INFECTION AND IMMUNITY, May 2007, p. 2260–2268
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
Vol. 75, No. 5
Modulation of Naive CD4?T-Cell Responses to an Airway Antigen
during Pulmonary Mycobacterial Infection?
Mursalin M. Anis,1,2Scott A. Fulton,2Scott M. Reba,2
Clifford V. Harding,1† and W. Henry Boom2,3†*
Department of Pathology,1Division of Infectious Diseases,2and Tuberculosis Research Unit,3Case Western Reserve University and
University Hospitals of Cleveland, Cleveland, Ohio
Received 25 October 2006/Returned for modification 28 November 2006/Accepted 2 February 2007
During pulmonary mycobacterial infection, there is increased trafficking of dendritic cells from the lungs to
the draining lymph nodes. We hypothesized that ongoing mycobacterial infection would modulate recruitment
and activation of antigen-specific naive CD4?T cells after airway antigen challenge. BALB/c mice were infected
by aerosol with Mycobacterium bovis BCG. At peak bacterial burden in the lungs (4 to 6 weeks postinfection),
carboxy-fluorescein diacetate succinimidyl ester-labeled naive ovalbumin-specific DO11.10 T cells were adop-
tively transferred into infected and uninfected mice. Recipient mice were challenged intranasally with soluble
ovalbumin (OVA), and OVA-specific T-cell responses were measured in the lungs, draining mediastinal lymph
nodes (MLN), and spleens. OVA challenge resulted in increased activation and proliferation of OVA-specific
T cells in the draining MLN of both infected and uninfected mice. However, only BCG-infected mice had
prominent OVA-specific T-cell activation, proliferation, and Th1 differentiation in the lungs. BCG infection
caused greater distribution of airway OVA to pulmonary dendritic cells and enhanced presentation of OVA
peptide by lung CD11c?cells. Together, these data suggest that an existing pulmonary mycobacterial infection
alters the phenotype of lung dendritic cells so that they can activate antigen-specific naive CD4?T cells in the
lungs in response to airway antigen challenge.
The lungs constantly come into contact with airborne par-
ticulates and pathogens. The interaction between inhaled par-
ticulate antigens, alveolar macrophages, and pulmonary den-
dritic cells (DCs) sets the stage for antigen-specific pulmonary
immune responses (3). In the absence of microbial stimuli,
immune responses in the lungs to soluble antigens are charac-
terized by either tolerance or Th2-like responses as seen in
murine models of asthma (29, 40). In contrast, acute viral
infections give rise to vigorous pulmonary T-cell responses
characterized by Th1 cytokine secretion (23, 32). These studies
have provided valuable insights into the dynamic role of the
pulmonary environment to discriminate airborne insults and
generate appropriate T-cell responses. However, few studies
have looked at the initiation of naive T-cell responses in the
lungs during prolonged mycobacterial infection.
Mycobacterium tuberculosis establishes a latent infection in
the vast majority of immunocompetent individuals. Upon in-
halation the mycobacteria are phagocytosed principally by al-
veolar macrophages. M. tuberculosis circumvents phagosomal
maturation and establishes a niche for intracellular survival
(14). In the ensuing adaptive response mycobacteria are con-
tained in granulomas within which M. tuberculosis persists la-
tently (6). The modulation of pulmonary immunity that per-
mits latency to develop is poorly understood. Mycobacterium
bovis bacillus Calmette-Guerin (BCG) is used as a vaccine to
prevent disseminated tuberculosis in children; BCG has been
used as a model organism to study the innate and adaptive
immune response to M. tuberculosis (15, 21, 22). After aerosol
BCG infection, pulmonary immune responses and bacterial
growth peak 4 to 6 weeks later, followed by gradual clearance
of BCG from the lungs (11, 12, 21).
It is thought that activation of naive CD4?T cells, in re-
sponse to airway antigens, occurs primarily in the mediastinal
lymph nodes (MLN) draining the lungs (8, 45). During pulmo-
nary influenza infections, virus-specific naive T cells divide in
the MLN, and only the most differentiated cells express the
appropriate adhesion molecules to migrate to the lungs (32).
In the lungs, differentiated, effector T cells colocalize with
antigen-carrying pulmonary DCs (4). However, recent evi-
dence suggests that primary activation of naive T cells can
occur in the lungs (24, 34). Mice lacking functional CCL19 and
CCL21 and mice lacking fucosyltransferases have impaired
localization of naive T cells to secondary lymphoid organs.
However, these mice are able to initiate naive T-cell responses
in the lungs against pulmonary pathogens. Pulmonary infection
may play a role in the apparent shift from draining lymph node
to the lung in priming of naive CD4?T cells. Few studies have
addressed this issue in their detailed analysis of naive CD4?
T-cell responses (16, 27, 44, 45).
In this report, we have used an adoptive-transfer technique
(19) to artificially increase the precursor frequencies of ovalbu-
min (OVA)-specific naive T cells in recipient mice that had
been previously infected with aerosolized BCG. Using flow
cytometry to track OVA-specific (KJ?) T cells, we found that
the naive KJ?T-cell response to intranasal OVA was localized
to the lungs and draining MLN. Both infected and uninfected
mice mounted vigorous OVA-specific T-cell responses in the
MLN, but only BCG-infected animals had marked activation,
* Corresponding author. Mailing address: Division of Infectious
Diseases, Biomedical Research Building, 1031, Case Western Reserve
University, 10900 Euclid Ave., Cleveland, OH 44106-4984. Phone:
(216) 368-4844. Fax: (216) 368-2034. E-mail: email@example.com.
† W.H.B. and C.V.H. share senior authorship.
?Published ahead of print on 12 February 2007.
proliferation, and differentiation of KJ?T cells in the lungs.
Infection caused greater distribution of OVA to pulmonary
DCs and enhanced presentation of OVA peptide by lung
CD11c?cells that led to local lung-resident KJ?T-cell activa-
tion and proliferation in vivo.
MATERIALS AND METHODS
Mice. Eight- to ten-week-old female BALB/c mice were purchased from The
Jackson Laboratory (Bar Harbor, ME). DO11.10 T-cell-receptor (TCR) trans-
genic mice that express TCRs specific for OVA323-339peptide presented in the
context of I-Ad(31) were a gift from Alan Levine (Case Western Reserve
University, Cleveland, OH). Mice were housed in specific-pathogen-free condi-
tions. All studies were approved by the Institutional Animal Care and Use
Committee at Case Western Reserve University.
Aerosol BCG infection. BALB/c mice were exposed to aerosol M. bovis BCG
in an inhalation exposure system (Glas Col, Terre Haute, IN) as previously
described (21). Day 1 colony counts consistently gave 3,200 ? 1,300 CFU
per mouse. Bacterial growth in the lungs peaked 4 to 6 wks afterwards with
190,000 ? 70,000 CFU per mouse. Bacterial growth in the lung-draining MLN
was determined to be 4,200 ? 1,300 CFU per mouse at 28 days after infection.
Infected mice were used as recipients in adoptive-transfer experiments 4 to 6
weeks postinfection. Uninfected mice in all of the experiments were not mock
Endotoxin depletion of OVA. Endotoxin was removed from OVA (Sigma-
Aldrich) by using the protocol of Aida and Pabst (1) with minor modifications.
OVA was dissolved in filtered, lipopolysaccharide-free water, and Triton-X-114
was added to yield a final concentration of 1% Triton X-114 in OVA solution.
The solution was chilled on ice for 10 min and then agitated gently at 4°C for 20
min. The solution was then warmed to 37°C for 10 min and spun at 20,000 ? g
for 20 min. The detergent phase was aspirated off, and the aqueous phase
containing OVA was subjected to seven more extractions with Triton X-114. The
endotoxin contamination was ?0.1 ng/ml, as determined by a Limulus amoebo-
cyte lysate assay (BioWhittaker).
DO11.10 T-cell isolation. Splenocytes from 9- to 14-week-old DO11.10 mice
were isolated, and red blood cells were lysed in hypotonic lysis buffer (10 mM
Tris-HCl and 0.83% ammonium chloride). The cells were plated in 100-mm petri
dishes and allowed to adhere for 1 h at 37°C. Nonadherent splenocytes were then
used to obtain untouched CD4?T cells by using the CD4?T-cell negative
selection kit (Miltenyi Biotec) according to the manufacturer’s instruction. In
most experiments, the resulting CD4?T cells were subsequently stained with
anti-CD62L and anti-CD44 monoclonal antibodies (MAbs) and fluorescence-
activated cell sorted (FACsorted) by gating on naive (CD62LhiCD44low) T cells
by using a BD Aria cell sorter. Purified CD4?T cells and flow-sorted naive
CD4?T cells were then used in adoptive-transfer experiments. FACsorted
CD4?T cells were ?98% CD44lowCD62Lhi, and 65 to 75% of these naive CD4?
T cells were OVA specific (KJ?).
Adoptive transfer and airway OVA challenge. Uninfected BALB/c mice and
BCG-infected mice were anesthetized intraperitoneally with a nonlethal dose of
tribromoethanol (180 mg/kg) and given 3 ? 106to 5 ? 106DO11.10 CD4?T
cells by retro-orbital injection. In some experiments DO11.10 T cells were la-
beled with 5 ?M carboxy-fluorescein diacetate succinimidyl ester (CFSE) (In-
vitrogen) for 10 min at 37°C in 0.1% bovine serum albumin (BSA)–phosphate-
buffered saline (PBS) and then washed three times in ice-cold PBS before
adoptive transfer in normal saline. Mice were allowed to rest for 2 days before
being challenged intranasally on day 0 with 500 ?g of endotoxin-depleted OVA
or BSA as the control antigen. On day 2, mice were challenged intranasally once
more with 500 ?g of endotoxin-depleted OVA, while control mice were not given
BSA again. On day 3, 5 days after DO11.10 CD4?T-cell transfer, the mice were
sacrificed and their spleens, lungs, draining MLN, and bronchoalveolar lavage
fluid (BALF) were collected.
Tissue isolation. For experiments involving CFSE, care was taken to minimize
exposure to light. Tissues were harvested and processed as previously described
(21). Briefly, mice were anesthetized with a lethal dose of tribromoethanol (240
mg/kg). For each animal, the abdominal cavity was incised, the spleen was
harvested, and the mouse was exsanguinated. The trachea was cannulated, and
the BALF was collected by three aspirations with 1 ml of PBS. Lungs were
perfused with 10 ml of PBS and harvested. The draining MLN were then har-
Spleens were homogenized and pressed through a 70-?m-pore-size nylon
filter. Red blood cells were lysed in red blood cell lysis buffer. Single cells were
resuspended in complete medium (Dulbecco modified Eagle medium, 10% fetal
bovine serum [FBS], 0.05 mM 2-mercaptoethanol, 2 mM HEPES, 1 mM sodium
pyruvate, 100 mM nonessential amino acids, 100 U of penicillin/ml, and 0.1 mg
of streptomycin/ml). Lungs were minced and digested with 125 U of type IV
collagenase and 30 U of DNase/ml for 90 min at 37°C. Lung aggregates were
drawn through a 18-gauge needle three times before being pressed through a
40-?m-pore-size nylon filter. The red blood cells were lysed, and the lungs were
resuspended in RPMI. Serial dilutions of lung suspension were plated onto 7H10
plates to determine the bacterial CFU counts. MLN were pressed through a
70-?m-pore-size nylon filter using the plunger of a 1-ml syringe and then resus-
pended in RPMI.
Cell staining and percentage of OVA-specific T cells that divided. Single-cell
suspensions of tissues were counted. Viability of cells was assessed by trypan blue
exclusion. A total of 5 ? 105to 1 ? 106viable lung, MLN, and spleen cells were
preincubated in a 1% BSA-PBS solution of FcBlock (BD Pharmingen) for 15
min at 4°C. The cells were then stained with the DO11.10 TCR clonotypic
antibody biotinylated KJ 1-26 (Invitrogen catalog number MM7515-3), along
with activation and adhesion markers anti-CD62L, anti-CD44, and anti-CD69
(eBioscience catalog numbers 25-0621, 12-0441, and 25-0691, respectively) and
anti-CD25 (BD Pharmingen catalog number 553075), for 30 min at 4°C. Cells
were washed once with 1% BSA, resuspended in streptavidin-Pacific Blue con-
jugate (Invitrogen), and incubated for an additional 30 min at 4°C. The cells were
washed once again with 1% BSA, and the pellets were resuspended in 0.3 ml of
1% paraformaldehyde in PBS. Stained samples were acquired by using a BD
LSR II flow cytometer. Flow cytometry results were analyzed with FlowJo (Tree
Star, Inc.) software.
The percentage of OVA-specific T cells that divided was calculated by using
the method used to determine the responder frequency as previously described
(43). Briefly, the number of events in each daughter cell generation N, charac-
terized by dimmer CFSE labeling, was divided by 2Nto arrive at the number of
precursors or responders that gave rise to those daughters in generation N. The
number of undivided CFSEhighKJ?T cells was used, along with the sum of the
responders, to calculate the fraction of KJ?T cells that divided or responded
after OVA challenge.
Intracellular cytokine staining. Lung cells were stimulated for 5 h with phor-
bol myristate acetate (PMA) at 50 ng/ml and 1 ?g/ml of ionomycin (Sigma-
Aldrich) in the presence of 10 ?g/ml of brefeldin A (Sigma-Aldrich). The cells
were collected and surface stained with KJ 1-26 in the presence of mouse
FcBlock (BD Pharmingen) in 2% FBS in 1? PBS staining solution at room
temperature. Cells were fixed with 4% paraformaldehyde and stained with allo-
phycocyanin anti-IFN-? or anti-interleukin-4 (IL-4) MAbs (eBioscience) in the
presence of saponin for 30 min. Cells were fixed in 1% paraformaldehyde and
acquired within 24 h with a BD LSR II flow cytometer.
BrdU incorporation. Recipient mice were challenged with OVA or BSA. After
3 days, mice received i.v. bromodeoxyuridine (BrdU; 2 mg/mouse) (Sigma-Al-
drich) 1 h prior to sacrifice (30). Tissues were harvested and single-cell suspen-
sions made. Cells were surface stained with the DO11.10 T-cell receptor anti-
body, KJ 1-26, at room temperature in the presence of mouse FcBlock (BD
Pharmingen) in 2% FBS in 1? PBS staining solution and then fixed with 4%
paraformaldehyde. Cells were permeabilized with saponin for 30 min at room
temperature and incubated with 50 U DNase I at 37°C for 1 h. Digested cells
were stained with anti-BrdU MAb in saponin solution. Cells were fixed in 1%
paraformaldehyde and acquired within 24 h with a BD LSR II flow cytometer.
ELISPOT assay. Enzyme-linked immunospot (ELISPOT) assay for IFN-? was
done as previously described (21). Briefly, sterile ELISPOT plates (Whatman)
were precoated with anti-IFN-? capture antibody (BD Pharmingen catalog no.
551216) overnight at 4°C at a concentration of 5 ?g/ml. The plates were blocked
with 1% BSA in PBS for 1 h and washed with PBS before the lung, spleen, and
MLN cells from OVA-challenged, BCG-infected, and uninfected mice were
added at 5 ? 105and 1 ? 106cells/well. Some wells received exogenous OVA
peptide (OVA323-339; 2 ?M), and the cells were incubated for 48 h at 37°C. Plates
were washed four times with PBS containing 0.05% Tween 20 and incubated for
4 h at room temperature with biotinylated anti-IFN-? (BD Pharmingen catalog
no. 554410) at a concentration of 2 ?g/ml. Plates were washed four times, and
bound IFN-? was detected by using streptavidin-alkaline phosphatase according
to the manufacturer’s instructions (R&D Elispot Blue Color Module). Plates
were dried at room temperature, and the spots were counted and analyzed by
using an immunospot reader and software (CTL Analyzers, LLC, Cleveland,
OH). ELISPOT assay for IL-4 was done with a mouse IL-4 ELISPOT kit
according to the manufacturer’s instruction (eBioscience). The same cell num-
bers were plated as described above.
Fluos-OVA preparation and intranasal challenge. Fluos-OVA was prepared
by using a fluorescein labeling kit (Roche). Briefly, OVA (Sigma-Aldrich) was
dissolved in PBS to make a 10-mg/ml solution. Fluos was dissolved in dimethyl
VOL. 75, 2007ACTIVATION OF NAIVE CD4?T CELLS DURING TUBERCULOSIS2261
sulfoxide to make a 2-mg/ml solution. A 95-?l portion of Fluos was added to 2
ml of the OVA solution (10 mg/ml), followed by incubation at room temperature
for 2 h with gentle mixing in the dark. Unbound Fluos was separated by using
PD-10 columns (GE Healthcare). Then, 450 ?g of Fluos-OVA was introduced
intranasally into BCG-infected and uninfected mice. After 18 to 24 h the mice
were sacrificed, and their lungs and MLN were harvested. Single-cell suspensions
were stained with anti-CD11c and anti-CD11b (BD Pharmingen) and either
anti-I-Ad, anti-CD80, or anti-CD86 (BD Pharmingen catalog numbers 553546,
553769, and 553691, respectively). The cells were fixed in 1% paraformaldehyde
and acquired by using a BD LSR II flow cytometer.
OVA peptide presentation by lung CD11c?cells. BCG-infected and unin-
fected mice were sacrificed, and their BALF, lungs, and MLN were harvested.
Lung cells were positively sorted for CD11c?cells by using N418 microbeads
(Miltenyi Biotec). CD11c?lung cells were plated in 96-well round-bottom plates
at various cell densities. Exogenous OVA323-339(2 ?M) was added to the wells,
along with 105DOBW T-cell hybridoma cells, which recognize OVA323-339–I-Ad
complexes and secrete IL-2. After 18 to 24 h, 100-?l portions of the supernatants
were collected from these wells, and the supernatants were assayed for IL-2 by
enzyme-linked immunosorbent assay (ELISA). Briefly, Immulon microtiter
plates (Thermo) were precoated overnight at 4°C with anti-IL-2 capture antibody
(eBioscience catalog no. 14-7022) at 1 ?g/ml. Plates were washed with PBS-
Tween and blocked with 10% FBS-PBS for 1 h at 37°C. Plates were incubated at
37°C for 2 h with supernatants from lung CD11c?/DOBW cell cultures. After a
washing step, the plates were incubated at room temperature with biotin-conju-
gated anti-IL-2 detection antibody (eBioscience catalog no. 13-7021) at 1 ?g/ml.
The plates were washed and incubated with avidin-alkaline phosphatase at room
temperature for 30 min. Substrate was added, and the plates were read after 20
to 30 min.
Statistical analysis. All statistical analyses were performed by using a one-
tailed Student t test. A P value of ?0.05 was considered statistically significant.
BCG infection causes enrichment and accumulation of
OVA-specific T cells in lungs and draining lymph nodes after
airway OVA challenge. CD4?T cells from DO11.10 OVA-
specific TCR transgenic mice were adoptively transferred into
uninfected BALB/c mice and mice that had been infected 4 to
6 weeks earlier. Recipient mice were challenged intranasally
on days 0 and 2 with either soluble endotoxin-depleted OVA
or BSA as a control antigen. After 3 days of OVA challenge,
OVA-specific T cells were identified with the clonotypic MAb
KJ 1-26 in the MLN, lung, and spleen. BCG-infected mice had
increased enrichment for KJ?T cells, i.e., a higher percentage
of KJ?T cells among CD4?T cells, compared to uninfected
mice challenged with OVA (P ? 0.02) (Fig. 1A). The enrich-
ment of KJ?T cells was antigen dependent because challenge
of infected and uninfected mice with an unrelated antigen,
BSA, did not result in an increase in the percentage of KJ?T
cells. In addition, the increase in the percentage of KJ?T cells
was found only in the lungs and MLN and not in the spleens of
mice challenged with intranasal OVA, indicating that the elic-
ited KJ?T-cell response was primarily localized to where the
antigen was delivered. The difference between the BCG?OVA
and the OVA groups was more pronounced when the total
number of KJ?T cells in the lungs and MLN was quantified
(P ? 0.004) (Fig. 1B). Thus, pulmonary inflammation induced
by BCG infection resulted in increased numbers of KJ?T cells
in the lung parenchyma and MLN after airway OVA challenge.
BCG infection enhances OVA-specific naive T-cell activa-
tion in the lungs after airway OVA challenge. The enhanced
accumulation of OVA-specific T cells in the lungs and MLN of
BCG-infected, OVA-challenged mice could be due to in-
creased recruitment, enhanced T-cell activation and prolifera-
tion, or both. To determine the contribution of T-cell activa-
tion, we adoptively transferred FACsorted naive (CD62Lhi
CD44low) CD4?T cells from DO11.10 mice into BCG-infected
and uninfected BALB/c mice and challenged them intranasally
with OVA. The expression of markers associated with T-lym-
phocyte activation (CD69 and CD25) on OVA-specific T cells
in the lungs and draining MLN was analyzed after OVA chal-
lenge (Fig. 2). The baseline levels of CD25 and CD69 on naive
KJ?T cells before adoptive transfer were not greater than
those of the isotype controls (Fig. 2A). The early T-cell acti-
vation marker CD69 remained highly expressed on KJ?T cells
present in the MLN of both BCG?OVA and OVA mice 24 h
after the last of two intranasal OVA exposures (Fig. 2A). The
rapid kinetics of CD69 expression after T-cell activation (27)
indicated that KJ?T cells were activated by OVA-presenting
antigen-presenting cells (APCs) in the draining MLN of both
BCG?OVA and OVA mice. In contrast, infected mice had a
higher percentage of KJ?T cells expressing CD69 in the lungs
than did uninfected mice given OVA (Fig. 2B and C). There
was no difference in the frequency of CD25-expressing KJ?T
cells in the lungs and MLN of mice between the two groups
(Fig. 2C). Generally, CD25, the high-affinity IL-2R alpha
chain, is upregulated later than CD69 and remains expressed
on the surface longer upon T-cell activation (27). The CD25?
CD69?phenotype of KJ?T cells in the lungs of infected mice
FIG. 1. BCG infection causes enrichment and accumulation of
OVA-specific T cells after airway OVA challenge. BALB/c mice were
infected with BCG, and 4 to 6 weeks later 3 ? 106to 5 ? 106CD4?
DO11.10 T cells were transferred into BCG-infected and uninfected
mice. Recipient mice were challenged with 500 ?g of endotoxin-de-
pleted OVA on days 0 and 2 or with BSA on day 0 and were sacrificed
on day 3. Single-cell suspensions were prepared from lungs, MLN, and
spleens and stained with anti-CD4 and KJ 1-26 to enumerate the
OVA-specific T cells. (A) The percentage of OVA-specific (KJ?) T
cells among the CD4?T cells present was determined in the MLN,
lungs, and spleens of the four different groups. (B) Total KJ?T-cell
numbers were calculated by multiplying viable cell counts, determined
by trypan blue exclusion, by the percentage of KJ?T cells among
viable cells gated from a forward-scatter (FSC) versus side-scatter
(SSC) plot during fluorescence-activated cell sorting analysis. This
experiment was repeated twice with similar findings. Three mice were
included in each group.*, P ? 0.02;**, P ? 0.01; #, P ? 0.004; ##,
P ? 0.02.
2262 ANIS ET AL.INFECT. IMMUN.
suggested recent T-cell activation. Thus, airway OVA chal-
lenge causes naive KJ?T-cell activation in the MLN of both
infected and uninfected mice but, during pulmonary BCG in-
fection, activation of naive KJ?T cells also occurs in the lungs.
BCG infection increases responder frequency of antigen-
specific T cells after airway antigen challenge. To determine
whether differences in activation markers measured in infected
and uninfected mice also resulted in differences in T-cell pro-
liferation, CFSE-labeled CD4?OVA-specific T cells were
transferred into infected and uninfected mice. At 3 days after
OVA challenge, CFSE dilution of KJ?T cells in lungs, MLN,
and spleens was used as a measure of T-cell proliferation in
vivo (25). CFSE dye dilution in KJ?T cells was antigen specific
because no KJ?T-cell division was observed in animals chal-
lenged with BSA (Fig. 3A). In draining MLN and lungs, in-
fected and OVA-challenged mice (BCG?OVA) had greater
numbers of divided KJ?T cells in each generation of daughter
cells compared to uninfected OVA challenged mice, even
though in both groups KJ?T cells underwent approximately
six cell divisions (Fig. 3A). Proliferation intermediates, corre-
sponding to the first few cell divisions, were prominent in the
MLN but not in the lungs of both OVA-challenged groups.
This suggested that priming of naive CD4?T cells occurs
primarily in the MLN but that KJ?T-cell activation can also
occur in infected lungs (Fig. 2B). Very little KJ?T-cell division
was apparent in the spleens, demonstrating again that the
antigen-specific T-cell response to an airway antigen was pri-
marily localized to the lungs and draining lymph nodes. We
calculated the percentage of OVA-specific T cells that divided by
using methods to derive responder frequencies (43) (Fig. 3B).
BCG?OVA group of mice had higher responder frequencies of
KJ?T cells than OVA-challenged mice. This difference was less
apparent in the MLN than the twofold difference observed in the
lungs (Fig. 3B). Enhanced CFSE dilution of KJ?T cells in the
lungs of BCG-infected mice could be due to (i) increased recruit-
ment to the lungs of KJ?T cells that had divided elsewhere or (ii)
local KJ?T-cell proliferation in the lungs.
BCG infection results in antigen-specific T-cell proliferation
in the lungs upon airway antigen challenge. To further deter-
mine the contribution of in situ pulmonary KJ?T-cell prolif-
eration in the accumulation of proliferating KJ?T cells in the
lungs (Fig. 3), we adoptively transferred naive (CD44low
CD62Lhi) KJ?T cells into recipient mice and challenged the
mice with 300 ?g of endotoxin-depleted OVA. After 3 days of
OVA challenge, recipient mice received 2 mg of BrdU intra-
venously 1 h prior to sacrifice (30). The short pulse allowed us
to measure T-cell division in situ with minimal chance for
migration. After this short BrdU pulse, BrdU?OVA-specific T
cells were found in the lungs of BCG-infected mice challenged
intranasally with OVA (Fig. 4B). In uninfected mice chal-
lenged with OVA, fewer BrdU?KJ?T cells were found in the
lungs (P ? 0.05). In animals infected with BCG and challenged
with BSA, no BrdU?KJ?T cells were detected. A small
percentage of BrdU?KJ?events within the lungs was due to
the short BrdU pulse. We did not detect BrdU?KJ?events in
the lungs earlier than 3 days after OVA challenge even in
BCG?OVA mice. BrdU incorporation was measured in drain-
ing MLN 2 days after OVA challenge in both BCG?OVA and
OVA mice (data not shown). Thus, BCG infection causes in
situ proliferation of antigen-specific T cells in the lungs after
airway antigen challenge.
BCG infection increases the frequency of antigen-specific
effector T cells that secrete IFN-? in the lungs of mice chal-
lenged with airway antigen. To determine whether pulmonary
BCG infection increased the differentiation of naive KJ?T
cells to effector T cells, flow-sorted naive (CD44lowCD62Lhi)
OVA-specific T cells were transferred into naive and BCG-
infected mice. Recipient mice were challenged with OVA as
described in Materials and Methods. We first determined the
fraction of divided KJ?T cells (CFSElow) in the lungs of
infected and uninfected OVA-challenged mice that attained an
effector phenotype characterized by loss of L-selectin,
CD62Llow(Fig. 5A). L-selectin is a lymph node homing mol-
ecule present on naive and central memory T cells but absent
on effector T cells (20, 38). Among KJ?T cells that had
divided more than two times (CFSElow), 49% were CD62Llow
in the lungs of BCG?OVA mice, whereas only 23% were
FIG. 2. BCG infection enhances T-cell activation in the lungs.
CD4?CD62LhiCD44lowFACsorted naive DO11.10 T cells were trans-
ferred into BCG-infected and uninfected mice. Mice were challenged
with OVA as described in Materials and Methods. Three days later,
the mice were sacrificed, and the draining MLN (A) and lungs
(B) were stained for TCR, CD25, and CD69. Histograms were gated
on KJ?T cells. The baseline levels of CD25 and CD69 on naive KJ?
T cells before adoptive transfer are shown in panel A. (C) Percentages
of CD25- and CD69-expressing KJ?T cells, above isotype staining,
from three mice per group.*, P ? 0.01. The data are representative of
three separate experiments.
VOL. 75, 2007 ACTIVATION OF NAIVE CD4?T CELLS DURING TUBERCULOSIS2263
CD62Llowin the lungs of uninfected OVA-challenged mice. In
contrast, 45% of CFSElowKJ?T cells were CD62Llowin the
MLN of uninfected mice, whereas 56% of CFSElowKJ?T cells
were CD62Llowin infected mice. The percentage of CD62Llow
KJ?T cells was greater in the MLN and lungs of infected mice
than in uninfected mice (Fig. 5B). Since effector T cells pref-
erentially migrate to sites of inflammation (38), the differences
between the two groups were magnified in BCG-infected
lungs. Thus, infection causes increased differentiation of acti-
vated T cells into effector cells, and this is more prominently
observed in the lungs than in the draining lymph nodes.
Next, to determine whether there are differences in cytokine
production by OVA-specific effector T cells from lungs of
BCG?OVA and OVA mice, we stimulated lung cells from
both groups of mice with PMA and ionomycin and measured
the intracellular IFN-? and IL-4 production. BCG infection
increased the frequency of IFN-?-producing KJ?T cells
among OVA-specific T cells present in the lungs of OVA-
challenged mice (Fig. 6A). We did not detect any IL-4-produc-
ing KJ?T cells in either group of mice by intracellular cytokine
staining (data not shown). However, stimulating lung T cells
with exogenous OVA peptide allowed detection of IL-4-pro-
ducing cells by ELISPOT assay. As shown in Fig. 6B, similar
numbers of IL-4 spot-forming units (SFU) were found between
infected and uninfected mice, whereas more IFN-? SFU were
observed in infected and OVA-challenged mice. This demon-
strates a Th1 effector phenotype of OVA-specific T cells in the
lungs of infected mice.
Lung CD11c?cells harbor airway-derived OVA and present
more OVA peptide in BCG-infected mice than in uninfected
mice. One possible explanation behind the increased activa-
tion, proliferation, and differentiation of antigen-specific T
FIG. 3. Pulmonary infection increases the frequency of antigen-specific T cells responding to an airway antigen. A total of 5 ? 106CFSE-
labeled CD4?DO11.10 T cells were transferred into BCG-infected and uninfected recipient mice. Recipient mice were challenged with 500 ?g
of endotoxin-depleted OVA on days 0 and 2 or with BSA on day 0. Mice were sacrificed on day 3, and single-cell suspensions from tissues were
stained and analyzed by flow cytometry to measure T-cell proliferation. (A) Representative dot plots showing KJ?staining and CFSE dye dilution
of four individual mice from the four groups. (B) The percentage of OVA-specific T cells that divided in each mouse was calculated as described
in Materials and Methods. This experiment was repeated twice with similar results (three mice per group).*, P ? 0.02;**, P ? 0.03.
FIG. 4. BCG infection induces OVA-specific T-cell proliferation in the lungs after airway OVA challenge. FACsorted naive (CD62Lhi
CD44low) CD4?T cells from DO11.10 mice were transferred into BCG-infected and uninfected recipient mice. Recipient mice were challenged
intranasally with 300 ?g of endotoxin-depleted OVA or BSA. At 3 days after challenge, mice were injected intravenously with 2 mg of BrdU. After
1 h of in vivo BrdU pulse, the mice were sacrificed, and single cell suspensions from the lungs were stained with KJ 1-26 and anti-BrdU as described
in Materials and Methods. (A) Isotype staining for anti-BrdU and KJ isotype control. (B) Representative dot plots of three mice from the three
groups. The numbers represent the percentage of BrdU?cells among the KJ?T cells. (C) Means ? the standard deviation of three mice per group.
*, P ? 0.01. The experiment was repeated twice with similar findings each time.
2264 ANIS ET AL.INFECT. IMMUN.
cells in the lungs of infected mice is activation of the pulmo-
nary innate immune cells responsible for initiating adaptive
responses. CD11c?lung DCs are efficient at presenting airway
antigens to both naive and effector T cells (4). To determine
whether CD11c?lung cells had a role in the increased prolif-
eration and differentiation of KJ?T cells in the lungs of
BCG?OVA mice, BCG-infected and uninfected mice were
given Fluos-labeled OVA. After 18 to 24 h, the lungs were
harvested and stained for phenotypic and maturation markers
of DCs. Almost all of the Fluos-OVA was sequestered in the
CD11c?lung cell population (Fig. 7A). This is in agreement
with other published findings where intranasally administered
antigens had been tracked in vivo (7, 42).
Thus, to determine whether infected and uninfected mice
differed in distribution of Fluos-OVA within the CD11c?lung
cell population, we gated on Fluos?CD11c?cells and mea-
sured the DC markers CD11b and CD11c (13). In BCG-in-
fected mice, 19% of cells harboring Fluos-OVA were DCs
(CD11bhiCD11chi), whereas in uninfected mice 8.2% of
Fluos?cells were DCs (Fig. 7B). There were greater numbers
of CD11bhiCD11chiDCs in the lungs of BCG-infected mice
(5%) than in uninfected mice (1%), and this may account for
more distribution of OVA to the DCs in infected mice (data
not shown). Greater numbers of Fluos-OVA-containing DCs
in infected mice could result in activation of more OVA-spe-
cific naive T cells (26).
The maturation of lung DCs in infected lungs is demon-
strated by high levels of major histocompatibility complex class
II (MHC-II; I-Ad) expression (Fig. 7C). Expression levels of
the costimulatory molecules CD80 and CD86 were marginally
elevated on DCs harboring OVA from infected mice versus
uninfected mice (data not shown). Upon infection, greater
numbers of CD11c?MHC-IIhicells acquired intranasal anti-
gen and accumulated in the lungs (Fig. 7D). To determine
whether increased expression of I-Adresulted in greater pre-
sentation of pOVA–I-Adcomplexes to KJ?T cells in infected
lungs, CD11c? lung cells were purified from both infected
and uninfected groups of mice using CD11c?magnetic beads.
Exogenous OVA peptide presentation by these CD11c?lung
FIG. 5. BCG infection increases the accumulation of effector anti-
gen-specific T cells in the lungs after airway antigen challenge. Flow-
sorted CD62LhiCD44lownaive DO11.10 T cells were labeled with
CFSE and transferred into BCG-infected and uninfected recipient
mice. Recipient mice were challenged with OVA as described in Ma-
terials and Methods. Mice were sacrificed 3 days after OVA challenge,
and single cell lung suspensions were prepared. (A) Lung cells were
stained with KJ 1-26 and anti-CD62L and analyzed by flow cytometry.
Plots were gated on KJ?T cells, and the expression of CD62L and
CFSE dilution was analyzed. The numbers represent the percentage of
CD62LlowT cells among KJ?T cells that have divided more than twice
(CFSElow). (B) Percentage of CD62LlowT cells among KJ?T cells
(three mice per group).*, P ? 0.02;**, P ? 0.03. The data are
representative of two separate experiments.
FIG. 6. Mycobacterial infection induces naive OVA-specific T cells
to differentiate into Th1 effectors in response to airway OVA chal-
lenge. Flow-sorted CD62LhiCD44lownaive DO11.10 T cells were
labeled with CFSE and transferred into BCG-infected and uninfected
recipient mice. Recipient mice were challenged with OVA as described
in Materials and Methods. Mice were sacrificed 3 days after OVA
challenge, and single cell lung suspensions were prepared. (A) Lung
cells were stimulated with PMA and ionomycin for 5 h in the presence
of brefeldin A. Cells were surface labeled with KJ 1-26, stained for
intracellular IFN-?, and analyzed by flow cytometry. Histograms were
gated on KJ?T cells. The isotype for anti-IFN-? is shown. The num-
bers represent the means ? the standard deviation of triplicate wells
for each group.*, P ? 0.03. (B) A total of 5 ? 105lung cells were
cultured for 48 h at 37°C with or without 2 ?M OVA323-339peptide.
The IL-4 and IFN-? SFU were quantified by ELISPOT assay.**, P ?
0.01. The data are representative of three separate experiments.
VOL. 75, 2007ACTIVATION OF NAIVE CD4?T CELLS DURING TUBERCULOSIS 2265
cells was assessed by using the DOBW T-cell hybridoma that
recognizes OVA323-339, the same epitope recognized by KJ?T
cells. The hybridoma response was monitored by measuring
IL-2 in culture supernatants using ELISA. The addition of
exogenous OVA peptide confirmed that lung CD11c?cells
from infected mice had more functional MHC-II than cells
from uninfected mice, as determined by their efficiency in
presenting OVA peptide to DOBW T-cell hybridomas (Fig.
7E). Together, these data suggest that ongoing pulmonary
mycobacterial infection increases the number and maturation
of lung DCs such that they can activate antigen-specific naive
CD4?T cells in the lungs themselves.
We determined here the effect of prolonged, pulmonary
mycobacterial infection on the initiation of naive CD4?T-cell
responses. The following specific questions were addressed. (i)
Does BCG infection change the distribution of naive CD4?T
cells in MLN and lung? (ii) Does pulmonary inflammation
promote naive CD4?T-cell recruitment and activation in the
lung? (iii) Finally, does inflammation affect CD4?effector
T-cell development? Exploring these questions in light of re-
cent reports suggesting that infection affects how and where
naive T cells are activated in the lungs was of particular interest
(24, 30, 34). BCG-infected mice after airway-OVA challenge
had greater accumulation, activation, and proliferation of na-
ive OVA-specific CD4?T cells in the draining MLN than did
uninfected mice. Prolonged pulmonary inflammation also was
associated with naive CD4?T-cell activation in lung tissue. In
contrast, in uninfected mice there was little to no naive CD4?
T-cell activation and proliferation in the lungs. Pulmonary
BCG infection was associated with decreased expression of
L-selectin (CD62L) and increased production of IFN-? by
activated OVA-specific CD4?T cells in the lungs. Enhanced
CD4?T-cell activation in infected lungs was associated with
increased maturation of lung DCs and presentation of exoge-
nous OVA peptide by lung CD11c?cells.
Pulmonary BCG infection allowed measurement of the na-
ive CD4?T-cell responses to an unrelated airway antigen in
mice in the presence or absence of prolonged pulmonary in-
flammation. Pulmonary infection and inflammation peak 4 to 6
weeks after aerosolized BCG infection (21). Naive OVA-spe-
cific CD4?T cells were adoptively transferred into uninfected
and 4- to 6-week-BCG-infected mice, and then the animals
were challenged intranasally with endotoxin-depleted OVA.
Previous studies have examined the role of mycobacterial in-
fection of in vitro-generated APCs on naive T-cell activation in
vivo (2). Our experimental system allowed for the character-
ization of naive CD4?T-cell activation by endogenous lung
APCs during mycobacterial infection. In addition, we could
FIG. 7. BCG infection causes upregulation of MHC-II on lung
CD11c?cells harboring intranasal Fluos-labeled OVA. BCG-infected
and uninfected mice were given 450 ?g of Fluos-labeled OVA intra-
nasally. After 18 to 24 h the mice were sacrificed, and single-cell lung
suspensions were prepared and stained with anti-CD11c, anti-CD11b,
anti-I-Ad, anti-CD80, and anti-CD86. (A) FSC versus SSC plots were
gated on live cells, and dot plots of individual mice from three groups
are shown examining CD11c and Fluos. The percentages represent live
cells (FSC versus SSC gate) harboring Fluos-OVA (three mice per
group). (B) The plots were gated on Fluos?CD11c?cells. The per-
centages represent the distribution of Fluos-OVA to DCs (CD11chi
CD11bhi) within the CD11c?population (n ? 3).*, P ? 0.01. (C) His-
togram plots show the surface expression of MHC-II on the gated
populations shown in panel A containing lung CD11c?cells that har-
bor Fluos-OVA. The geometric mean fluorescence intensities of I-Ad
staining are 31,000 ? 5,300 and 1,700 ? 450 for the BCG?OVA and
OVA groups, respectively (n ? 3). P ? 0.03. (D) Total numbers of
Fluos?CD11c?cells expressing high I-Ad(mean fluorescence inten-
sity ? 3,000; n ? 3).**, P ? 0.04. (E) CD11c?cells were isolated from
the lungs of infected and uninfected mice and plated out at different
cell densities along with 2 ?M OVA peptide and 105DOBW cells. IL-2
in the culture supernatants was assayed by ELISA. Similar data were
obtained in two separate experiments.
2266 ANIS ET AL.INFECT. IMMUN.
control the amount of antigen and the precursor frequency of
transferred naive CD4?TCR transgenic T cells in infected and
uninfected mice. Mice were challenged with a high concentra-
tion of antigen to maximize access to antigen and minimize
clonal competition (5). Our experimental system specifically
addressed infection-induced pulmonary modulation of naive
CD4?T-cell priming without measuring pathogen-specific T-
cell responses. The latter will vary as pathogen burden and
pathogen-derived antigens change during chronic infection
(18, 23, 32, 35).
Pulmonary BCG infection increased the trafficking of CD4?
T cells of all antigen specificities to the MLN, as seen in other
infection models, but only T cells recognizing cognate antigens
remained and proliferated within the MLN (39). In vivo pro-
liferation as measured by CFSE demonstrated that infected
mice had a higher T-cell responder frequency than did unin-
fected mice. CD4?T-cell proliferation could occur elsewhere,
and pulmonary inflammation recruited activated T cells to the
MLN. However, the proliferation of OVA-specific T cells was
evident in the MLN but not in the lungs or spleens at earlier
time points (i.e., fewer than 3 days) after OVA challenge (data
not shown). Thus, a greater proliferation of OVA-specific T
cells in the MLN of infected mice was responsible for the
enhanced accumulation of KJ?T cells in the MLN.
Our results also suggest that the increased accumulation of
KJ?T cells in the lungs of BCG-infected mice was due to naive
CD4?T-cell activation and proliferation in the lungs. First,
BCG-infected mice had CD69?KJ?T cells in the lungs. CD69
expression by T cells has a role in inhibiting their egress from
lymphoid organs (36). The presence of CD69?KJ?T cells in
the lungs suggests that these cells may have been activated
without migration through the MLN. There were no CD69?
KJ?T cells in the lungs of uninfected mice even though both
infected and uninfected groups had comparable percentages of
CD69?KJ?T cells in the MLN. Second, short pulses of BrdU
in vivo demonstrated that KJ?T cells proliferated in the lungs
of infected mice. Evidence for KJ?T-cell activation and pro-
liferation in the lungs does not rule out that OVA-specific
naive T cells could have been activated elsewhere, migrated to
the lungs, and then re-encountered OVA there. However,
other researchers have shown that influenza virus infection
leads to formation of lymphoid aggregates, bronchus-associ-
ated lymphoid tissue, within the lungs of infected mice that
facilitate naive T-cell activation and proliferation in the ab-
sence of peripheral lymphoid organs (30). In addition, lungs
from M. tuberculosis-infected mice express CCL21 and CCL19,
naive T-cell chemoattractants, and contain organized neolym-
phoid structures (34). Secondary lymphoid structures induced
by chronic infection could serve as sites of naive CD4?T-cell
priming in the lung.
BCG infection also resulted in increased numbers of effector
CD4?T cells in the MLN and lungs. This difference was more
pronounced in the lungs. Increased migration and effector
CD4?T-cell differentiation could explain this finding (7). BCG
infection may enhance T-cell activation and effector cell de-
velopment by causing a greater influx of naive T cells into
lymphoid tissues in the mediastinum and lung, as has been
demonstrated during herpes simplex virus type 2 infection
(37). This increases the likelihood that during BCG infection
greater numbers of naive T cells are exposed to cognate anti-
gen presented by DCs in lymphoid tissues. Increased exposure
to cognate antigen results in increased naive T-cell activation
and division as observed in our experiments (28). In addition,
Catron et al. have shown that the presence of greater numbers
of naive T cells in lymph nodes at the time of antigen challenge
facilitates the generation of effector T cells (5).
The two different pulmonary environments responsible for
generating T-cell responses in infected and uninfected mice
gave rise to divergent maturation states of lung DCs. BCG can
cause DC maturation in vitro by interacting with Toll-like
receptors 2 and 4 present on these cells (17, 41). Our studies
indicate that pulmonary BCG infection results in the matura-
tion of lung CD11c?cells harboring airway OVA. Lung
CD11c?cells from infected mice were more capable of pre-
senting exogenous OVA peptide ex vivo than lung CD11c?
cells from uninfected mice. Concomitant with DC maturation,
the activation of Toll-like receptors on DCs enhances migra-
tion from peripheral sites to draining lymph nodes by upregu-
lating CCR7 (17). The distribution of airway OVA to greater
numbers of DCs expressing high levels of MHC-II suggests
that mature DCs from infected lungs could migrate to MLN
and initiate robust naive OVA-specific T-cell responses. In
addition, TLR engagement on DCs induces the production of
cytokines such as IL-12 that promote the differentiation of
responding T cells to Th1 cells, with the balance between Th1
and Th2 being determined by the strength of the TLR stimu-
lation (9). BCG infection enhanced the differentiation of
OVA-specific naive CD4?T cells in the lungs into Th1 effec-
tors. This is in agreement with previous studies that examined
the role of BCG infection in promoting allergen-specific Th1
responses (10, 33).
Therefore, we propose two complementary mechanisms for
the initiation of naive CD4?T-cell responses in the lungs
during mycobacterial infection. Pulmonary infection causes
lung DC maturation and increases DC trafficking to MLN to
initiate robust naive CD4?T-cell responses in the lymph node.
Alternatively, BCG infection causes lung DC maturation and
in situ activation of the naive CD4?T cells in the lungs.
Additional studies will determine where naive CD4?T-cell
activation occurs within different lung compartments during
chronic inflammation: the alveolar space, the bronchus-associ-
ated lymphoid tissue, or the lung parenchyma.
We thank Melanie Campbell for mouse husbandry. Nicole Pecora
advised us on the choice of fluorochromes and general flow cytometry
techniques. Robert Mahon helped with cytokine assays. David Cana-
day lent his expertise in depleting endotoxin from ovalbumin and
labeling ovalbumin with Fluos. Roxana Rojas provided guidance and
suggestions for assays examining DCs.
This study was supported by National Institutes of Health grants
AI27243 and HL55967 to W.H.B., grants AI34343 and AI35726 to
C.V.H., and grant T32 GM07250 to M.M.A.
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Editor: R. P. Morrison
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