+ T cells binding the Kd/p82 tetramer were determined. Figure 1 gives an example of the sensitivity and specificity of the tetramer reagent used in these experiments. The low background staining allowed clear identification of down to 0.3% of RSV-specific among total CD8+ T cells. To achieve a clear population, at least 200 000 cells from the spleen and 50–100 000 cells from the other compartments were analyzed. For FACS analysis, lymphocytes from 2–3 mice were pooled and each data point in Figure 2 therefore represents the mean of 2–3 mice. The results show that independent of the route of priming infection, 50 days later the majority of virus-specific T cells localized to the spleen (Fig. 2). Similar results were obtained 28 days after infection (data not shown) and when RSV-specific T cells were identified by intracellular cytokine staining after p82 peptide stimulation (data not shown). As expected (8,13), the difference between spleen and lung was no more than 5-fold after pulmonary RSV infection (Fig. 2A). However, after intravenous or cutaneous immunization, 10–30-fold more memory CD8+ T cells localized to the spleen than to the other compartments analyzed (Fig. 2B and C).



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Fig. 1. Detection of RSV-specific memory CD8+ T cells by tetramer staining. Groups of BALB/c mice were infected with 2 x 106 pfu RSV i.n. or with 2 x 106 pfu rVV RSV M2 i.v. or by scarification. Mice infected with 2 x 106 pfu rVV LCMV NP i.v. served as a specificity control. Fifty days later, the percentage of CD8+ T cells binding the H-2Kd p82 tetramer was determined by flow cytometry. At least 50 000 lung lymphocytes and 200 000 spleen lymphocytes were gated for analysis. The representative dot plots are gated on CD3+ live lymphocytes.

 


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Fig. 2. The main reservoir of memory CD8+ T cells is the spleen. The experimental setup is described in Fig. 1. Fifty days after infection, lymphocytes were isolated from spleen (S), lung (L), pooled respiratory tract lymph nodes (RTLN), pooled peripheral lymph nodes (PLN), liver (L) and bone marrow (BM). The obtained cells were pooled from 2–3 mice. The absolute number of RSV-specific T cells was determined by multiplying the percentage of tetramer-binding CD8+ T cells among total live cells as determined by a live gate in combination with microscopic cell counting. Each data point represents the mean of 2–3 mice. The data were obtained in two independent experiments involving four and six mice per group.

 
Recruitment and reactivation of memory T cells to the lung is not impaired in splenectomized memory mice
Having confirmed the spleen as the main reservoir of virus-specific memory CD8+ T cells, we addressed the question whether splenectomy would impair the memory T cell response to a challenge infection. Mice immunized against RSV were splenectomized 4 weeks after infection, rested for 3 weeks and were then challenged with 2 x 106 pfu RSV i.n. Intranasal priming with RSV was also performed in µMT mice to avoid rapid neutralization of the challenge infection by antibodies. This control was not needed for vvM2 immunized mice, since this recombinant virus does not induce RSV-specific antibodies. Five days after challenge infection, a significant number of RSV-specific memory T cells was recruited to the lung of RSV-primed BALB/c and µMT mice (Fig. 3A and B). At this early time point, few RSV-specific T cells could be found in control mice undergoing primary RSV infection (Fig. 3, ‘primary’), indicating that the identified tetramer binding CD8+ T cells truly represented memory T cells. Splenectomy had no measurable impact on the number of RSV-specific T cells recruited to the lung in response to challenge infection, irrespective of whether neutralizing antibodies were present (Fig. 3A) or not (Fig. 3B).



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Fig. 3. Recruitment of memory CD8+ T cells to the lung is not impaired in splenectomized mice. Groups of BALB/c or BALB/µMT mice (B) were infected with 2 x 106 pfu RSV i.n. (A and B) or with 2 x 106 pfu rVV RSV M2 i.v. (C and E) or i.c. (D and F). Four weeks later, half of the mice were splenectomized (open symbols) and 7 weeks after priming infection all immune mice as well as naive control mice (primary; closed squares) were challenged with 2 x 106 pfu RSV i.n. Five days later, inflammatory cells were eluted by BAL. The absolute number of RSV-specific CD3+CD8+ T cells was determined by tetramer staining (A–D) or intracellular cytokine staining after short-term stimulation with p82 (E and F). Similar data were obtained in two independent experiments.

 
Splenic memory T cells may not be required for a pulmonary recall response in intranasally immunized mice since recruitment and expansion of memory T cells persisting in the lung and local lymphoid tissue may be sufficient (8,13). By contrast, after pulmonary challenge infection of mice primed via intravenous or intracutaneous routes, most memory T cells have to be recruited from systemic compartments. Most of these reside in the spleen (Fig. 2). Surprisingly, however, splenectomy of mice immunized i.v. or i.c. did not reduce their ability to recruit virus-specific memory T cells in response to pulmonary challenge infection (Fig. 3C and D). Similar results were obtained when functional assays were used to study memory T cells ex vivo. Irrespective of the route of priming infection, no difference could be detected in RSV-specific IFN-{gamma} production (Fig. 3E and F) or RSV-specific target cell lysis (Fig. 4) among T cells obtained from the BAL of unmanipulated versus splenectomized memory mice after RSV challenge.



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Fig. 4. Reactivation of memory CD8+ T cells is not impaired in splenectomized mice. The experimental setup is described in Fig. 3. RSV-specific lytic activity of BAL cells was tested in a 51Cr release assay on P815 cells labelled with p82. Data represent means and SD of four mice/group. Closed squares: primary infection; closed symbols: infection of immune mice; open symbols: infection of splenectomized mice. Maximal lysis of unlabelled targets was 15%, spontaneous release was <20% in all assays. The experiment was repeated twice with similar results.

 
Protective antiviral T cell memory is not impaired in splenectomized mice
The key function of antiviral memory T cells is their ability to provide protective immunity by controlling virus replication in vivo. To address the question whether the spleen is required to maintain protective CD8+ T cell immunity after various routes of priming infection, groups of unmanipulated or splenectomized memory mice were challenged with RSV and pulmonary virus titers were determined 4 days after infection. Virus was eliminated below the detection limit in µMT mice immunized with 2 x 106 pfu RSV and in BALB/c mice primed with 2 x 106 pfu vvM2 i.v., independent of the presence of an intact spleen (Fig. 5A and B). It is possible that priming infection under these conditions induced such an abundance of memory T cells, that the effect of splenectomy was not detectable. To study protective immunity under more limiting conditions, the priming dose was therefore reduced to 105 pfu i.v. and mice immunized intracutaneously with 2 x 106 pfu rVVM2 were analyzed. After these immunization protocols, control of RSV challenge infection was variable, but all mice were partly protected by at least 1–2 logs (Fig. 5C and D). Nevertheless, even under these limiting conditions, no difference in memory T cell-mediated virus control could be detected between splenectomized and control mice.



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Fig. 5. T cell-mediated protective antiviral immunity is not impaired in splenectomized mice. The experimental setup is described in Fig. 3. Infectious doses for priming infection were used as indicated. Three weeks after splenectomy, immune and naive control mice were challenged with 2 x 106 pfu RSV i.n. and lung virus titers were determined 4 days later. Pooled data from two independent experiments are shown involving four and five mice/group. Closed squares: primary infection; closed symbols: infection of immune mice; open symbols: infection of splenectomized mice.

 
Memory T cell differentiation status in lung, spleen and respiratory tract lymph nodes
To address the question whether the lack of importance of the spleen during secondary responses was linked to effector or central memory differentiation in different anatomic compartments, we investigated the expression of CCR7 and CD62L on RSV-specific T cells. Eight weeks after intranasal infection with RSV, the proportion of CCR7low and of CD62Llow (‘effector memory’) cells among RSV specific CD8 T cells was highest in the lung (Fig. 6). In spleen and lymph node, a relatively higher proportion had a ‘central memory’ phenotype. There was no significant difference between the latter two compartments (Fig. 6). Thus, mostly T cells with a ‘central memory’ phenotype were lost after splenectomy, but cells with this differentiation state remained present both in the lung and in extrasplenic lymphoid tissue.



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Fig. 6. Memory T cell differentiation state in different anatomic compartments. A group of six mice was infected with 106 pfu RSV i.n. Eight weeks after infection, lungs, respiratory tract lymph nodes and the spleen were harvested and the obtained cells were pooled from two mice. The dot plots show Kd p82 tetramer binding and CD62L (A) or CCR7 expression (B) of CD8+ T cells. The experiment was repeated twice with similar results.

 

    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
This study shows that although the majority of antiviral memory CD8+ T cells can be found in the spleen after primary infection or immunization of mice with RSV or vaccinia virus, this important lymphoid organ is dispensable for efficient T cell-mediated antiviral recall responses in vivo.

The anatomic distribution of antiviral memory CD8+ T cells has been characterized in several previous studies. The most comprehensive study has been presented by Marshall et al., who have meticulously analyzed the localization of CD8+ memory T cells after intranasal influenza reinfection of mice primed by intraperitoneal influenza inoculation (5). In that study, most specific memory T cells were found in the spleen followed by bone marrow, lung, liver and lymph nodes. A factor of 8–10-fold more T cells were found in the spleen than in the other organs analyzed. Haanen et al. studied the localization of virus-specific memory T cells after primary influenza infection and reported 50-fold more T cells in the spleen than in lung or lymph nodes (4). Masopust et al. studied memory CD8+ T cell distribution after systemic infection with vesicular stomatitis virus and found ~2-fold more T cells in the spleen compared to the cumulative number of all other tissues analyzed (7). In all of these studies, including our own, the number of virus-specific memory CD8+ T cells in non-splenic compartments was probably underestimated. We did not analyze NALT, Peyer's patches, intraepithelial lymphocytes, some lymph nodes and other sites of bone marrow production. Nevertheless, it is unlikely that memory T cells in these organs will account for the 10–30-fold difference we observed between the spleen and non-splenic compartments. Our experiments with local and systemic infections with RSV and vaccinia virus thus confirm that the spleen is the major reservoir of memory CD8+ T cells after viral infections in mice.

The surprising finding of this study was that this large reservoir of antiviral memory T cells is apparently dispensable for CD8+ T cell-mediated protective antiviral memory responses in vivo. Our experimental set-up required that memory T cells not only underwent proliferation and reactivation, but also migrated to the lung to perform their antiviral effector function. The accumulation of virus-specific memory T cells in the lung proceeded unimpaired in splenectomized mice, since, irrespective of the route of primary immunization, protective immunity was not impaired. Even under limiting conditions, when T cell immunity induced by the immunization mediated only partial protection, splenectomy did not impair the recall response. In the particular context of RSV infection, these data also provide further evidence for the significant contribution of T cell memory to protective immunity against this respiratory virus (14,15) and argue against previously postulated T cell immunosuppressive properties of RSV (16).

It has been shown that the activation state of antiviral memory T cells isolated from non-lymphoid tissue is higher than that of spleen cells (7). In our experiments, the largest proportion of CD62low CCR7low RSV-specific CD8+ T cells could be found in the lung, although—similar to CD4+ T cells in influenza infection (17)—a significant proportion retained expression of these two markers. This preferential localization of ‘effector memory’ (18) T cells to non-lymphoid tissue could explain why splenic memory T cells are dispensable for protective immunity in the lung. However, although it has been shown that pulmonary effector memory T cells can contribute to protective immunity (19), it is not clear how important this contribution really is. In addition, our experiments show that the spleen is also dispensable after extrapulmonary priming infection with recombinant vaccinia viruses, when much less RSV specific ‘effector memory’ T cells persist in the lung. Alternatively, virus control by memory T cells in the lung could be mainly mediated by ‘central memory’ T cells recruited from lymphoid tissue, which rapidly proliferate and differentiate in draining lymph nodes in response to re-exposure to viral antigen. Adoptive transfer experiments have shown that effective pulmonary T cell responses can be rapidly amplified from systemic memory T cells in the absence of memory T cells persisting in the lung from a previous infection (8,9). The rapid proliferation dynamics of such cells present in extrasplenic lymphoid compartments may well be sufficient to compensate for the loss of the spleen. Splenectomy may also affect the distribution and mobility of the recirculating lymphocyte pool, leading to enhanced accumulation in lymph nodes (20). Such changes could also compensate for the numerically significantly reduced memory T cell pool.

The clinical question behind this study is whether T cell immunity to viral infections acquired by wild-type infection or vaccination is impaired in patients who underwent splenectomy. Although increased rates of viral reinfection or reactivation have not been reported in splenectomized patients (21), this question has not been addressed in detail. Most viral reinfections are controlled by neutralizing antibodies (22) such that contributions of memory T cells may be difficult to assess (23). Wolf et al. have shown that there is a long-term decrease in the percentage of CD4+ CD45RA+ lymphocytes accompanied by impaired primary responses to a T cell-dependent viral antigen (24). However, levels of CD4+ CD45RO+ T cells were not influenced by splenectomy and changes in circulating antigen-specific T cells have not been analyzed. Using the RSV mouse model, we were not only able to quantify a virus-specific memory T cell population at the site of challenge infection in splenectomized versus control mice, but we were also able to test their ability to mediate protective immunity. Using three different routes of priming with two different viruses, we mimicked memory T cell induction by local primary infection or by distant local or systemic vaccine inoculation. Our data show that, at least in mice, protective antiviral T cell memory is not impaired after splenectomy.


    Acknowledgements
 
We thank Matthias Brandis (Children's Hospital, Freiburg) for continuous support and Nicole Wehrle and Annette Schult for excellent technical assistance. We gratefully acknowledge the NIH tetramer facility for providing MHC I tetramers and Heike Unsoeld for providing COS cell supernatant containing CCL19-Ig. This work was supported by the Deutsche Forschungsgemeinschaft (SFB 620/A4).


    Abbreviations
 
BAL   bronchoalveolar lavage
p82   RSV matrix protein (M2) peptide 82–90
RSV   respiratory syncytial virus
rVV   recombinant vaccinia virus

    Notes
 
Transmitting editor: P. Ohashi

Received 13 May 2004, accepted 1 October 2004.


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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