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. Author manuscript; available in PMC: 2013 Sep 1.
Published in final edited form as: J Immunol. 2012 Jul 23;189(5):2432–2440. doi: 10.4049/jimmunol.1200799

CD8 T cells are essential for recovery from a respiratory vaccinia virus infection

John Goulding 1,2, Rebecka Bouge 1, Vikas Tahiliani 1,2, Michael Croft 1, Shahram Salek-Ardakani 1,2
PMCID: PMC3496758  NIHMSID: NIHMS389052  PMID: 22826318

Abstract

The precise immune components required for protection against a respiratory Orthopoxviridae infection, such as human smallpox or monkey pox, remain to be fully identified. In this study we utilized the virulent Western Reserve strain of vaccinia virus (VACV-WR) to model a primary respiratory Orthopoxviridae infection. Naïve mice infected with VACV-WR mounted an early CD8 T cell response, directed against dominant and subdominant VACV-WR antigens, followed by a CD4 T cell and immunoglobulin (Ig) response. In contrast to other VACV-WR infection models that highlight the critical requirement for CD4 T cells and Ig, we found that only mice deficient in CD8 T cells presented with severe cachexia, pulmonary inflammation, viral dissemination and 100 % mortality. Depletion of CD8 T cells at specified times throughout infection highlighted that CD8 T cells perform their critical function between days four and six post infection and that their protective requirement is critically dictated by initial viral load and virulence. Finally, the capacity of adoptively transferred naïve CD8 T cells to protect RAG -/- mice against a lethal VACV-WR infection demonstrated that CD8 T cells are both necessary and sufficient in protecting against a primary VACV-WR infection of the respiratory tract.

Keywords: CD8 T cells, Antibodies, Vaccinia Virus, Poxviruses, Lung

Introduction

Orthopoxviridae genus members, including Variola (VARV), Ectromelia (ECTV), Monkeypox (MPXV) and Vaccinia virus (VACV), are large double-stranded DNA viruses that encode transcription and replication machinery that facilitates their life cycle within the cytoplasm of the host cell (1). Despite the global eradication of VARV, the etiological agent of human smallpox, the continuing threat of intentional or accidental release of VARV and a growing number of human respiratory MPXV and natural VACV infections has propelled a renewed interest in Orthopoxviridae research (2, 3) (4-6). Historically, in vivo models have utilized both VACV and ECTV virus to simulate a primary infection and vaccination regimes to understand the essential immune requirements needed for protection. Collectively these studies emphasized the necessity for CD4 T cells and B cell derived immunoglobulin (Ig) in protecting against a primary and secondary Orthopoxviridae infection (7-11). However, the majority of these studies utilized infection models that do not represent the natural route of a primary Orthopoxviridae infection in humans. To address this a number of respiratory Orthopoxviridae infection models were developed (12-17).

In many respects intranasal (i.n.) infection with the highly virulent mouse adapted VACV Western reserve (VACV-WR) strain simulates the spread of smallpox virus throughout the respiratory tract resulting in severe disease that is associated with pulmonary inflammation, intra-alveolar edema, hemorrhage, peribronchial and perivascular inflammation and subsequent death (16, 18, 19). Initial VACV-WR replication is thought to occur in the respiratory epithelium and alveolar macrophages before the development of a transient viremia that spreads virus to peripheral reticuloendothelial cells throughout the host (12, 20). In the past several years a number of studies have established an important role for innate immune cells such as NK cells (21, 22), monocytes (23, 24), and dendritic cells (25) during primary VACV infections, however, the relative contribution of adaptive immune cells in particular CD8 T cells remains unclear.

Herein we show that a respiratory VACV-WR infection elicits a robust antigen specific CD8 T cell response that is shortly followed by the development of a VACV-specific humoral response that is completely dependent on CD4 T cell help. Unexpectedly we found that CD8 but not CD4 T cell deficient or depleted mice failed to recover from infection and presented with elevated viral titers and 100 % mortality by day nine post infection. Selective depletion of CD8 T cells at specified times throughout infection highlighted a critical role for CD8 T cells between days four and six post infection. Finally, as evident by the capacity for adoptively transferred naïve CD8 T cells to protect RAG -/- mice against a lethal VACV-WR infection, we reveal that CD8 T cells are both necessary and sufficient in protecting against a primary VACV-WR infection of the respiratory tract.

Materials and Methods

Mice

The studies reported here conform to the animal welfare ACT and the National Institutes of Health guidelines for the care and use of animals in the biomedical research. All experiments were completed in compliance with the regulations of the La Jolla Institute Animal Care Committee and in accordance with the guidelines of the Association for Assessment and Accreditation of Laboratory Animal Care. Eight to 12 week old female C57BL/6J (WT), RAG-/-, CD8-/-, MHC I-/-, MHC II-/-, and μMT mice were purchased from the Jackson Laboratory, USA.

Vaccinia virus and stock preparation

Vaccinia virus Western Reserve (VACV-WR), New York City Board of Health (NYCBOH) and Lister strains were purchased from the American Type Culture Collection (ATCC), grown in HeLa cells and subsequently titered on VeroE6 cells as described previously (26).

Respiratory VACV-WR infection model

Naïve mice were anesthetized by isoflurane inhalation and infected intranasally (i.n.) with 1.25 × 103 – 2 × 106 PFU of the indicated VACV strain, with daily measurements of body weight, lung pathology, and viral titers (as described before (27)). No animals were allowed to die of natural causes; therefore, the time of death indicated on the survival curves is the time at which an animal was euthanized due to severe disease (weight loss of > 25 %).

VACV titer assay

After i.n. infection, specified tissues from individual mice were homogenized and sonicated for 1 minute with a pause every 10 seconds using an ultrasonic cleaner (1210 Branson). Serial dilutions were made, and virus titers were determined by plaque assay on confluent VeroE6 cells.

Flow cytometric analysis

Spleens, lungs and lymph nodes were aseptically removed from euthanized mice and single-cell suspensions were prepared by mechanically dispersing the tissues through 70-μm cell strainers (Falcon BD Labware) into Hanks balanced salt solution (HBSS). In addition, before mechanical disruption the lung tissue was treated for 1 hr at 37°C with 250 μg Collagenase D (Roche) followed by treatment for 10 minutes at 4°C with 100mM EDTA supplemented media. Following red blood cell lysis (Sigma Aldrich), cells were re-suspended in RPMI 1640 medium (Invitrogen) supplemented with 10 % FCS (Omega Scientific). 1 % l-glutamine (Invitrogen), 100 μg/ml streptomycin, 100 U/ml penicillin, and 50 μM 2-ME (Sigma-Aldrich) and enumerated using a BD automated Vicell counter.

T cell, NK cell, NKT cell and germinal cell staining

Cells were washed with FACS buffer (PBS and 2 % FCS) and stained with anti-Fc II/III receptor monoclonal antibody 2.4G2 for 15 min at 4°C. After an additional FACS buffer wash, the following antibodies were incubated with the cells for 30 mins at 4°C: CD4 (RM4-5, BD Pharmingen), CD8 (53-6.7, BD Pharmingen), CD3 (145-2C11, eBioscience), NK1-1 (PK136, eBioscience) DX5 (CD49b) (DX5, eBioscience) were used to determine T cell, NK and NKT cell subsets, whilst anti-mouse IgD (11-26; Southern Biotech), CD138 (281-2, RDI) and PNA (FITC; Vector Laboratories), FAS (Jo2; BD Pharmingen) delineated plasma cell and germinal center B cells, respectively (28). All samples were acquired on a FACS Calibur flow cytometer or Canto II (BD Bioscience) and analyzed using FlowJo (Tree Star).

VACV-WR specific IFN-γ cytokine production

CD8 and CD4 T lymphocyte VACV-WR specific cytokine production was assessed as previously described (26, 28). Briefly, after lysing RBCs, splenocytes and lung cells from infected mice were plated in round-bottom 96 well micro-titer plates in 200 μl of RPMI 1640 medium (Invitrogen) supplemented with 10 % FCS (Omega Scientific), 1 % l-glutamine (Invitrogen), 100 μg/ml streptomycin, 100 U/ml penicillin and 50 μM 2-ME (Sigma-Aldrich). 10 μg/ml of the indicated MHC class I or II restricted VACV peptide was then added and incubated for 1 h at 37°C. The VACV peptide epitopes used in this study to identify virus specific T cells were predicted and synthesized as described previously (29-31). GolgiPlug (BD Biosceinces) was then added to the cultures according to the manufacturer's instructions and the incubation was continued for 9 h. Cells were then stained with anti-CD4 (RM4-5, BD PharMingen) or anti-CD8 (53-6.7, BD Pharmingen) and anti-CD62L (MEL-14; BD PharMingen) followed by fixation with Cytofix-Cytoperm (BD Biosceinces) for 20 min at 4°C. Fixed cells were subjected to intracellular cytokine staining in perm/wash buffer (BD Biosciences) for 30 min at 4°C. Cells were stained with anti-IFN-γ (XMG1.2, eBioscience) for 30 min at 4°C. Samples were analyzed for their proportion of cytoplasmic cytokines after gating on CD4 or CD8 CD62L-low T cells by a FACS Calibur flow cytometer using CellQuest and FlowJo software.

In vivo CD8 and CD4 T cell cell depletion

Hybridomas were cultured in protein-free hybridoma medium-II (Invitrogen), and mAbs were isolated by dialysis of supernatant. Groups of VACV-WR infected mice were depleted of CD8 and/or CD4 T cells with anti-CD8 (clone 2.43; 200 μg/mouse) and anti-CD4 (GK1.5; 200 μg/mouse) given in one i.v. injection 3 days before, and i.p. injections on days -1 and every 3 days thereafter until the termination of the experiment. T cell depletion was confirmed by flow cytometry of peripheral blood and lung tissue.

In vivo naïve CD8+ T cell transfer

Naïve CD8 T cells (CD3+ CD8+ CD44lo) were isolated from naïve wild type C57BL/6J mice. Briefly, naive spleens were homogenized to a single cell suspension as described above, anti-CD8 microbeads (Miltenyi) were subsequently added following manufacturers instructions. Following CD8 T cell MACS column enrichment the naïve CD8 T cells were further purified using CD3+ CD44lo populations and FACS sorted with a BD Aria. Subsequently 5 × 106 naïve CD8 T cells/mouse were transferred into aged matched RAG-/- via the retro orbital plexus.

Measurement of serum VACV-WR specific IgG isotype titers

Serum was obtained after centrifugation of blood samples collected with a heparinized capillary pipette from the retro-obital plexus. All samples were stored at - 20°C until analyzed for antibody titre. The level of specific antibodies against VACV-WR in serum was quantitated by an enzyme-linked immunosorbent assay as previously described (32-34).

In vivo passive VACV-WR immunization

VACV-WR immune or control serum was prepared from naïve (WT) mice that were previously infected (i.n) with VACV-WR (1 × 104 PFU/mouse). Aged matched naïve C57BL/6J mice were injected i.p with 250 μl of serum from the indicated mice. The following day, mice were anesthetized by inhalation of isoflurane and infected i.n. with 1 × 104 PFU VACV-WR. Mice were weighed daily for 2 weeks following infection and were euthanized at a pre-determined time point or if they lost > 25 % of their initial body mass.

Statistical analysis

Tests were performed using Prism 4.0 (GraphPad, San Diego, CA). Statistics were done using two-tailed, unpaired Student's t test with 95 % confidence intervals unless otherwise indicated. Two-way ANOVA was used to determine differences in weight loss profiles and the Mantel-Cox test was utilized for survival analysis. Unless otherwise indicated, data represent the mean ± one SEM, with p < 0.05 considered statistically significant.

Results

Respiratory VACV-WR infection results in severe lung inflammation

To study the role of CD8 T cells on the susceptibility to respiratory infection with vaccinia virus (VACV), naïve C57BL/6 wild-type (WT) controls were infected via the intranasal (i.n.) route with a low (1 × 104 PFU), medium (5 × 104 PFU), or high (1 × 106 PFU) dose of VACV-WR, with daily measurements of body weight, lung pathology, and viral titers (Fig. 1A). Mice infected with 5 × 104 PFU of VACV-WR or greater had lost greater than 25 % of their initial weight by day 7 post-infection (Fig. 1A), when they were euthanized for humane reasons. These mice were hunched, had significantly labored breathing, ruffled fur, and were minimally responsive to manipulation. Hematoxylin and eosin (H & E) stained sections of lungs were evaluated for the presence of inflammation, hemorrhage, edema, and necrosis. Alterations were not observed in any lung samples obtained at 1 and 2 days post infection (not shown). However by day 6 (Fig. 1B; middle panel), the majority of virus-infected mice had moderate to severe, multifocal, mixed (predominantly mononuclear) inflammation that tended to be focused around the bronchioles and blood vessels. The degree of bronchiolar epithelial hyperplasia and necrosis was moderate to severe with several bronchioles being affected. Generally, there was mild to moderate, multifocal, vascular necrosis and occasionally mild to moderate hemorrhage and edema. Thus, the lesion characteristics were consistent with a diagnosis of moderate to severe viral bronchopneumonia characterized by mononuclear inflammation, bronchiolar hyperplasia, and necrosis.

Figure 1. Respiratory VACV-WR infection results in severe lung inflammation.

Figure 1

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Wild type C57BL/6J mice (WT) were intranasally (i.n.) infected with increasing plaque forming units (PFU) of VACV-WR. Body mass was monitored daily (a) and lung inflammation was determined by staining with H & E (magnification × 10) (b). Three representative micrographs (i-iii) are shown for two separate PFU doses. At specified time points following infection with 1×104 pfu VACV-WR tissue specific viral titers were measured (c). Body mass and viral titers are presented as the mean ± one SEM of three separate experiments containing 5-12 mice per group; or as a representative micrograph of lung tissue.

Mice infected i.n. with 1 × 104 PFU VACV-WR produced a less severe disease, reduced weight loss (Fig. 1A), reduced lung inflammation and pathology (Fig. 1Bbottom row) at day 6 post-infection. All mice started to recover from disease starting at day 7, returning to their original mass by day 14 p.i. (Fig. 1A). The titer of infectious virus in lung tissue and airways of VACV-WR infected mice followed the pattern observed in the weight loss and histological assessment (Fig. 1C). While a large amount of infectious virus was recovered from VACV-WR-infected lungs at 4 and 6 days p.i., by 10 days p.i. little or no virus could be detected. Thus infection with 1 × 104 PFU VACV-WR promoted a robust immune response that led to virus clearance from the lung within 10 days and protected all animals from death; therefore 1 × 104 PFU was utilized to further characterize the protective components of a primary respiratory infection with VACV-WR.

Mice lacking adaptive immunity fail to control a respiratory VACV-WR infection

To determine the importance of T and B cells we studied infection in RAG-/- mice deficient in mature T and B cells due to an inability to perform V(D)J recombination (35). Following infection with 1 × 104 PFU VACV-WR the initial weight loss and illness score was comparable to wild type mice, however progressive weight loss and illness in the RAG -/- mice resulted in 100 % mortality by day 12 p.i, suggesting an important contribution of B and/or T cells in controlling VACV-WR infection. (Fig. 2A). Histo-pathological analysis of the lung tissue on day 7 and 10 p.i highlighted extensive lung pathology, cellular infiltrate, alveoli destruction and pulmonary edema in the RAG -/- mice compared to WT controls (Fig. 2B). RAG -/- mice also failed to contain initial viral titers in the lung and displayed signs of viral dissemination as determined by the significant levels of virus in their ovaries (Fig. 2C).

Figure 2. Adaptive immunity is necessary to control a respiratory VACV-WR infection.

Figure 2

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Wild type C57BL/6J (WT) and RAG-/- mice were infected i.n. with 1×104 pfu VACV-WR and weighed daily (a). Lung inflammation was assessed by H & E (magnification × 10) staining on days seven and ten post infection (b) and viral titers were measured in the lung and ovaries also on day ten post infection (c). Body mass and viral titers are presented as the mean ± one SEM of four independent experiments with 4-8 mice per group; or as a representative micrograph of lung tissue. Students t test with Bonferroni's correction was used to determine statistical significance * p<0.05, ** p<0.01

Robust mucosal and systemic T cell responses to a respiratory VACV-WR infection

To determine if recovery from a primary respiratory VACV-WR infection in WT mice correlated with the presence of T cells at the site of infection, groups of age-matched naïve WT mice were infected i.n. with VACV-WR and the kinetics of lymphocyte recruitment to the lung and spleen were determined. The number of cells recovered from the lung increased between days 3 and 7, peaked between days 7 and 10, and decreased by day 15 (Fig. 3A). The majority of the lymphocytes infiltrating the lung and spleen were CD8 T cells (Fig. 3A & B). Between day 7 and day 10 p.i., the number of CD8 T cells in the lung was at least 10-20 fold higher than that of CD4 T cells. To determine the specificity and functionality of the T cell response at the peak of the primary response, total lung and spleen cells were isolated on 10 p.i. and stimulated ex vivo with different VACV peptides. As the extent of CD8 and CD4 T cell VACV responses is large, with a total of 49 CD8 and 14 CD4 antigens recognized thus far in total (29-31), we utilized eight immune-dominant antigens to interrogate the diversity of VACV-specific T cell response. Interferon-γ (IFN-γ) producing CD8 T cells specific for B8R, B16R, J3R and A8R viral antigens dominated both the lung and splenic T cell response, whilst VACV-specific CD4 T cells targeted multiple epitopes with similar magnitude (Fig. 3C & D). The emergence of VACV-specific T cells in the lung tissue, and their secretion of IFN-γ in these sites paralleled the clearance of virus from the lungs of infected mice and the cessation of weight loss, suggesting that effector CD4 and/or CD8 T cells are playing critical roles in recovery from infection.

Figure 3. Lung and splenic T cell responses to a respiratory VACV-WR infection.

Figure 3

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Wild type C57BL/6J mice (WT) were infected i.n. with 1×104 PFU of VACV-WR. Total lung (a) and spleen (b) CD8 and CD4 T cell numbers were measured at various time points post infection. The percentage of VACV-WR specific CD8 (c) and CD4 (d) IFN-γ producing T cells in the lung and spleen on day 10 post infection were also enumerated. T cell numbers are presented as the mean ± one SEM of three separate experiments containing 5-8 mice per group; or as a representative FACS plot from the represented tissue. Students t test with Bonferroni's correction was used to determine statistical significance * p<0.05, ** p<0.01

B cell and immunoglobulin responses to a respiratory VACV-WR infection

IgG and IgM class-specific endpoint ELISA was used to investigate the kinetics of VACV-specific Ig in the serum of mice infected i.n. with VACV-WR. After infection, VACV-specific serum IgG was still undetectable on day 10, a time at which the animals were already recovering from disease (Fig. 1A & C). Low levels of VACV-specific IgG were measured on day 15 and increased thereafter reaching maximal levels by day 148. Previously we and others have shown that almost all of the anti-VACV IgG response after i.p infection is CD4 T cell help dependent (7, 33, 36). Therefore, we used MHC class II deficient mice, which lack CD4 T cells, to evaluate the T cell help requirement for production of anti-VACV IgG after an i.n. infection. Similar to B cell deficient mice, no anti-VACV IgG was found in MHC II -/- mice (Fig. 4B). Analysis of the relative contribution of virus specific IgG isotypes indicated that the levels of IgG1 (not shown) and IgG2c also peaked between day 30 and 148 p.i. and that IgG2c was the most abundantly produced isotype (Fig. 4C). IgM was also produced in response to respiratory VACV-WR infection, however it had more rapid kinetics than IgG and peaked in titer between day 7 and 10 p.i before diminishing by day 30 p.i. (Fig. 4D).

Figure 4. B cell and immunoglobulin responses following a respiratory VACV-WR infection.

Figure 4

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Wild type C57BL/6J mice (WT) were infected i.n. with 1×104 PFU of VACV-WR. Absolute serum concentrations of VACV-WR specific immunoglobulin (Ig) G (a), IgG2c (c) and IgM (d) were determined in wild type mice by ELISA at specific days post infection. Total IgG levels were also measured on day 30 post infection in MHC II-/- (CD4 T cell deficient) and μMT (B cell deficient) mice (b). End point Ig titers equates to the dilution needed to produce an absorbance value of 0.2. Lung draining lymph node germinal center (GC) (B220+FAS+PNA+) and plasma cell (B220+CD138+IgD-) development were also measured at specified times following infection (e). All immunoglobulin levels are presented as the mean ± one SEM of three independent experiments with 5-8 mice per group, or as a representative FACS plot. Students t test with Bonferroni's correction was used to determine statistical significance * p<0.05, ** p<0.01

Mediastinal lymph nodes (MLN) drain the lung and are a site where mucosal immune responses are initiated against antigens reaching the lung. Indeed, we found high frequencies of germinal center (GC) B cells (PNA+FAS+) within these LN (Fig. 4ETop panel), consistent with the presence of IgG Ab in the serum (Fig. 4A). Frequencies of GC B cells declined over time, reaching basal levels by day 30 (not shown). The development of MLN B cells positive for the plasma cell differentiation marker, CD138, followed similar kinetics to that of GC B cells (Fig. 4EBottom panel). Together these data highlighted that the development of germinal center and Ig secreting plasma cells following a respiratory VACV infection occurred after the peak of the T cell response but was maintained over the course of several months.

CD4 T cells and Ig are not required for protection against a respiratory VACV-WR infection

To assess the contribution of CD4 effector T lymphocytes in virus clearance and recovery, we infected MHC II -/- mice, which are devoid of CD4 T cells, with VACV-WR and monitored their weight loss over time. MHC II -/- mice displayed comparable weight loss and recovery profiles to that observed in wild type mice (Fig. 5A), even though Ig production was inhibited (Fig. 4B). Also, in vivo depletion of CD4 T cells in WT mice did not substantially modify either survival (Fig. 5A) or virus clearance from the respiratory tract (not shown). Consistent with this, lung pathology and lung infiltrate was similar between CD4 T cell deficient and WT isotype treated (Ig) control mice (Fig. 5B).

Figure 5. CD4 T cells and Ig are not required for protection against a respiratory VACV-WR infection.

Figure 5

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Wild type C57BL/6 (WT) and CD4 T cell deficient mice (CD4 depleted and MHC II-/-) were infected i.n. with 1×104 pfu VACV-WR. Their body mass (a) and lung inflammation was assessed on specified days post infection (b). In a separate experiment 200ul of control (naïve) and hyper-immune serum, isolated from WT mice on day four, seven, ten, 15 and 148 post i.n. infection with 1×104 pfu VACV-WR, was injected i.p. into naïve WT mice (c). 24 hours later the passively immunized mice were infected i.n. with 1×104 PFU VACV-WR and their body mass monitored daily (d and e). Data are presented as the mean ± one SEM of three independent experiments with 4-8 mice per group; or as a representative micrograph of lung tissue. Students t test with Bonferroni's correction was used to determine statistical significance * p<0.05, ** p<0.01

To address the role of Ig during a primary respiratory infection with VACV-WR more directly, serum was prepared from mice that were infected with VACV-WR 4, 7, 10, 15, or 148 days prior to the start of the experiment and transferred i.p into naïve WT mice. Serum from uninfected (naïve) mice was used as control. The following day, all mice were challenged with a sublethal i.n. inoculum of VACV-WR and weight loss was monitored for 15 days (Fig. 5C). All mice that received serum from day 4 and day 7 VACV-infected mice (anti-VACV IgG negative, anti-VACV-IgM low) displayed comparable weight loss and recovery to mice that received control serum (Fig. 5D). In contrast, mice that received hyper-immune serum isolated after day 10 of a respiratory VACV infection (anti-VACV IgG positive) exhibited a significant reduction in weight loss (Fig. 5E) and accelerated recovery. These data together with the kinetics of GC B cell development and viral clearance suggest that humoral responses develop too late to be of significant value in the initial containment of VACV replication in the lung.

An early CD8 T cell response is necessary for protection during a respiratory VACV-WR infection

One likely explanation for recovery of CD4-depleted or MHC II -/- mice after a respiratory VACV-WR infection is the ability of CD8 T cells to compensate for the lack of CD4 and antibody responses. To evaluate this MHC II -/- mice were infected with VACV-WR and primary ex vivo CD8 T cell responses were measured at day 10, the peak time of anti-VACV response (Fig. 6A). In agreement with our previous data in the i.p infection model (27, 36), we found that CD4 T cell help is not required for the generation of VACV-specific effector CD8 T cells after i.n. infection. Indeed, the frequency of IFN-γ producing virus-specific CD8 T cells were slightly elevated in the lung and spleens of VACV-infected MHC II -/- mice as compared with WT controls.

Figure 6. An early CD8 T cell response is necessary for protection during a respiratory VACV-WR infection.

Figure 6

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Total lung and splenic IFN-γ and TNF-α producing CD8 T cells in wild type C57BL/6 (WT) (black bars) and MHC II-/- (open bars) mice 8 days following an i.n. infection with 1×104 pfu VACV-WR was assessed after ex-vivo stimulation with B8R-peptide (a). Subsequently, WT, CD4 and CD8 (b – left panel) and CD8 alone (b – right panel) T cell deficient mice were infected i.n. with 1×104 pfu VACV-WR and monitored for weight loss. Lung inflammation (c) and tissue viral titers (d) in mice deficient of CD8 T cells were assessed at day eight post infection. Lastly, weight loss and survival (e) was measure in WT mice depleted, or not, of CD8 T cells at the same time as, 3 days or 6 days after i.n. infection with 1×104 pfu VACV-WR. In addition, the percentage (f) and total number (g) of lung B8R tetramer positive and IFN-γ producing CD8 T cells was enumerated on day 3.5, 4.5 and 5.5 p.i. Data are presented as the mean ± one SEM of three independent experiments with 4-8 mice per group; or as a representative micrograph of lung tissue. Students t test with Bonferroni's correction was used to determine statistical significance * p<0.05, ** p<0.01.

To test if CD8 T cells can protect in the absence of CD4 T cells we depleted both subsets of T cells simultaneously, starting before infection and continuing during the observation period. Strikingly, depletion of CD8 T cells in CD4-depleted mice resulted in 100 % mortality by day nine p.i (Fig. 6B, left panel), implying that the CD8 subset was indeed required for the survival of the CD4-deficient mice. This result was phenocopied in MHC I -/-, CD8 -/- and WT mice depleted of CD8 T cells throughout the course of a VACV-WR infection (Fig. 6B, right panel). All mice deficient of CD8 T cells presented with profoundly altered lung architecture, multiple foci of perivascular and peribronchial inflammation consisting of polymorphonuclear and some mononuclear cells (Fig. 6C). Consistent with this, VACV clearance was markedly impaired in CD8 T cell depleted mice, which showed a 1000-fold and 10000-fold increased viral load at day seven after infection as compared with WT controls (Fig. 6D). Virus titers in the ovaries were also highly elevated (1000- to 10000-fold) after infection with VACV-WR, which was similar to titers found in RAG -/- infected mice (Fig. 2C). A concern with systemically administered depleting antibodies is an unexpected affect on other cell types important for protection. To verify this we measured the proportion and total number of lung and splenic NK and NK T cells and demonstrated no reduction in mice transiently depleted of CD8α+ cells (Supplementary Fig 1).

To determine the time at which CD8 T cells provide their protective function we depleted CD8 T cells in WT mice starting on days -1, 3, or 6 after a primary infection. Mice depleted of CD8 T cells throughout the course of infection or on day three onwards failed to control the primary VACV-WR infection and resulted in 100 % mortality by day eight p.i (Fig. 6E). Both groups of mice presented with similar lung pathology, systemic inflammation and viral dissemination similar to that observed in MHC I -/- and CD8 -/- mice (not shown). Mice depleted of CD8 T cells after day six post VACV-WR infection demonstrated comparable weight loss and survival to the isotype treated (Ig) control mice (Fig. 6E). This time dependent importance for CD8 T cells was further supported by the observation that VACV-specific CD8 T cells, that are capable of producing IFNγ, began entering the lung tissue on day 4.5 days p.i before increasing significantly by day 5.5 p.i. (Fig. 6F & G). Thus, our results demonstrate that an early CD8 T cell response to respiratory VACV infection is crucial for host defense, whereas CD4 T cells appear neither required for virus clearance nor for the induction of this protective response.

Initial viral load and virulence critically determines the requirement for CD8 T cells in protection against a respiratory VACV-infection

Next we hypothesized that the protective requirement for CD8 T cells during a primary anti-VACV immune response in the lung may vary with the infective inoculum and the virulence of strain used for infection. First we assessed the role of CD8 T cells during a respiratory VACV-WR infection with a reduced inoculum bolus. WT mice infected with 1 × 103 PFU of VACV-WR, a log less than our determined sub-lethal dose, experienced moderate weight loss (10 – 15 %) but quickly recovered and went on to maintain normal weight after day 10 (Fig. 7A). Strikingly, mice depleted of CD8 T cells throughout a low dose VACV infection presented comparable weight loss and survival to the control treated mice (Fig. 7A). To investigate the requirement for CD8 T cell immunity during a less virulent VACV infection we utilized two live-attenuated VACV variants that differ in their expression of several virulence factors that determine their replicative capacity and virulence in mice (37, 38). This reduced virulence is revealed by the need to utilize a significantly higher infectious dose (2 × 106 PFU/mouse) to provoke a measurable immune response and the lack of significant weight loss or outward signs of illness throughout infection. Similar to a low dose VACV-WR infection the depletion of CD8 T cells throughout infection with Lister or NYCBOH did not affect weight loss, illness or recovery (Fig. 7B and C). Therefore CD8 T cells are necessary for protection against a respiratory VACV-WR infection but are not required when the infectious inoculum is reduced or a less virulent VACV strain is used.

Figure 7. Initial viral load and virulence critically determines the requirement for CD8 T cells in protection against a respiratory VACV-WR infection.

Figure 7

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Wild type C57BL/6 (WT) mice were depleted of CD8 T cells and infected i.n. with 1.25×103 pfu VACV-WR (low dose) (a), 2×106 pfu VACV Lister (b) or 2×106 pfu VACV NYCBOH (c) and monitored for weight loss. Data are presented as the mean ± one SEM of three independent experiments with 4-8 mice per group. Students t test with Bonferroni's correction was used to determine statistical significance * p<0.05, ** p<0.01.

CD8 T cells are sufficient for protection during a respiratory VACV-WR infection

We further addressed the ability of CD8 T cells to mediate protective immunity by adoptive transfer into naïve RAG -/- mice. Splenic naïve CD8 T cells (CD8+CD44-low) were enriched by magnetic bead purification and then sorted by FACS to a purity ranging from 99-100 %. Twenty-four hours later these mice along with two groups of control mice (RAG -/- and WT mice with no CD8 T cell transfer) were infected i.n. with 1 × 104 PFU VACV-WR and monitored for signs of weight loss and illness (Fig. 8A). As previously demonstrated RAG -/- mice fail to survive infection and succumb by day 10 p.i. (Fig. 8B and 8C). In marked contrast, RAG -/- mice that received naïve wild type CD8 T cells were protected and exhibited comparable survival rates to wild type controls (Fig. 8B and 8C). These results suggest that CD8 T cells can act independently of a humoral immune response in order to confer resistance to respiratory VACV-WR infection.

Figure 8. CD8 T cells are sufficient for protection during a respiratory VACV-WR infection.

Figure 8

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5×106 naïve (CD3+ CD8+ CD44 low) wild type CD8 T cells were transferred i.v. into RAG-/- mice that were infected i.n. 24 hours later with 1×104 pfu VACV-WR (a). Body mass (b), illness and survival (c) were followed throughout the experiment. Data are presented as the mean ± one SEM of three independent experiments with 4-5 mice per group. Two way ANOVA and Mantel-Cox tests were used to determine statistical significance * p<0.05, ** p<0.01, **p<0.001.

Discussion

To our knowledge, the present study is the first systematic evaluation of the host adaptive immune response to a respiratory VACV infection. We demonstrate that VACV infection results both in a cell-mediated immune response with the induction of virus-specific CD8 CTLs and CD4 T cells, and a humoral response with the production of VACV specific Ig. Contrary to alternative routes of VACV infection, we found that VACV-specific CD8 T cells, rather than CD4 T cells and Ig, are necessary and sufficient for clearing virus from the respiratory tract and protection against VACV-induced lung pathology and death. Most significantly, the protective requirement for CD8 T cells was critically dictated by the initial infection load and VACV strain virulence. These experiments help define the precise immune mechanisms that govern the efficient generation of protective immunity against acute respiratory poxvirus infections.

For many years the antiviral function of CD8 T cells in resistance against a primary infection with VACV has remained controversial. An early study by Spriggs et al. (8) showed that β2m-/- mice, which are devoid of CD8 T cells, were able to survive a subcutaneous infection with VACV-WR, even at doses exceeding 108 PFU, and experienced little if any outward signs of viremia or illness. These results demonstrated directly that CD8 T cells are not required for the clearance of a VACV infection as long as humoral immunity is intact. Similarly, Xu et al. (7) demonstrated that despite a robust cytotoxic CD8 T cell response in C57BL/6 mice following an intra-peritoneal (i.p) VACV-WR infection, depletion of CD8 T cells by mAb or deficiency in CD8 cells did not alter virus clearance or survival in comparison to control treated mice. On the contrary, CD4 T cells were shown to be essential for clearing an i.p. VACV infection. Accordingly, CD4-depleted and MHC class II-deficient mice were compromised in their ability to clear virus at day 14 p.i. and harbored 100-1000 fold higher titers than WT mice at day 20 p.i. B cell-deficient mice showed a similar inability to clear VACV as CD4-depleted mice. This failure of CD4 T cell-depleted, MHC class II -/- and Igh -/- mice to clear a primary i.p. VACV infection was attributed to their inability to mount an effective Ig response. Thus following an i.p infection with VACV, CD4-dependent virus-specific Ig responses appear to be the most important effector mechanism required for clearing the virus from infected tissues.

Our data now extend these observations by demonstrating that in the context of a respiratory VACV-WR infection CD8 T cells are both necessary and sufficient for recovery from disease. Interestingly, despite a robust CD4 T cell and IgG response, we found that both CD4-depleted WT and MHC class II -/- mice, which are unable to elicit effective CD4 T cell or Ig responses, cleared the virus form their lungs at similar rates as WT mice. Detailed kinetic analysis indicated that VACV replication in the lung is under control before serum IgG can be detected. Our data indicated that i.n. VACV infected mice did not develop circulating anti-VACV IgG until day 15, with a strong IgG response present by day 30. This is significantly later than the time point at which the presence of VACV in the lung can be detected. Consistent with this, the passive transfer of serum, collected from day four and seven infected mice, into naïve recipients failed to protect them from disease caused by an i.n. VACV challenge. In contrast, mice that received hyper-immune serum isolated after day 10 exhibited a significant reduction in weight loss and accelerated recovery following an i.n. VACV challenge. These data provide further evidence to suggest that the development of a virus specific Ig response occurs too slowly to limit VACV replication in the respiratory tract following a primary infection with VACV-WR. Rather, they suggest that the dominant role of a VACV-specific IgG response is to protect against secondary infections. However, the small delay in weight gain between day 10 and 18 post infection observed in MHC II-/- mice suggests a possible role for CD4 T cells and/or Ig during the recovery period.

The current study provides several lines of evidence that suggest that trafficking of CD8 T cells into a VACV-infected mouse lung is critical for viral clearance, survival and recovery. First, the emergence of IFN-γ-producing VACV-specific CD8 T cells in the lung tissue paralleled the clearance of virus from the lungs of infected mice and the cessation of weight loss. Second, by depleting CD8 T cells with mAb treatment throughout the course of infection, we found that in the absence of Ig, CD8 T cells are able to provide protection against an i.n. VACV infection in CD4-depleted mice. Third, the absence of CD8 T cells in MHC class I -/-, CD8 -/- mice or mice transiently depleted of this subset failed to clear VACV which resulted in severe lung immunopathology and 100 % mortality by day nine p.i.. Fourth, our kinetic studies indicated that CD8 T cells perform their protective role between days three and six post i.n. VACV infection. Interestingly, this was prior to the peak (between days seven and ten) of the CD8 T cell response, suggesting that relatively small numbers of CD8 T cells are capable of controlling VACV infection in the lung. Lastly, we observed that the adoptive transfer of large numbers of naïve polyclonal CD8 T cells into RAG 1 -/- recipient mice, containing presumably a very low frequency reactive with VACV, resulted in significantly reduced viral load in the lungs and fully protected mice from death following challenge with a lethal dose of VACV-WR. Collectively, these results demonstrate that CD8 T cells are a critical component of the adaptive immune response that act independently of CD4 T cell and humoral immune responses to control an acute respiratory VACV infection.

As briefly described above, much of the existing literature investigating the protective components of primary VACV infections utilize i.p., i.d., or s.c. infectious routes in order to simulate human vaccination and understand Ig development. This critical difference may explain the conflicting evidence for the requirement for CD8 T cells in protection against primary VACV infections. It is plausible that the inoculation of different tissues affects both the propensity of viral replication and dictates the kinetics, and thus the relative importance, of an ensuing cellular or humoral immune response. We highlight that despite infecting with 100 – 1000 times less VACV than that used during an i.p or s.c. infection, CD8 T cells are necessary for protection following a respiratory VACV-WR infection. This suggests that the route of infection dictates either the virulence of VACV or its capacity to replicate rapidly and thus the requirement for a protective CD8 T cell response. Essentially the higher the virulence or capacity for viral replication the greater the requirement for CD8 T cells for protection. This concept is supported by evidence generated using the mouse adapted and highly virulent ECTV. Analogous to a respiratory VACV infection, ECTV infection requires a rapid CD8 T cell response to contain initial viral titers before virus specific Ig is generated and assists in eliminating virus from infected tissues (39, 40). We also show that a reduction in the primary infectious dose, a parameter of replicative capacity, or decreased VACV virulence abrogated the requirement for CD8 T cells for protection. Clearly an important component of replicative capacity is the amount of time available for viral replication before a protective immune response is elicited. We demonstrate that CD8 T cell mediated immunity in the lung occurs between day three and six post infection. This suggests the existence of a time dependent titer threshold. One can consider that if the titer of VACV in the lung, or infected tissue, does not reach a specific threshold CD8 T cells are not required for protection. However, if this threshold is surpassed (by a greater inoculum dose or increased rate of viral replication) then CD8 T cells are required and essential for protection. In addition to viral titer, the time taken to reach this threshold or for alternative mechanisms of immunity to develop might also explain the differential requirement for CD8 T cells during other primary VACV infection models. We demonstrate that despite the generation of a long-lived virus specific IgG response following a respiratory VACV-WR infection, significant germinal center B cell number or Ig titers were not detected until day 10 p.i. In contrast, following an i.p VACV-WR infection, germinal center and plasma B cells are readily observed in the spleen as early as day five, implying the speed of antigen mobilization, presentation and Ig production might negate the requirement for CD8 T cells when infected via this route (33, 34). The ability for Ig to gain access to the site of infection might also impact the effectiveness of alternative protective immune mechanisms. For instance, it has been shown in mice and in humans that a neutralizing IgM response following VACV immunization is present within the first five days post infection and likely contributes to virus control before the development of IgG (33). However, our serum transfer data suggests that the presence of IgM on day seven following an i.n. VACV infection provides little if any protection, again highlighting the requirement for an early CD8 T cell response. Although pentameric IgM isotypes contain a functional J chain that allows trans-epithelial secretion, its protective role during a respiratory VACV infection remains to be determined.

In summary, this study adds to the current literature by providing evidence that CD8 T cells are both necessary and sufficient at protecting against a primary VACV-WR infection of the respiratory tract. These results highlight the plasticity of the immune system in combating VACV infections administered via different routes. Moreover they have important implications in furthering our understanding of host-pathogen interactions and also in the development of novel vaccines and therapeutics for human respiratory Orthopoxviridea infections.

Supplementary Material

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Acknowledgments

This work was supported by NIH grants AI77079 and AI087734 to S.S.-A., and AI67341 to M.C., and by a fellowship from the Center for Infectious Disease at the La Jolla Institute for Allergy and Immunology to S.S.-A. This is publication #1501 from the La Jolla Institute for Allergy and Immunology.

References

  • 1.Roberts KL, Smith GL. Vaccinia virus morphogenesis and dissemination. Trends Microbiol. 2008;16:472–479. doi: 10.1016/j.tim.2008.07.009. [DOI] [PubMed] [Google Scholar]
  • 2.Weinstein RS. Should remaining stockpiles of smallpox virus (variola) be destroyed? Emerg Infect Dis. 2011;17:681–683. doi: 10.3201/eid1704.101865. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Henderson DA, Inglesby TV, Bartlett JG, Ascher MS, Eitzen E, Jahrling PB, Hauer J, Layton M, McDade J, Osterholm MT, O'Toole T, Parker G, Perl T, Russell PK, Tonat K. Smallpox as a biological weapon: medical and public health management. Working Group on Civilian Biodefense. Jama. 1999;281:2127–2137. doi: 10.1001/jama.281.22.2127. [DOI] [PubMed] [Google Scholar]
  • 4.Rimoin AW, Mulembakani PM, Johnston SC, Lloyd Smith JO, Kisalu NK, Kinkela TL, Blumberg S, Thomassen HA, Pike BL, Fair JN, Wolfe ND, Shongo RL, Graham BS, Formenty P, Okitolonda E, Hensley LE, Meyer H, Wright LL, Muyembe JJ. Major increase in human monkeypox incidence 30 years after smallpox vaccination campaigns cease in the Democratic Republic of Congo. Proceedings of the National Academy of Sciences of the United States of America. 2010;107:16262–16267. doi: 10.1073/pnas.1005769107. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Megid J, Borges IA, Abrahao JS, Trindade GS, Appolinario CM, Ribeiro MG, Allendorf SD, Antunes JM, Silva-Fernandes AT, Kroon EG. Vaccinia virus zoonotic infection, sao paulo state, Brazil. Emerg Infect Dis. 2011;18:189–191. doi: 10.3201/eid1801.110692. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Breman JG, Arita I. The confirmation and maintenance of smallpox eradication. N Engl J Med. 1980;303:1263–1273. doi: 10.1056/NEJM198011273032204. [DOI] [PubMed] [Google Scholar]
  • 7.Xu R, Johnson AJ, Liggitt D, Bevan MJ. Cellular and humoral immunity against vaccinia virus infection of mice. J Immunol. 2004;172:6265–6271. doi: 10.4049/jimmunol.172.10.6265. [DOI] [PubMed] [Google Scholar]
  • 8.Spriggs MK, Koller BH, Sato T, Morrissey PJ, Fanslow WC, Smithies O, Voice RF, Widmer MB, Maliszewski CR. Beta 2-microglobulin-, CD8+ T-cell-deficient mice survive inoculation with high doses of vaccinia virus and exhibit altered IgG responses. Proceedings of the National Academy of Sciences of the United States of America. 1992;89:6070–6074. doi: 10.1073/pnas.89.13.6070. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Panchanathan V, Chaudhri G, Karupiah G. Protective immunity against secondary poxvirus infection is dependent on antibody but not on CD4 or CD8 T-cell function. J Virol. 2006;80:6333–6338. doi: 10.1128/JVI.00115-06. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Panchanathan V, Chaudhri G, Karupiah G. Correlates of protective immunity in poxvirus infection: where does antibody stand? Immunol Cell Biol. 2008;86:80–86. doi: 10.1038/sj.icb.7100118. [DOI] [PubMed] [Google Scholar]
  • 11.Belyakov IM, Earl P, Dzutsev A, Kuznetsov VA, Lemon M, Wyatt LS, Snyder JT, Ahlers JD, Franchini G, Moss B, Berzofsky JA. Shared modes of protection against poxvirus infection by attenuated and conventional smallpox vaccine viruses. Proceedings of the National Academy of Sciences of the United States of America. 2003;100:9458–9463. doi: 10.1073/pnas.1233578100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Williamson JD, Reith RW, Jeffrey LJ, Arrand JR, Mackett M. Biological characterization of recombinant vaccinia viruses in mice infected by the respiratory route. J Gen Virol. 1990;71(Pt 11):2761–2767. doi: 10.1099/0022-1317-71-11-2761. [DOI] [PubMed] [Google Scholar]
  • 13.Schriewer J, Buller RM, Owens G. Mouse models for studying orthopoxvirus respiratory infections. Methods Mol Biol. 2004;269:289–308. doi: 10.1385/1-59259-789-0:289. [DOI] [PubMed] [Google Scholar]
  • 14.Chapman JL, Nichols DK, Martinez MJ, Raymond JW. Animal models of orthopoxvirus infection. Vet Pathol. 2010;47:852–870. doi: 10.1177/0300985810378649. [DOI] [PubMed] [Google Scholar]
  • 15.Briody BA. Response of mice to ectromelia and vaccinia viruses. Bacteriol Rev. 1959;23:61–95. doi: 10.1128/br.23.2.61-95.1959. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Turner GS. Respiratory infection of mice with vaccinia virus. J Gen Virol. 1967;1:399–402. doi: 10.1099/0022-1317-1-3-399. [DOI] [PubMed] [Google Scholar]
  • 17.Hayasaka D, Ennis FA, Terajima M. Pathogeneses of respiratory infections with virulent and attenuated vaccinia viruses. Virol J. 2007;4:22. doi: 10.1186/1743-422X-4-22. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Reading PC, Smith GL. A kinetic analysis of immune mediators in the lungs of mice infected with vaccinia virus and comparison with intradermal infection. J Gen Virol. 2003;84:1973–1983. doi: 10.1099/vir.0.19285-0. [DOI] [PubMed] [Google Scholar]
  • 19.Nelson JB. The Behavior of Pox Viruses in the Respiratory Tract : I. The Response of Mice to the Nasal Instillation of Vaccinia Virus. J Exp Med. 1938;68:401–412. doi: 10.1084/jem.68.3.401. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Luker KE, Hutchens M, Schultz T, Pekosz A, Luker GD. Bioluminescence imaging of vaccinia virus: effects of interferon on viral replication and spread. Virology. 2005;341:284–300. doi: 10.1016/j.virol.2005.06.049. [DOI] [PubMed] [Google Scholar]
  • 21.Martinez J, Huang X, Yang Y. Direct TLR2 signaling is critical for NK cell activation and function in response to vaccinia viral infection. PLoS Pathog. 2010;6:e1000811. doi: 10.1371/journal.ppat.1000811. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Kawakami Y, Tomimori Y, Yumoto K, Hasegawa S, Ando T, Tagaya Y, Crotty S, Kawakami T. Inhibition of NK cell activity by IL-17 allows vaccinia virus to induce severe skin lesions in a mouse model of eczema vaccinatum. J Exp Med. 2009;206:1219–1225. doi: 10.1084/jem.20082835. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Lehmann MH, Kastenmuller W, Kandemir JD, Brandt F, Suezer Y, Sutter G. Modified vaccinia virus ankara triggers chemotaxis of monocytes and early respiratory immigration of leukocytes by induction of CCL2 expression. J Virol. 2009;83:2540–2552. doi: 10.1128/JVI.01884-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Sanchez-Puig JM, Sanchez L, Roy G, Blasco R. Susceptibility of different leukocyte cell types to Vaccinia virus infection. Virol J. 2004;1:10. doi: 10.1186/1743-422X-1-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Bonduelle O, Duffy D, Verrier B, Combadiere C, Combadiere B. Cutting edge: Protective effect of CX3CR1+ dendritic cells in a vaccinia virus pulmonary infection model. J Immunol. 2012;188:952–956. doi: 10.4049/jimmunol.1004164. [DOI] [PubMed] [Google Scholar]
  • 26.Salek-Ardakani S, Moutaftsi M, Crotty S, Sette A, Croft M. OX40 drives protective vaccinia virus-specific CD8 T cells. J Immunol. 2008;181:7969–7976. doi: 10.4049/jimmunol.181.11.7969. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Salek-Ardakani S, Flynn R, Arens R, Yagita H, Smith GL, Borst J, Schoenberger SP, Croft M. The TNFR family members OX40 and CD27 link viral virulence to protective T cell vaccines in mice. The Journal of clinical investigation. 2011;121:296–307. doi: 10.1172/JCI42056. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Zhao Y, De Trez C, Flynn R, Ware CF, Croft M, Salek-Ardakani S. The adaptor molecule MyD88 directly promotes CD8 T cell responses to vaccinia virus. J Immunol. 2009;182:6278–6286. doi: 10.4049/jimmunol.0803682. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Moutaftsi M, Bui HH, Peters B, Sidney J, Salek-Ardakani S, Oseroff C, Pasquetto V, Crotty S, Croft M, Lefkowitz EJ, Grey H, Sette A. Vaccinia virus-specific CD4+ T cell responses target a set of antigens largely distinct from those targeted by CD8+ T cell responses. J Immunol. 2007;178:6814–6820. doi: 10.4049/jimmunol.178.11.6814. [DOI] [PubMed] [Google Scholar]
  • 30.Tscharke DC, Karupiah G, Zhou J, Palmore T, Irvine KR, Haeryfar SM, Williams S, Sidney J, Sette A, Bennink JR, Yewdell JW. Identification of poxvirus CD8+ T cell determinants to enable rational design and characterization of smallpox vaccines. J Exp Med. 2005;201:95–104. doi: 10.1084/jem.20041912. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Moutaftsi M, Peters B, Pasquetto V, Tscharke DC, Sidney J, Bui HH, Grey H, Sette A. A consensus epitope prediction approach identifies the breadth of murine T(CD8+)-cell responses to vaccinia virus. Nat Biotechnol. 2006;24:817–819. doi: 10.1038/nbt1215. [DOI] [PubMed] [Google Scholar]
  • 32.Davies DH, McCausland MM, Valdez C, Huynh D, Hernandez JE, Mu Y, Hirst S, Villarreal L, Felgner PL, Crotty S. Vaccinia virus H3L envelope protein is a major target of neutralizing antibodies in humans and elicits protection against lethal challenge in mice. J Virol. 2005;79:11724–11733. doi: 10.1128/JVI.79.18.11724-11733.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Moyron-Quiroz JE, McCausland MM, Kageyama R, Sette A, Crotty S. The smallpox vaccine induces an early neutralizing IgM response. Vaccine. 2009;28:140–147. doi: 10.1016/j.vaccine.2009.09.086. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Salek-Ardakani S, Choi YS, Rafii-El-Idrissi Benhnia M, Flynn R, Arens R, Shoenberger S, Crotty S, Croft M. B cell-specific expression of B7-2 is required for follicular Th cell function in response to vaccinia virus. J Immunol. 2011;186:5294–5303. doi: 10.4049/jimmunol.1100406. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Schatz DG, Ji Y. Recombination centres and the orchestration of V(D)J recombination. Nat Rev Immunol. 2011;11:251–263. doi: 10.1038/nri2941. [DOI] [PubMed] [Google Scholar]
  • 36.Salek-Ardakani S, Arens R, Flynn R, Sette A, Schoenberger SP, Croft M. Preferential use of B7.2 and not B7.1 in priming of vaccinia virus-specific CD8 T cells. J Immunol. 2009;182:2909–2918. doi: 10.4049/jimmunol.0803545. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Dimier J, Ferrier-Rembert A, Pradeau-Aubreton K, Hebben M, Spehner D, Favier AL, Gratier D, Garin D, Crance JM, Drillien R. Deletion of major nonessential genomic regions in the vaccinia virus Lister strain enhances attenuation without altering vaccine efficacy in mice. J Virol. 2011;85:5016–5026. doi: 10.1128/JVI.02359-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Ober BT, Bruhl P, Schmidt M, Wieser V, Gritschenberger W, Coulibaly S, Savidis-Dacho H, Gerencer M, Falkner FG. Immunogenicity and safety of defective vaccinia virus lister: comparison with modified vaccinia virus Ankara. J Virol. 2002;76:7713–7723. doi: 10.1128/JVI.76.15.7713-7723.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Karupiah G, Buller RM, Van Rooijen N, Duarte CJ, Chen J. Different roles for CD4+ and CD8+ T lymphocytes and macrophage subsets in the control of a generalized virus infection. J Virol. 1996;70:8301–8309. doi: 10.1128/jvi.70.12.8301-8309.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Fang M, Sigal LJ. Antibodies and CD8+ T cells are complementary and essential for natural resistance to a highly lethal cytopathic virus. J Immunol. 2005;175:6829–6836. doi: 10.4049/jimmunol.175.10.6829. [DOI] [PubMed] [Google Scholar]

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