Our understanding of memory CD8+ T cells has been largely derived from acute, systemic infection models. However, memory CD8+ T cells generated from mucosal infection exhibit unique properties and, following respiratory infection, are not maintained in the lung long term. To better understand how infection route modifies memory differentiation, we compared murine CD8+ T cell responses to a vesicular stomatitis virus (VSV) challenge generated intranasally (i.n.) or i.v. The i.n. infection resulted in greater peak expansion of VSV-specific CD8+ T cells. However, this numerical advantage was rapidly lost during the contraction phase of the immune response, resulting in memory CD8+ T cell numerical deficiencies when compared with i.v. infection. Interestingly, the antiviral CD8+ T cells generated in response to i.n. VSV exhibited a biased and sustained proportion of early effector cells (CD127loKLRG1lo) akin to the developmental program favored after i.n. influenza infection, suggesting that respiratory infection broadly favors an incomplete memory differentiation program. Correspondingly, i.n. VSV infection resulted in lower CD122 expression and eomesodermin levels by VSV-specific CD8+ T cells, further indicative of an inferior transition to bona fide memory. These results may be due to distinct (CD103+CD11b+) dendritic cell subsets in the i.n. versus i.v. T cell priming environments, which express molecules that regulate T cell signaling and the balance between tolerance and immunity. Therefore, we propose that distinct immunization routes modulate both the quality and quantity of antiviral effector and memory CD8+ T cells in response to an identical pathogen and should be considered in CD8+ T cell–based vaccine design.

Respiratory infections, including influenza, continue to be a major cause of morbidity and mortality globally (1). Current influenza vaccines target protective strain-specific Ab responses, which prevent viral entry into the host cell (2). However, because of mutation and evolution of the targeted influenza hemagglutinin Ags, these vaccines lose efficacy and do not provide long-term protection against infection. Evidence in mouse models and human studies not only implicates CD8+ T cells as requisite for viral clearance but also as protective against heterologous challenge with novel influenza strains (3). To date, no approved vaccine has been developed to specifically generate CD8+ memory T cells (Tmem) despite the fact that the intranasal (i.n.) influenza vaccine FluMist elicits a larger effector CD8+ T cell (Teff) response than inactivated or subunit vaccines in mice and humans (4). Although the contribution of CD8+ Tmem to vaccine efficacy has not been tested comprehensively or longitudinally in the latter population, studies in mice have shown that CD8+ T cell responses are generated from a single i.n. dose of FluMist. However, these cells are lost within 30 d and are not protective against lethal heterogeneous challenge (5). Correspondingly, in June of 2016, the Centers for Disease Control and Prevention’s Advisory Committee on Immunization Practices voted that FluMist should not be using during the 2016–2017 influenza season because of a lack of a protective benefit compared with the inactivated influenza vaccine (3% versus 63%, respectively) (6). Together, these studies suggest that CD8+ Teff generated following respiratory infection or delivery of live-attenuated influenza vaccines either do not form or retain Tmem at protective levels.

Over the last few decades, several laboratories have delineated pathways important in CD8+ Tmem development and defined the attributes and molecules that support robust T cell memory long term. This gold standard for CD8+ T cell memory has been defined in murine models of acute viral infection whereby the pathogen of interest was delivered via the i.v. route (5). However, it is becoming increasingly clear that the formation of Tmem is a dynamic process, with memory potential influenced by a variety of factors, including cytokines (6), the type of APCs involved (7), and the strength and duration of Ag exposure (8), all of which can be differentially affected by inherent properties of both the pathogen and the exposure site. Indeed, our laboratory and others have shown that mucosally derived antiviral CD8+ T cells acquire properties incongruent with memory formation as defined from the systemic infection models (9, 10). For example, by simply altering the route of viral acquisition, from i.v. to i.n., CD8+ Tmem are not only less abundant overall, but those that do develop are maintained independent of the cytokine IL-15 (10). In contrast, IL-15 deficiency results in Tmem decay after systemic infection (1012). Thus, as CD8+ Tmem generation does not appear to be a one model fits all scenario, it is important to understand how and why the respiratory Tmem program is offset from the benchmark Tmem derived in systemic model systems to improve vaccine formulation.

To date, the sole contribution of a respiratory infection on CD8+ Tmem potential has not been determined because many respiratory pathogens, like influenza, do not generate a productive infection outside of this environment. In this study, we used differential routes of inoculation of the same virus, vesicular stomatitis virus (VSV), to determine how the properties of the respiratory environment influence CD8+ Tmem development. VSV is highly suitable to our studies, as the CD8+ T cell response after systemic i.v. infection is well characterized and generates long-lived Tmem. In addition, unlike influenza virus, VSV is transmitted via multiple routes in both mice and its natural bovine host, including the respiratory tract (13). Importantly, we have shown that Tmem derived from i.n. VSV infection numerically and phenotypically resemble anti-influenza CD8+ Tmem (14), validating the use of this model for evaluating how the respiratory environment affects Tmem formation. In this article, we report that although pulmonary VSV infection resulted in higher CD8+ Teff responses early postinfection, this numerical advantage was lost rapidly through the contraction phase of the CD8+ T cell response, resulting in a quantitatively reduced Tmem pool. This loss is facilitated by altered programming whereby i.n. infection favors the development of Teff with delayed CD127 expression and a subsequent reduced ability of these cells to express eomesodermin (Eomes) and CD122 as emerging Tmem. Coincidently, i.n. infection promotes the accumulation of a distinct population of CD11c+CD103+CD11b+CCR2+ monocyte/dendritic cells (moDCs) in the respiratory tract draining lymph nodes (dLNs) during CD8 T cell priming, which may regulate CD8+ Tmem differentiation. In summary, our data suggest that the environments encountered in distinct immunization routes are sufficient to modulate both the quality and quantity of antiviral Teff and CD8+ Tmem in response to an identical pathogen and should be considered in CD8+ T cell–based vaccine design.

C57BL/6 mice were purchased from Charles River Laboratories (Wilmington, MA) through the National Cancer Institute program and bred in house. For VSV infections, age- and sex-matched mice were infected with 104 PFU of VSV-Indiana serotype or VSV-Indiana-OVA (originally obtained from Dr. L. Lefrançois), either i.n. in 50 μl PBS or i.v. through the tail vein in 200 μl PBS. VSV stocks were maintained and isolated by growth in BHK cells, while viral titers were determined by plaque assay. The influenza virus A/HK-x31(x31, H3N2) was generously provided by Dr. S.M. Tompkins (University of Georgia, Athens, GA). For influenza experiments, age- and sex-matched animals were infected i.n. with 103 PFU x31 in 50 μl PBS. For respiratory syncytial virus (RSV) infections, mice were i.n. infected with 2 × 103 PFU of the RSV A2 strain (generously provided by Dr. B. He) delivered in 50 μl PBS. All animal experiments were approved by the Institutional Animal Care and Use Committee of the University of Georgia.

Single-cell suspensions from tissues were obtained as previously described (10). Briefly, cells were isolated from the lung parenchyma after first perfusing the lungs with ∼10 ml PBS/heparin. Lungs were excised, minced, and incubated with 1.25 mM EDTA at 37°C for 30 min followed by a 1 h incubation with 150 U/ml collagenase (Life Technologies, Grand Island, NY). After passage through 40-μM cell strainers, lymphocytes were resuspended in 44% Percoll, underlaid with 67% Percoll, centrifuged, and the cellular interface was collected. Lymph nodes (LNs) and splenic tissues were mechanically disrupted, then passed through a cell strainer. Erythrocytes were depleted from the spleen samples using Tris-buffered ammonium chloride. Blood samples were obtained either by retro-orbital eye bleeding or by cardiac puncture at time of sacrifice. Erythrocytes were depleted from blood samples by two serial treatments (10 min each at 37°C) with Tris-buffered ammonium chloride. Cell numbers were determined using a Z2 Coulter Particle Counter (Beckman Coulter, Fullerton, CA). For lymphocyte numbers obtained from blood samples, 250 μl blood was counted and expressed as lymphocytes per milliliter.

The VSV nucleoprotein (N) MHC class I (MHC I) [H-2Kb/RGYVYQGL] tetramer and the influenza nuclear protein (NP) MHC I [H-2Db/ASNENMETM] tetramer (conjugated to APC) were obtained from the National Institutes of Health Tetramer Core Facility (Emory University, Atlanta, GA). Staining was carried out at room temperature for 1 h in conjunction with other surface staining. Abs (clones) used for staining include the following: PE-conjugated αCD127 (A7R34), αEomes (Dan11mag), or αCD103 (2E7); FITC-conjugated αCD11b (M1/70) or αCD44 (IM7); PerCP-Cy5.5–conjugated αCD8a (53-6.7), αCD19 (1D3), αNK1.1 (PK136), αCD3e (145-2C11), or αT-bet (eBio4B10); PE-Cy7–conjugated αCD44 (IM7), αCD11c (N418), αKLRG-1 (2F1), αPD-L1 (MIH5), or αCD86 (GLI); violetFluor 450–conjugated αCD8a (2.43) or αCD11c (N418); APC-conjugated αCD80 (16-10A1) or αSIi.n.FEKL-H-2Kb (25-D1.16); APC-eFluor 780–conjugated αCD62L (MEL-14) or αMHC II (M5/114.15.2); and APC-eFluor 647–conjugated phospho-S6 ribosomal protein (Ser235/236) (D57.2.2E) (Cell Signaling Technology, Danvers, MA). All other Abs were purchased from eBioscience, Tonbo Biosciences (both San Diego, CA), or BD Biosciences (San Jose, CA). When tetramer was not used, cells were surface stained for 20 min at 4°C. Tissue-resident memory cells were identified by intravascular staining, in which mice were injected i.v. with 3 μg of FITC-conjugated αCD45.2 (104) in 200 μl PBS 3 min prior to sacrifice; cells were isolated and subsequently stained ex vivo. For analysis of intracellular proteins (T-bet and Eomes), cells were fixed, permeabilized using eBioscience Fix & Perm, and intracellularly stained according to the manufacturer’s instructions (eBioscience). Following staining, cells were fixed in 2% paraformaldehyde, flow cytometric analysis was performed using a BD LSR II, and data were acquired with FACSDiva software (BD Biosciences). For sorting experiments, FACS was performed on a Beckman Coulter MoFlo (XDP) (Indianapolis, IN). For analysis of p-S6, LNs and spleens were removed from mice and immediately disassociated as previously described in ice-cold methanol. Cells were then washed twice in BD Perm/Wash (San Jose, CA) and stained in BD Perm/Wash for 45 min at room temperature. After washing, cells were resuspended in BD Perm/Wash, and flow cytometric analysis was immediately performed. Data were analyzed using FlowJo software version 9.6.2 from Tree Star. In all analyses, cells were first gated on single cells and, where indicated, followed by a lymphocyte gate as determined by forward and side scatter. Subsequent gating strategies are noted in the figure legends.

Statistical analysis was carried out using GraphPad Prism version 5 or 6 using the specific analysis indicated in the individual figure legends. Significance was determined when the p value was <0.05.

Respiratory infections produce CD8+ Tmem responses, which are limited both in number and lifespan compared with systemic infections (10, 14). This is believed to be due, at least in part, to the heightened level of immune regulation at these surfaces, which limit inadvertent immunopathology. However, the mechanisms linked to the development of substandard Tmem have been poorly understood, mainly because of inequitable comparisons made using two different pathogens with distinct tissue tropisms and inflammatory signals. Therefore, we sought to modify route of infection alone to address how the respiratory environment affects the development CD8+ Tmem.

To first validate that the respiratory route of infection modifies the development of CD8+ Tmem, mice were infected either i.n. or i.v. with a sublethal dose of VSV. This virus produces a replicating viral infection via multiple routes (15) and thus provides a model to compare the emergence of CD8+ Tmem arising from systemic and respiratory infection. Ag-specific CD8+ T cell responses were assessed at 35 d postinfection (dpi) using MHC I tetramers against the immunodominant epitope of the VSV-N (N-tet+ cells). As previously reported (10), i.n. delivery of VSV resulted in a lower frequency of VSV N-tet+ CD8+ Tmem when compared with systemic infection using the same viral dose (data not shown). Moreover, i.n. infection also resulted in numerically deficient CD8+ Tmem in sites proximal (lung) and distal (spleen) to the respiratory tract (Fig. 1A). Thus, in a system where an identical pathogen is used, i.n. infection resulted in a quantitatively reduced Tmem.

FIGURE 1.

Respiratory infection results in quantitatively deficient CD8 Tmem pools despite higher effector responses. (A) Number of VSV N-tet+ CD8 T cells assessed at 35 dpi in the lung and spleen postinfection with 104 PFU VSV via the i.v. (black bar) or i.n. (open bar) route. (B) Mice were infected i.n. or i.v. with 104 PFU VSV, and CD8+ T cell responses were monitored in the blood 5–9 dpi. Data are displayed as frequency of N-tet+ CD8 T cells (n = 3 mice per group and data represent two independent experiments). (C and D) Mice were infected with 104 PFU VSV by either route, and lymphocytes were assessed for N-tet reactivity at 8 dpi in the indicated tissues and displayed as total frequency of CD8+ T cells (C) or quantified (D) (n = 10 mice per group and data are representative of three independent experiments). Significance between groups was assessed using a two-tailed Student t test (*p ≤ 0.05, **p < 0.01, ***p < 0.001).

FIGURE 1.

Respiratory infection results in quantitatively deficient CD8 Tmem pools despite higher effector responses. (A) Number of VSV N-tet+ CD8 T cells assessed at 35 dpi in the lung and spleen postinfection with 104 PFU VSV via the i.v. (black bar) or i.n. (open bar) route. (B) Mice were infected i.n. or i.v. with 104 PFU VSV, and CD8+ T cell responses were monitored in the blood 5–9 dpi. Data are displayed as frequency of N-tet+ CD8 T cells (n = 3 mice per group and data represent two independent experiments). (C and D) Mice were infected with 104 PFU VSV by either route, and lymphocytes were assessed for N-tet reactivity at 8 dpi in the indicated tissues and displayed as total frequency of CD8+ T cells (C) or quantified (D) (n = 10 mice per group and data are representative of three independent experiments). Significance between groups was assessed using a two-tailed Student t test (*p ≤ 0.05, **p < 0.01, ***p < 0.001).

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The size of the CD8+ Tmem pool is often correlated with the overall size of the corresponding effector population (16). Given that the lung has multiple innate barriers and may promote immune tolerance (17), we speculated that the CD8+ Teff response may also be suppressed following respiratory infection, resulting in the observed Tmem reduction. To test this hypothesis, we assessed the CD8+ Teff responses to i.v. and i.n. VSV infection. Surprisingly, the quantitative deficiency of i.n. derived N-tet+ CD8+ Tmem was in direct contrast to the peak Ag-specific CD8+ T cell response, in which the i.n. derived CD8+ Teff were more prominent. i.n. VSV infection produced greater CD8+ T cell activation in the blood, as assessed by CD44 expression (data not shown) and increased frequencies of circulating VSV-specific CD8+ T cells at the peak of the response (∼7 dpi; Fig. 1B). Moreover, a higher Ag-specific CD8+ Teff response was also observed in the spleen, lung, and lung draining mediastinal LN (MdLN) 8 dpi (Fig. 1C, 1D). Together, these data demonstrate that the reduced CD8 Tmem responses observed following i.n. infection versus i.v. infection are not merely the result of numerically reduced Teff responses.

i.n. VSV infection results in heightened CD8+ Teff responses yet quantitatively decreased memory pools (Fig. 1). This indicates that respiratory-derived CD8+ T cells, compared with their systemically derived counterparts, are either unable to transition to Tmem or are not maintained. To address the former of these scenarios, we monitored Ag-specific CD8+ T cell responses throughout the contraction phase of immune response. Comparison of the kinetics of CD8+ T cell contraction between i.n. and i.v. VSV-infected mice indicated that Ag-specific CD8+ T cells that develop following i.n. infection are lost more rapidly during the contraction phase (8–15 dpi) in all analyzed tissues (Fig. 2). The average loss of Teff during this period was 1.7-fold (in the MdLN), 3-fold (in the spleen), and 3.5-fold (in the lung) greater in i.n. versus i.v. infected animals (Fig. 2C). Although it has been known for some time that respiratory-derived CD8+ T cells are not maintained in the lung long term, this loss is thought to occur months following infection (14). Our data suggest that the instability of respiratory Tmem may not be solely due to simple attrition but is more complex, involving a programmatic modification of respondent CD8+ T cells much earlier in the Teff response.

FIGURE 2.

Respiratory-derived CD8+ T cell responses contract more rapidly, and to a greater extent, than those derived from systemic infection. (A) Representative flow plots of splenic lymphocytes isolated from mice infected with 104 VSV i.n. or i.v. and sacrificed at 8, 12, and 15 dpi. Plots show populations previously gated on CD8+ lymphocytes. Frequencies (B) or total cell number (C) of VSV-N-tet+ CD8+ T cells isolated from the indicated issues 8, 12, and 15 dpi (n = 5–10 mice per group and data are representative of three independent experiments). Significance between groups was assessed using a two-tailed Student t test (*p ≤ 0.05, **p < 0.01, ***p < 0.001).

FIGURE 2.

Respiratory-derived CD8+ T cell responses contract more rapidly, and to a greater extent, than those derived from systemic infection. (A) Representative flow plots of splenic lymphocytes isolated from mice infected with 104 VSV i.n. or i.v. and sacrificed at 8, 12, and 15 dpi. Plots show populations previously gated on CD8+ lymphocytes. Frequencies (B) or total cell number (C) of VSV-N-tet+ CD8+ T cells isolated from the indicated issues 8, 12, and 15 dpi (n = 5–10 mice per group and data are representative of three independent experiments). Significance between groups was assessed using a two-tailed Student t test (*p ≤ 0.05, **p < 0.01, ***p < 0.001).

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One of the key transcription factors identified in promoting memory cell development is Eomes (18). Eomes-deficient CD8+ T cells undergo primary clonal expansion but are defective in long-term survival (19). As respiratory-derived N-tet+CD8+ T cells expanded normally, yet developed substandard memory responses, we tested the hypothesis that N-tet+ CD8+ Teff derived from a respiratory infection fail to initiate Eomes expression and the subsequent memory cell program. As early as 6 dpi, i.n. derived N-tet+CD8+ T cells isolated from the lung and spleen expressed less Eomes than those Ag-specific cells derived from systemic infection (Fig. 3B). By 35 dpi, this deficiency in Eomes expression was exacerbated and reduced to 25–50% of Tmem derived following systemic infection (Fig. 3A, 3C). These data indicate that a known initiator of memory cell programming was considerably less in Tmems, which are also numerically reduced after respiratory infection.

FIGURE 3.

Respiratory infection results in N-tet+ CD8+ T cells that have reduced Eomes and CD122 expression. (A) Representative flow of Eomes and CD122 expression of CD8 T cells isolated from the spleen 35 d following i.v. or i.n. VSV infection. Frequency of Eomes+ N-tet+ CD8+ T cells following i.v. (black bar) or i.n. (white bar) VSV infection in the lung or spleen was assessed by intracellular staining at 6 (B) and 35 (C) dpi (n = 3 mice per group; data are representative of three independent experiments). Frequency (D) and number (E) of CD122hi of N-tet+CD8+ T cells following i.v. (black bar) or i.n. (white bar) VSV infection in the lung or spleen was assessed at 35 dpi (n = 3 mice per group; data are representative of three independent experiments). (F) Average proportion of TCM (CD62Lhi, black), TEM (CD62Llo, gray), and TRM (CD62Llo, intravascular CD45, white) populations in the lung and of TCM (CD62Lhi, black) and TEM (CD62Llo, gray) in the spleen 35 d after i.v. or i.n. VSV infection is depicted. Significance between groups was assessed using a two-tailed Student t test (*p ≤ 0.05, **p < 0.01, ***p < 0.001).

FIGURE 3.

Respiratory infection results in N-tet+ CD8+ T cells that have reduced Eomes and CD122 expression. (A) Representative flow of Eomes and CD122 expression of CD8 T cells isolated from the spleen 35 d following i.v. or i.n. VSV infection. Frequency of Eomes+ N-tet+ CD8+ T cells following i.v. (black bar) or i.n. (white bar) VSV infection in the lung or spleen was assessed by intracellular staining at 6 (B) and 35 (C) dpi (n = 3 mice per group; data are representative of three independent experiments). Frequency (D) and number (E) of CD122hi of N-tet+CD8+ T cells following i.v. (black bar) or i.n. (white bar) VSV infection in the lung or spleen was assessed at 35 dpi (n = 3 mice per group; data are representative of three independent experiments). (F) Average proportion of TCM (CD62Lhi, black), TEM (CD62Llo, gray), and TRM (CD62Llo, intravascular CD45, white) populations in the lung and of TCM (CD62Lhi, black) and TEM (CD62Llo, gray) in the spleen 35 d after i.v. or i.n. VSV infection is depicted. Significance between groups was assessed using a two-tailed Student t test (*p ≤ 0.05, **p < 0.01, ***p < 0.001).

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CD122 is a downstream target of Eomes (20), expressed at high levels on Tmem (12), and confers IL-15 reactivity, which is necessary for the maintenance of Tmem following systemic infection (11). In contrast, IL-15 is dispensable for the development of memory following respiratory infections (10). Because Eomes expression was higher on N-tet+ CD8+ T cells following i.v. infection compared with i.n. infection, we assessed whether the i.v. memory cell program is responsible for the differences in IL-15 dependency between infection routes. Indeed, the proportion of N-tet+ CD8+ Tmems that express CD122 following i.n. infection was reduced in both the spleen and the lung, compared with i.v. VSV infection (Fig. 3D). This loss in CD122hi cells is even more apparent when overall numbers of CD122hi cells were quantified (Fig. 3E). Thus, the reduction in Eomes and CD122 expression along with the more significant contraction of CD8+ Teff indicate that early signals following i.n. infection may suppress the transition of CD8+ Teff to bona fide long-lived, IL-15–dependent Tmem.

We next wished to determine whether the reduction of Eomes and CD122 expression in Ag-specific Tmem was simply due to differences in the development of specific Tmem subsets postinfection by the two routes. Therefore, we assessed the contribution of central Tmem (TCM; CD62Lhi) and effector Tmem (TEM; CD62Llo) (21) to the overall Tmem pool in the lung and spleen 35 dpi with VSV delivered i.n. and i.v. We additionally examined the proportion of tissue-resident Tmem (TRM; CD62Llo and negative for intravascular staining; see Ref. 22) in the lung. At 35 d after i.v. VSV infection, TEM were found at a 28% greater number in the lung compared with i.n. infected animals, whereas TCM and TRM were represented more in the lung following i.n. (versus i.v.) VSV infection (Fig. 3F). Because TEM express the highest level of Eomes compared with the other Tmem subsets (23), more Eomes expression in the lung following i.v. VSV (Fig. 3) infection could be due to increased representation of this population within the tissue. However, the Tmem pools generated in the spleen were identical irrespective of infection route (Fig. 3F), whereas Eomes and CD122 expression was globally reduced in splenic Tmem after i.n. infection (Fig. 3). Together, these data indicate that the early memory cell programming and subsequent development of the Tmem pools are distinct following respiratory VSV infection, the former favoring a global reduction in Tmem potential.

At the proliferative peak of the antiviral CD8+ T cell response, Tmem can be identified within the effector cell pool using the IL-7 receptor α-chain (CD127) and killer cell lectin-like receptor G1 (KLRG1) (2426). These markers have been used extensively in CD8+ Tmem studies and can predict which cells will survive the contraction phase, largely in the context of systemic viral infections (25, 26). Memory precursor effector cells (MPECs) are CD127hiKLRG1lo and will dominate the Ag-specific CD8 T cell pool over time based on the enhanced survival conferred by IL-7 (24, 27). These MPECs differentiate from ancestral clones, referred to as early effector cells (EECs; CD127loKLRG1lo). EECs have the greatest developmental plasticity, with the potential to develop into any of the other phenotypes (28), but are generally thought not to persist into memory because of their lack of CD127 expression. Short-lived effector cells (SLECs) (CD127loKLRG1hi) constitute the majority of the early antiviral CD8+ T cell responses during systemic infection (26) yet are terminally differentiated and lost during CD8+ T cell contraction (24). Because respiratory-derived Ag-specific CD8+ T cells become activated but do not appear to phenotypically or quantitatively match their systemically derived counterparts (Figs. 1, 3), we sought to determine whether memory cell differentiation was stalled following i.n. infection, aborting the development of MPECs and affecting Tmem development.

To this end, we monitored the emergence and persistence of the aforementioned effector CD8+ T cell subsets in the blood following i.n. and i.v. VSV infection using the CD127 and KLRG1 markers. Early after i.v. infection (Fig. 4A), both SLEC and EEC N-tet+ cells predominated until ∼11 dpi, when MPECs surpassed these subsets as the dominant phenotype. The survival advantage of these MPECs is very apparent by 50 dpi, when this subset prevails. In contrast, N-tet+ Teff derived from an i.n. infection (Fig. 4B) harbor predominately EECs with sustained persistence compared with i.v. infection. Moreover, MPECs do not emerge as the dominant subset until ∼15 dpi. Direct comparison of the composition of the effector pools between the two routes of infection at 12 dpi support the conclusion that the prolonged frequency of EECs observed after i.n. infection is largely at the expense of the generation of SLECs and MPECs (Fig. 4C). Furthermore, the pattern of effector CD8+ T cell distribution following respiratory VSV infection is similar to that observed following influenza infection, in which EECs are observed for a sustained period of time in the blood (Fig. 4D).

FIGURE 4.

Respiratory infection results in skewed populations of effector cells based on CD127 and KLRG1 expression. Ag-specific CD8+ T cell responses were measured in the blood over time; populations were assessed for their expression of CD127 and KLRG1 and identified as previously defined effector or memory cell subsets using these markers. Phenotypes expressed after i.v. (A) or i.n. (B) VSV infection over time were compared between the two routes specifically at 12 dpi (C). (D) The frequency of NP-specific memory CD8 T cell phenotypes in the blood over time following infection with 103 PFU of influenza A/HKx31 (n ≥ 3 mice per group per time point and repeated twice).

FIGURE 4.

Respiratory infection results in skewed populations of effector cells based on CD127 and KLRG1 expression. Ag-specific CD8+ T cell responses were measured in the blood over time; populations were assessed for their expression of CD127 and KLRG1 and identified as previously defined effector or memory cell subsets using these markers. Phenotypes expressed after i.v. (A) or i.n. (B) VSV infection over time were compared between the two routes specifically at 12 dpi (C). (D) The frequency of NP-specific memory CD8 T cell phenotypes in the blood over time following infection with 103 PFU of influenza A/HKx31 (n ≥ 3 mice per group per time point and repeated twice).

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We next wanted to confirm that the antiviral Teff and developing Tmem in the tissues were equally impacted by route of infection. Thus, we monitored the kinetics of the appearance and contribution of the various Teff subsets to the overall CD8+ T cell pool in the lung, spleen, and MdLN at 8, 12, and 15 dpi (Fig. 5). At 8 dpi, SLECs dominate the i.v. derived N-tet+ CD8+ T cell response, totaling up to 50% of the overall Ag-specific Teff pool (Fig. 5A, 5B). In contrast, after i.n. infection, ∼50% or more of the VSV-specific CD8 T cells are EECs in all tissues at 8 and 12 dpi and numerically total twice as many compared with i.v. infection (Fig. 5C). Although EECs are lost during contraction in both i.v. and i.n. VSV infected animals (Fig. 5C), the fold loss is greater after i.n. infection (Fig. 5D). These data, combined with the earlier emergence of MPECs after i.v. infection, have consequences for Tmem development, as MPECs have a survival advantage because of expression of CD127 (24). The inability of the enhanced numbers of EECs to transition to MPECs as efficiently after i.n. infection could mechanistically explain the steep and persistent decline of CD8+ T cells during contraction, resulting in numerically reduced CD8+ Tmem.

FIGURE 5.

Respiratory infection results in increased and sustained proportions of EEC, which impact MPEC numbers. (A) Representative flow plots of splenic lymphocytes following i.v. or i.n. infection with 104 PFU VSV at 8, 12, and 15 dpi. Data were previously gated on CD8+ lymphocytes, then VSV N-tet+ cells. (B) N-tet+ responses in the indicated tissues following i.v. (top) or i.n. (bottom) infection. Size of bar indicates the total frequency of VSV N-specific CD8+ T cells, and the different shading within indicates the proportion of these cells of the indicated phenotype. Statistical significance is noted for an increased frequency of EECs (*), SLECs (†), and MPECs (#) between the given route and tissue as indicated. (C) Total numbers of N-tet+ cells with an EEC (top) or MPEC (bottom) phenotype in i.v. (black bar) or i.n. (open bar) infected animals at 8, 12, and 15 dpi. (D) Fold change in N-tet+ EEC (top) and MPEC (bottom) numbers was calculated from 8 to 15 dpi in i.v. (black bar) and i.n. (open bar) infected animals (data are shown as log2 and n = 5–10 mice per group). Data is representative of three independent experiments. Significance between groups was assessed using a two-tailed Student t test (*p ≤ 0.05, **p < 0.01, ***p < 0.001).

FIGURE 5.

Respiratory infection results in increased and sustained proportions of EEC, which impact MPEC numbers. (A) Representative flow plots of splenic lymphocytes following i.v. or i.n. infection with 104 PFU VSV at 8, 12, and 15 dpi. Data were previously gated on CD8+ lymphocytes, then VSV N-tet+ cells. (B) N-tet+ responses in the indicated tissues following i.v. (top) or i.n. (bottom) infection. Size of bar indicates the total frequency of VSV N-specific CD8+ T cells, and the different shading within indicates the proportion of these cells of the indicated phenotype. Statistical significance is noted for an increased frequency of EECs (*), SLECs (†), and MPECs (#) between the given route and tissue as indicated. (C) Total numbers of N-tet+ cells with an EEC (top) or MPEC (bottom) phenotype in i.v. (black bar) or i.n. (open bar) infected animals at 8, 12, and 15 dpi. (D) Fold change in N-tet+ EEC (top) and MPEC (bottom) numbers was calculated from 8 to 15 dpi in i.v. (black bar) and i.n. (open bar) infected animals (data are shown as log2 and n = 5–10 mice per group). Data is representative of three independent experiments. Significance between groups was assessed using a two-tailed Student t test (*p ≤ 0.05, **p < 0.01, ***p < 0.001).

Close modal

CD8+ Tmem population size is ultimately governed by the rate of cell death and conversion between phenotypes. We would therefore expect that as EECs (and SLECs) numerically decrease through contraction, MPECs would remain numerically stable or increase because of EEC → MPEC conversion. Any accelerated loss of EECs or stalled conversion to MPEC would impact the resultant MPEC/Tmem pool. Interestingly, although the overall numbers of MPECs remain relatively stable in the spleen or even increase in the lung through contraction after i.v. VSV infection, there is a numerical attrition of MPECs in all tissues monitored after i.n. VSV infection, with an enhanced fold loss compared with those MPECs generated after i.v. infection (Fig. 5C, 5D). Importantly, the number of MPECs recovered from the lung 15 dpi is significantly reduced after i.n. infection. Together, these data suggest that inherent defects in EEC programming lead to stalled CD127 expression, which is requisite for survival and adequate seeding of the respiratory MPEC pool.

Our data thus far indicate that early events following respiratory (versus systemic) infection likely result in a different developmental program, ultimately resulting in numerically deficient CD8+ Tmem development. By using an identical virus (VSV) in our studies, we have eliminated pathogen-specific pattern recognition receptor bias and the events underlying disparate pattern recognition receptor signaling pathways. Furthermore, differences in Ag availability were not observed between either route of infection at a time coordinate with Tmem programming (Supplemental Fig. 1), suggesting that other extrinsic environmental factors are likely responsible for modified Tmem programming.

Two populations of migratory CD11c+ dendritic cells (DCs) exist in the lung: CD103+ (CD11b) and CD11b+ (CD103) respiratory DCs (RDCs). Upon activation, both CD103+ and CD11b+ RDCs migrate to the lung dLNs (7, 29). CD103+ RDC and LN-resident DCs (CD8α+) are believed to primarily participate in CD8 T cell priming after influenza infection both indirectly and via cross-presentation, respectively, whereas the role of CD11b+ RDC in this process is controversial (3033). Nonetheless, CD11b+ RDC are a source of inflammatory chemokines and thus are likely to impact T cell activation and programming (34, 35). CD103+ RDCs can drive effector function in activated CD8+ T cells, increasing Teff transcriptional profiles and migratory capabilities while diverting cells from a central memory fate (7). Therefore, we hypothesized that the preferential activation by respiratory CD103+ RDCs following i.n. infection may inherently favor the development of CD8 Teff, resulting in a delayed and substandard transition to memory.

To first assess if CD103+ RDCs were activated and migrating to LNs following respiratory VSV infection, DC populations were identified in LNs at 3 dpi. DCs were defined as lineage-negative (Lin) cells (CD3, CD19, NK.1.1) expressing CD11c and MHC class II (MHC II) (Fig. 6A). As previously reported, following influenza infection (30), mice infected i.n. with VSV harbored high numbers of CD11c+ DCs in their lung dLNs (MdLN and cervical LN [cLN]), with a significant proportion of these cells expressing CD103 (Fig. 6A, 6B). Interestingly, three quarters of the CD103+ DCs coexpressed CD11b and had a reduced expression of MHC II compared with the other DC populations (Table I). These cells were not found either in analogous LNs after i.v. infection or in the lung after VSV infection by either route (Fig. 6C, 6G, data not shown). Importantly, these cells were also identified following respiratory infection with influenza (Fig. 6C) and RSV (Fig. 6C). The morphological (cytoplasm/nucleus ratio, kidney-shaped nucleus) (Fig. 6D) and phenotypical (CCR2+GR1+MHC class IIlo) attributes of these cells (Table I), as well as their transient appearance in the LNs after i.n. VSV infection (Fig. 6G), suggest these cells are derived from circulating monocyte precursors (36). From this point on, we will refer to these cells as moDCs.

FIGURE 6.

Respiratory infection results in the accumulation of distinct DC populations at sites of T cell priming. (A) Representative flow samples showing the initial gating of lineage (CD3,CD19, NK1.1 cells) DCs based on CD11c and MHC II (left) followed by CD103 (right). (B) Total numbers of DCs (height of bar) and CD103+ DCs (gray portion of bar) isolated from indicated tissues. The color of the asterisks indicate significant differences between the corresponding populations within i.v. and i.n. VSV infected animals. (C) Representative contribution of classical CD103+CD11b and CD103CD11b+ RDC and CD103+CD11b+ moDC in the cLN postinfection with the indicated pathogen. (D) The cytospin image of Giemsa-stained CD11c+MHC II+ cells FACS sorted based on CD11b and CD103 expression is shown (original magnification ×400). (E) Representative OVA-H2Kb staining of indicated DC subsets isolated from the cLN 3 d after i.n. VSV-OVA (white histogram) or i.n. VSV (gray histogram) infection compared with the fluorescence minus one control for each subset (black histogram). (F) Average adjusted mean fluorescence intensity of OVA-H2Kb by indicated DC subsets. Adjusted mean fluorescence intensity calculated as the average OVA-H2Kb expression (VSV-OVA) with average background OVA-H2Kb expression (VSV) subtracted out; data pooled from three independent experiments. (G) The lung tissue and pooled cLN and mLN were isolated on the indicated dpi with i.n. VSV, and the frequency and total number of CD103+CD11b+ moDCs was determined by flow cytometry. (H) cLNs, iLNs, and spleens were isolated at the indicated hour postinfection with i.v. (black bar) or i.n. (open bar) VSV, and the levels of S6 phosphorylation at serine 235 and 236 in CD44hi CD8+ T cells was determined by flow cytometry. Significance was determined using a two-tailed Student t test to test between i.v. and i.n. infected mice (n = 3 mice per group, representative of two independent experiments; *p ≤ 0.05, **p < 0.01, ***p < 0.001).

FIGURE 6.

Respiratory infection results in the accumulation of distinct DC populations at sites of T cell priming. (A) Representative flow samples showing the initial gating of lineage (CD3,CD19, NK1.1 cells) DCs based on CD11c and MHC II (left) followed by CD103 (right). (B) Total numbers of DCs (height of bar) and CD103+ DCs (gray portion of bar) isolated from indicated tissues. The color of the asterisks indicate significant differences between the corresponding populations within i.v. and i.n. VSV infected animals. (C) Representative contribution of classical CD103+CD11b and CD103CD11b+ RDC and CD103+CD11b+ moDC in the cLN postinfection with the indicated pathogen. (D) The cytospin image of Giemsa-stained CD11c+MHC II+ cells FACS sorted based on CD11b and CD103 expression is shown (original magnification ×400). (E) Representative OVA-H2Kb staining of indicated DC subsets isolated from the cLN 3 d after i.n. VSV-OVA (white histogram) or i.n. VSV (gray histogram) infection compared with the fluorescence minus one control for each subset (black histogram). (F) Average adjusted mean fluorescence intensity of OVA-H2Kb by indicated DC subsets. Adjusted mean fluorescence intensity calculated as the average OVA-H2Kb expression (VSV-OVA) with average background OVA-H2Kb expression (VSV) subtracted out; data pooled from three independent experiments. (G) The lung tissue and pooled cLN and mLN were isolated on the indicated dpi with i.n. VSV, and the frequency and total number of CD103+CD11b+ moDCs was determined by flow cytometry. (H) cLNs, iLNs, and spleens were isolated at the indicated hour postinfection with i.v. (black bar) or i.n. (open bar) VSV, and the levels of S6 phosphorylation at serine 235 and 236 in CD44hi CD8+ T cells was determined by flow cytometry. Significance was determined using a two-tailed Student t test to test between i.v. and i.n. infected mice (n = 3 mice per group, representative of two independent experiments; *p ≤ 0.05, **p < 0.01, ***p < 0.001).

Close modal
Table I.
Phenotypic characterization of RDC subsets
CD11b+CD103CD11bCD103+CD11b+CD103+
CCR2 ++ − +++ 
Gr1 +++ − ++ 
MHC II ++ +++ 
MHC I ++ ++ +++ 
B7-1 ++ ++ +++ 
B7-2 ++ +++ 
PD-L1 ++ ++ +++ 
Ag presentation +++ 
CD11b+CD103CD11bCD103+CD11b+CD103+
CCR2 ++ − +++ 
Gr1 +++ − ++ 
MHC II ++ +++ 
MHC I ++ ++ +++ 
B7-1 ++ ++ +++ 
B7-2 ++ +++ 
PD-L1 ++ ++ +++ 
Ag presentation +++ 

Phenotypic analysis of DC populations in the cLN 3 d following i.n. VSV infection. Isolated Lin (CD3e, CD19, NK1.1), CD11c+MHCII+ cells were gated into the three DC lineages based on CDllb and CD103 expression. Ag presentation was determined via reactivity to a fluorescently labeled SIi.n.FEKL-H-2Kb Ab. The phenotype of each of the subsets was determined based on mean fluorescence intensity of individual cell surface markers relative to naive control animals (CD11c+MHCII+ cells). Expression was determined as follows: −, negative, +, low, ++, intermediate, +++, high (n = 4 mice per group and data are representative of two separate experiments).

The moDCs were present in the dLNs only after respiratory infection, coordinate with T cell priming, and the skewed development Teff. To begin to decipher how these moDCs could be influencing CD8+ Tmem development, we sought to determine whether these cells compromise Teff–Tmem programming via modified Ag presentation or costimulation and/or inhibition. Thus, we infected animals i.n. with VSV or VSV-OVA and measured the surface expression of the OVA epitope SIi.n.FEKL presented by H-2Kb. Whereas CD103+CD11b cells expressed the highest level of Ag/MHC I complexes (Fig. 6E, 6F) and the costimulatory molecule B7-2 (Table I), the moDCs expressed low levels of Ag (Fig. 6E), low levels of B7-2, high levels of B7-1, and high levels of the inhibitory ligand programmed death ligand 1 (PD-L1) (Table I). Activated CD8+ T cells express PD-1 and CTLA-4 (37, 38). Engagement of PD-1 by PD-L1 or preferential signaling of CTLA-4 in the context of B7-1/2 expression can affect signaling via the PI3K/Akt pathway (39), which is known to regulate Tmem programming via mTOR (40). Early in priming (48 h), we observed an increase in the phosphorylation of ribosomal protein S6 (p-S6), which would occur via either the MAPK- (41) or mTOR-dependent pathway (42, 43), specifically in the respiratory dLN after i.n. infection (Fig. 6H), suggesting hyperactivation of Teff after respiratory infection.

Overall, the data presented in this paper show that there is a common developmental pathway for CD8+ Tmem generated following respiratory infection, resulting in the prolonged survival of EEC with concomitant reduced conversion to MPEC. The overall consequence of this is the reduced size and durability of the memory pool (Supplemental Fig. 2). To our knowledge, our data demonstrate for the first time that i.n. infection preferentially recruits moDCs to the dLN, which expressed PD-L1 and increased levels of B7-1 over traditional RDCs (Table I). This inhibitory phenotype, coupled with rapid p-S6 suppression within the dLN, suggests an intrinsic modulation of respiratory CD8+ T cell responses. In conclusion, these results offer emerging insight as to why CD8+ Tmem development following i.n. vaccines and natural respiratory infections is neither as robust nor long-lasting as those generated in systemic models of infection.

Systemic infection models have dominated the field of CD8+ Tmem development since its infancy (44). This is likely because of the ability to consistently produce large, traceable pools of Tmem using acute viral infections delivered by this route (45). However, neither vaccines nor most naturally transmitted infections are acquired through an i.v. route. Therefore, the possibility arises that the systemic models we use to study CD8+ Tmem development may not fully encapsulate the properties of CD8+ Tmem derived from physiologically relevant infection routes. Indeed, Mueller et al. (9) showed that infection via a mucosal route (i.n. influenza infection) results in qualitatively deficient CD8+ Tmem with reduced protective capacity compared with those acquired via a systemic route (i.v. lymphocytic choriomeningitis virus). How the respiratory environment regulates this response was unclear in these studies, as distinct inflammatory and cytokine profiles elicited by the divergent priming viruses could not be eliminated as a confounding variable. By using VSV infection in our studies, we directly tested the impact of the respiratory environment on the developmental pathways responsible for antiviral CD8+ T cell development.

Respiratory VSV infection recapitulates many features of antiviral CD8+ T cells derived from a native influenza infection, including the generation of a robust effector cell pool yet reduced frequencies of Tmem (Fig. 1). The rate of attrition was most pronounced immediately after the peak number of CD8+ Teff were detected in all tissues (Fig. 2), which led us to speculate that respiratory infection may differentially program antiviral CD8+ T cells in a way that favors short- over long-term protection. Indeed, EECs, which maintain the plasticity to differentiate into either MPECs or SLECs (28), were enriched and selectively maintained within the Ag-specific CD8+ T cell pool after respiratory VSV infection, where they constituted approximately one-third of the Ag-specific response out to 15 dpi (Figs. 4, 5). The sustained EEC phenotype is also observed following influenza infection (Fig. 4D), providing evidence that infection via the respiratory route may uniformly contribute to a developmental stall, preventing full transition to memory. Indeed, the sustained population of EECs generated following i.n. infection came at the expense of generating MPECs (Fig. 4C, 4D) and correlated with the timing of the greatest loss of the i.n. VSV-specific CD8+ T cells (Fig. 2). Prior to our study, it was unclear whether EEC maintained beyond contraction could convert to MPECs. Our data suggest that they do not (Fig. 5D); however, this remains a possibility. Nonetheless, the delayed appearance of CD127+ (MPECs) after respiratory infection likely accounted for the greater loss of antiviral CD8+ T cells during contraction.

Limitations on Tmem in the respiratory tract makes teleological sense given the plethora of respiratory assaults an individual encounters over its lifetime and the limited space to harbor accumulating Tmem without compromising tissue function. However, reductions in Tmem were also observed in the spleen (Fig. 1A, 1B), suggesting Tmem development was not selectively suppressed in the respiratory tract when exposure was by the respiratory route. Taking this into account, it is quite possible that respiratory-derived restrictions may directly or indirectly lead to the selective development of certain subsets of Tmem. Our data suggest this is true, at least in part, as the lung parenchyma of i.n. infected mice harbor more TRM and TCM but less TEM than i.v. infected animals, although pools are equivalent in the spleen after VSV infection by either route (Fig. 3F). However, respiratory VSV infection resulted in lower expression of Eomes (as well as the Eomes-regulated IL-15 receptor), CD122 by both splenic and lung Tmem (Fig. 3), and MPECs (data not shown), suggesting that differences in Tmem Eomes expression between the two infection routes is likely cell intrinsic and not necessarily Tmem subset specific. Eomes expression is a key factor relevant to maintaining systemically derived CD8+ Tmems, partially because of its ability to upregulate CD122 expression (20). Therefore, respiratory viral infections may favor memory cells that are IL-15 independent. Intriguingly, TRM, defending in mucosal sites such as the lung, express less CD122 than other Tmem subsets (46) and are two times more abundant after i.n. VSV infection. Additionally, a subset of TRM isolated from the LNs of mice was found to develop independent of IL-15 signaling (47). Although only a subset of Tmem have reduced CD122 expression, we have shown that equivalent numbers of Tmem are isolated from the lung in IL-15–deficient and replete mice (10). Therefore, CD8+ T cell programming after respiratory infection may favor the development of specific subsets of Tmem, many of which will provide protection at the site of infection (perhaps with reduced longevity) over large pools of classical memory cells, which are considered dependent on IL-15. Because the role of IL-15 in CD8+ Tmem generation and maintenance has not been well studied after oral or intravaginal infection, it is possible that these results are not exclusive to respiratory environment; other mucosal immunizations may provoke similar changes in Tmem formation.

The difference in Tmem programming after respiratory infection is likely associated with events occurring very early postinfection, as VSV is a promiscuous virus (48), eventually resulting in a brief systemic infection, even after i.n. delivery (49). Respiratory-resident CD103+ DCs are particularly important in the priming of naive CD8+ T cells after influenza infection (30, 35). Given that these DCs can influence effector cell differentiation and migration (7), we hypothesized that these migratory CD103+ DCs were also responsible for the altered developmental phenotypes observed following i.n. infection. On first glance, our data seemed to confirm this hypothesis, as CD103+ T cells were specifically enriched in the respiratory tract dLNs following i.n. infection (Fig. 6). However, upon further analysis, we found that only ∼10% of the total dLN DCs were classic CD103+ RDCs, whereas a striking proportion (∼30%) of the DCs coexpressed CD103 and CD11b and had low expression of MHC II, a phenotype more indicative of a monocyte-derived CD103+ population of DCs (50). This DC phenotype is not observed after i.v. infection. Because many classical tissue-resident CD103+ will die in the lung dLN after priming CD8+ T cells (36), perhaps this newly recruited CD103+CD11b+ pool develops to replace the tissue-resident CD103+ DCs as a result of emergency myelopoiesis. Unlike lymphocytes, which can undergo rapid proliferation, innate immune cells like DCs must be replenished through expansion of hematopoietic stem cell populations in the bone marrow (51), which is crucial for controlling infection in many infection models (52, 53). It is possible that i.n. infection induces either selectively or at a greater level certain inflammatory signals, which trigger myelopoiesis and the development of moDCs. Inflammatory cytokines IFN-α/β and IFN-ɣ can stimulate proliferation of hematopoietic stem cell populations in the bone marrow (54), both of which are produced in abundance following viral infections (5557). It remains unclear why the moDCs are preferentially recruited to the lung dLN (and not the lung) after i.n. infection. One possibility is that CCR2 ligands are projected to the dLN, selectively recruiting the CCR2+ moDC (58). Nonetheless, once there, moDCs have the potential to regulate the CD8 Tmem program by delivering inhibitory signals via PD-1 or CTLA-4 engagement because of their high expression of PD-L1 and B7-1 (Table I). Both PD-1 and CTLA-4 engagement can result in T cell inhibition by directly blocking TCR signaling (59, 60) as well as via the mTOR signaling pathway (39). Indeed, ribosomal S6 protein is phosphorylated following both i.v. and i.n. VSV infection yet reduced only in i.n. infected animals 72 h postinfection in the lung dLN and coordinate with moDC arrival. Therefore, it is possible that the developmental stall in memory formation observed following respiratory infection could be due to suppressive signals delivered directly to respondent T cells by moDCs during T cell priming. Future experiments will examine the role of myelopoiesis and specific immunosuppressive pathways in regulating Tmem development.

By comparing VSV infection delivered by the i.n. or i.v. route, we showed that the respiratory environment results in Tmem that is skewed from the archetypical memory developmental programs defined in systemic models of infection, resulting in numerically deficient memory (Supplemental Fig. 2). The implications of this work suggest that the induction of protective memory CD8+ T cells should be studied in the context of appropriate infection route, as the developmental pathways and requirements for memory vary between routes of priming. As there continues to be a growing interest in developing CD8+ T cell–based vaccines (particularly those that will induce respiratory-specific responses) (61), it is imperative that we continue to improve our understanding regarding the mechanism of how the respiratory environment modifies Tmem. Fine-tuning of the local respiratory environment via targeting specific DC pools or perhaps induction of responses via other routes of mucosal infection may be necessary to secure the desired Tmem outcome.

We thank Dr. Ralph Tripp for access to an LSR II, Julie Nelson for assistance with cell sorting, Dr. Shannon Pham for infecting animals with RSV, and Drs. Katherine Verbist and Dave Rose for assistance with experiments.

This work was supported by grants from the National Institutes of Health (AI081800 and AI131093 to K.D.K.).

The online version of this article contains supplemental material.

Abbreviations used in this article:

cLN

cervical LN

DC

dendritic cell

dLN

draining lymph node

dpi

day postinfection

EEC

early effector cell

Eomes

eomesodermin

i.n.

intranasal(ly)

KLRG1

killer cell lectin-like receptor G1

LN

lymph node

MdLN

mediastinal LN

MHC I

MHC class I

MHC II

MHC class II

moDC

monocyte/dendritic cell

MPEC

memory precursor effector cell

N

nucleoprotein

PD-L1

programmed death ligand 1

RDC

respiratory DC

RSV

respiratory syncytial virus

SLEC

short-lived effector cell

TCM

central Tmem

Teff

effector T cell

Tem

effector Tmem

Tmem

memory T cell

TRM

tissue-resident Tmem

VSV

vesicular stomatitis virus.

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The authors have no financial conflicts of interest.

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