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Mucosally Delivered Dendritic Cells Activate T Cells Independently of IL-12 and Endogenous APCs¹

Department of Pathology and Molecular Medicine, Centre for Gene Therapeutics, and M. G. DeGroote Institute for Infectious Disease Research, McMaster University, Hamilton, Ontario, Canada

	Abstract

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In vitro manipulated dendritic cells (DC) have increasingly been used as a promising vaccine formulation against cancer and infectious disease. However, improved understanding of the immune mechanisms is needed for the development of safe and efficacious mucosal DC immunization. We have developed a murine model of respiratory mucosal immunization by using a genetically manipulated DC vaccine. Within 24 h of intranasal delivery, the majority of vaccine DCs migrated to the lung mucosa and draining lymph nodes and elicited a significant level of T cells capable of IFN- secretion and CTL in the airway lumen as well as substantial T cell responses in the spleen. And such T cell responses were associated with enhanced protection against respiratory mucosal intracellular bacterial challenge. In comparison, parenteral i.m. DC immunization did not elicit marked airway luminal T cell responses and immune protection regardless of strong systemic T cell activation. Although repeated mucosal DC delivery boosted Ag-specific T cells in the airway lumen, added benefits to CD8 T cell activation and immune protection were not observed. By using MHC-deficient vaccine DCs, we further demonstrated that mucosal DC immunization-mediated CD8 and CD4 T cell activation does not require endogenous DCs. By using IL-12-deficient vaccine DCs, we also observed that IL-12^–/– DCs failed to migrate to the lymph nodes but remained capable of T cell activation. Our observations indicate that mucosal delivery of vaccine DCs represents an effective approach to enhance mucosal T cell immunity, which may operate independent of vaccine IL-12 and endogenous DCs.

	Introduction

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Dendritic cells (DC)³ are highly specialized APCs that form a network over both the interior and exterior surfaces of the body, constituting an important part of host defense against invading pathogens (1). DCs are the most potent APC critically involved in the initiation of innate and adaptive immune responses to pathogens, particularly at mucosal surfaces (2). Thus, in vitro manipulation of DCs and subsequent in vivo administration have been exploited as an important strategy for immunization against cancer or infectious disease.

In vitro/ex vivo-manipulated DCs have been used to activate T cells in vivo in the field of immunotherapy. For instance, autologous patient-derived DCs loaded ex vivo with tumor Ags were demonstrated to be able to break tolerance and activate T cells in tumor-bearing patients (3, 4, 5, 6). Furthermore, DC-based immunotherapy is also being explored to enhance pathogen-specific immune responses in immunocompromised individuals, such as those infected with HIV (7). In this regard, there are two advantages of using in vitro-manipulated DC vaccine; 1) to restore anti-HIV immunity, the in vivo otherwise deactivated DCs can be reactivated and loaded with HIV epitopes ex vivo and thus enabled to activate T cells upon being readministered back to the same host; and 2) the same DCs can be potentially manipulated to carry or express Ags of multiple opportunistic pathogens such as mycobacteria and Aspergillus that often cause lethal secondary infection in HIV hosts (8, 9). Although both experimental and clinical studies suggest that such DC-based vaccines are safe and promising, their immune-activating and protective efficacy has remained questionable.

Like any other immunization strategies, both the mode of Ag expression and the route of delivery play an important role in determining the efficacy of DC-based immunization strategy. We and others have shown that virally transduced or gene-modified DCs are superior at in vivo T cell activation over peptide- or whole-protein-loaded DCs (10, 11, 12, 13). Because many of the infectious diseases are acquired via the mucosal route, it is believed that compared with the systemic or parenteral route, mucosal immunization at the site of pathogen entry may elicit the most protective immunity. Indeed, mounting experimental evidence supports this notion (13, 14, 15, 16). Although live organism-based vaccines including genetically modified viruses or attenuated pathogens may be directly used for the purpose of mucosal immunization, they could be unsafe and ineffective, particularly in immunocompromised hosts. In this regard, DC immunization represents an attractive approach which is not only safe and repeatable for mucosal application but is also able to effectively deliver Ags to activate adaptive immune cells, thus overcoming the limitations of impaired mucosal immunity in immunocompromised hosts.

To date, a small number of in vitro-manipulated DC formulations have been explored for the purpose of respiratory mucosal immunization only with limited success. For instance, only a short term of protection against Mycobacterium tuberculosis was accomplished following intranasal (i.n.) mucosal immunization with the DCs infected with live Mycobacterium bovis bacillus Calmette-Guérin (BCG; Ref. 17). Likewise, DCs loaded in vitro with heat-killed Bordetella pertussis were only modestly effective in priming lung mucosal immune responses (18). Furthermore, i.n. delivery of the DCs genetically modified with plasmid DNA encoding a microbial Ag, only minimally enhanced T cell responses and protection from Coccidioides challenge (8, 19). Such limited success in mucosal DC immunization is believed to be attributable at least in part to our lack of knowledge in DC trafficking, immunogenic differences by mucosal and parenteral routes of delivery, and the mechanisms involved in DC mucosal immunization.

In this study, we have used the in vitro genetically manipulated DCs as a tool and murine i.n. mucosal immunization as a model system to specifically address: 1) where vaccine DCs travel postmucosal delivery; 2) how well such delivered DCs may activate Ag-specific T cells both locally and systemically and how it compares to the parenteral route of delivery; 3) whether DC-activated T cell responses translate to immune protection; and 4) the role of DC-associated MHC class I (MHCI) and II (MHCII) molecules and IL-12 in T cell activation.

	Materials and Methods

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Mice and reagents

Female BALB/c and C57BL/6 mice (6–10 wk old) were purchased from Harlan Laboratories. Congenic CD45.1⁺ mice were purchased from The Jackson Laboratory. C2DBN12 (MHCII^–/–) mice were purchased from Taconic. β₂-Microglobulin-CD8 double-knockout (MHCI^–/–) and IL-12 p40 knockout mice (IL-12^–/–) were bred in house at McMaster University (Hamilton, Ontario, Canada). All animals were housed in a specific-pathogen-free facility in the Central Animal Facility at McMaster University in accordance with the McMaster Animal Research Ethics Board. The construction and amplification of a replication deficient (E1/E3 deleted) recombinant adenovirus encoding the gene for Ag 85A has been previously described (11) and was used to transduce bone marrow derived dendritic cells. All viruses were purified and stored at –70°C until needed. Purified M. tuberculosis Ag 85 complex and M. tuberculosis culture filtrate proteins were provided by Colorado State University (Fort Collins, CO) through funds from the National Institute of Allergy and Infectious Disease (Contract 1-A1-75320). Two synthetic Ag85A peptides on a BALB/c background (H-2^d) were used. The MHCI-specific peptide (MPVGCQSSF) and MHCII-specific peptide (LTSELPGWLQANRHVKPTGS) were synthesized by Dalton Chemical Laboratories (Toronto, Ontario, Canada). All proteins were dissolved in DMSO and stored at –20°C until needed.

Generation of bone marrow-derived DCs and preparation of DC-based vaccine

Bone marrow cells were harvested from the femurs and tibiae of naive BALB/c mice as previously described (11) and cultured in RPMI 1640 containing 10% FBS, 100 U/ml penicillin, 100 µg/ml streptomycin, 2 mM L-glutamine, 0.1 mM nonessential amino acids, 1 mM sodium pyruvate, and 40 pg/ml recombinant murine GM-CSF. Three and six days after initial culture, cells were replenished with fresh medium supplemented with GM-CSF. On day 7, DCs were infected with AdAg85A (DCAdAg85A) at a multiplicity of infection of 100. After 4 h of incubation at 37°C, all DCs were harvested by scraping the bottom of culture plates with a spatula, washed thee times with PBS, and used to immunize mice.

Immunization, preparation of DC vaccines, and in vivo trafficking

Vaccines were prepared immediately before immunization and kept on ice until use. All DCs were delivered either i.n. or i.m. Immunizations were conducted by using 1.0 x 10⁶ DCs/mouse in 30 µl of PBS for i.n. injections and 0.5 x 10⁶ DCs/mouse in 100 µl of PBS for i.m. injections. A higher dose was used for i.n. delivery considering the loss of some of i.n. delivered materials to the gastrointestinal tract. Anesthetized mice were allowed to breath in the 30 µl containing vaccine DCs, or DC vaccine suspension was injected into the hind legs (50 µl each leg). For in vivo trafficking studies, either autologous bone marrow-derived DCs or congenic CD45.1⁺ DCs were prepared as described and labeled with 5 µM CFSE. Cells were counted by trypan exclusion, and 1 x 10⁶ live cells were delivered i.n. in 30 µl of PBS. At 24-h intervals, mice were sacrificed and bronchoalveolar lavage (BAL), lung interstitium, cervical lymph nodes (cLN), mediastinal lymph nodes (mLN), popliteal LN (pLN), and spleen were isolated. Lung tissue, cLN, mLN, and pLN were digested with collagenase for 1 h to release APC populations from the connective tissue and processed to single-cell suspensions. Cells were stained for CD45.1 and/or CD11c and analyzed for the presence of CD45.1⁺ or CFSE-labeled CD11c⁺ populations by flow cytometry. Live M. bovis BCG (0.5 x 10⁶ CFU/mouse; Connaught Laboratories) was injected s.c. around both hind legs, as a positive control of vaccination in M. tuberculosis challenge experiments.

In vivo i.v. and intratracheal (i.t.) CTL assays

The in vivo CTL assay was conducted as previously described (11, 14). Briefly, splenocytes from naive female BALB/c mice were isolated the night before each in vivo cytotoxicity assay. Spleens were removed into complete medium (RPMI 1640 supplemented with 10% FBS, 100 U/ml penicillin, 100 µg/ml streptomycin, 2 mM L-glutamine, and 50 µM 2-ME). Splenocyte suspension was filtered through a 55-µm pore size nylon membrane before being centrifuged at 1500 rpm for 5 min. Pellets were resuspended in 2 ml/spleen of RBC lysis buffer (R&D Systems) and incubated at room temperature for 12 min. Approximately 30 ml of PBS were added after the 12-min incubation to stop the RBC lysis. The whole splenocytes were filtered through a 55-µm pore size nylon membrane, centrifuged at 1500 rpm for 5 min, and resuspended at 20 x 10⁶ cells/ml in complete medium. For the in vivo cytotoxicity assay, the splenocytes were pulsed with either the CD8 or CD4 T cell Ag85A peptide (10 µg/ml) and incubated overnight at 4°C. Such splenocytes used as CTL target cells were then resuspended at 20 x 10⁶ cells/ml in PBS containing 5% FBS. The Ag85A peptide-pulsed splenocytes were labeled with 5 µM CFSE and denoted CFSE^high, and the unpulsed splenocytes were labeled with 0.5 µM CFSE (CFSE^low) for 5 min at room temperature in unlit conditions. The cells were washed twice to remove any free CFSE with 5% FBS-PBS and then once with PBS. Both CFSE^high and CFSE^low cell populations (5 x 10⁶ of each per mouse for i.v. CTL, 1 x 10⁶ of each per mouse for i.t. CTL) and were mixed in a 1:1 ratio (in a total of 200 µl volume for i.v CTL and 40 µl for i.t. CTL) and injected into DC-vaccinated mice. A naive mouse was also injected with peptide-pulsed, CFSE-labeled cells and used as unprimed control for calculation (see equation at end of paragraph). After 6 h (for the CD8 CTL assay) or 24 h (for the CD4 CTL assay) following target cell injection, either splenocytes (i.v CTL) or BAL cells (i.t. CTL) were isolated. The in vivo lysis of the target cells was determined according to the extent of loss of CFSE dye by flow cytometry. Up to 1 x 10⁶ events were collected for analysis. To calculate Ag-specific, CD8 or CD4 T cell-mediated lysis, the formula used was: percentage of specific lysis = [1 – (ratio unprimed/ratio primed) x 100], where ratio is (percentage CFSE^low/percentage CFSE^high).

Cell isolation for characterization of immune activation following vaccination

Immunized mice were sacrificed 2, 6, or 12 wk postimmunization to examine immunogenicity. Spleens and lungs were removed aseptically and the intra-airway luminal cells were removed from the lung by exhaustive lavage as previously described (14, 15). Briefly, the mouse lung was lavaged five times with 1.8 ml of PBS through a polyethylene tube cannulated into the trachea to ensure maximal recovery. After lavage, the lungs were perfused through the left ventricle with Hanks’ buffer to remove RBC from the vasculature. The lungs were then cut into small piece (>1 mm x 1 mm) and incubated with collagenase type 1 (Sigma-Aldrich) for 1 h at 37°C. Lung fragments were then crushed through a 100-µm pore size filter (18). Cells were collected and enumerated on a hemocytometer after dilution in Turks white blood cell counting buffer. Spleen cells were isolated as previously described (12). All isolated cells (spleen, lung, and airway-luminal) were then resuspended in RPMI 1640 supplemented with 10% FBS, 1% L-glutamine, and 1% penicillin and streptomycin.

FACS, intracellular cytokine staining, and tetramer staining

These procedures were conducted as previously described (11, 14, 15, 20). Briefly, single-cell suspensions from immunized mice of spleen, lung, and airway-luminal cells were obtained as described above. Cells were cultured in a U-bottom 96-well plate at a concentration of 20 x 10⁶ cell/ml, and airway-lumen-derived cells were cultured at a concentration of 5 x 10⁶ cells/ml. Cells were cultured in the presence of Golgi plug (10 µg/ml brefeldin A; BD Pharmingen) and either no stimulation, Ag85A CD4 or CD8 T cell peptides at a concentration of 1 µg/well for 6 h. Cells cultured with Ag85 complex and M. tuberculosis culture filtrate at a concentration of 10 µg/ml in the absence of Golgi plug for the first 18 h followed by 6 h in the presence of Golgi plug. Cells were then washed and blocked with CD16-CD32 in 0.5% BSA-PBS for 15 min on ice and then stained with the appropriate surface Abs. Cells were then washed, permeabilized, and stained according to the manufacturer’s instructions included in the ICCS kit (BD Pharmingen). The following Abs were used: CD3-CyChrome (BD Pharmingen); CD4-PE-Cy7 (BD Pharmingen); CD8a-allophycocyanin-Cy7 (BD Pharmingen); and IFN--allophycocyanin (BD Pharmingen). Stained cells were then run on a FACS Canto, and 250,000 events were collected per sample (BD Pharmingen) and analyzed on FlowJo software (version 6.3.4; Tree Star). Tetramer flow cytometric analysis was conducted using the immunodominant CD8 T cell peptide (MPVGGQSST) of Ag85A bound to the BALB/c MHCI allele H-2L^d which was ordered from Texas A&M University (College Station, TX). Cells were washed and blocked with CD16-CD32 in 0.5% BSA-PBS for 15 min on ice, then stained with tetramer for 1 h in the dark at room temperature, and then washed and stained with surface Abs. Stained cells were then run on a FACS Canto, and 250,000 events are collected per sample (BD Pharmingen) and analyzed on FlowJo software.

Immune protection against pulmonary M. tuberculosis challenge

M. tuberculosis (H37Rv strain) (ATCC 27294) was grown in Middlebrook 7H9 broth supplemented with Middlebrook OADC enrichment (Invitrogen), 0.002% glycerol, and 0.05% Tween 80 for 10–15 days and then aliquoted and stored in –70°C until needed as previously described (14, 15, 20). Before each use, M. tuberculosis bacilli were washed with PBS containing 0.05% Tween 80 twice and passed through a 27-gauge needle 10 times to disperse clumps. Immunized and non-immunized mice were infected i.n. with 10,000 CFU of M. tuberculosis at either 4 or 6 wk postimmunization in the Level III Containment Facility of McMaster University. The level of bacterial burden was determined 4 wk postchallenge in the lung and spleen by plating serial dilutions of tissue homogenates in triplicates onto Middlebrook 7H10 agar plates containing Middlebrook OADC enrichment. Plates were incubated at 37°C for 21 days in semisealed plastic bags. Colonies were then counted, calculated, and presented as log₁₀ CFU per organ.

Detection of M. tuberculosis Ag85A by Western blotting

Western blot was used to detect both intracellular and secreted forms of Ag85A from AdAg85A-infected cells as previously described (21). Briefly, bone marrow-derived DCs were infected with a multiplicity of infection of 100 AdAg85A for 6 h and were washed with 5% FBS-PBS to remove free virus. Cells were cultured in 12-well plates for up to 7 days. Medium was replenished every other day as needed to ensure that cells remained viable. On the day of harvest, cell supernatants were collected and spun to remove any cellular contamination. Cells were lysed with 0.05% Triton in water and immediately frozen. A 10% SDS gel was cast, and samples were loaded immediately. Membranes were probed with a mouse anti-Ag85A mAb and HRP-conjugated anti-mouse IgG.

Statistical analysis

All statistics were performed with Microsoft Excel using a two-tailed t test assuming equal variances. Individual p values are included in the text.

	Results

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 
In vivo distribution of vaccine DCs following i.n. administration

Although cell-based mucosal immunization strategies are an increasingly popular approach to generating protective immunity against pathogens, there is a lack of information on the distribution of vaccine cells in the mucosal and nonmucosal compartments following i.n. mucosal delivery. To this end, we prepared CD45.1⁺ congenic vaccine DCs transduced to express an immunogenic M. tuberculosis Ag Ag85A by using a recombinant adenoviral vector (DCAdAg85A) for i.n. delivery. At various times post-DC delivery, cells isolated from various tissue compartments were analyzed for the frequency of CD45.1+ DCs. Maximal numbers of vaccine DCs were detected in the airway lumen (Fig. 1A) and the lung interstitium (Fig. 1B) at 24 h. These airway luminal and lung mucosa-associated populations underwent contraction over the 14-day period examined (Fig. 1, A and B), and a small population of residual vaccine DCs could be detected 14 days after i.n. delivery. Vaccine DCs could be detected as early as 24 h following i.n. delivery in the cLNs (7.5% of total recovered DCs), which drain the nasal mucosa and in the mLNs (16% of total recovered DCs) which drain the lung (Fig. 1, C and D). Vaccine DCs migrated to these lymph nodes (LN) contracted rapidly, and the total number of vaccine DCs declined sharply by 72 h. A small but stable population of vaccine DCs could be detected up to 14 days, particularly in mLNs. In sharp contrast, no appreciable CD45.1⁺ congenic vaccine DCs could be detected in the distal lymphoid organs including the spleen and pLNs at any time point (data not shown). Similar kinetics of vaccine DC trafficking were also observed by using CFSE-labeled autologous DCs (data not shown). These data suggest that some of the i.n. delivered vaccine DCs have the ability to migrate rapidly from the airway lumen to the local draining LNs and lung interstitium while a portion of these cells remain within the airway lumen.

View larger version (16K): [in this window] [in a new window]   FIGURE 1. Localization of virally transduced vaccine DCs following i.n. administration. Congenic CD45.1-expressing DCs were transfected with AdAg85A (multiplicity of infection, 100), and 1 x 106 cells were delivered i.n. to BALB/c mice. At 24-h intervals, mice were sacrificed, and the cells isolated from BAL (A), lung interstitium (B), cLN (C), and mLN (D) were examined by flow cytometry for quantification of CD45.1+CD11c+ DCs. Data are expressed as the mean value of two mice per time, representative of two independent experiments. d, Day.

Having examined the distribution of vaccine DCs, we set out to determine the kinetics of activation, phenotype, and pulmonary distribution of Ag-specific T cells in response to i.n. DC immunization. We also compared i.n. with parenteral i.m. route of immunization. We paid special attention to the number of airway luminal T cells that have been shown to be essential in providing protection against pathogenic mucosal challenge (14, 22). Mice were immunized i.n. or i.m. with DCAdAg85A and sacrificed at 2, 6, and 12 wk postimmunization. We first analyzed the number of total CD4⁺ and CD8⁺ T cells in the airway lumen by FACS. The levels of CD4⁺ and CD8⁺ T cells in the airway lumen of i.m. DCAdAg85A-immunized mice were very small and insignificant at various time points (Table I). In contrast, i.n. DCAgAg85A induced much more total CD4⁺ and CD8⁺ T cells into the airway lumen than i.m. immunization (8.5- and 3-fold more CD4⁺ and CD8⁺ T cells at 2 wk; 3- and 5.4-fold more at 6 wk; and 5.8- and 3.3-fold more at 12 wk, respectively; Table I). Next, the number of Ag-specific T cells in the airway lumen was quantified by tetramer staining and ICCS for IFN-. DCAdAg85A immunization i.n. elicited a significant population of CD8⁺tetramer⁺ T cells in the airway lumen at 2 wk postimmunization (Fig. 2A). This population contracted between 2 and 6 wk postimmunization (from 2.18 x 10⁴ cells to 0.48 x 10⁴ cells/BAL; >5.5-fold reduction). However, it remained stable up to 12 wk (0.33 x 10⁴ cells/BAL; Fig. 2A). To understand the functional capacity of such Ag-specific airway luminal T cells elicited by i.n. DC immunization, the number of IFN--producing CD8⁺ T cells was enumerated by ICCS. The kinetic pattern of IFN--secreting cells mirrored the tetramer⁺ cells, although the absolute number of IFN--secreting cells was lower than tetramer⁺ cells. CD8⁺IFN-⁺ T cell responses peaked at 2 wk (1.62 x 10⁴ cells/BAL) declined markedly by 6 wk (0.25 x 10⁴ cells/BAL) and further contracted by 12 wk (0.02 x 10⁴ cells/BAL) after immunization (Fig. 2B). An i.t. CTL was conducted as previously described to further characterize the function of airway luminal CD8⁺ T cells (14, 22). CD8 T cell-mediated CTL activity was greatest 2 wk (17.1%) after i.n. DC immunization, and low levels of CTL activity could still be detected up to 12 wk (2.5%; Fig. 2C). We also determined the level of CD4 T cell activation in the airway lumen following i.n. DC immunization. A significant population of CD4⁺IFN-⁺ T cells could be detected in the airway at all time points examined (Fig. 2D). Similar to CD8⁺ T cells, the number of CD4⁺IFN-⁺ T cells peaked at 2 wk postimmunization and declined by 6 wk; only a small population could be detected at 12 wk post-i.n. DCAdAg85A delivery. Compared with T cell responses in the airway lumen, different kinetics of T cell responses were observed in the lung interstitium post-i.n. DC vaccine delivery. Similar numbers of total CD3⁺CD4⁺ and CD3⁺CD8⁺ T cells were found in the lung interstitium between the two immunization regimens at all time points examined (data not shown). The levels of CD8⁺tetramer⁺ cells, IFN--secreting CD8⁺ T cells and IFN--secreting CD4⁺ T cells in the lung interstitium peaked at 12, 6, and 6 wk, respectively (Fig. 3). Also, similar to CD4 T cell responses in the airway lumen (Fig. 2D), CD4 T cell responses in the lung interstitium almost vanished at 12 wk (Fig. 3C).

View larger version (24K): [in this window] [in a new window]   FIGURE 2. T cell responses in the airway lumen following i.m. or i.n. DCAdAg85A immunization. BALB/c mice were immunized and scarified 2, 6, or 12 wk later. Total airway luminal cells were obtained by exhaustive BAL. Cells were stained directly with tetramer (A) or cultured in the presence of Ag85A CD8 T cell peptide (B) or CD4 T cell peptide (D) for 6 h and in the presence of Golgi plug. Cells were then stained and analyzed by flow cytometry. CD8 T cell-mediated i.t. CTL activity (C) was measured by transferring CFSE-labeled Ag85A CD8 T cell peptide-loaded target splenocytes to the lungs of vaccinated mice. Lungs were lavaged 6 h later, and the relative loss of CFSE-labeled cells was examined by flow cytometry. The representative histograms of week 2 CTL analysis are presented. Data represent the means ± SEM of six to nine mice per time per group from two to three independent experiments. *, p < 0.05; **, p < 0.01.

View larger version (25K): [in this window] [in a new window]   FIGURE 3. T cell responses in the lung following i.m. or i.n. DCAdAg85A immunization. BALB/c mice were immunized and scarified 2, 6, or 12 wk later. Total lung cells were obtained by collagenase digestion and mechanical disruption of lung tissue. Cells were stained directly with tetramer (A) or cultured in the presence of Ag85A CD8 T cell peptide (B) or CD4 T cell peptide (C) for 6 h and in the presence of Golgi plug. Cells were then stained and analyzed by ICCS and flow cytometry. Data represent the means ± SEM of six to nine mice per time per group from two to three independent experiments.

Ag-specific T cell responses in the distant lymphoid organs following i.n. administration of vaccine DCs

It is believed that besides the elicitation of T cell responses at the mucosa, an effective vaccine is also expected to generate Ag-specific T cells in the systemic lymphoid organs to effectively control the systemic dissemination of pathogens from the mucosa as well as to provide a rich peripheral reservoir of Ag-specific T cells that may be mobilized to the mucosal site upon mucosal pathogen exposure. For these reasons, we also examined the level of systemic T cell responses following i.n. DC immunization. We first quantified the total CD4⁺ and CD8⁺ T cells in the spleen following i.n. and i.m. DCAdAg85A vaccination (Table II). Compared with i.n. DC delivery, i.m. DC immunization resulted in more total CD4⁺ and CD8⁺ T cells at both 2 and 6 wk but similar T cell numbers in the spleen at 12 wk (Table II). We then quantified the number of Ag-specific T cells in the spleen following i.n. DC immunization. A population of CD8⁺tetramer⁺ T cells were detected in the spleen at 2 wk and steadily increased in numbers from 2 to 6 wk and maintained between 6 and 12 wk (Fig. 4A). Ag-specific CD8⁺IFN-⁺ T cells were also readily detectable in the spleen of i.n. DC-immunized mice at various time points following immunization (Fig. 4B). We further assessed CD8 T cell-mediated CTL activity in the spleens of i.n. DC immunized mice by using a systemic in vivo CTL assay (14, 22). Low levels of CD8 CTL were measurable in the spleens by 2 wk which steadily increased between 2 and 12 wk post-i.n. immunization (Fig. 4C). We also observed a relatively small yet stable population of Ag-specific CD4⁺IFN-⁺ T cells in the spleen at all time points examined (Fig. 4D) and similar low levels of CD4 T cell-mediated CTL activity in the spleen (data not shown).

View larger version (27K): [in this window] [in a new window]   FIGURE 4. T cell responses in the spleen following i.m. or i.n. DCAdAg85A immunization. BALB/c mice were immunized and sacrificed 2, 6, or 12 wk later. Whole splenocytes were processed to single-cell suspensions. Cells were stained directly with tetramer (A) or cultured in the presence of Ag85A CD8 T cell peptide (B) or CD4 T cell peptide (D) and Golgi plug for 6 h. Cells were then stained and analyzed by ICCS and flow cytometry. Systemic CTL responses were measured by i.v. delivery of CFSE-labeled, Ag85A CD8 T cell peptide-pulsed splenocytes (C). The representative histograms of week 2 CD8 CTL analysis are presented. Mice were sacrificed 6 h later, and the splenocytes were analyzed for the relative loss of CFSE-labeled cells. Data represent the means ± SEM of six to nine mice per group from two to three independent experiments. *, p < 0.05; **, p < 0.01.

Enhanced local and systemic immune protection from pulmonary bacterial challenge by i.n. administration of vaccine DCs

To establish the relevance of improved T cell responses by i.n. mucosal DC immunization, we assessed the immune protective capacity by i.n. DC immunization and compared this with i.m. parenteral DC immunization. Because the model microbial Ag expressed by adenoviral vector-transduced vaccine DCs in this study is an immunogenic M. tuberculosis Ag, the i.n. or i.m. DC-immunized mice were challenged with virulent M. tuberculosis. Compared with naive controls, i.m. DC immunization provided little protection against pulmonary pathogenic challenge as measured by bacterial colony assay in the lung and spleen (Fig. 5A). In contrast, i.n. DC immunization provided an enhanced level (half-log) of protection against challenge, although it does not appear to be as potent as BCG immunization, a standard positive control set up in parallel in this study (Fig. 5A). Furthermore, i.n. DC immunization also led to an enhanced level of systemic immune protection in the spleen, and this level of protection was comparable with that by BCG control (Fig. 5B). These data demonstrate that the Ag-specific T cell responses elicited by mucosally administered, gene-modified vaccine DCs can lead to enhanced host defense both at local and systemic tissue compartments.

View larger version (14K): [in this window] [in a new window]   FIGURE 5. Immune protection against pulmonary M. tuberculosis (M.tb) challenge by i.n. delivery of vaccine DCs. Mice were immunized with DCAdAg85A i.n. or i.m. or with BCG s.c. (positive control) or PBS (naive control). Mice were challenged i.n. with virulent M. tuberculosis 6 wk postimmunization. Four weeks after challenge, mice were sacrificed, and the level of M. tuberculosis infection in the lung (A) and spleen (B) was enumerated by colony-forming assay. Data represent the means ± SEM of three to five mice per group. *, p < 0.05; **, p < 0.001.

Repeated systemic delivery of vaccine DCs has been attempted for cancer immunotherapy with some success in activating tumor-associated Ag-specific T cells (7, 23). Here, after having investigated the immunogenicity and protective efficacy of a single mucosal DC delivery, we examined the possibility of repeated administrations of vaccine DCs by using a homologous prime-boost immunization regimen. To this end, mice were immunized either i.n. or i.m. with vaccine DCs as described above. After 2 wk, mice were boosted i.n. with the same vaccine DCs. Four weeks after i.n. boosting, mice were sacrificed and the number of tetramer⁺ CD8⁺ T cells in the airway lumen and spleen was quantified by flow cytometry. DC boosting i.n. of i.m. or i.n. primed mice resulted in significant expansion of the number of tetramer⁺ CD8 T cells in the airway lumen, and such enhanced levels of tetramer⁺ CD8 T cell responses were comparable in i.m. and i.n. primed mice. However, i.n. DC boosting did not significantly increase the number of CD8⁺IFN-⁺ T cells in the airway lumen of i.m. primed mice, and it even decreased such cells in i.n. primed mice (Fig. 6B). In comparison, i.n. DC boosting increased CD4⁺IFN-⁺ cells in the airway lumen of either i.m.- or i.n.-primed mice (Fig. 6C). In the lung interstitium, i.n. DC boosting did not increase tetramer⁺ CD8 T cells (Fig. 6D) and as in the airway lumen, it led to significantly lower numbers of CD8⁺IFN-⁺ cells (Fig. 6E), whereas it had little enhancing effect on CD4⁺IFN-⁺ cells (Fig. 6F). In the spleen, whereas i.n. DC boosting substantially increased tetramer⁺ CD8⁺ T cells above the levels generated by either i.m. or i.n. prime immunization, it led to decreased CD8⁺IFN-⁺ and CD4⁺IFN-⁺ T cells (data not shown).

View larger version (25K): [in this window] [in a new window]   FIGURE 6. Expansion of Ag-specific T cells by repeated homologous DC immunization. Mice were immunized either i.n. or i.m. with DCAdAg85A; 2 wk later, some of the mice were boosted i.n. with DCAdAg85A. Total airway luminal cells were obtained by exhaustive BAL, and lung interstitial cells were digested with collagenase before mechanical disruption to single-cell suspensions. BAL (A) and lung interstitial cells (D) were stained directly with tetramer or cultured in the presence of Ag85A CD8 T cell peptide (B and E, respectively) or CD4 T cell peptide (C and F, respectively) and Golgi plug for 6 h. Cells were then stained and analyzed by ICCS and flow cytometry. Mice boosted with DCAdAg85A were compared with single immunization controls. Data represent the mean value ± SEM of three to five mice per group from two independent experiments. *, p < 0.05; **, p < 0.01.

View larger version (14K): [in this window] [in a new window]   FIGURE 7. Immune protection against pulmonary M. tuberculosis (M.tb) challenge by single i.n. DCAdAg85A immunization or by DCAdAg85A prime-boost immunization. Mice were immunized either i.m. or i.n. with DCAdAg85A; 2 wk later, some of these mice were boosted i.n. with DCAdAg85A. Additional mice were set up as controls with BCG s.c. or PBS at the time of priming. Mice were challenged i.n. with virulent M. tuberculosis 4 wk postimmunization. Four weeks after challenge, mice were sacrificed, and the level of M. tuberculosis infection in the lung (A) and spleen (B) was determined by colony-forming assay. Data represent the means ± SEM of five to six mice per group. *, p < 0.05; **, p < 0.001 compared with all other groups.

Experimental evidence has suggested that endogenous DCs may be required for optimal T cell, particularly CD4 T cell, activation following the systemic delivery of vaccine DCs (24). Whether this may be the case for mucosally delivered vaccine DC-mediated CD4⁺ T cell activation still remains unclear. To address this question, we first examined whether AdAg85A-infected vaccine DCs could release the soluble form of Ag85A protein, considering that soluble microbial Ags could easily be picked up by endogenous DCs which may then activate T cells. We found that both cell-associated and soluble Ag85A protein could be detected by Western blotting as early as 24 h after AdAg85A infection of DCs (Fig. 8A) and as late as 7 days (data not shown). Because both cell-associated and -secreted forms of Ag85A could be detected from the cultures of vaccine DCs, we sought to examine whether endogenous APCs could be involved in vaccine DC-mediated CD8⁺ and CD4⁺ T cell activation. To address this, C57BL/6 background MHCI-deficient or MHCII-deficient DCs were prepared, infected with AdAg85A, and used to immunize naive wild-type (wt) C57BL/6 mice i.n.. In this case, if the endogenous APCs were significantly involved in T cell activation, we would still expect to see unchanged CD8 and CD4 T cell activation, respectively. To first examine the role of endogenous DCs in CD8 T cell activation, MHCI-deficient DC vaccine was delivered i.n. to naive wt mice. Compared with wt DC vaccination, CD8 T cell activation by MHCI-deficient DC vaccination was severely impaired as we observed sharply reduced numbers of CD8⁺IFN-⁺ T cells in both the airway lumen (Fig. 8B) and lung interstitium (Fig. 8C) of these mice. The number of such Ag-specific activated CD8 T cells was also significantly reduced in the spleen (Fig. 8D). In contrast, the numbers of CD4⁺IFN-⁺ T cells in all tissue compartments of MHCI^–/– DC-immunized mice was comparable with those in wt DC-immunized mice (Table III). We next examined the role of endogenous DCs in CD4 T cell activation following i.n. delivery of MHCII-deficient DC vaccine. Similar to impaired immunogenicity by MHCI-deficient DC vaccination, we observed no detectable CD4⁺IFN--producing T cells in the airway lumen of wt mice immunized i.n. with MHCII-deficient vaccine DCs, in sharp contrast to enhanced CD4 T cell responses by wt DC vaccination (Fig. 8E). Similarly, few CD4⁺IFN-⁺ T cells could be detected in the lung interstitium (Fig. 8F) and in the spleen (Fig. 8G). However, the numbers of CD8⁺IFN--producing T cells in all examined tissue sites of these mice remained comparable with those by wt DC vaccination (Table III). These data indicate that mucosally delivered wt vaccine DCs can directly activate both Ag-specific CD8 and CD4 T cells by and large independently of endogenous DCs.

View larger version (15K): [in this window] [in a new window]   FIGURE 8. Impaired CD8+ and CD4+ T cell activation by i.n. immunization with MHC-deficient vaccine DCs. DCs were infected with AdAg85A for 24 h, and both cell-associated and soluble Ag85A proteins were detected by Western blotting (A). Purified Ag85 complex protein was run as a positive control in Western blot. In separate experiments, AdAg85A-infected wt and MHCI or MHCII–/– vaccine DCs were delivered i.n. to wt C57BL/6 mice, and mice were sacrificed 2 wk later. Total airway luminal cells (B and E) were obtained by exhaustive BAL, whole-lung interstitial cells were obtained by collagenase digestion and mechanical disruption to single-cell suspensions (C and F), and whole spleen cells were processed to single-cell suspensions (D and G). Cells were cultured with M. tuberculosis Ags for 12 h followed by an additional 6 h in the presence of Golgi plug and then immunostained for ICCS. Ag-specific IFN--producing CD4 and CD8 T cells were analyzed by flow cytometry. The representative histograms of CD8+IFN-+ (B) and CD4+IFN-+ (E) T cell responses in BAL are also displayed. Data represent the means ± SEM of seven to nine mice per group from two independent experiments. *, p < 0.01; **, p < 0.001.

View this table: [in this window] [in a new window]   Table III. Comparison of CD4+IFN-- and CD8+IFN--producing T cell responses following i.n. immunization with wt, MHCI–/–, or MHCII–/– DCAdAg85Aa

Little is known about the role of vaccine DC-derived cytokines in T cell activation. However, endogenous APC-derived IL-12 has recently been shown to be critically involved in APC migration and type 1 T cell activation (25, 26); alternatively, vaccine DCs engineered to overexpress IL-12 was found to be more immunogenic to CD4⁺ T cell activation (27). We have previously shown that virally transduced vaccine DCs displayed increased amounts of IL-12 (11). To investigate whether vaccine DC-derived IL-12 was required for in vivo T cell activation, DCs deficient in IL-12 (IL-12^–/– DCs) isolated from C57BL/6 IL-12^–/– mice were transduced with AdAg85A and compared with wt C57BL/6 DCs for their transmigration to the local LNs and their ability to subsequently activate CD4 and CD8 T cells following i.n. administration to naive C57BL/6 mice. IL-12^–/– DCs showed no impairment in the ability to up-regulate MHCII and costimulatory B7 molecules as well as to produce proinflammatory cytokine TNF- (data not shown). In agreement with the data by Khader et al. (25), we found that whereas similar to wt controls, IL-12^–/– vaccine DCs could readily be detected in the airway and lung interstitium (Fig. 9, A and B), these cells in stark contrast to their wt counterparts, failed to migrate to the local LNs draining the nasal mucosa (cLN) and the lung (mLN; Fig. 9, C and D). Moreover, no CFSE⁺ vaccine DCs could be detected in the distal lymphoid organs including the spleen and pLNs at any time (data not shown). In light of these findings, we further examined the ability of IL-12^–/– DCAdAg85A to prime naive T cells in vivo. Thus, 2 wk after i.n. delivery, mice were sacrificed, and the number of Ag-specific IFN--secreting CD8⁺ and CD4⁺ T cells in the BAL and spleen was quantified by ICCS. To our surprise, similar to wt controls, IL12^–/– vaccine DCs activated Ag-specific CD8⁺IFN-⁺ and CD4⁺IFN-⁺ T cells in both the airway lumen (Fig. 9, E–G) and the spleen (Fig. 9, F–H). Similar numbers of Ag-specific CD8⁺ and CD4⁺ T cells were also detected in the lung and draining LNs (data not shown). These data suggest that although mucosally delivered IL-12-deficient vaccine DCs demonstrate an impaired ability to migrate to the draining LNs, the local microenvironment is still permissible for them to prime Ag-specific CD8⁺ and CD4⁺ T cell responses.

View larger version (24K): [in this window] [in a new window]   FIGURE 9. Immune activation by IL-12–/– vaccine DCs. wt C57BL/6 and IL12–/– DCs were transfected with AdAg85A and CFSE labeled. Vaccine DCs were delivered i.n. to wt C57BL/6 mice. At 24 and 96 h, mice were sacrificed and the BAL (A), lung interstitium (B), cLN (C), and mLN (D) were examined by flow cytometry for CFSE+CD11c+ DCs. Data represent the means ± SEM of two mice per group. In separate experiments, mice immunized with wt or IL-12–/– DCAdAg85A were sacrificed 2 wk after i.n. immunization. Total airway luminal cells were obtained by exhaustive BAL (E and G), and whole spleens were processed to single-cell suspensions (F and H). Cells were cultured in the presence of M. tuberculosis Ags for 12 h, followed by an additional 6 h with Golgi Plug, and immunostained for ICCS and CD8+IFN-+ and CD4+IFN-+ T cells were analyzed by flow cytometry. Data represent the means ± SEM of three to six mice per group.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

In this study, we addressed the outstanding issues in the field of mucosal DC vaccination including vaccine formulation, localization, kinetic T cell activation, protection, and mechanisms of immune activation. In addition, for the first time we conducted a direct comparison of parenteral and mucosal DC immunization strategies. The DCs virally transduced with a recombinant adenovirus expressing an M. tuberculosis Ag, which has previously been demonstrated to elicit superior immune responses over the peptide- or protein-pulsed counterparts (11, 28), is used as a model vaccine for immunization against an invasive intracellular pathogen. It has been known that resistance to many respiratory pathogens requires strong type 1 polarized immune responses characterized by IFN- and CTL responses in the airway lumen. Therefore, to profile the immunity and protective efficacy of i.n. and i.m. DC immunization, the immune responses elicited in the airway lumen, lung, and spleen by both parenteral and mucosal vaccination strategies were directly compared. DC immunization i.n. effectively elicited the sustained CD4⁺ and CD8⁺ T cell populations capable of IFN- and CTL responses in the airway lumen. Parenteral immunization primed a very small and short-lived CD8 T cell population in the airway lumen with only low CD8 CTL activities. Both i.n. and i.m. DC immunization activated CD4⁺ and CD8⁺ T cell responses in the spleen although i.m. immunization did tend to prime higher levels of T cell activation at this site than i.n. immunization. Associated with the higher magnitude of T cell responses within the respiratory mucosa by i.n. immunization was enhanced host defense against intracellular bacterial challenge, in contrast to the lack of local protection in the lung by parenteral immunization despite slightly improved protection in the spleen. Our study thus provides the first evidence that respiratory mucosal DC immunization results in effective mucosal T cell responses and improved host resistance against an intracellular pathogen. We admit that compared with 0.7- to 0.8-log reduction of bacterial burden in the lung by a live organism-based BCG vaccine, a standard control for TB vaccine evaluation used in this study, respiratory mucosal DC immunization led to a smaller reduction in bacterial burden (0.5 log). Although further studies are warranted to improve its efficacy, DC-based vaccine is safer and may be potentially used for immunizing HIV-infected hosts to which BCG vaccine is unsafe.

Considering repeated immunization is a common strategy to boost pre-existing T cell responses, we further evaluated the expansion of Ag-specific T cells in the airway lumen and spleen following either parenteral or mucosal prime-mucosal boost immunization. Mucosal boosting efficiently boosted Ag-specific tetramer⁺ CD8 T cells as well as IFN--secreting CD4 T cells in the airway lumen after either parenteral or mucosal DC prime immunization. However, we found that mucosal DC boosting either did not further expand the number of IFN--secreting CD8 T cells by i.m. DC priming or even reduce such activated CD8 T cells following i.n. DC priming. A similar phenomenon was also observed in the lung. Thus, the lack of further enhanced CD8 T cell activation by mucosal DC boosting represents a mechanism underlying its inability to further enhance immune protection offered by single mucosal DC immunization. We have recently documented a critical role by airway luminal CD8 T cells in immune protection against respiratory intracellular bacterial pathogens (14, 22). Although at this point, it still remains to be fully understood as to why i.n. DC boosting fails to enhance CD8 T cell activation as judged by Ag-stimulated CD8 T cell IFN- responses, it is possible that repeated DC immunization causes a shift of airway luminal CD8 T cells from a predominantly effector phenotype to a memory pool. This speculation is supported by our finding that although DC boosting increases tetramer⁺ CD8 T cells, it does not enhance or even reduce CD8 IFN- producers in the airway lumen. We further observed that repeated DC immunization does not increase the number of regulatory CD4⁺FoxP3⁺ or IL-10-secreting T cells (data not shown). These findings together suggest that many factors regulate secondary T cell responses and that pre-existing T cell immunity can modulate the effectiveness of boosting vaccine likely by altering APC Ag expression and T cell-activating capacities (29, 30). In recent years repeated DC immunization strategies have been attempted for cancer immunotherapy with only limited success (31, 32, 33, 34). Our current study not only provides mechanistic insights into such DC immunization modality but also highlights the need to further fine tune DC-based vaccination strategies.

Because the immunocompromised conditions, particularly HIV infection, are often associated with seriously diminished DC number and function (35, 36, 37, 38, 39), immunization with viral or bacterial vectored vaccines will not be an effective strategy. However, DC-based vaccines may be catered in vitro, before in vivo immunization, to potentially overcome the impaired function of the mucosal immune system in such hosts by delivering target Ags and costimulation signals for T cell activation. Nonetheless, such a strategy may still offer only a limited benefit if the endogenous DCs are heavily involved in vaccine DC-mediated immune activation. On the basis of this consideration, our current study investigated whether virally transduced vaccine DCs were still able to activate CD8⁺ and CD4⁺ T cells independently of endogenous APC populations in the lung and draining LNs. We observed that mucosal immunization with MHC-deficient vaccine DCs failed to lead to detectable CD8⁺ and CD4⁺ T cell activation. This observation suggests that mucosally delivered vaccine DCs are the primary driver of T cell priming and that endogenous APC populations, even if they do acquire soluble Ags released from wt vaccine DCs or the fragments of apoptotic/necrotic vaccine DCs, play a minor role in T cell priming. That mucosally delivered vaccine DCs activate T cells independently of endogenous APCs contrasts the previously observed significant contribution by endogenous APCs following parenteral DC immunization (23, 24, 40, 41, 42).

Following a similar line of clinical consideration, we investigated whether the vaccine DCs with some level of immunodeficiency, i.e., cytokine deficiency, could still go on to activate T cells after mucosal administration. We found that although the vaccine DCs deficient in type 1 cytokine IL-12 failed to migrate to the local draining LNs, consistent with a previous study (25), surprisingly both CD8⁺ and CD4⁺ T cell responses remained unimpaired. As the failure of IL-12^–/– DCs to migrate to LNs resulted in impaired T cell activation as previously shown (25), our current study draws a different conclusion that migration of DCs to LNs is not required for T cell activation. Such a discrepancy might be due to the fact that we analyzed directly the activation of endogenous naturally generated T cells whereas adoptively transferred OVA-specific transgenic T cells were analyzed in the previous report (25). The fact that we demonstrated the presence of only a small portion (1%) of transferred vaccine DCs in LNs, in keeping with our previous study (43) and the presence of a significant portion of these APCs in the lung mucosal tissue suggests that vaccine DC-mediated T cell activation might have occurred in lung mucosal lymphoid tissues outside of the LNs, thus lending further support to the conclusion from some previous studies by others. However, our study for the first time derives such conclusion from a wt host with intact draining LNs, different from the previous studies conducted in the mice lacking draining LNs (44, 45). Furthermore, considering the importance of IL-12 in the development of Th1 development (26, 46), our current finding with IL-12^–/– vaccine DCs also suggests that this cytokine may be provided by bystander cells in the respiratory mucosa for T cell activation.

In summary, our study has established that respiratory mucosal delivery of genetically modified vaccine DCs is an effective immunization modality for the generation of antimicrobial T cell immunity. We report critical findings that mucosally delivered vaccine DCs can activate both CD8 and CD4 T cells independent of endogenous DCs and vaccine DC-derived IL-12. These features, together with their efficiency in transmigrating to the draining LNs as well as mucosal tissue, support its great promise for immunization in immunocompromised hosts.

	Acknowledgments

 
The authors acknowledge the invaluable technical assistance of Anna Zganiacz, Duncan Chong, Xueya Feng, and Elizabeth Roediger; and the kind provision of anti-Ag85A mAb by Dr. Kris Huygen and of M. tuberculosis Ags by TB Research Center, Colorado State University.

	Disclosures

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 
The authors have no financial conflict of interest.

	Footnotes

 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ This study is supported by funds from the Canadian Institutes for Health Research.

² Address correspondence and reprint requests to Dr. Zhou Xing, Room 4012-MDCL, Department of Pathology and Molecular Medicine, McMaster University, 1200 Main Street West, Hamilton, Ontario, Canada L8N 3Z5. E-mail address: xingz{at}mcmaster.ca

³ Abbreviations used in this paper: DC, dendritic cell; BCG, Bacillus Calmette-Guérin; i.n., intranasal; MHCII, MHC class II; MHCI, MHC class I; BAL, bronchoalveolar lavage; LN, lymph node; cLN, cervical LN; mLN, mediastinal LN; pLN, popliteal LN; i.t., intratracheal; ICCS, intracellular cytokine staining; wt, wild type.

Received for publication September 12, 2007. Accepted for publication June 10, 2008.

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Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 

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