MODULATION OF NAÏVE CD4+ T CELL ACTIVATION
AND DENDRITIC CELL FUNCTION IN THE LUNGS
DURING PULMONARY MYCOBACTERIAL INFECTION
by
MURSALIN M. ANIS
Submitted in partial fulfillment of the requirements
for the degree of Doctor of Philosophy
Thesis Advisors:
Dr. W. Henry Boom, M.D.
Dr. Clifford V. Harding, M.D., Ph.D.
Department of Pathology and Division of Infectious Diseases
CASE WESTERN RESERVE UNIVERSITY
August 2007
CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the dissertation of
______
candidate for the Ph.D. degree *.
(signed)______(chair of the committee)
______
______
______
______
______
(date) ______
*We also certify that written approval has been obtained for any proprietary material contained therein.
DEDICATION
For my mother and father
TABLE OF CONTENTS
Signature Sheet ii CWRU Waiver iii Dedication iv Table of Contents v List of Tables vii List of Figures viii Acknowledgements x List of Abbreviatioins xi Abstract xii
Chapter 1: Introduction 1 Lung environment 2 Chemokines 3 MTB infection in the lungs 6 Adaptive immunity to MTB 6 Formation of tertiary lymphoid structures 8 Cytokines and T cell differentiation 11 Naïve and effector T cells 13 Biology of TCR gene rearrangement 14 TCR transgenic mice – a tool 16 Ag85B-specific TCR transgenic mouse 18
Chapter 2: Activation of naïve CD4+ T cells 20 during pulmonary mycobacterial infection Summary 21 Introduction 22 Materials and Methods 24 Results 32 Discussion 46
Chapter 3: Generation of a MTB-specific T cell receptor 52 transgenic mouse Summary 53 Introduction 54 Materials and Methods 55 Results 59 Discussion 70
Chapter 4: Modulation of pulmonary dendritic-cell function 72 during mycobacterial infection Summary 73 Introduction 74 Materials and Methods 75 Results 81
v Discussion 91
Chapter 5: Discussion 96 Notebook table 114
Works Cited 115
vi List of Tables
3.1 Sequence of primers used to identify and clone Vα and Vβ 56 segments of Ag85B-specific T cell receptor
5.1 List of the experiment numbers corresponding to the presented figures 114
vii List of Figures
1.1 GPCR signaling during chemotaxis of leukocyte 5
1.2 Architecture of secondary and tertiary lymphoid organs 10
1.3 Cytokines and T-cell differentiation 12
1.4 Gene rearrangements that yield a productive T cell receptor 15
2.1 BCG infection causes enrichment and accumulation of 33 OVA-specific T cells after airway OVA challenge
2.2 BCG infection enhances T cell activation in the lungs 35
2.3 Pulmonary infection increases the frequency of antigen-specific 37 T cells responding to an airway antigen
2.4 BCG infection induces OVA-specific T cell proliferation in the 39 lungs after airway OVA challenge
2.5 BCG infection increases accumulation of effector antigen-specific 41 T cells in the lungs after airway antigen challenge
2.6 Mycobacterial infection induces naïve OVA-specific T cells 43 to differentiate into Th1 effectors in response to airway OVA challenge
2.7 BCG infection causes up-regulation of MHC-II on lung 45 CD11c+ cells harboring intranasal Fluos-labeled OVA
3.1 Vβ surface expression on Ag85B reactive T cell hybridomas 60
3.2 Vα cloning scheme 62
3.3 BLAST search identifying Vα5 leader sequence 64
3.4 Colony PCR of genomic Vα and Vβ sequences of BB7 65
3.5 Vα5Jα30 and Vβ11DJβ2.1 sequences of BB7 TCR 66
3.6 Restriction digests of pTβ-Vβ colonies 68
3.7 Restriction digests of pTα-Vα colonies 69
viii 4.1 Accumulation and maturation of dendritic cells in the lung 82 during BCG infection
4.2 CCR7 mRNA and surface expression on lung DCs are not 84 increased during peak and late stages of BCG infection
4.3 BCG infection upregulates expression of CCL19 in lung in 86 a MyD88-dependent manner
4.4 BCG infection modulates CCR7-mediated chemotaxis of 88 lung DCs
4.5 BCG infection alters ability of lung CD11c+ cells to present 90 peptide to naïve CD4+ T cells in vitro
5.1 Activation of naïve CD4+ T cells by lung DCs 101
5.2 Chemokine receptor CCR7-signaling pathway 106
ix ACKNOWLEDGEMENTS
My parents have always taught me to develop a deep respect for my teachers. I
would like to thank all of my teachers who have motivated me to pursue a career in
medicine and science. I am indebted to Mr. Joseph Tarello, Ms. Mary Sceppa, Dr.
Andrew Souers and Dr. Peter Radkowski for their guidance and inspiration during high
school and college.
The work here would not be possible without the mentorship of Dr. W. Henry
Boom and Dr. Clifford V. Harding. They had given me considerable freedom to explore
and pursue topics in Immunology that interested me most. At critical junctures they have
provided insights that transformed the nature of the research to one of understanding
important biological phenomena. Henry’s encouragement always fought off despair.
I would like to thank past and present members of Henry Boom’s Laboratory:
Roxana Rojas, Scott Fulton, Scott Reba, David Canaday, Adam Gehring, Robert Mahon,
Martha Torres, Jeremy Thomas, Leola Jones, Scott Mahan and Keith Chervenak. Due to
our close collaboration with Cliff Harding’s I would like to thank members of his lab
who have helped me tremendously: Nancy Nagy, Meghan Pennini, Yi Liu (for qRT-PCR
studies), Nicole Pecora, Reginald Gray, Michael Drage, Aaron Tobian, Rish Pai, Peter
Chefalo, Gareth Hardy, David Askew, and Lakshmi Ramachandra. I would also like to
thank Melanie Campbell in Alan Levine’s lab and members of Eric Arts’ lab.
Lastly, but most importantly I would like to acknowledge my mother and father; their love and prayers kept me company during long nights in the lab. I am grateful for
my loving sisters, Rehnuma and Soniya and my dear wife Nasreen for their love and support. My brother in faith, Khalifah Alyy’s friendship and support kept me going.
x ABBREVIATIONS
Ab antibody
Ag, antigen
APC antigen presenting cell
BALT bronchus-associated lymphoid tissue
BCG bacillus Calmette Guérin
BCG+OVA BCG-infected mice given OVA intranasally
BLC B lymphocyte chemoattractant
BMDC bone marrow-derived dendritic cells
DC dendritic cells
ELC EBV-induced molecule 1 ligand chemokine,
CCL19, MIP3β
G protein guanine nucleotide binding protein
GeoMFI geometric mean fluorescent intensity
GPCR G-protein coupled receptor
mAb monoclonal antibody
MHC major histocompatibility complex
MTB Mycobacterium tuberculosis
PAMP pathogen associated molecular pattern
PCR polymerase chain reaction
pOVA OVA peptide 323-339 a.a.
SLC secondary lymphoid chemokine, CCL21
xi SLO secondary lymphoid organ
TCR T cell receptor
TLO tertiary lymphoid organ
TLR Toll-like receptor
xii Modulation of naïve CD4+ T cell activation and dendritic cell function
in the lungs during pulmonary mycobacterial infection
Abstract
By
MURSALIN M. ANIS
Initiation of CD4+ T cell responses is critical for a successful host response
against the intracellular pathogen Mycobacterium tuberculosis (MTB). Naïve CD4+ T
cells are activated by dendritic cells (DCs) that sample antigen in the periphery,
upregulate chemokine receptor CCR7, and migrate to secondary lymphoid organs to
encounter naïve CD4+ T cells. Pulmonary mycobacterial infection with the attenuated
strain, Mycobacterium bovis BCG is used as a model to study host immune responses in
the lungs. During BCG infection, lung bacterial burden peaks 4-6 wks post-infection and declines to undetectable levels by 12-14 wks.
Inflammation caused by peak BCG infection (4-6 wks) led to enhanced naïve
CD4+ T cell responses directed against a model airway antigen, ovalbumin (OVA). BCG infection caused accumulation, activation, and proliferation of OVA-specific CD4+ T cells in lung and draining mediastinal lymph node (MLN). Compared to uninfected mice, infected mice had greater proliferation in lung and lymph node but only infected mice had detectable in situ proliferation of OVA-specific CD4+ T cells in the lungs.
BCG-infection induced expression of CCL19 in the lungs; CCL19 is a CCR7 ligand and naïve T cell chemoattractant. Lung inflammation, during infection, caused accumulation
xiii and maturation of lung DCs that could present peptide antigens ex vivo. OVA-specific
CD4+ T cells from the lungs of infected mice were more likely to differentiate into
effector T cells and produce IFN-γ than T cells from uninfected mice.
BCG infection caused accumulation and maturation of DCs in infected lungs even
as the mycobacterial burden declined. Lung DCs from infected mice expressed increased
amounts of MHC-II but comparable amounts of CCR7 relative to uninfected mice. Gene
expression of a CCR7 ligand, CCL19 progressively increased throughout BCG infection
and the expression was MyD88-dependent. Lung CD11c+ cells from BCG-infected mice
activated OVA-specific naïve CD4+ T cells more than lung CD11c+ cells from uninfected mice. Therefore, our findings suggest that during BCG infection, inflammation and sustained expression of CCL19 recruit and retain mature DCs in the lung where they can activate naïve CD4+ T cells.
xiv
CHAPTER 1
Introduction
1 Overview
Ever since the emergence of HIV, the intracellular pathogen Mycobacterium
tuberculosis (MTB) has come into the radar of concerned public health officials and
researchers in the developed nations. Even in immunocompetent individuals MTB is
rarely eradicated; rather, in the majority of cases MTB establishes a latent infection.
Latently infected people have a life-time risk of 5-10% for developing pulmonary reactivation tuberculosis. CD4+ T-cell responses are critical in the host response to MTB.
Dendritic cells (DCs) are regarded as the primary antigen presenting cells (APCs)
responsible for activating naïve CD4+ T cells. Modulation of DC function could lead to diminished immunity to MTB and allow latent infection to develop (Flynn 2004; Marino,
Pawar et al. 2004).
Lung Environment
As primary organ for gas exchange the lungs come into contact with both innocuous particulates and pathogens. If the ciliary motion of the respiratory tract is unable to clear such foreign material, then the body’s next line of defense is phagocytic cells in the alveolar spaces – mainly alveolar macrophages (AM). The interaction between the inhaled particulate, AM, and dendritic cells, which are below the epithelial lining but near the AM, sets the stage for development of antigen-specific immune responses (Bingisser and Holt 2001).
In the absence of microbial stimuli, immune responses in the lungs to soluble antigens are characterized by either tolerance or Th2-like responses as seen in murine
models of asthma. In such models, mice are periodically injected intraperitoneally with
2 soluble OVA and 3-4 weeks later OVA sensitized mice are challenged by an aerosol of
1% OVA in PBS. Lung and spleen cells from these OVA challenged mice produce Th2 cytokines IL-4 and IL-5 in response to OVA (Mojtabavi, Dekan et al. 2002). To prevent such Th2 driven asthma development, soluble antigen challenge through the airways also leads to the development of regulatory T cells that secrete IL-10 and diminish responsiveness to the soluble antigen (Tsitoura, DeKruyff et al. 1999; Akbari, Freeman et al. 2002). In addition to IL-10, AM in the lungs secrete a variety of immunomodulating molecules such as nitric oxide, prostaglandin E2, and TGF-β. All of these immunomodulating molecules serve to down regulate T-cell responses and maintain the lungs in an ‘immune-privileged’ state (Bingisser and Holt 2001).
Chemokines
Chemokines are small (8-14 kDa) mostly basic secreted proteins that together with their receptors form a complex cellular communication network. Chemokines share a motif of 4 cysteine residues, with the position of the cysteine residues relative to each other being chosen as a distinguishing feature to establish the current classification scheme. The two main classes are CC and CXC chemokines. In CC chemokines the first two N-terminal cysteines are adjacent while in CXC chemokines there is an amino acid in between. Upon engaging seven transmembrane G-protein coupled receptors (GPCR), chemokines can direct various cell processes some of which are chemotaxis, maturation, proliferation, differentiation and apoptosis. The cellular process directed by all chemokines regardless of cell type is chemotaxis (Zlotnik and Yoshie 2000).
3 Chemotaxis is an observable phenomenon that involves both migratory speed
(random motion) and unidirectional movement toward a chemical gradient. Upon ligand
binding GPCRs change conformation in an allosteric fashion to activate heterotrimeric G
proteins which then exchange GDP for GTP and dissociate into Gαi and Gβγ dimers (Fig.
1.1). Gαi, which is inhibited by pertussis toxin, inhibits adenylate cyclase, thus decreasing cAMP concentration. However, Gβγ subunits initiate signaling important for chemotaxis by activating PI3K (phosphoinositide-3 kinase) which localizes to the leading
edge of a cell and generates PIP3 (phosphotidylinositol-3,4,5,-tripohosphate). PIP3
activates the Rho family GTPase Rac and that eventually leads to focal actin
polymerization at the leading edge and development and extension of a pseudopod. A
parallel pathway initiated by Gβγ subunits leads to activation of Cdc42, and F-actin
nucleation and exclusion of PIP3 inhibitors (phosphatases) from the pseudopod. GPCR
binding also activates an entirely opposite pathway that occurs at the trailing edge
(uropod) and causes contraction of the uropod by the action of Rho, another small
GTPase. The opposite functions of Rho and Rac are therefore spatially segregated in the cell at the uropod and pseudopod respectively to coordinate unidirectional migration (Rot and von Andrian 2004). There also seems to be a link between the mitogen activated protein kinase (MAPK) pathways and chemokine-directed chemotaxis based on pharmacologic inhibition studies, although the mechanism has not been established (Riol-
Blanco, Sanchez-Sanchez et al. 2005). Furthermore, chemokine responsiveness is fine- tuned with G-protein receptor kinases and regulators of G protein signaling which confer greater sensitivity to chemokine gradients than a simple “on/off” switch (Shi, Harrison et al. 2004).
4
Figure 1.1. GPCR signaling during chemotaxis of leukocyte (Rot and Andrian 2004)
5 MTB infection in the lungs
Upon inhalation, droplets of aerosolized MTB reach the alveolar spaces of the lungs where they are phagocytosed principally by alveolar macrophages (AM). Once phagocytosed, MTB circumvents natural phagosomal maturation and acidification and the bacilli survive within vacuoles inside the AM (Honer zu Bentrup and Russell 2001).
The bacilli interfere with phagosomal maturation by retention of TACO, an actin binding protein, and Rab5, a GTPase present in early endosomes (Chan and Flynn 2004).
In addition to phagosomal maturation arrest, MTB evades nitric oxide and other reactive nitrogen intermediates (RNIs) generated by the macrophage enzyme nitric oxide synthase 2 (NOS2). The MTB genes noxR1 and noxR3 confer resistance to RNIs by undefined mechanisms. In spite of these MTB resistance genes, RNIs play a prominent role in the host defense against MTB in mice. This is evident in the increased susceptibility of mice deficient in RNIs to MTB infection. Therefore it is thought that
NOS2 is critical in containing acute and chronic tuberculosis but not for total eradication
of the bacilli (Flynn and Chan 2003; Chan and Flynn 2004).
Adaptive Immunity to MTB
Once MTB evades the innate anti-microbial defenses of macrophages, the bacilli
replicate intracellularly. At this point adaptive immunity becomes critical in controlling
the infection. In response to TLR stimulation by MTB products, infected AM, DCs, and
lung macrophages produce inflammatory cytokines and chemokines. Inflammatory
chemokines recruit DCs into the lungs. Influx of DCs into the lungs mediated by
chemokine receptor CCR2 is important for the early response against MTB (Peters, Scott
6 et al. 2001). Once PAMPs engage TLRs on DCs, they undergo a well-characterized switch in chemokine receptor expression as they mature, down-regulating CCR2 and other inflammatory chemokine receptors and up-regulating lymph node homing receptor
CCR7 (Sallusto, Schaerli et al. 1998). It is postulated that mature lung DCs transport
MTB antigens to the draining lymph nodes to initiate T-cell responses (Bhatt, Hickman et al. 2004; Humphreys, Stewart et al. 2006). Migration of DCs to the draining MLN is key for timely initiation of T-cell responses (Chackerian, Alt et al. 2002; Marino, Pawar et al.
2004). Any delay in cell-mediated immune responses leads to impaired control of MTB infection (Tian, Woodworth et al. 2005). A report that influx of DCs into draining MLN is delayed relative to acquisition of DTH responses suggests that MTB may modulate DC trafficking to delay adaptive immune responses in the lungs (Garcia-Romo, Pedroza-
Gonzalez et al. 2004).
Activated CD4+ and CD8+ T cells migrate back to foci of infection in the lungs to secrete IFN-γ and cytotoxic molecules to activate or lyse MTB infected macrophages.
The infected macrophages are walled off from the lung parenchyma by T cells, B cells, fibroblasts, and other lung parenchymal cells in a structure known as a granuloma. While serving to contain the infection, the granuloma also serves as a reservoir of live bacilli that set up latent infection (Chan and Flynn 2004). A mechanism that may contribute to the survival of the bacilli in chronic infection is the secretion of lipoproteins such as the
19kDa lipoprotein that interacts with TLR-2 to inhibit IFN-γ mediated enhancement of
MTB antigen processing and presentation to T cells (Gehring, Rojas et al. 2003; Pecora,
Gehring et al. 2006). This may allow MTB to evade the surveillance of T cells inside granulomas.
7 Adaptive immunity is critical in controlling MTB infection. In mouse models of
tuberculosis, depletion or total absence of CD4+ T cells causes mice to succumb to the
infection earlier than wild-type mice (Mogues, Goodrich et al. 2001). In humans the loss
of CD4+ T cells due to HIV infection leads to fatal primary and reactivation tuberculosis.
One important role of CD4+ T cells during the immune response to MTB is IFN-γ
secretion. IFN-γ activates infected macrophages to kill intracellular MTB. In addition,
IFN-γ enhances the processing and presentation of MTB antigens to T cells. IFN-γ
knockout mice are very susceptible to tuberculosis infection. Aside from their role in
IFN-γ secretion, CD4+ T cells enhance the cytotoxic effects of MTB specific CD8+ T
cells. CD4+ T cells can also exert their own cytotoxic effect by inducing apoptosis of
MTB infected macrophages via Fas/Fas ligand interaction. Finally, CD4+ T cells are responsible for stimulating macrophages to produce IL-12, IL-10, and IL-15 among other immunomodulatory cytokines (Tufariello, Chan et al. 2003).
Formation of tertiary lymphoid organs (TLOs)
Even with robust cell-mediated immune responses, mycobacterial infections are rarely completely cured without antibiotics. The bacteria initiate latent infection in the
host that continually activates an infected host’s innate and adaptive immune responses.
Inflammatory infiltrates into the sites of bacterial persistence eventually begin to organize
into distinct areas of T cells and B cells aided by the induction of homeostatic
chemokines CCL19, CCL21, and CXCL12, CXCL13 in these sites (Aloisi and Pujol-
Borrell 2006). CCL19 (MIP-3β) and CCL21 (SLC) are CCR7 ligands and attract naïve T
cells and mature DCs to T zones of TLOs and secondary lymphoid organs (SLOs).
8 CXCL12 (stromal cell-derived factor 1) is a CXCR4 ligand and attracts naïve T cells, B cells, DCs, and plasma cells. CXCL13 (BLC, B lymphocyte chemoattractant) is a ligand for CXCR5 and attracts B cells to the follicular compartments of TLOs and SLOs
(Luther, Bidgol et al. 2002). CXCL13 and CCL21 have been shown to be important for the compartmentalization of T and B cells in SLOs and TLOs. The induction of these critical homeostatic chemokines depends upon TLR stimulation and/or lymphotoxin
(LTα1β2) stimulation. A positive feedback mechanism exists where homeostatic chemokines induce expression of LTα1β2 and LTα1β2 stimulation can further induce expression of these homeostatic chemokines. Additionally, even though B cells serve a key role in the generation of TLOs, naïve B cells alone don’t give rise to mature TLOs, which arise in the presence of activated B cells. Lastly cytokines such as IL-4 and IL-7 have also been shown to promote LTα1β2 expression on T cells. However the organization of tertiary lymphoid structures or BALT during lung inflammation varies from the structure of secondary lymphoid organs (Fig. 1.2) (Aloisi and Pujol-Borrell
2006). Two main points of difference are: 1) SLOs are surrounded by a capsule whereas
TLOs are not implying that TLOs are directly influenced by the inflammatory milieu of the tissue parenchyma within which they reside, and 2) TLOs lack afferent lymphatics that coordinate the entry and organized distribution of immune cells in SLOs. The absence of organization may result in abnormal T and B-cell responses within TLOs
(Aloisi and Pujol-Borrell 2006). Finally the lungs, liver, and gut represent some of the tissues that are especially permissive in the formation of TLOs. Therefore, it is not surprising that mycobacterial infection induces expression of CCR7 ligands (Schreiber,
Ehlers et al. 2006; Kahnert, Hopken et al. 2007).
9
Figure 1.2. Architecture of secondary (a), and tertiary(b) lymphoid organs (Aloisi and Pujol-Borrell 2006)
10 Cytokines and T cell differentiation
The cytokines IL-12 and IL-4 are important in the differentiation of naïve T cells
to Th1 and Th2 effector T cells respectively. In vitro studies have shown that MTB
infected bone marrow derived dendritic cells (BMDC) secrete IL-12. Furthermore, in
vivo experiments involving adoptive transfer of MTB infected BMDCs have shown that
such DCs migrate to regional lymph nodes to initiate Th1 type T-cell responses (Bhatt,
Hickman et al. 2004). However, late stages of intratracheal virulent MTB infection in
Balb/c mice give rise to high levels of IL-4 as well as IFN-γ, and TGF-β (Hernandez-
Pando, Aguilar et al. 2004). Signals triggered by IL-12 and IL-4 are relayed by signal
transducer and activator of transcription (STAT) proteins. STAT6 and the transcription
factor GATA-3 are essential for Th2 differentiation, while STAT4 and the transcription
factor T-bet is essential for Th1 differentiation (Ansel, Lee et al. 2003). In addition, T-bet
induction depends upon STAT1 activity. Synergy among IL-12, IL-18, and IL-27 also
directs early Th1 differentiation events (Robinson and O'Garra 2002). The complex
processes underlying T cell differentiation is influenced by MTB infection. MTB cell
extracts induce DCs to upregulate CD86 (B7-2) as well as the immunomodulatory
cytokines IL-10 and TGF-β. Both IL-10 and TGF-β downregulate Th1 responses specific for cell extract and decrease pro-inflammatory cytokines (Balkhi, Sinha et al. 2004).
Furthermore, costimulation through CD86 (B7-2) is necessary for differentiation into Th2 cells in the lungs (Tsuyuki, Tsuyuki et al. 1997). The significance of these findings is that in general the inhibition or absence of Th1 differentiation promotes Th2
differentiation (Robinson and O'Garra 2002). The interplay between these cytokine
pathways is illustrated in Fig. 1.3.
11
Figure 1.3. Cytokines and T-cell differentiation (Tato, Laurence et al. 2006).
12 Naïve & Effector T cells
Once naïve CD4+ T cells encounter antigens presented by MHC Class II molecules, the naïve T cells can either differentiate into Th1 and Th2 effector and memory cells or undergo deletion. Presentation of antigens by mature DCs, expressing high levels of peptide:MHC Class II complexes and co-stimulatory molecules like CD28, activates naïve T cells to proliferate and differentiate. Immature DCs express low levels of antigen and co-stimulatory molecules and induce anergy and deletion of naïve T cells. Critical to the survival of activated CD4+ T cells in the absence of antigenic stimulation is their
ability to respond to homeostatic cytokines such as IL-7 and the expression of anti-
apoptotic molecules such as B cell lymphoma-2 (Bcl-2). The strength of the stimulation
received by naïve T cells enables them to express receptors for homeostatic cytokines and
anti-apoptotic molecules. TGF-β weakens the signal delivered to naïve T cells such that
these T cells express less IL-7 receptors (Gett, Sallusto et al. 2003)
Once naïve T cells have become activated, they expand and differentiate into a
heterogeneous population of effector Th1 and Th2 cells. The expansion phase of activated
T cells is followed by the death of almost 90% of effector T cells. Mechanisms involving
Fas/Fas ligand, CD40/CD40 ligand, and TNF receptors are responsible for the
contraction of the T cell population. In addition, IFN-γ, which is significant in the control of MTB infections, also induces apoptosis of effector T cells by a caspase 8 dependent pathway. This fact may explain the inability of IFN-γ producing Th1 cells to
become resting memory T cells. The remaining pool of effector T cells differentiate to
become pools of central memory and effector memory T cells (Seder and Ahmed 2003).
13 In contrast to naïve T cells, effector immune responses are more rapid, secrete more cytokines such as IFN-γ, and effector T cells are able to localize to non-lymphoid tissues in greater numbers (Garcia, DiSanto et al. 1999; London, Lodge et al. 2000).
Effector and memory T cells are able to migrate to non-lymphoid tissues much better than naïve T cells; whereas naïve T cells are better able to home into lymph nodes (Iezzi,
Scheidegger et al. 2001). Furthermore, using TCR transgenic mice, Kimachi and others have demonstrated that effector T cells respond to low affinity ligands while naïve T cells do not – corroborating the fact that effector and memory T cells are more sensitive to stimulation than naïve T cells (Kimachi, Sugie et al. 2003).
Biology of TCR gene rearrangement
Antigen receptor genes in both B cells and T cells undergo recombination to generate the tremendous diversity of receptors capable of binding antigens to initiate adaptive immune responses. The T cell receptor is a heterodimer composed of α and β chains. Both the TCRα and TCRβ chain genes undergo recombination during the development of T cells.
First, the TCRβ chain gene is recombined and after successful recombination, the
TCRα chain gene is recombined (Fig. 1.4). The process of rearranging α and β chain genes generates combinatorial diversity by mixing and matching variable α regions with joining (Jα) regions for the α chain. For the β chain of the TCR, first the diversity (Dβ) regions combine with joining (Jβ) regions. The DJβ rearrangement undergoes further recombination by attaching to variable Vβ regions. In mice there are 100 Vα regions, 50
Jα regions, 25 Vβ regions, 2 Dβ regions, and 12 Jβ regions. The genetic rearrangements
14
Figure 1.4. Gene rearrangements that yield a productive T cell receptor
Immunology. Goldsby RA, Kindt TJ, Osbourne BA and Kuby J. (Freeman & Co. New York) 4th Edition, 2002.
15 are made possible by the recombination machinery – two key players of which are
recombination activating gene -1 (RAG-1) and RAG-2. These enzymes recognize
recombination signal sequences (RSS), conserved heptamers and nonamers which flank
the different V, D, and J regions (Jung and Alt 2004).
Aside from combinatorial diversity due to mixing and matching different V and J
regions, other sources of possible diversity in mouse TCR genes are junctional flexibility,
“N” nucleotide additions, and association of α and β chains. This rearrangement at both
β and α chain loci is regulated in terms of development, cell lineage, and allelic exclusion. Allelic exclusion applies to β chain gene rearrangements but not α chain genes. α chain allelic exclusion is considered to be phenotypic – majority of peripheral T cells express one single TCR. This occurs by a post-translational mechanism involving signaling through two possible TCRs (1 β and 2 α chains). It is thought that the αβ pair delivering the most positive signal to the T cell is selectively retained on the surface of mature T cells (Niederberger, Holmberg et al. 2003). However, around 8% of peripheral
T cells express both α chains on the surface (Corthay, Nandakumar et al. 2001).
TCR transgenic mice – a tool
Much information regarding the in vivo activation and migration of naïve and effector T cells during infection has been gathered using TCR transgenic mice. Until recently, the study of antigen-specific T-cell responses was hampered due to the low precursor frequency of naïve antigen specific T cells in naïve animals. TCR transgenic mice have large numbers of T cells specific for distinct antigen epitopes. With the advent
16 of MHC-tetramer and TCR transgenic mice, studies examining antigen-specific immune
responses during infection have become routine (Pape, Kearney et al. 1997).
Adoptive transfer of antigen specific T cells from TCR transgenic mice into
syngeneic wild-type mice represents a powerful tool to model the kinetics of naïve
antigen specific T cell activation and differentiation. Although this method artificially
increases the precursor frequency of naïve antigen specific T cells, studies have shown
that the kinetics of naïve T cell activation remains dependent upon antigen:MHC
complex recognition. Still, increasing the precursor frequency decreases the clonal
expansion of the antigen specific T cells due to possible limited access to MHC, growth factor and physical space (Laouar and Crispe 2000). In recent years, several studies have
shown that competition for limited number of peptide:MHC complexes have profound
influence on the make-up of the responding T cells (Marzo, Klonowski et al. 2005;
Catron, Rusch et al. 2006). T cells that receive strong stimulation develop into effector
and effector-memory T cells while those that receive weak stimulation in the proper
context develop into central-memory T cells (Marzo, Klonowski et al. 2005; Hataye,
Moon et al. 2006). Adoptively transferring less antigen-specific T cells lessens the
contraction phase after antigen-specific T cell proliferation in vivo. As a result, more
transferred T cells develop memory phenotypes (Hataye, Moon et al. 2006).
In the context of studying T-cell responses during various infections, investigators
have generally used TCR transgenic mice in two principal ways. First, researchers have
genetically engineered bacteria and viruses to express heterologous antigens. Then, scientists have used well characterized TCR transgenic mice, specific for those heterologous antigens, to study T-cell responses during infection with recombinant
17 pathogens (Smith, Dudani et al. 2002). While such non-pathogen specific TCR transgenic mice provide versatility when studying different recombinant pathogens, they invariably study T-cell responses directed against heterologous antigens that are not pathophysiologically relevant.
Investigators have tried to overcome these problems, associated with heterologous antigen-expressing recombinant pathogens, by developing TCR transgenic mice specific for native pathogen-derived antigens. Due to the ready availability of MHC Class I tetramers, such studies have mostly used viral antigen specific TCR transgenic mice to study immune responses to viral infections. Two recently generated TCR transgenic mice specific for bacterial pathogens are SM1 TCR transgenic mice (McSorley, Asch et al. 2002), specific for a salmonella derived flagellin peptide and NR1 TCR transgenic mice, specific for an epitope of Chlamydia trachomatis (Roan, Gierahn et al. 2006).
Ag85B specific TCR transgenic mouse
The 30kD Ag85B and the 6kD early and secreted antigenic target (ESAT-6) are being used in clinical trials to assess their efficacy as vaccines for tuberculosis. Ag85B is part of a family of proteins involved in mycobacterial cell wall synthesis. These proteins, including Ag85B, are often the most common proteins found in MTB culture fluid. More importantly, Ag85B elicits strong humoral and cell-mediated immune responses (Wiker and Harboe 1992). Cohort studies of people with MTB-infected household contacts have shown that lymphocytes from newly infected individuals respond vigorously to the mycobacterial antigen, Ag85B. Studies have also shown that actively replicating MTB
18 have high Ag85B transcript levels that drop as adaptive immunity to the bacilli develops
(Shi, North et al. 2004).
Using the unique Ag85B241-256 TCR transgenic mouse we will be able to address questions regarding MTB-specific immune responses during MTB infection. Other
investigators have tried to address this issue using MHC tetramers to track ESAT-6 specific T cells. However, due to the low precursor frequency of these MTB specific T cells, that study could only address the persistence of such MTB specific cells during chronic infection (Winslow, Roberts et al. 2003). Consequently, we will use the Ag85B
TCR transgenic mice in adoptive transfer experiments to increase the precursor frequency of MTB specific T cells in wild-type mice. We will then observe the effect of varying precursor frequency on the ensuing immune response to MTB. Since vaccines based on
Ag85B are also designed to generate a small pool of effector Ag85B specific T cells, these experiments will shed insight on the efficacy of such vaccines.
19
CHAPTER 2
Activation of naïve CD4+ T cells during pulmonary mycobacterial infection
Adapted from
Anis, M. M., S. A. Fulton, S. M. Reba, C. V. Harding, and W. H. Boom (2007).
“Modulation of Naive CD4+ T-Cell Responses to an Airway Antigen during Pulmonary
Mycobacterial Infection.” Infect Immun 75:2260-8.
20 Summary
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 naïve 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 post-infection), CFSE-labeled naïve ovalbumin-specific DO11.10 T cells were
adoptively transferred into infected and uninfected mice. Recipient mice were challenged
intranasally with soluble ovalbumin (OVA) and OVA-specific T-cell responses were measured in lung, draining mediastinal lymph node (MLN) and spleen. OVA challenge resulted in increased activation and proliferation of OVA-specific T cells in the draining
MLNs 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 naïve CD4+ T cells in the
lungs in response to airway antigen challenge.
21 Introduction
The lungs constantly come into contact with airborne particulates and pathogens.
The interaction between inhaled particulate antigens, alveolar macrophages, and
pulmonary dendritic cells (DCs) sets the stage for antigen-specific pulmonary immune
responses (Bingisser and Holt 2001). In the absence of microbial stimuli, immune responses in the lungs to soluble antigens are characterized by either tolerance or Th2-like responses as seen in murine models of asthma (Tsitoura, DeKruyff et al. 1999;
Mojtabavi, Dekan et al. 2002). In contrast, acute viral infections give rise to vigorous
pulmonary T-cell responses characterized by Th1 cytokine secretion (Roman, Miller et al.
2002; Lawrence and Braciale 2004). 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 naïve T-cell responses in the lungs during prolonged mycobacterial infection.
Mycobacterium tuberculosis establishes a latent infection in the vast majority of immunocompetent individuals. Upon inhalation the mycobacteria are phagocytosed principally by alveolar macrophages. M. tuberculosis circumvents phagosomal
maturation and establishes a niche for intracellular survival (Honer zu Bentrup and
Russell 2001). In the ensuing adaptive response mycobacteria are contained in granulomas within which M. tuberculosis persists latently (Chan and Flynn 2004). The modulation of pulmonary immunity that permits 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 (Ladel, Daugelat et al. 1995;
22 Humphreys, Stewart et al. 2006; Kuchtey, Fulton et al. 2006). 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 (Fulton, Martin et al. 2000; Fulton,
Reba et al. 2002; Kuchtey, Fulton et al. 2006).
It is thought that activation of naïve CD4+ T cells, in response to airway antigens,
occurs primarily in the mediastinal lymph nodes (MLN) draining the lungs (Xia, Pinto et
al. 1995; Curtis 2005). During pulmonary influenza infections, virus-specific naïve T cells divide in the MLN, and only the most differentiated cells express the appropriate adhesion molecules to migrate to the lungs (Roman, Miller et al. 2002). In the lungs, differentiated, effector T cells colocalize with antigen-carrying pulmonary DCs
(Byersdorfer and Chaplin 2001). However, recent evidence suggests that primary activation of naïve T cells can occur in the lungs (Lund, Partida-Sanchez et al. 2002;
Schreiber, Ehlers et al. 2006). Mice lacking functional CCL19 and CCL21 and mice lacking fucosyltransferases have impaired localization of naïve T cells to secondary lymphoid organs. However, these mice are able to initiate naïve 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 naïve CD4+ T cells. Few studies
have addressed this issue in their detailed analysis of naïve CD4+ T-cell responses (Xia,
Pinto et al. 1995; Itano, McSorley et al. 2003; Mempel, Henrickson et al. 2004;
Wikstrom, Batanero et al. 2006).
In this report, we have used an adoptive-transfer technique (Jenkins, Khoruts et al.
2001) to artificially increase the precursor frequencies of ovalbumin (OVA)-specific naïve T cells in recipient mice that had been previously infected with aerosolized BCG.
23 Using flow cytometry to track OVA-specific (KJ+) T cells, we found that the naïve 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, 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 activation 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) transgenic mice that
d express TCRs specific for OVA323-339 peptide presented in the context of I-A (Murphy,
Heimberger et al. 1990) were a gift from Alan Levine (Case Western Reserve University,
Cleveland, Ohio). Mice were housed in specific-pathogen-free conditions. 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 (Kuchtey, Fulton et al. 2006).
Day 1 colony counts consistently gave 3,200 ± 1,300 CFU per mouse. Bacterial growth
in the lungs peaked 4 to6 weeks afterwards with 190,000 ± 70,000 CFU per mouse.
24 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 post-infection. Uninfected mice in all of the experiments were not mock infected.
Endotoxin depletion of OVA
Endotoxin was removed from OVA (Sigma-Aldrich) by using the protocol of
Aida and Pabst with minor modifications (Aida and Pabst 1990). OVA was dissolved in filtered, lipopolysaccharide-free water, and Triton-X114 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
amoebocyte 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 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
25 with anti-CD62L and anti-CD44 monoclonal antibodies and fluorescence-activated cell
sorted (FACsorted) by gating on naïve (CD62Lhi CD44low) T cells by using a BD Aria
cell sorter. Purified CD4+ T cells and flow-sorted naïve CD4+ T cells were then used in
adoptive-transfer experiments. FACsorted CD4+ T cells were >98% CD44low CD62Lhi,
and 65-75% of these naïve 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-
5×106 DO11.10 CD4+ T cells by retro-orbital injection. In some experiments DO11.10 T
cells were labeled with 5 μM CFSE (Invitrogen) for 10 min at 37˚C in 0.1% bovine
serum albumin in phosphate buffered saline (BSA/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 μg of endotoxin-depleted
OVA or BSA as the control antigen. On day 2, mice were challenged intranasally once
more with 500 μg 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)
collected.
Tissue isolation
For experiments involving CFSE, care was taken to minimize exposure to light.
Tissues were harvested and processed as previously described (Kuchtey, Fulton et al.
26 2006). 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 harvested.
Spleens were homogenized and pressed through a 70-μm-pore-size nylon filter.
Red blood cells were lysed in red blood cell (RBC) lysis buffer. Single cells were
resuspended in complete medium (Dulbecco modified Eagle medium, 10% fetal bovine
serum [FBS], 0.05 mM 2-mercaptoethanol, 2mM HEPES, 1mM sodium pyruvate, 100 mM non-essential amino acids, 100 U of penicillin/ml and 0.1 mg of streptomycin/ml).
Lungs were minced and digested with 125 units of type IV collagenase and 30 units 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-μm-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-μm-pore-size nylon filter using the plunger of a 1-ml syringe
and then resuspended 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 5×105 to 1×106 viable lung, MLN, and spleen cells
were pre-incubated 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
27 biotinylated KJ 1-26 (Invitrogen catalog number MM 7515-3), along with activation and
adhesion markers anti-CD62L, anti-CD44, 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. The cells were washed once with 1% BSA,
resuspended in streptavidin-Pacific Blue conjugate (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 (Wells,
Gudmundsdottir et al. 1997). Briefly, the number of events in each daughter cell
generation N, characterized by dimmer CFSE labeling, was divided by 2N to 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 phorbol myristate acetate (PMA) at 50
ng/ml and 1 μg/ml ionomycin (Sigma-Aldrich) in the presence of 10 μg/ml 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 1X PBS staining solution at
28 room temperature. Cells were fixed with 4% paraformaldehyde and stained with allophycocyanin anti-IFN-γ 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-Aldrich) 1 h prior to sacrifice
(Moyron-Quiroz, Rangel-Moreno et al. 2004). Tissues were harvested and single-cell suspensions made. Cells were surface stained with the DO11.10 T-cell receptor antibody, 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-γ was done as previously described (Kuchtey, Fulton et al. 2006). Briefly, sterile ELISPOT plates (Whatman) were precoated with anti-IFN-γ capture antibody (BD Pharmingen catalog number
551216) overnight at 4˚C at a concentration of 5 μg/ml. The plates were blocked with
1% BSA in PBS for 1 h and washed with PBS before the lung, spleen, and MLN cells
29 from OVA-challenged, BCG-infected and uninfected mice were added at 5×105 and
6 1×10 cells/well. Some wells received exogenous OVA peptide (OVA323-339; 2 μM), 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-γ (BD Pharmingen catalog number 554410) at a concentration of 2 μg/ml.
Plates were washed four times, and bound IFN-γ 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 numbers 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 sulfoxide to make a 2-mg/ml solution. A 95 μl 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 μg 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-
30 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 uninfected 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 μM) was added to the wells,
5 d along with 10 DOBW T-cell hybridoma cells, which recognize OVA323-339:I-A
complexes and secrete IL-2. After 18 to 24 h, 100-μl portions of 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 number
14-7022) at 1 μg/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-conjugated anti-IL-2 detection antibody (eBioscience catalog
number 13-7021) at 1 μg/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.
31 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.
Results
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 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. 2.1A). The enrichment 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 elicited 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. 2.1B).
32 A.
BCG+OVA OVA BCG+BSA BSA 30 * s ll
e 25 20 15 CD4+ T c 10 ** 5 % KJ+ / 0 MLN lungs spleen MLN lung spleen
B. 3 25 # ) ## 5 2.5 10
X 20 2 ls ( 15 1.5 T cel + 10 1
5 0.5 Total KJ 0 0 MLN lung
Figure 2.1. BCG infection causes enrichment and accumulation of OVA-specific T cells after airway OVA challenge. BALB/c mice were infected with BCG and 4-6 wks later, 3-5×106 CD4+ DO11.10 T cells were transferred into BCG infected and uninfected mice. Recipient mice were challenged with 500 μg of endotoxin-depleted OVA on Days 0 and 2 or with BSA on Day 0, and were sacrificed on Day 3. Single cell suspensions were prepared from lung, MLN, and spleen and stained with anti-CD4 and KJ 1-26 to enumerate OVA-specific T cells. A. % of OVA-specific (KJ+) T cells among CD4+ T cells present, was determined in the mediastinal lymph node (MLN), lungs, and spleens of the 4 different groups. B. Total
KJ+ T cell numbers were calculated by multiplying viable cell counts, determined by trypan blue exclusion, by the % of KJ+ T cells among viable cells gated from FSC vs. SSC plot during FACS acquisition. This experiment was repeated twice with similar findings. n=3 mice per group. *, p=0.02; **, p=0.01; #, p=0.004; ##, p=0.02.
33 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 naïve T-cell activation 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 increased recruitment, enhanced T- cell activation and proliferation, or both. To determine the contribution of T-cell activation, we adoptively transferred FACsorted naïve (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-lymphocyte activation (CD69, CD25) on OVA-specific T cells in the lungs and draining MLN was analyzed after OVA challenge (Fig. 2.2). The baseline levels of CD25 and CD69 on naïve KJ+ T cells before adoptive transfer were not greater than those of the isotype
controls (Fig. 2.2A). The early T-cell activation 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. 2.2A). The rapid kinetics of CD69
expression after T- cell activation (Mempel, Henrickson et al. 2004) 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. 2.2B and 2.2C). 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.
34 Pre-transfer
Isotype OVA BCG+OVA A. MLN
0 102 103 104 105 0 102 103 104 105 CD25 CD69 B. lungs
02 103 104 105 0 102 103 104 105 CD25 CD69
C. BCG+OVA OVA
s 60 s 30 cell cell * 20 40 9+/ KJ+ T
5+/ KJ+ T 10 20 % CD6
% CD2 0 0 MLN1 lungs2 MLN lungs
Figure 2.2. BCG infection enhances T cell activation in the lungs. CD4+CD62LhiCD44low FACSorted naïve DO11.10 T cells were transferred into BCG infected and uninfected mice. Mice were challenged with OVA as described in Materials and Methods. 3 days later, mice were sacrificed and draining mediastinal lymph nodes (MLN) (A) and lungs (B), stained for TCR, CD25, and CD69. Histograms were gated on KJ+ T cells. Baseline levels of CD25 and CD69 on naïve KJ+ T cells before adoptive transfer is shown in (A). Percentages of CD25 and CD69-expressing KJ+ T cells, above isotype staining, from 3 mice per group is shown in (C). *, p<0.01. Data is representative of 3 separate experiments.
35 2.2C). 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 (Mempel,
Henrickson et al. 2004). The CD25+ CD69+ phenotype of KJ+ T cells in the lungs of
infected mice suggested recent T-cell activation. Thus, airway OVA challenge causes
naive KJ+ T-cell activation in the MLN of both infected and uninfected mice but, during
pulmonary BCG infection, activation of naïve 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 proliferation, 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 (Lyons 2000). CFSE dye dilution in KJ+ T cells
was antigen-specific because no KJ+ T-cell division was observed in animals challenged with BSA (Fig. 2.3A). In draining MLN and lungs, infected 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. 2.3A). Proliferation intermediates, corresponding to the first few cell divisions, were prominent in the MLN but not in lungs of both OVA-challenged groups. This suggested that priming of naïve CD4+ T cells
occurs primarily in the MLN but that KJ+ T-cell activation can also occur in infected lungs (Fig. 2.2B). Very little KJ+ T-cell division was apparent in the spleens,
36 Figure 2.3. Pulmonary infection increases the frequency of antigen-specific T cells responding to an airway antigen. 5x106 CFSE labeled CD4+ DO11.10 T cells were transferred into BCG-infected and uninfected recipient mice. Recipient mice were challenged with 500 µg of endotoxin-depleted OVA on
Days 0 and 2 or with BSA on Day 0. Mice were sacificed 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 4 individual mice from the 4 groups. B. % 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, n=3 mice per group; *, p=0.02; **, p=0.03.
37 demonstrating again that the antigen-specific T-cell response to an airway antigen was
primarily 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
(Wells, Gudmundsdottir et al. 1997), (Fig. 2.3B). 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. 2.3B).
Enhanced CFSE dilution of KJ+ T cells in the lungs of BCG-infected mice could be due
to i) increased recruitment 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 determine the contribution of in situ pulmonary KJ+ T-cell proliferation in the accumulation of proliferating KJ+ T cells in the lungs (Fig. 2.3), we adoptively transferred naïve (CD44low CD62Lhi) KJ+ T cells into recipient mice and challenged the mice with 300 μg of endotoxin-depleted OVA. After 3 days of OVA challenge, recipient mice received 2 mg of BrdU intravenously 1 h prior to sacrifice (Moyron-Quiroz,
Rangel-Moreno et al. 2004). 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. 2.4B). In uninfected mice challenged 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
38 A. B. Isotypes BCG+OVA OVA BCG+BSA
5 5 5 10 10 105 10 6.6% 2.3% 0%
4 4 10 104 104 10
3 3 10 103 103 10 KJ 1-26 2 2 10 102 102 10
0 0 0 0
2 3 4 5 2 3 4 5 2 3 4 5 2 3 4 5 0 10 10 10 10 0 10 10 10 10 0 10 10 10 10 0 10 10 10 10
BrdU
C. BCG+OVA OVA 10
8 *
6
4
2 % BrdU+/ KJ+ T cells
0 Lungs
Figure 2.4. BCG infection induces OVA-specific T cell proliferation in the lungs after airway OVA challenge. FACS-sorted naïve (CD62LhiCD44low) CD4+ T cells from DO11.10 mice were transferred into
BCG infected and uninfected recipient mice. Recipient mice were challenged intranasally with 300 μg of endotoxin-depleted OVA or BSA. 3 days after challenge, mice were injected i.v. with 2 mg of BrdU.
After 1 h of in vivo BrdU pulse, mice were sacrificed and single cell suspensions from lungs 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 3 mice from the 3 groups; numbers represent percentage of
BrdU+ cells among KJ+ T cells. C. Means ± SD of 3 mice per group; *, p=0.01. Experiment repeated twice with similar findings.
39 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 draining 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-γ in the lungs of mice challenged with airway antigen.
To determine whether pulmonary BCG infection increased the differentiation of naïve KJ+ T cells to effector T cells, flow-sorted naïve (CD44low CD62Lhi) OVA-specific
T cells were transferred into naïve 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. 2.5A). L-selectin is a lymph node homing molecule present on naïve and
central memory T cells but absent on effector T cells (Tietz and Hamann 1997; Kipnis,
Irwin et al. 2005). Among KJ+ T cells that had divided more than two times (CFSElow),
49% were CD62Llow in lungs of BCG+OVA mice, whereas only 23% were CD62Llow in lungs of uninfected OVA-challenged mice. In contrast, 45% of CFSElow KJ+ T cells were
CD62Llow in the MLN of uninfected mice, whereas 56% of CFSElow KJ+ T cells were
CD62Llow in infected mice. The percentage of CD62Llow KJ+ T cells was greater in the
MLN and lungs of infected mice than in uninfected mice (Fig. 2.5B). Since effector T
cells preferentially migrate to sites of inflammation (Tietz and Hamann 1997), the
40 A. OVA BCG+OVA
5 5 10 10 76543210 76543210 10 4 10 4
MLN 10 3 10 3
2 2 10 10 0 45% 0 56% 2 3 4 5 2 3 4 5 0 10 10 10 10 0 10 10 10 10
5 5 10 10 76543210 76543210 10 4 10 4
10 3 10 3 lung 2 2 10 10 CD62L 0 23% 0 49% 2 3 4 5 2 3 4 5 0 10 10 10 10 0 10 10 10 10 CFSE
B. BCG+OVA OVA 80
60 **
+ T cells *
/ KJ 40 low
D62L 20 C %
0 MLN lungs
Figure 2.5. BCG infection increases accumulation of effector antigen-specific T cells in the lungs after airway antigen challenge. Flow-sorted CD62LhiCD44low naïve 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 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 expression of CD62L and CFSE dilution analyzed.
Numbers represent percentage of CD62Llow T cells among KJ+ T cells that have divided more than twice
(CFSElow). B. Percentage of CD62Llow T cells among KJ+ T cells, n=3 mice per group; *, p=0.02; **, p=0.03. Data representative of 2 separate experiments.
41 differences between the two groups were magnified in BCG-infected lungs. Thus,
infection causes increased differentiation of activated 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-γ and IL-4 production. BCG infection increased the frequency of IFN-γ producing
KJ+ T cells among OVA-specific T cells present in the lungs of OVA-challenged mice
(Fig. 2.6A). We did not detect any IL-4-producing 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-producing cells by ELISPOT assay.
As shown in Fig. 2.6B, similar numbers of IL-4 spot-forming units (SFU) were found between infected and uninfected mice, whereas more IFN-γ SFU were observed in infected and OVA-challenged mice. This demonstrates 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 activation, proliferation, and differentiation of antigen-specific T cells in the lungs of infected mice is activation of the
pulmonary innate immune cells responsible for initiating adaptive responses. CD11c+
lung DCs are efficient at presenting airway antigens to both naïve and effector T cells
(Byersdorfer and Chaplin 2001). To determine whether CD11c+ lung cells had a role in
42 A. Isotype OVA BCG+OVA 30 30 30 8.8±1.4% 19±3.5%* 20 20 20
10 10 10
0 0 0 0 102 103 104 105 2 3 4 5 0 102 103 104 105 0 10 10 10 10
IFN-γ
B. BCG+OVA + pep OVA + pep BCG+OVA OVA
160 ells ** g g c 120 Lun 5 80 5X10 40 SFU/ 0 IL-4 IFN-γ
Figure 2.6. Mycobacterial infection induces naïve OVA-specific T cells to differentiate into Th1 effectors in response to airway OVA challenge. Flow-sorted CD62LhiCD44low naïve 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 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-γ and analyzed by flow cytometry. Histograms were gated on KJ+ T cells. Isotype for anti-IFN-γ is shown. Numbers represent means ± SD of triplicate wells for each group; *, p=0.03. B.
5x105 ˚lung cells were cultured 48 h at 37 C with or without 2 μM OVA323-339 peptide. IL-4 and IFN-γ spot forming units (SFU) were quantified by ELISPOT. **, p<0.01. Figure is representative of 3 separate experiments.
43 the increased proliferation 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.
2.7A). This is in agreement with other published findings where intranasally
administered antigens had been tracked in vivo (Vermaelen, Carro-Muino et al. 2001;
Constant, Brogdon et al. 2002).
Thus to determine if infected and uninfected mice differed in distribution of
Fluos-OVA within the CD11c+ lung cell population, we gated on Fluos+ CD11c+ cells and measured the DC markers CD11b and CD11c (Gonzalez-Juarrero and Orme 2001).
In BCG-infected mice, 19% of cells harboring Fluos-OVA were DCs (CD11bhiCD11chi)
whereas in uninfected mice 8.2% of Fluos+ cells were DCs (Fig. 2.7B). There were
greater numbers of CD11bhi CD11chi DCs 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-specific naïve T cells (MartIn-
Fontecha, Sebastiani et al. 2003).
The maturation of lung DCs in infected lungs is demonstrated by high levels of
major histocompatibility complex class II (MHC-II; I-Ad) expression (Fig. 2.7C).
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-IIhi cells acquired intranasal antigen and accumulated in the lungs (Fig. 2.7D). To determine whether increased
44 A. No OVA BCG+OVA OVA
105 105 105
4 10 104 104
3 10 103 103 2.8±0.1% 3.0±0.1% 2 10 102 102
0 0 0 11c
2 3 4 5 0 10 10 10 10 0 102 103 104 105 0 102 103 104 105 CD
Fluos-OVA 2µM p O VA E. 60 B. BCG+OVA OVA BCG+OVA 5 5 50 10 10 OVA 2 (ng/ml)
4 10 4 10 40 IL-
3 10 3 19±1.7%* 10 8.2±3.0% 30
2 10 2 10 0 0 20
D11c 2 3 4 5 0 10 2 10 3 10 4 10 5 0 10 10 10 10 C 10 CD11b DOBW response: 0 10000 100000 D. BCG+OVA Lung CD11c+ cells C. BCG+OVA OVA hi 16 OVA ) 4 isotype **
(×10 12 s c+MHC-II 8 cell c 4 s+CD11
0 102 103 104 105 dendriti
Fluo 0 I-Ad lungs1
Figure 2.7. BCG infection causes up-regulation of MHC-II on lung CD11c+ cells harboring intranasal Fluos-labeled OVA. BCG infected and uninfected mice were given 450μg of Fluos-labeled OVA intranasally. 18-24 hrs. later 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 vs. SSC plots were gated on live cells and dot plots of individual mice from 3 groups are shown examining CD11c and Fluos. Percentages represent live cells (FSC vs. SSC gate) harboring Fluos-OVA, n=3 mice per group. B. Plots have been gated on Fluos+ CD11c+ cells. Percentages represent distribution of Fluos-OVA to dendritic cells (CD11chiCD11bhi) within the CD11c+ population; n=3; *, p=0.01 C. Histogram plots show surface expression of MHC-II on the gated populations shown in (A) containing lung CD11c+ cells that harbor Fluos-OVA. Geometric mean fluorescent intensities of I-Ad staining are 31000 ± 5300 and 1700 ± 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 (MFI>3000); 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 2μM OVA peptide and 1×105 DOBW cells. IL-2 in the culture supernatants was assayed by ELISA. Similar data obtained in 2 separate experiments.
45 expression of I-Ad resulted in greater presentation of pOVA-I-Ad complexes 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 cells was assessed 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. 2.7E).
Together, these data suggest that ongoing pulmonary mycobacterial infection increases
the number and maturation of lung DCs such that they can activate antigen-specific naïve
CD4+ T cells in the lungs themselves.
Discussion
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 naïve CD4+ T cells in MLN
and lung? (ii) Does pulmonary inflammation promote naïve 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 recent reports suggesting that
infection affects how and where naïve T cells are activated in the lungs was of particular
interest (Lund, Partida-Sanchez et al. 2002; Moyron-Quiroz, Rangel-Moreno et al. 2004;
Schreiber, Ehlers et al. 2006). BCG-infected mice after airway-OVA challenge had
46 greater accumulation, activation, and proliferation of naïve OVA-specific CD4+ T cells in
the draining MLN than did uninfected mice. Prolonged pulmonary inflammation also
was associated with naïve CD4+ T-cell activation in lung tissue. In contrast, in uninfected mice there was little to no naïve 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-γ 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 exogenous OVA peptide by lung
CD11c+ cells.
Pulmonary BCG infection allowed measurement of the naïve CD4+ T-cell
responses to an unrelated airway-antigen in mice in the presence or absence of prolonged
pulmonary inflammation. Pulmonary infection and inflammation peak 4 to 6 weeks after
aerosolized BCG infection (Kuchtey, Fulton et al. 2006). Naïve OVA-specific 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 infection of in vitro-generated
APCs on naïve T-cell activation in vivo (Bhatt, Hickman et al. 2004). Our experimental
system allowed for the characterization of naïve CD4+ T-cell activation by endogenous
lung APCs during mycobacterial infection. In addition, we could control the amount of
antigen and the precursor frequency of transferred naïve CD4+ TCR transgenic T cells in infected and uninfected mice. Mice were challenged with a high concentration of antigen to maximize access to antigen and minimize clonal competition (Catron, Rusch et al.
2006). Our experimental system specifically addressed infection-induced pulmonary
47 modulation of naïve 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 (Roman, Miller et al. 2002; Lawrence and Braciale 2004; Shi,
North et al. 2004; Jelley-Gibbs, Brown et al. 2005).
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 (Topham, Castrucci et al.
2001). In vivo proliferation as measured by CFSE demonstrated that infected mice had a
higher T- cell responder frequency than did uninfected 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 naïve 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 (Shiow,
Rosen et al. 2006). 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.
48 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 proliferation in the lungs
does not rule out that OVA-specific naïve 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-associated lymphoid tissue, within the lungs of infected mice that facilitate
naïve T-cell activation and proliferation in the absence of peripheral lymphoid organs
(Moyron-Quiroz, Rangel-Moreno et al. 2004). In addition lungs from M. tuberculosis- infected mice express CCL21 and CCL19, naïve T-cell chemoattractants, and contain organized neolymphoid structures (Schreiber, Ehlers et al. 2006). Secondary lymphoid structures induced by chronic infection could serve as sites of naïve 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 (Constant, Brogdon et
al. 2002). BCG infection may enhance T-cell activation and effector cell development by
causing a greater influx of naïve T cells into lymphoid tissues in the mediastinum and
lung, as has been demonstrated during herpes simplex virus type 2 infection (Soderberg,
Payne et al. 2005). This increases the likelihood that during BCG infection, greater
numbers of naïve T cells are exposed to cognate antigen presented by DCs in lymphoid
tissues. Increased exposure to cognate antigen results in increased naïve T-cell activation
and division as observed in our experiments (Miller, Safrina et al. 2004). In addition,
Catron et. al. have shown that the presence of greater numbers of naïve T cells in lymph
49 nodes at the time of antigen challenge facilitates the generation of effector T cells
(Catron, Rusch et al. 2006).
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 (Uehori, Matsumoto et al. 2003; Iwasaki and Medzhitov 2004).
Our studies indicate that pulmonary BCG infection results in the maturation of lung
CD11c+ cells harboring airway OVA. Lung CD11c+ cells from infected mice were more
capable of presenting exogenous OVA peptide ex vivo than lung CD11c+ cells from uninfected mice. Concomitant with DC maturation, activation of Toll-like receptors on
DCs enhances migration from peripheral sites to draining lymph nodes by upregulating
CCR7 (Iwasaki and Medzhitov 2004). 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 stimulation
(Eisenbarth, Piggott et al. 2002). BCG infection enhanced the differentiation of OVA-
+ specific naïve CD4 T cells in the lungs into Th1 effectors. This is in agreement with
previous studies that examined the role of BCG infection in promoting allergen-specific
Th1 responses (Erb, Holloway et al. 1998; Sano, Haneda et al. 1999).
Therefore, we propose two complementary mechanisms for the initiation of naïve
CD4+ T-cell responses in the lungs during mycobacterial infection. Pulmonary infection
50 causes lung DC maturation and increases DC trafficking to MLN to initiate robust naïve
CD4+ T-cell responses in the lymph node. Alternatively, BCG infection causes lung DC maturation and in situ activation of naïve CD4+ T cells in the lungs. Additional studies will determine where naïve CD4+ T-cell activation occurs within different lung compartments during chronic inflammation: the alveolar space, the bronchus-associated lymphoid tissue, or the lung parenchyma.
51
CHAPTER 3
Generation of a MTB-specific T cell receptor transgenic mouse
52 Summary
To study modulation of MTB-specific naïve CD4+ T-cell responses during the
course of mycobacterial infection, we set out to generate a MTB-specific T cell receptor
(TCR) transgenic mouse specific for the antigen Ag85B of MTB. Initial characterization of the TCR present on the Ag85B-specific T cell hybridoma, BB7, identified its Vβ chain as Vβ11 using monoclonal antibody. To identify the Vα chain, total RNA from BB7 was extracted and used to perform RT-PCR using primers specific for Vα families. The amplified product was cloned, sequenced, and searched using BLAST and IMGT/V-
QUEST. The TCR specific for Ag85B was identified as Vβ11-D-Jβ2.1 and Vα5-
Jα30. Genomic DNA was extracted from BB7 and amplified using primers designed for
Vα5 and Vβ11. The product was being cloned into 20kb pTα and pTβ expression
vectors, when it was learned that another laboratory had made an Ag85B-specific TCR
transgenic mouse called P25 TCR tg. The TCR chains used to construct the P25 TCR tg
mice are Vα5-Jα15 and Vβ11-Jβ2.3. The mice were shipped to our laboratory. P25 TCR
tg mice will be a valuable tool to understand MTB-specific naïve CD4+ T-cell responses
during different stages of pulmonary infection.
53 Introduction
The previous chapter demonstrated that ongoing pulmonary mycobacterial infection leads to enhancement of naïve CD4+ T-cell responses directed against an airway
model antigen. In an effort to understand how applicable those observations are to naïve
MTB-specific CD4+ T-cell responses during mycobacterial infection, we set out to
generate a TCR transgenic mouse whose T cells recognize a defined Ag of MTB.
Most of the TCR transgenic mice currently available are specific for model
antigens or a limited number of viral epitopes. TCR transgenic mice specific for bacteria
include Salmonella-specific SM1 mice and Chlamydia-specific NR1 mice (McSorley,
Asch et al. 2002; Roan, Gierahn et al. 2006). This chapter reports our pursuit of a
tuberculosis-specific TCR transgenic mouse. We were fortunate by the generation in our
lab of a highly sensitive T cell hybridomas BB7 with specificity towards an epitope
present in Ag85B, a mycolyl transferase of MTB (Wiker and Harboe 1992). The epitope,
as presented by the MCH Class II molecule I-Ab, was determined to be around amino
acids 241-256 of Ag85B (Noss, Pai et al. 2001).
The TCR of BB7 was cloned and sequenced. The data in this chapter show that
the variable regions of the TCR α and β chains of BB7 are Vα5-Jα30 and Vβ11-D-Jβ2.1,
respectively. Cloning of these genes into pTα and pTβ expression vectors proved to be
difficult due to the natural tendency of the homologous sequences in these vectors to
recombine at a high rate. It was learned at this time that an Ag85B-specific TCR transgenic mouse called P25 TCR tg had been made by others. Presently, the P25 TCR tg mice have been provided to us. P25 mice will be a useful tool in adoptive transfer
54 experiments to measure MTB-specific naïve CD4+ T-cell responses at different stages of pulmonary mycobacterial infection.
Materials and Methods
Cell staining
5×105 T cell hybridomas were stained in each well of a V-bottom plate with a panel of FITC-conjugated anti-Vβ monoclonal antibodies (BD Pharmingen) diluted in staining buffer (1% BSA in 1X PBS) for 30 mins at 4˚C. The cells were washed twice and fixed with 1% paraformaldehyde before being acquired in a BD FACS Scan flow cytometer.
RNA extraction and RT-PCR
For the β chain: total RNA was extracted from BB7 using RNeasy Kit (Qiagen).
First strand cDNA was synthesized using oligo dT primers and SuperScript II reverse transcriptase (Invitrogen). There are two possible β chain constant regions (Cβ1 and
Cβ2) in mice. Primers for both β chain constant regions were made and they were used along with the Vβ11 primer. The Vβ11 primer, was obtained from literature (Kouskoff,
Signorelli et al. 1995). These primers (Table 3.1) were then used to PCR amplify the β chain from BB7 cDNA using Platinum pfx DNA polymerase (Invitrogen). PCR cycling parameters were: 30 s melting at 94˚C, 1min. annealing at 65˚C, and 2 min. extension at
68˚C.
For the α chain: cDNA was made as mentioned above. RNA ligase mediated rapid amplification of cDNA ends, RLM-RACE PCR (Invitrogen) was done according to
55 Table 3.1 Primer sequences used for RT-PCR screens and genomic cloning of Vα and Vβ regions of T cell receptor on BB7 hybridoma.
cDNA amplification primers
Vβ11 5’-AGCCACTCTCGAGTCTTGCCATGGCCCC-3’
Cβ1 5’- GCCAAGCACACGAGGGTAGCCTTT-3’ Cβ2 5’-AGCTCCACGTGGTCAGGGAAGAAGC-3’
Vα primer pool sequences are from literature (Kraig, Pierce et al. 1996) Cα 5’-CCTCGGTCTCAGGACAGCACC-3’
Genomic PCR primers
ALPHA5 5’-ATGAAGACAGTGACTGGACCTTTGTTC-3’
ALPHA3 5’-CCTGCCTGCCTTGGTCACGAT-3’
BETA5 5’-AGCCACTCTCGAGTCTTGCCATGGCCCC-3’
BETA3 5’-GGGGTCTTTGTGGCTGACTGT-3’
56 manufacturer’s instruction. As an alternative for identifying the Vα region of BB7, pools of consensus primers for Vα families generously provided by Dr. Ellen Kraig (Kraig,
Pierce et al. 1996), were used in conjunction with Cα primer (Table 3.1) to amplify the cDNA of BB7 using Platinum pfx DNA polymerase. PCR cycling parameters were: 30 s melting at 94˚C, 1 min. annealing at 55˚C, and 2 min. extension at 68˚C.
Cloning and Sequencing
Immediately after the RT-PCR using Platinum pfx, native Taq DNA polymerase was added to add A and T overhangs at the ends of the amplicon. The amplified α and β chains were separately cloned into pCR2.1TOPO vector (Invitrogen) using TA cloning according to manufacturer’s instruction. The ligation reaction was transformed into
TOP10 (Invitrogen) chemically competent cells. Ligations were plated on LB-Amp plates and incubated at 37˚C overnight. Colonies were picked and screened by PCR for the Vα and Vβ inserts before being plated again for plasmid isolation using Plasmid Spin
Miniprep Kit (Qiagen). Plasmid DNA was sent for sequencing.
Colony PCR to check Vα and Vβ inserts
Autoclaved toothpicks are used to dab colonies on a plate and place toothpick in
20 μl of sterile water in 0.5 ml tubes. Toothpick is used to mix dab of colony into water.
Mixture is heated at 94˚C for 10 min and then placed on ice. Aliquots of this mixture is used as template to perform PCR using primers for full-length and nested products to confirm specificity of amplified product.
57 Genomic cloning into pTα and pTβ vectors
Genomic DNA was isolated from BB7. The genomic sequence of the TCR was deduced by performing BLAST and IMGT/V-QUEST (Giudicelli, Chaume et al. 2004) searches using the sequences derived from RT-PCR as queries. This was done to design primers positioned in the leader sequence upstream of the Vα and Vβ regions and in the
introns following the specific J regions for insertion into the pTα and pTβ cassette vectors
(Kouskoff, Signorelli et al. 1995), respectively. The cassette vectors were generously
provided by Dr. Stephen McSorley. For the α chain a forward primer ALPHA5 was
designed for the leader sequence of Vα5 that had an artificial XmaI restriction site and a
reverse primer ALPHA3, incorporating a NotI site, was designed specific for the putative
intron 188 bp following the Jα30 exon. The ALPHA5 and ALPHA3 primers (Table 3.1)
were used to amplify the genomic rearranged product from total BB7 DNA. For the β
chain, the primer BETA5 specific for the Vβ11 leader sequence was obtained from
literature (Kouskoff, Signorelli et al. 1995). BETA5 had a XhoI restriction site. The
BETA3 primer was designed in the intron region 61 bp downstream of the Jβ2.1 exon, incorporating a SacII restriction site. BETA5 and BETA3 (Table 3.1) were used to amplify the genomic DNA of BB7. Cycling parameters for both Vα and Vβ genomic
PCR were identical to that used for RT-PCR.
Amplified genomic VJα and VDJβ were cloned into pCR2.1TOPO vector and sent for sequencing. Colonies of VJα and VDJβ, whose sequences matched those found in the literature and BLAST searches, were used for subsequent cloning. Plasmid DNA from these colonies were isolated with Plasmid Mini Kit (Qiagen) and digested with
XmaI and NotI for VJα colonies and XhoI and SacII for VDJβ colonies. Likewise pTα
58 vector was digested with XmaI and NotI and pTβ vector digested with XhoI and SacII.
Digested vectors were treated with shrimp alkaline phosphatase (Promega) and used for
ligation reactions. VJα and VDJβ inserts were gel purified and used in ligation reactions
with dephosphorylated corresponding pT vectors using T4 DNA ligase (NEB). Ligation
reactions were transformed into STBL-2 cells (Invitrogen) and grown overnight at 30˚C.
Resulting pTαVα and pTβVβ colonies were screened by PCR. STBL-2 colonies with the
correct inserts were further grown overnight at 30˚C in minimal TB media with ampicilin
(50 ug/ml). Plasmids were isolated using PureYield Plasmid Midiprep System
(Promega). Integrity of the unwieldy pT vectors was determined by restriction digests
with BamHI and EcoRI (NEB).
Results
b The Ag85B241-256-specific, I-A -restricted BB7 T cell hybridoma expresses Vβ11
The general procedure for making a TCR transgenic mouse involves identification
of the α and β chains of the TCR and then isolation and cloning of the genetic message
encoding the α and β chains. Due to unavailability of many Vα antibodies, monoclonal
Vβ antibodies were used to find the Vβ chains. BB7 expresses Vβ11 (Fig. 3.1). Another
T cell hybridoma AD9, with specificity towards the Ag85B241-256 epitope expresses Vβ8
(Fig. 3.1). The Vβ8 monoclonal antibody does not distinguish between Vβ8.1 and 8.2.
BB7 expresses Vα5-Jα30 and Vβ11-D-Jβ2.1
To identify the α chain of BB7 the technique of RNA ligase mediated rapid amplification of cDNA ends, RLM-RACE PCR was used. The method involves ligating
59
BB7 AD9 98% 93% M1 M1
anti-Vß11 anti-Vß8.1/8.2
Figure 3.1. Vβ surface expression on Ag85B-reactive T cell hybridomas. 5 5.0×10 T cell hybridomas were stained in the dark at 4˚C for 30mins with monoclonal antibodies diluted in staining buffer (1%BSA in PBS). Cells were fixed in 1% paraformaldehyde and analyzed with a BD FACS Scan flow cytometer.
60 a known RNA oligonucleotide sequence upstream (5’) of mRNAs and PCR amplifying the message of interest (TCR α chain mRNA) with a gene-specific 3’ primer (Cα primer).
Both times the procedure was done, a truncated α chain mRNA was amplified. BLAST searches confirmed the cryptic Vα message as that of BB7’s fusion partner, BW5147
(White, Blackman et al. 1989). Due to the abundance of the cryptic message, the Ag85B- specific TCR’s Vα message was unable to be identified using this method.
A new Vα cloning scheme was undertaken, Fig. 3.2. PCR was done using BB7 cDNA as template, the constant region Cα primer as the reverse primer, and Vα consensus primer pools as forward primers (Kraig, Pierce et al. 1996). The primer pools were designed to be specific for sequences near the VαJα junctional region. Forward primers ‘Vα123’, ‘Vα102’, ‘Vα105’, and ‘Vα116’ gave amplified products. However primer ‘Vα116’ was known to be specific for the cryptic Vα product. Amplified products from the ‘Vα123’, ‘Vα102’, and ‘Vα105’ PCRs that were greater than the ‘Vα116’ product in size, were further used as templates in secondary PCRs with ‘Vα116’ as the forward primer and Cα primer as reverse primer. Only the amplified products that failed to give a product in the secondary PCRs were cloned and sent for sequencing. The sequences were queried against mouse mRNA sequences using BLAST. It was found that one of the amplified products showed sequence homology to the Vα region of a mouse TCR mRNA that was not the cryptic BW5147 Vα. No ‘hits’ were found to be identical in Vα and Jα to the amplified BB7 product. Since the primer pools used were specific for the Vα-Jα junctional region, the amplified and cloned product did not contain upstream Vα sequence information, which was needed for cloning into pTα vector, Fig
3.2.
61
Full-length mRNA Vα Vα consensus primer
Step 1 m G-p-p-p 7 L V J C α AAAA
C α primer
PCR amplify, Sequence, and BLAST against genomic DNA to identify Vα, Jα and intron sequences in leader region and after the Jα region
Step 2 L primer w/ Xma-I site Genomic DNA TCR α chain 3’ 5’ L Vα5 Jα30 Jn Cα En Intron primer w/ Not-I site
PCR amplify, Sequence, Insert this into pTα vector Insert fragment into Step 3 pTα
Xma-I Not-I pTα 3’ J Cα En pTα vector contains endogenous TCR promoter
Figure 3.2. Vα cloning scheme. V alpha consensus primers were obtained from Dr. Ellen Kraig. pT alpha vector was obtained from Dr. Stephen McSorley.
62 To find the Vα upstream sequence, only the Vα portion of the amplified sequence was searched against mouse genomic and mRNA databases. Genomic and mRNA ‘hits’ were found that identified the Vα of the amplified product as Vα5. Comparing the genomic and longest mRNA ‘hits’, as shown in Fig 3.3, the very beginning of the Vα5 locus was identified by the presence of a leader sequence. A forward primer ALPHA5 was designed for the Vα leader sequence that had an artificial XmaI restriction site to enable cloning into a similar site in pTα vector. The Jα sequence of the ‘Vα123’ product was ‘BLAST’ed against genomic sequences and the ‘hit’ was to Jα30. The α chain mRNA lacks the intron following Jα30. Comparison of genomic and mRNA ‘hits’ for
Jα30 revealed a putative intron sequence. A reverse primer ALPHA3, incorporating a
NotI site, was designed specific for the putative intron 188 bp following the Jα30 exon.
The ALPHA5 and ALPHA3 primers were used to amplify the genomic rearranged product from total BB7 DNA. PCR cycling parameters were: 30 s melting at 94˚C, 1 min. annealing at 55˚C, and 2 min. extension at 72˚C. The amplified product along with a nested PCR product are shown in Fig 3.4B. The full length product was cloned into pCR2.1 vector for sequencing Fig 3.5A.
The Vβ cloning was made much simpler knowing that BB7 expressed Vβ11, Fig.
3.1. Total RNA was isolated from the subclones and RT-PCR was done using published
Vβ11 and Cβ primers (Kouskoff, Signorelli et al. 1995). The sequence of the amplified product revealed the TCR β chain to be Vβ11-D-Jβ2.1. The sequence of the amplified
Vβ product was used to generate genomic ‘hits’. A genomic ‘hit’ was used to find a putative sequence for the intron lying downstream of the Jβ2.1 exon. A primer was designed 61 bp downstream of the Jβ2.1 exon, incorporating a SacII restriction site. PCR
63
A. >gi|14253141|emb|AJ311366.1|MMU311366 Mus musculus mRNA for T-Cell receptor alpha chain (TRAV gene), 5'UTR Length = 478
Score = 325 bits (164), Expect = 1e-86 Identities = 257/288 (89%) Strand = Plus / Plus
Query: 1 atgaagacagtgactggacctttgttcctgtgcttctggctgcagctgaact 52 |||||||||||||||||||||||||| ||||||||||||||||||||||||| Sbjct: 50 atgaagacagtgactggacctttgttgctgtgcttctggctgcagctgaact 101
Query: 212 gtgtgagcagaggcgagcaggtggagcagcgcccccctcacctgagtgtccgggag 267 |||||||||||||||||||||||||||||||||| ||||||||||||||||||||| Sbjct: 102 gtgtgagcagaggcgagcaggtggagcagcgccctcctcacctgagtgtccgggag 157
B. ATGAAGACAGTGACTGGACCTTTGTTCCTGTGCTTCTGGCTGCAGCTGAACTGTGAGTGAGAGCTTTTGGG GAACAAGGTGTTTGTCAGTGTTCATTATCAGTCTGTGGAGCAGACTGGACTTCTCTAGATTGCCTGAAATC ACTGTAGGGAGGAAAGCAATGTCTGGAGACTGGCAGCCTGATGGTCTTCTGCTGTTTCCTTTTGCACAGGT GTGAGCAGAGGCGAGCAGGTGGAGCAGCGCCC
Figure 3.3. BLAST search identifying Vα5 leader sequence. Genomic and mRNA ‘hits’ were obtained for the ‘Vα123’ product. A. Comparing the genomic ‘hit’ (Query) with the longest mRNA ‘hit’, a region extending from 53-211bp was missing from the mRNA and thus deduced to be intronic DNA separating leader sequence from Vα region. Therefore the shared exon region 1-52bp in the genomic ‘hit’ and 50- 101bp in mRNA ‘hit’ was presumed to be the leader sequence of Vα5. B. The intron separating leader sequence from Vα5 exon is underlined.
64 Vβ11Jβ2.1 Vα5Jα30 10 100 0 5-3N 5-3N bp 5N-I 5N-I 5-I 5-I 5-I bp Φ 5-I Φ 5-I 5-I Φ Φ
760 500 540 520 500 350
Colony #46 #47 #38 Colony #14 #15 #21
Figure 3.4. Colony PCR of genomic Vα and Vβ sequences of BB7 The lanes “5-I” are the full length inserts cloned into the respective vectors. Lanes “5N-I” and “5-3N” denote nested PCR products of the beta and alpha chains respectively to confirm specificity of bands .
65 A. ATGAAGACAGTGACTGGACCTTTGTTCCTGTGCTTCTGGCTGCAGCTGAACT GTGAGTGAGAGCTTTTGGGGAACAAGGTGTTTGTCAGTGTTCATTATCAGTCTGTG GAGCAGACTGGACTTCTCTAGATTGCCTGAAATCACTGTAGGGAGGAAAGCAATGT CTGGAGACTGGCAGCCTGATGGTCTTCTGCTGTTTCCTTTTGCACAGGTGTGAGC AGAGGCGAGCAGGTGGAGCAGCGCCCCCCTCACCTGAGTGTCCGGGAGGGA GACAGTGCCGTTATCACCTGCACCTACACAGACCCTAACAGTTATTACTTCTT CTGGTACAAGCAAGAGCCGGGGGCAAGTCTTCAGTTGCTTATGAAGGTTTTC TCAAGTACGGAAATAAACGAAGGACAAGGATTCACTGTCCTACTGAACAAGA AAGACAAACGACTCTCTCTGAACCTCACAGCTGCCCATCCTGGGGACTCAGC CGCGTACTTCTGCGCAGTCA{JCGCCAGGCAATACCGGAAAACTCATCTTTGG ACTGGGGACAACTTTACAAGTGCAACCAG}GTGGGTCTGAAGGTTCCCACGTG AACCAGCTGTCGCTTATGCTCATTTTAAACTAGTCTTCTTCATCTTTTCTAACTAGAG ACTCTTGGTTTCAGGCTTTCCTTTCTTTTTAAGAAGCAGAGTCTGTAGAGTAAGTAA CAAGACCTCCTCTGCCTGTTGAGTTTCATATCGTGACCAAGGCAGGCAGG
VALPHA5: XmaI site underlined TCTCCCGGGCTTCTCACTGCCTAGCCATGAAGACAGTGACTGGACCT
VALPHA3: NotI site underlined TAAGCGGCCGCCTGCCTGCCTTGGTCACGAT
B. AGCCACTCTCGAGTCTTGCCATGGCCCCCAGGCTCCTTTTCTGTCT GGTTCTTTGCTTCTTGAGAGCAGGTGAGCTTCCAGGAATAAAGAAGCTGCAGCTT TTGATCATTGTTTGTCATAGCTACAATTACATTCTTTATTCTCTATTCTGTTTCTTTCT AGAACCAACAAATGCTGGTGTCATCCAAACACCTAGGCACAAGGTGACAGG GAAGGGACAAGAAGCAACTCTGTGGTGTGAGCCAATTTCAGGACATAGTGCT GTTTTCTGGTACAGACAGACCATTGTGCAGGGCCTGGAGTTCCTGACTTACTT TCGAAATCAAGCTCCTATAGATGATTCAGGGATGCCCAAGGAACGATTCTCA GCTCAGATGCCCAATCAGTCGCACTCAACTCTGAAGATCCAGAGCACGCAAC CCCAGGACTCAGCGGTGTATCTTTGTGCAAGCAGCTTAGAAGC{JTGCTGAGC AGTTCTTCGGACCAGGGACACGACTCACCGTCCTAG}GTAAGAAGGCAGAGGC CATACAGGTGGGAGAGAAGGTAGGACAGTCAGCCACAAAGACCCC
VBETA5: XhoI site underlined AGCCACTCTCGAGTCTTGCCATGGCCCC
VBETA3: SacII site underlined TGCTCCGCGGGTCTTTGTGGCTGACTGT
Figure 3.5. Vα5Jα30 and Vβ11DJβ2.1 sequences of BB7 TCR A. Vα sequence along with primers. B. Vβ sequence along with primers. In the sequences the underlined portions are the introns. “{J” denote J segment. Bold letters in sequence are complementary to the primers.
66 cycling parameters were: 30 s melting at 94˚C, 1 min. annealing at 65˚C, and 2 min.
extension at 72˚C. The amplified Vβ product was run on an electrophoretic gel along
with a nested PCR of the product to confirm size, Fig. 3.4A. Then the product was
cloned into pCR2.1 for sequencing, Fig. 3.5B.
Vα5-Jα30 and Vβ11-D-Jβ2.1 of BB7 cloned into pTα and pTβ vectors fail to be expressed
Vα5-Jα30 and Vβ11-D-Jβ2.1 segments were cloned into the TCR expression vectors pTα and pTβ respectively. The Vβ cloning was done first because pTβ had lower
propensity to recombine. Several pTβ-Vβ colonies were screened with PCR using
BETA5 and BETA3 primers. Colonies with Vβ11 inserts were grown and their plasmids
purified. Restriction digests with EcoRI and BamHI were done to ensure the pTβ-Vβ
colonies with inserts retained plasmid integrity, Fig. 3.6. Three pTβ-Vβ colonies were
selected for electroporation: #3, #11, and #12. For the pTα-Vα cloning, many more
colonies needed to be screened by PCR due to the high likelihood for pTα to recombine.
Integrity of colonies with Vα inserts was ascertained with BamHI restriction digests, Fig.
3.7. pTα-Vα colony #29 was chosen having retained restriction digest pattern as native
pTα. The cloned products were transformed into STBL-2 cells which minimize
recombination events. Resulting colonies were checked by ‘Colony PCR’ using primers
for the inserts only. As we learned later, this cloning step into the pTα and the pTβ
vectors leads to the greatest number of recombination events in the unwieldy 20kb pTα
and pTβ vectors. Subsequently, we tried in vain to transfect T cell hybridoma cells with
these ‘successfully’ cloned pTαVα and pTβVβ products.
67 EcoRI BamHI
M pTβ #3 #11 #12 #13 pTβ #3 #11 #12 #13 M
Figure 3.6. Restriction digests of pTβ-Vβ colonies. Plasmids were isolated from pTβ-Vβ colonies (numbered) with Vβ inserts (as confirmed by colony PCR). Isolated pTβ-Vβ plasmids were digested with EcoRI and BamHI to ascertain integrity of the plasmids. Colonies 3, 11, and 12 retained the same restriction digest pattern as native pTβ vector while colony 13 had rearranged.
68 A. pTα M pTα pTα pTβ pTβ pTβ * *
B.
M 11 19 29 91 37 24 64 U M
Figure 3.7. Restriction digests of pTα-Vα colonies. A. BamHI digestion of native pTα and pTβ vectors (2 different gel loading). * lanes show patterns of band. B. BamHI restriction digest of pTα-Vα colonies confirmed to have Vα insert by colony PCR. Colony 29 still retains same banding pattern as native pTα. Lane U is uncut plasmid.
69 Discussion
In this chapter, the attempt to generate Ag85B-specific T cell receptor transgenic mouse was described. The identification of the variable, diversity, and junctional regions of the β chain of the TCR was straightforward but the identification of the variable and junctional regions of the α chain consumed more time. Once the identities of the two chains of the TCR were known, the sequences were amplified from genomic DNA and cloned into sequencing vectors before undertaking the difficult cloning into the unwieldy pTα and pTβ TCR expression plasmids. The cloning into the expression plasmids did not work out.
Our strategy to use the genomic sequences of the alpha and beta chain under the control of natural TCR promoters has been employed successfully by others (McSorley,
Asch et al. 2002; Roan, Gierahn et al. 2006). In addition, an earlier report had indicated that using cDNA based constructs under the control of heterologous promoters and enhancers led to defective expression of the transgenic TCR (Barnden, Allison et al.
1998). However, we learned that another group of researchers had generated a TCR transgenic mouse specific for the same epitope of Ag85B as the BB7 hybridoma
(Tamura, Ariga et al. 2004). They had used cDNA-based constructs under the control of
H-2Kb promoter. The P25 TCR has the same Vα and Vβ regions as BB7’s TCR but differs from BB7’s TCR in Jα and Jβ segments. As a result, the two TCRs have different
CDR3 regions based on amino acid sequence but may share similar biochemical property. Further work needs to be done to determine if the two TCRs differ in their affinity and specificity for P25:I-Ab.
70 The P25 TCR transgenic mouse was generously provided to us by Dr. Takatsu.
Future experiments will use the transgenic mice in adoptive transfer experiments to understand how naïve CD4+ MTB-specific T-cell responses are modulated during
pulmonary mycobacterial infection. The study designs will be similar to the one used in
Chapter 2. One caveat is that the expression of Ag85B varies during the course of mycobacterial infection (Shi, North et al. 2004) and thus experiments will need to be designed that examine P25 T-cell responses during peak expression.
71
CHAPTER 4
Modulation of pulmonary dendritic-cell function
during mycobacterial infection
72 Summary
We have previously reported that during mycobacterial infection naïve CD4+ T
cell activation in the lungs is enhanced. Here we investigated the role of CD11c+ dendritic cells (DCs) in the activation of naïve CD4+ T cells during pulmonary infection
with M. bovis bacillus Calmette-Guerin (BCG). BCG infection caused accumulation and
maturation of DCs in infected lungs even as the mycobacterial burden declined. Lung
DCs from infected mice expressed increased amounts of MHC-II but comparable
amounts of CCR7 relative to uninfected mice. Gene expression of a CCR7 ligand,
CCL19 progressively increased throughout BCG infection and the expression was
MyD88-dependent. Lung CD11c+ cells from BCG-infected mice activated OVA-specific
naïve CD4+ T cells more than lung CD11c+ cells from uninfected mice. Interestingly,
during peak bacterial burden lung DCs had slightly decreased chemotaxis towards CCR7
ligand CCL21 and were less efficient in activating naive CD4+ T cells compared to mice harboring few bacilli in the lung. Taken together, these findings suggest that during BCG infection, inflammation and sustained expression of CCL19 recruit and retain mature
DCs in the lung where they can activate naïve CD4+ T cells. The duration of pulmonary
mycobacterial infection may affect encounters between DCs and naïve CD4+ T cells in
the lung and influence subsequent naive T cell activation.
73 Introduction
During pulmonary mycobacterial infection, migration of dendritic cells (DCs) and
dissemination of mycobacteria to draining lymph nodes are thought to be important for a
successful cell-mediated immune response (Chackerian, Alt et al. 2002; Marino, Pawar et
al. 2004; Tian, Woodworth et al. 2005). Upon engagement of toll like receptors (TLRs)
on DCs by mycobacteria-derived pathogen-associated molecular patterns (PAMPs), DCs
undergo a coordinated maturation program that upregulates expression of the chemokine
receptor CCR7. Chemokine receptor CCR7 expression is essential for migration of DCs
to lymph nodes to coordinate adaptive immune responses following TLR stimulation
(Dieu, Vanbervliet et al. 1998; Sallusto, Schaerli et al. 1998; Sallusto, Palermo et al.
1999; Lande, Giacomini et al. 2003; MartIn-Fontecha, Sebastiani et al. 2003).
CCR7 and its ligands CCL19 (ELC, EBV-induced molecule 1 ligand chemokine)
and CCL21 (SLC, secondary lymphoid chemokine) are important for both the initiation
and regulation of adaptive immunity by directing migration of mature dendritic cells,
naïve CD4+ T cells, and central memory T cells to secondary lymphoid organs (SLO)
(Randolph, Angeli et al. 2005; Randolph, Sanchez-Schmitz et al. 2005). CCL19 and
CCL21-ser are produced by stromal cells within the T cell zones of SLO. CCL21-ser is also expressed on high endothelial venules while CCL21-leu is expressed in lymphatic endothelium. Upon maturation DCs also express CCL19 (Ngo, Tang et al. 1998). Both
CCL19 and CCL21 are involved in the organization of lymphoid structures under normal and chronic inflammatory conditions by facilitating encounters among stromal cells, T cells, B cells and dendritic cells (Luther, Bidgol et al. 2002; Aloisi and Pujol-Borrell
2006; Marinkovic, Garin et al. 2006).
74 CCR7 may have a restricted role in immune responses to airborne pathogens.
Mice lacking expression of CCL19 and CCL21-ser (plt/plt) are able to mount delayed but protective immune responses to influenza infection (Moyron-Quiroz, Rangel-Moreno et al. 2004). Furthermore, CCR7-/- mice are able to control virulent mycobacterial infection similar to their wild-type counterparts (Kahnert, Hopken et al. 2007). However, in both models deficient in CCR7 signaling the role of lung DCs have not been explored. Since a common attraction for CCR7 ligands within SLOs facilitates interactions between naïve
T cells and mature DCs (Luther, Bidgol et al. 2002), we examined if pulmonary BCG infection affects CCR7-dependent interactions between lung DCs and naïve CD4+ T cells. This study was conducted to shed light on naive CD4+ T cell activation in the lungs. Using an attenuated strain of mycobacteria, Mycobacterium bovis BCG, we report that peak and late stages of infection differ in their modulation of CCR7-mediated DC-T cell interaction and subsequent naive CD4+ T cell activation in the lungs.
Materials and Methods
Mice
Eight- to ten-week-old female BALB/c and C57BL/6 mice were purchased from
The Jackson Laboratory (Bar Harbor, ME). MyD88-/- mice were obtained from Drs.
Osamu Takeuchi and Shizua Akira (Osaka University, Osaka, Japan) and backcrossed to
BL6 background for 7 generations. DO11.10 TCR transgenic mice that express TCRs
d specific for OVA323-339 peptide presented in the context of I-A (Murphy, Heimberger et al. 1990) were obtained from Alan Levine (Case Western Reserve University, Cleveland,
OH) and bred on-site. Mice were housed in specific pathogen-free conditions. All
75 studies were approved by the Institutional Animal Care and Use Committee at Case
Western Reserve University.
Aerosol BCG infection
Mice were exposed to aerosol M. bovis BCG in an Inhalation Exposure System
(Glas Col, Terre Haute, IN) as previously described (Kuchtey, Fulton et al. 2006).
Uninfected mice served as controls.
Tissue isolation
Tissues were harvested and processed as previously described (Kuchtey, Fulton et
al. 2006). Briefly, mice were anesthetized with a lethal dose of tribromoethanol (240
mg/kg). Abdominal cavity was incised, spleen harvested and the mouse exsanguinated.
The trachea was cannulated and bronchoalveolar lavage fluid (BALF) collected by
aspirating 3 times with 1 ml of PBS. Lungs were perfused with 10 ml of PBS and harvested. Then draining mediastinal lymph nodes (MLN) were harvested. Single cells were resuspended in complete medium (DMEM, 10% fetal bovine serum, 0.05 mM 2- mercaptoethanol, 2mM HEPES, 1mM sodium pyruvate, 100 mM non-essential amino acids, 100 U/ml penicillin and 0.1 mg/ml streptomycin). Lungs were minced and digested with 125 units/ml of type IV collagenase and 30 units/ml DNase for 90 min at
37˚C. Lung aggregates were drawn through a 18 G needle 3X before being pressed through a 40 μm nylon filter. Red blood cells were lysed and lungs resuspended in
RPMI. Serial dilutions of lung suspension were plated on 7H10 plates to determine bacterial CFU. Lung cells were then positively sorted for CD11c+ cells using N418
76 microbeads (Miltenyi Biotec). MLNs were pressed through a 70 μm nylon filter using the plunger of a 1 ml syringe and resuspended in RPMI.
Cell staining
Single-cell suspensions of tissues and sorted cells were counted and viability was assessed by trypan blue exclusion. A total of 5 × 105 to 1 × 106 viable 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 anti-CCR7 (4B12), anti-CD11c, anti-CD11b
(eBioscience), and anti-I-Ad (BD Pharmingen catalog number 553546). Cells were
incubated for 30 min at 37˚C according to manufacturer’s staining protocol
(eBioscience). Cells were washed once with 1% BSA and pellets were resuspended in
1% paraformaldehyde in PBS. Stained samples were acquired using a BD LSR II flow
cytometer. Flow cytometry results were analyzed with FlowJo (Tree Star, Inc.) software.
Lung CD11c+ cell chemotaxis assay
Lungs were sorted with magnetic CD11c (N418) beads (Miltenyi) as described
above. In 24-well transwell plates with 8 μm pore polycarbonate inserts (Corning), a
total of 4×105 viable lung CD11c+ cells were added to inserts and 600 μl of chemotaxis
medium (0.1% BSA in serum-free HL-1 medium) containing 100 ng/ml of CCL21
(R&D) was added to lower chambers. After 5 h of incubation at 37˚C, cells from the
lower chambers were collected and pelleted. Migrated cells were stained with anti-
CD11c, anti-CD11b, and anti-I-Ad and counted in a BD LSR II flow cytometer along
with 10 μm reference beads (Polysciences). Chemotaxis index was calculated by
77 dividing the number of mature lung DCs that migrated from the input DC population in
response to CCL21 by the number of DCs that migrated to medium alone.
RNA isolation and quantitative real-time PCR
Right lower lobes of murine lungs were excised and flash frozen on dry ice. Lung tissues were stored in -80˚C until RNA extracted using the RNAeasy Kit (Qiagen) following the manufacturer’s protocol. Freshly bead-sorted lung CD11c+ cells were
lysed and RNA extracted using RNAeasy Kit (Qiagen) according to manufacturer’s
instructions.
Yi Liu, a graduate student in Dr. Harding’s laboratory, performed the
quantitative-real time PCR assays. Total RNA yield was determined by
spectrophotometer, and 1 μg of total RNA was used in a reverse transcriptase reaction
(SuperScript First Strand Synthesis System, Invitrogen, Carlsbad, CA) to convert RNA to
single-stranded cDNA. One tenth of the resulting cDNA template was used for real time
PCR analysis with SYBR Green and the Bio-Rad iCycler fluorescence detection system.
Gene amplification was done at 95˚C for 15 s, 59˚C for 15 s, and 72˚C for 30 s for 40
cycles. A standard curve for each gene was generated by serial dilution of amplified
product standard of known starting concentration. The following primers were used:
CCR7 sense, 5’ TGT ACG TCA GTA TCA CCA GC 3’; CCR7 antisense, 5’ TTT
TCC AGG TGT GCT TCT GC 3’; CCL19 sense, 5’ TGT GGC CTG CCT CAG ATT AT
3’; CCL19 antisense, AGT CTT CCG CAT CAT TAG CAC 3’; CCL21 sense, 5’ TCC
AAG GGC TGC AAG AGA 3’; CCL21 antisense, 5’ TGA AGT TCG TGG GGG ATC
T; IL-10 sense, 5’ ATT TGA ATT CCC TGG GTG AGA AG 3’; IL-10 antisense 5’
78 CAC AGG GGA GAA ATC GAT GAC A 3’; GAPDH sense, 5’ CCA GGT TGT CTC
CTG CGA CT 3’; GAPDH antisense, 5’ ATA CCA GGA AAT GAG CTT GAC AAA
GT 3’. Quantification was determined based on a standard curve of known
concentrations for each gene and normalized to GAPDH as previously described
(Pennini, Pai et al. 2006).
Lung CD11c+ cell culture
Lung CD11c+ bead-sorted cells, enriched for DCs, were cultured at 1×106/ml in
HL-1 medium supplemented with 1% FBS for 18 h at 37˚C. Culture supernatants were harvested and spun down to eliminate cells. Supernatants were frozen at -20˚C. Pelleted cells were stained as described above to examine CCR7 surface expression.
DO11.10 T cell isolation
Splenocytes from 9-14 wks. old DO11.10 mice were isolated and red blood cells lysed in hypotonic lysis buffer (10 mM Tris-HCl and 0.83% ammonium chloride).
Splenocytes were used to obtain untouched CD4+ T cells using the CD4+ T cell negative
selection kit (Miltenyi Biotec) according to manufacturer’s instruction. The resulting
CD4+ T cells were subsequently stained with anti-CD62L and anti-CD44 mAbs and
FACsorted by gating on naïve (CD62LhiCD44low) T cells using a BD Aria cell sorter.
FACsorted naïve CD4+ T cells were then used in antigen presentation assays. FACsorted
CD4+ T cells were >98% CD44lowCD62Lhi and 65-75% of these naïve CD4+ T cells were
OVA-specific CD4+ T cells, as determined by staining with TCR-clonotypic antibody KJ
1-26 (Invitrogen).
79
Antigen presentation assay
+ CD11c sorted lung cells were incubated with different concentrations of OVA323-
339 peptide at 37˚C for 90 mins. Cells were washed and plated along with FACsorted
naïve (CD44lowCD62Lhi) CD4+ T cells (5×104) from spleens of DO11.10 mice. After 48
h of incubation at 37˚C, supernatants were harvested and IL-2 levels measured by
ELISA. Briefly, Immulon microtiter plates (Thermo) were pre-coated overnight at 4˚C
with anti-IL-2 capture antibody (eBioscience #14-7022) at 1 μg/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 IL-2-containing supernatants. After washing, plates were incubated
at room temperature with biotin-conjugated anti-IL-2 detection antibody (eBioscience
#13-7021) (1 μg/ml). Plates were washed and incubated with avidin-alkaline
phosphatase at room temperature for 30 min. Substrate was added and the plates were
read after 20-30 min at 405 nm.
Statistical analysis
All statistical analyses were performed using Student’s t test. A value of p<0.05
was considered statistically significant.
80 Results
BCG infection results in the accumulation and maturation of lung dendritic cells beyond
peak bacterial burden in the lungs.
Mice were infected with BCG by aerosol. BCG burden in the lungs peaked 4 to 6
weeks after infection and declined to <100 bacilli/lung by 12 to 14 wks (Fig. 4.1A). To
determine the effect of BCG infection on number and maturity of dendritic cells in the
lungs, mice were divided into 3 groups: uninfected mice, mice infected for 4 to 6 wks and
mice infected for 12 to 14 wks. Mice were sacrificed and their lungs harvested. Lung cells were stained for CD11b, CD11c, and I-Ad to measure the maturation of myeloid
dendritic cells (CD11bhi CD11chi) as described by Gonzalez et. al. (Gonzalez-Juarrero
and Orme 2001). Expression of MHC-II on myeloid DCs increased during infection and
remained elevated up to 12 to14 wks post-infection compared to uninfected mice (Fig.
4.1B). BCG infection also led to >8-fold increase in numbers of mature lung DCs
(CD11chi MHChi) in infected mice compared to uninfected mice (Fig. 4.1C).
Interestingly, maturation and accumulation of DCs in the lung persisted up to 12 to 14
wks after infection when lung bacterial burden had declined to the limit of detection (i.e.
<100 CFU/lung). Thus, BCG infection causes maturation of lung DCs. Furthermore,
activated lung DCs are retained in the lung well beyond peak mycobacterial burden in the
lungs.
81 A. B. C.
6 3 10 7000 # 700 5 10 6000 ## 600 ** hi 4 10 5000 500 hi * 3 4000 400 10 2 3000 300 10 hi ( Geometric MFI ) 2000 200 1 MHC-II on CD11c cells MHC-II on CD11c lung CFU per mouse lung CFU per <100 CFU/lung 10 1000 100
0 MHC cells (x10 ) CD11c mouse per 10 0 0 0 4-6 12-14 Uninf. 4-6 wks 12-14 wks Uninf. 4-6 wks 12-14 wks Time, post-infection (wks) Time, post-infection Time, post-infection
Figure 4.1. Accumulation and maturation of dendritic cells in the lung during BCG infection. A. Lungs from BALB/c mice at 1 day, 4-6 wks, and 12-14 wks after aerosol-BCG infection were processed and aliquots plated out to enumerate lung CFUs. Mean CFUs ± SD from a single BCG infection of 3 mice per time-point are shown. Similar bacterial burdens were measured in 3 separate experiments. Lung CD11c+ sorted cells from uninfected mice, mice infected for 4-6 wks, and mice infected for 12-14 wks were stained for MHC-II (I-Ad), CD11b, and CD11c. B. Maturation of myeloid (CD11bhiCD11chi) DCs was assessed by gating on these cells and examining geometric MFI of surface MHC-II staining. C. After gating on live cells the number of mature (CD11chiMHChi) lung cells was calculated by multiplying the percentage of
CD11chiMHChi cells with the total number of live cells, as assessed by trypan blue exclusion. Data in B and C are mean ± SD of 4 independent experiments with 3-4 mice per experiment; #, ##, *, **, p<0.02 compared to uninfected mice.
82 BCG infection does not increase surface expression of CCR7 on lung DCs during peak
and late stages of infection.
Upon maturation, tissue dendritic cells upregulate CCR7 expression and migrate
to secondary lymphoid organs (SLO) and encounter naïve CD4+ T cells (Sallusto,
Schaerli et al. 1998). To determine if maturation of lung DCs during the course of
mycobacterial infection resulted in increased expression of CCR7, lungs were harvested
from uninfected mice and mice that had been infected with aerosolized BCG 4 to 6 weeks
(peak-infection group) or 12 to 14 weeks earlier (late-infection group). Lung cells from
these different groups of mice were sorted with CD11c beads to enrich for lung DCs.
Total RNA was extracted from a total of 2-3×106 CD11c+ sorted cells from the three
groups of mice and used to measure CCR7 mRNA transcript levels by quantitative RT-
PCR (qRT-PCR). BCG infection induced 1.5 to 2 fold increase in CCR7 gene expression in lung CD11c+ cells compared to uninfected mice (Fig. 4.2A). In addition, CCR7 gene
expression on lung macrophages (CD11chi CD11b-) and lung myeloid DCs (CD11chi
CD11bhi) was assessed using flow cytometry. Although lung macrophages from the three
groups of mice did not stain for CCR7, lung DCs expressed small amounts of CCR7 (Fig.
4.2B). However, there was minimal to no change in CCR7 surface expression on lung
DCs from BCG-infected and uninfected mice. Thus, accumulation of DCs in the lung during infection increased the percentage of CCR7+ cells in the lung as seen by qRT-PCR
without causing significant shift in mean fluorescent intensity (MFI) as determined by
flow cytometry. Surface expression of CCR7 was also determined by staining with
CCL19-Fc (eBioscience); however, no evident shift in MFI was noted among the 3
groups using this staining reagent as well (data not shown). Thus activated, mature lung
83 A. B. 0.4 hi hi hi - CD11c CD11b CD11c CD11b 100 100 0.3 80 80 isotype Uninf. 60 60 0.2 BCG % of Max 40 40 % of Max 4-6 wks CCR7 mRNA 0.1 20 20 BCG (Fold over GAPDH) 12-14 wks 0 0 2 3 4 5 2 3 4 5 0 0 10 10 10 10 0 10 10 10 10 Uninf. 4-6 wks 12-14 wks CCR7 CCR7 Time, post-infection
Figure 4.2. CCR7 mRNA and surface protein expression on lung DCs are not increased during peak and late stages of BCG infection. A. Lung CD11c+ cells were sorted from uninfected mice and infected mice either 4-6 wks or 12-14 wks after BCG infection. Total RNA was extracted from the CD11c+ cells and used to perform quantitative RT-PCR for CCR7 gene expression relative to GAPDH. Results are representative of 2 independent experiments (n=3 mice per group) B. Lung CD11c+ sorted cells were stained for CCR7 (clone 4B12), CD11b and CD11c and gated on live cells. CCR7 expression on DCs
(CD11chiCD11bhi) and macrophages (CD11chiCD11b-) are shown. Representative plots from 3-5 experiments.
84 DCs did not upregulate CCR7 surface expression during early and late stages of BCG
infection.
BCG infection upregulates expression of CCL19 in lung in a MyD88-dependent manner.
Like other chemokine receptors, CCR7 surface expression is modulated by its cognate ligands CCL19 and CCL21 via ligand-induced down-regulation (Bardi, Lipp et al. 2001; Otero, Groettrup et al. 2006). To determine if lack of up-regulation of CCR7 surface protein expression during BCG infection was due to expression and binding of its cognate ligands in infected lungs, RNA from lungs of uninfected and BCG-infected mice was reverse transcribed, and CCL19 and CCL21 measured by quantitative RT-PCR.
BCG infection induced gene expression of CCL19 but not CCL21 in the lungs (Fig.
4.3A). Further, mice infected for 12 to 14 wks expressed higher levels of CCL19 mRNA compared to mice infected for 4 to 6 wks (p=0.01). Thus, expression of CCL19 may have inhibited up-regulation of CCR7 on lung DCs during BCG infection as suggested by results.
CCL19 expression in mouse lungs occurs after infection with virulent
Mycobacterium tuberculosis as well (Schreiber, Ehlers et al. 2006; Kahnert, Hopken et al.
2007). In both studies lung mycobacterial burden remained around 106/mouse for the
duration of the endpoints examined. It was surprising that BCG infection upregulated
expression of CCL19 mRNA 12 to 14 wks after infection, when the lung bacterial burden
had been markedly reduced (Fig. 4.1A). To determine if recognition of whole
mycobacteria or mycobacterial products by Toll-like receptors (TLRs) had a role in
85 A. 1.2 Uninf. BCG 4-6 wks 1 * BCG 12-14 wks
0.8
0.6
0.4 Fold Over GAPDH 0.2
0 CCL19 CCL21
B. 50 # CCL19 40
30
20
10 Fold Over Uninfected Mice 0 wt MyD88-/-
Figure 4.3. BCG infection upregulates expression of CCL19 mRNA in lung in a MyD88-dependent manner. RNA was extracted from the right lower lobe of lungs from uninfected and infected mice. Yi Liu, graduate student in Dr. Harding’s lab, performed quantitative RT-PCR using 1 μg of total RNA and transcript levels were normalized to GAPDH. A. BALB/c mice were infected with aerosolized BCG.
Induction of CCL19 and CCL21 in lungs; *, p=0.01 between the two infected groups 4-6 wks and 12-14 wks. B. C57BL/6 mice and MyD88-/- were infected with aerosol of BCG. 4-6 wks post-infection right lower lung lobes were harvested and used to quantitate CCL19 induction over that of uninfected mice; #, p=0.04 between wt and MyD88-/-. For all qRT-PCR data gene expressions of individual mice from each group were used to calculate mean ± SD, (n=3-5 mice per group). Representative of 2 independent experiments. 86 increased expression of CCL19 mRNA in BCG-infected mice, wild-type C57Bl/6 and
MyD88-/- mice were infected with BCG. MyD88-/- mice are deficient in recognition of bacteria and bacterial products through multiple TLRs including TLR2 and TLR9, shown to play a role in mycobacterial infection (Bafica, Scanga et al. 2005). BCG-induced
CCL19 mRNA expression was MyD88-dependent (Fig. 4.3B). These data support the importance of MyD88 for CCL19 expression during mycobacterial infection in light of recent evidence that MyD88 along with other TLR adaptors and cytokines can regulate
CCL19 promoter (Pietila, Veckman et al. 2007).
BCG infection modulates CCR7-mediated chemotaxis of lung DCs.
The lack of CCR7 upregulation on the surface of DCs and the enhanced expression of CCL19 in infected mice suggested that CCR7-mediated migration of lung
DCs to the draining lymph node could be affected by BCG infection. To measure migration, we used transwell assays to examine the ability of lung DCs from infected and uninfected mice to migrate in response to the CCR7 ligand, CCL21. CCL19 and CCL21 have similar affinity for CCR7. Since CCL19 causes greater down-regulation of CCR7 and could interfere with CCR7-mediated chemotaxis of lung DCs we used CCL21 to measure CCR7 function (Bardi, Lipp et al. 2001; Otero, Groettrup et al. 2006).
CD11chiMHChi mature DCs from BCG-infected lungs migrated to CCL21 in greater numbers compared to lung DCs from uninfected mice (Fig. 4.4A). Interestingly, lung
DCs from mice infected 12 to 14 wks earlier migrated even more than lung DCs from mice infected for 4 to 6 wks (p=0.04), even though the lung DCs expressed similar levels of CCR7 and MHC-II (Fig. 4.1-4.2). In addition, the accumulation of mature lung DCs
87 A.
CCL21 Medium 450 400 350
hi 300 250 200 + 150 100 # of transmigrated
CD11c MHC cells CD11c 50 0 Uninf 4-6 wks 12-14 wks B. Duration of BCG infection
5
4
3 #
2 Chemotaxis Index 1
0 (normalized to % of input lung DCs) Uninf 4-6 wks 12-14 wks Duration of BCG infection
Figure 4.4. BCG infection modulates CCR7-mediated chemotaxis of lung DCs. A. Lung CD11c+ cells
(4×105) were placed on 8 μm polycarbonate inserts and allowed to transmigrate towards CCL21 (100 ng/ml) for 5 h at 37˚C. Migrated cells were collected from the bottom chamber and stained for CD11b,
CD11c, and MHC-II (I-Ad). Number of transmigrating cells was calculated using the formula: (Total number of reference beads acquired) × (% of transmigrating cells) / (% of beads acquired). B. Data from
Fig. 4.4A was expressed as a chemotaxis index normalized to the % of input CD11chiMHChi lung cells:
(CD11chiMHChi cells migrating to CCL21) / ( CD11chiMHChi cells migrating to medium alone).
Chemotaxis assays representative of 3 similar experiments; #, p=0.03 (n=3-4 mice per group).
88 in infected mice could be responsible for the greater numbers of DCs migrating to CCL21
in infected mice.
To compare the chemotactic response of individual mature lung DCs (CD11chi
MHChi) among the three groups of mice, data in Fig. 4.4A were normalized to a
chemotaxis index as described in Materials and Methods (Fig. 4.4B). Mature lung DCs
isolated during peak BCG-infection had a lower chemotaxis index compared to mature
lung DCs from mice infected for 12 to 14 wks (p=0.03). Thus, during peak BCG
infection (4 to 6 wks) CCR7-mediated chemotactic responses of mature lung DCs were
slightly blunted at an individual-cell basis compared to uninfected mice even though
accumulation of mature DCs in the lung led to greater numbers of migrating DCs.
BCG infection alters ability of lung CD11c+ cells to present peptide to naïve CD4+ T cells in vitro.
The increased number of mature DCs in BCG-infected lungs along with expression of naïve T cell chemoattractant, CCL19 suggests that during infection naïve T cell-DC interaction could occur in infected lungs and result in naïve T cell activation. To measure the antigen presenting cell (APC) function of lung DCs and their ability to activate naïve CD4+ T cells, lung CD11c+ cells from BCG-infected and uninfected mice
low were pulsed with OVA323-339 peptide and incubated with FACsorted naïve (CD44
CD62Lhi) CD4+ T cells specific for OVA from DO11.10 TCR transgenic mice. Naïve T
cell activation was assessed by measuring IL-2 production. Lung CD11c+ cells from
BCG infected mice caused greater naïve CD4+ T cell activation (Fig. 4.5A). T cell
activation was greater using lung CD11c+ cells from mice infected for 12 to 14 wks than
89 A. uninf BCG-6wks BCG-12wks 1400 1200 1000 800 600 IL-2 (pg/ml) 400 200 0 100 1000 10000 100000 # lung CD11c+ cells B.
350 300 250 200 150 IL-2 (pg/ml) 100 50 0 10 100 1000 10000 + hi # CD11c MHC cells
Figure 4.5. BCG infection alters the ability of lung CD11c+ cells to present peptide to naïve CD4+ T cells in vitro. Lung CD11c+ cells from uninfected mice and mice infected with BCG for 4-6 wks or 12-14 wks
o were pulsed with OVA323-339 peptide (1 μM) for˚ 90 min at 37 C. Cells were washed with culture medium and incubated with FACSorted naïve (CD44lowCD62Lhi) CD4+ T cells (5×104) from DO11.10 mice. After
48 h culture supernatants were tested for IL-2 production by ELISA. (A) Data expressed as increased numbers of lung CD11c+ cells added. Wells without naïve OVA-specific T cells had undetectable levels of
IL-2 (data not shown). (B) Percentage of CD11chiMHChi cells present within the lung CD11c+ cells from the 3 groups of mice was multiplied by number of CD11c+ lung cells added per well to obtain number of
CD11chiMHChi cells added to each well. Data is representative of 2 independent experiments. 90 using lung CD11c+ cells from uninfected mice and mice infected 4 to 6 wks earlier,
p=0.01 and p=0.02 respectively. Interestingly, the number of mature CD11chi MHChi lung DCs in mice infected for 4 to 6 wks is comparable or greater than the number in mice infected for 12 to 14 wks (Fig. 4.1B). Thus, to normalize APC function of lung
CD11c+ cells at 4 to 6 wks post infection, the data in Fig 4.5A was expressed to show the
number of mature (CD11chi MHChi) lung DCs added to the in vitro T cell activation
assay. During peak BCG infection (4 to 6 wks) mature lung DCs activated naïve CD4+ T
cells less than during late infection (12 to 14 wks), p=0.01 (Fig. 4.5B). The same pattern
of T cell activation was observed when the data in Fig 4.5A were normalized to the
number of myeloid (CD11bhi CD11chi) DCs added (data not shown). Our data suggest
that BCG infection makes the lung permissive to naïve CD4+ T cell activation in situ.
Increased numbers of mature lung DCs during infection lead to enhanced activation of naïve CD4+ T cells, although naïve T cell priming is modulated by the stage of
mycobacterial infection (Fig. 4.5B).
Discussion
Our prior study of naïve CD4+ T-cell responses during pulmonary mycobacterial
infection led us to focus on the role of lung dendritic cells in priming naïve CD4+ T cells
(Anis, Fulton et al. 2007). Immune responses ordinarily involve activation of naïve T cells in the organized microenvironment of secondary lymphoid organs (SLO) (Jenkins,
Khoruts et al. 2001; Luther, Ansel et al. 2003). Recently, investigations have demonstrated that mycobacterial and influenza infections lead to the development of neolymphoid structures in the lung that could serve as potential sites for priming naïve T
91 cells (Moyron-Quiroz, Rangel-Moreno et al. 2004; Schreiber, Ehlers et al. 2006; Kahnert,
Hopken et al. 2007). We report here that pulmonary mycobacterial infection caused
accumulation and maturation of DCs in the lung that persisted beyond peak bacterial
burden for up to 12 to 14 wks post-infection. Accumulation of mature DCs in infected
lungs was associated with reduced CCR7-mediated chemotaxis of mature lung DCs
during peak infection (4 to 6 wks post-infection) while surface expression of CCR7
remained similar between infected and uninfected mice. Mycobacterial infection induced
expression of the CCR7 ligand, CCL19 in the lungs through a MyD88-dependent
pathway. OVA-specific naïve CD4+ T cells were activated more by lung CD11c+ cells
from BCG-infected mice. Lung DCs during peak infection (4 to 6 wks post-infection)
caused less naïve T cell activation compared to DCs during late infection (12 to 14 wks post-infection).
Stimulation of Toll-like receptors by mycobacterial products leads to maturation of dendritic cells (Dieu, Vanbervliet et al. 1998; Pecora, Gehring et al. 2006). BCG infection caused accumulation and maturation of lung DCs throughout infection even as lung bacterial burden declined. Although BCG infection induced higher surface expression of MHC-II on lung myeloid DCs, it minimally influenced CCR7 surface expression. Moreover, CCR7-mediated chemotaxis of mature lung DCs was reduced when bacterial burden peaked in the lungs. Reduction in CCR7-mediated chemotaxis may result in decreased migration of mature lung DCs to draining lymph nodes and increased retention in the lungs.
Naïve CD4+ T cells are present in both lymphoid and non-lymphoid organs such
as the lungs (Cose, Brammer et al. 2006). Formation of bronchus-associated lymphoid
92 tissue (BALT) during pulmonary mycobacterial infection may facilitate encounters
between such naïve T cells and lung dendritic cells (Schreiber, Ehlers et al. 2006;
Kahnert, Hopken et al. 2007). Expression of CCR7 ligands, CCL19 and CCL21, is
associated with maturation and organization of neolymphoid follicles (Fukuyama,
Nagatake et al. 2006). Pulmonary BCG infection induced expression of CCL19 in the
lung. However, CCR7-signaling is not essential since mice deficient in CCR7 or its
ligands CCL19 and CCL21-ser (plt/plt) develop BALT during mycobacterial and influenza infections (Moyron-Quiroz, Rangel-Moreno et al. 2004; Kahnert, Hopken et al.
2007). Recently it has been demonstrated that formation of BALT in mice and the T-cell responses elicited in such animals are directly affected by the lack of CCR7-mediated migration of regulatory T cells (Kocks, Davalos-Misslitz et al. 2007).
We had previously shown that lung inflammation during peak BCG infection
caused enhanced pulmonary antigen-specific CD4+ T-cell responses in vivo (Anis, Fulton
et al. 2007). The enhanced CD4+ T-cell responses in infected lungs could have occurred
due to naïve T cell priming in situ secondary to increased naïve T cell recruitment to the lung and retention of mature DCs in the lung. In the present study, OVA-specific naïve
CD4+ T cell activation by lung DCs was less during peak infection (4 to 6 wks) compared
to late infection (12 to 14 wks). This may represent modulation of DC function during
peak mycobacterial infection that subsides as the infection is controlled. Schreiber et. al.
have also reported that T cells, which are unable to migrate to lymph nodes, are recruited
to the lungs during pulmonary mycobacterial infection where they become activated and
acquire effector functions after a delay (Schreiber, Ehlers et al. 2006). Perhaps, delayed
93 acquisition of effector function by T cells in the latter study reflect mycobacteria-induced modulation of lung DC function.
Modulation of CCR7-mediated migration of dendritic cells may occur at multiple levels. Ligand-induced receptor down-regulation is a common mechanism used by chemokine receptors, although CCR7 is more resistant to this form of regulation compared to other chemokine receptors (Sallusto, Palermo et al. 1999). Once bound by its ligand, CCR7 is endocytosed and recycled back while bound ligand dissociates and gets degraded (Otero, Groettrup et al. 2006). The expression of CCL19 in the lungs of
BCG-infected mice indicated that this mode of receptor down-regulation might play a role. However, reduction in chemotaxis during peak infection was not associated with reduced surface expression of CCR7. In addition, mice had higher expression of CCL19 in the lungs during 12 to 14 wks but mature lung DCs from these mice had greater responsiveness to CCL21 compared to mice infected for 4 to 6 wks. Another possible mechanism may involve the CCX-CKR decoy receptor that efficiently scavenges CCL19,
CCL21, and CCL25 (Comerford, Milasta et al. 2006). CCX-CKR does not couple to down-stream signaling components; instead, it acts as a chemokine sink. BCG infection may modulate the expression of CCX-CKR and in an in vitro transwell assay could limit the amount of CCL21 available for CCR7 to bind and respond to.
In summary, BCG infection caused accumulation and maturation of dendritic cells in the lungs. Surprisingly, chemotaxis of mature lung DCs was reduced during peak bacterial burden. Our results do not exclude the possibility that acute stages of mycobacterial infection increases migration of lung DCs to the lymph node as others have seen with bone-marrow derived DCs (Bhatt, Hickman et al. 2004). Mycobacteria
94 have been found inside lymph node dendritic cells but the origin of such DCs was not determined (Humphreys, Stewart et al. 2006). The ability of infection to impede migration of DCs is not exclusive to mycobacterial infection. Legge et. al. have shown that lung DCs during pulmonary virus infection become refractory to migration even as inflammation persists and viral replication continues to occur (Legge and Braciale 2003).
Therefore, the modulation seen in mature lung DC chemotaxis during mycobacterial infection may involve a common set of pathways that recognize PAMPs from different
pathogens. Additional studies will determine how DCs in the lung become refractory to
CCR7-mediated migration and activate naïve CD4+ T cells during mycobacterial
infection.
95
CHAPTER 5
Discussion
96 The studies presented in this dissertation suggest that inflammation during
pulmonary infection may overcome mycobacterial modulation of lung dendritic cell
function to cause adequate naïve CD4+ T-cell responses in the lungs. The studies in this
thesis employed in vivo and in vitro approaches to examine the initiation of naïve CD4+
T-cell responses in the lungs during pulmonary Mycobacterium bovis BCG infection.
BCG is an attenuated mycobacterial strain that establishes an infection in mice with lung
bacterial burden peaking at 4 to 6 wks and declining by 12 to 14 wks. Due to the
temporal nature of an infection, pathogen associated molecular patterns (PAMPs) that
stimulate the immune system at the onset of infection may suppress it in the setting of
established infection. We were thus interested in examining immune responses in the
lungs during prolonged pulmonary infection. Lungs are classically considered to be
immune-privileged organs that suppress pulmonary inflammation although recent
investigations suggest otherwise.
Our investigations in Chapter 2 demonstrated that naïve CD4+ T-cell responses
could be generated in the lungs during mycobacterial infection based on in situ proliferation of antigen-specific CD4+ T cells specific for an airway antigen. To establish
the relevance of our observations to pathogen-specific T-cell responses during infection,
we described our efforts to generate a MTB-specific T cell receptor transgenic mouse in
Chapter 3. Finally, in chapter 4 we indirectly demonstrated that there were increased
encounters between mature lung dendritic cells and naïve CD4+ T cells during BCG infection although mature lung DCs had reduced chemotaxis and APC function. In light of our findings we deduce that lung inflammation may overcome modulation of mature
97 lung DC function during BCG infection by a yet unknown mechanism to cause adequate
CD4+ T-cell responses in the lungs as described in Chapter 2.
Inflammation is essential for generation of naïve CD4+ T-cell responses that
further differentiate into effector and memory T cells and confers protective cell mediated
immunity to host (Jenkins, Khoruts et al. 2001). To understand role of lung
inflammation, we adoptively transferred naïve OVA-specific CD4+ T cells into BCG-
infected and uninfected mice during peak infection (4 to 6 wks). Recipient mice were
subsequently challenged with intranasal OVA. The ensuing OVA-specific T cell response was measured in lung, draining mediastinal lymph node (MLN), and spleen.
OVA-specific T cell response was most prominent in MLN and least prominent in spleen
three days after intranasal OVA challenge. However, the difference in OVA-specific T
cell response between BCG-infected and uninfected mice was dramatic in the lungs.
Upon OVA challenge, adoptively-transferred naïve OVA-specific CD4+ T cells became
activated and proliferated more in the lungs of infected mice than uninfected mice. This proliferation occurred in draining lymph node and lung over the course of 3 days (Fig.
2.3) and was detected in situ in the lungs of mostly infected animals (Fig. 2.4).
Detection of in situ proliferation in the lungs of infected mice indicated that naïve
CD4+ T cells may become activated there during infection. Priming of naïve CD4+ T cells in the lungs is postulated to occur during mycobacterial and other lung infections due to the presence of neolymphoid BALT-like structures in infected lungs (Moyron-
Quiroz, Rangel-Moreno et al. 2004; Schreiber, Ehlers et al. 2006; Kahnert, Hopken et al.
2007). To determine which antigen presenting cells in the lung may be responsible for priming, Fluos-OVA was introduced intranasally into BCG-infected and uninfected mice.
98 The labeled OVA was taken up primarily by CD11c+ lung cells and distributed more to lung myeloid DCs (CD11bhiCD11chi) in infected mice than in uninfected mice (Fig.
2.7B). Additionally, ex vivo OVA presentation was better using lung CD11c+ cells from infected mice (Fig. 2.7E). This indicated that during infection airway antigens are presented to naïve CD4+ T cells by larger numbers of CD11chiMHChi lung DCs than in uninfected mice (Fig. 2.7D), leading to more T cell proliferation in infected lungs. As a result, in situ proliferation of OVA-specific T cells in infected lungs was within limit of detection of the BrdU assay (Fig. 2.4). Additionally, expression of CCL19, a naïve T cell chemoattractant, is induced in the lungs upon mycobacterial infection (Fig. 4.3A)
(Kahnert, Hopken et al. 2007) thereby recruiting naïve CD4+ T cells to encounter antigen-laden mature (CD11chiMHChi) DCs in the lungs to become activated. As mentioned, a fundamental effect of inflammation on immune responses is differentiation of T cells once they have become activated. In BCG-infected lungs OVA-specific T cells were twice as likely to change their phenotype to that of activated effector T cells
(CD62Llow) that produced more IFN-γ (Fig. 2.5, 2.6).
The enhancements in naïve CD4+ T-cell responses were associated with increased numbers of mature dendritic cells in infected lungs during peak infection (Fig. 4.1B).
The accumulation of mature lung DCs persisted even as lung bacterial burden declined
1000 fold between 4 to 6 wks and 12 to 14 wks of infection (Fig. 4.1A). Presumably, continued recognition of mycobacterial PAMPs by innate immune cells was responsible for the heightened state of maturation of APCs even as bacterial burden declined.
Likewise, others have shown that continued presentation of pathogen-derived antigens, long after pathogen clearance, contributes to priming naïve CD4+ T-cell responses
99 (Jelley-Gibbs, Brown et al. 2005). Thus, even after 12 to 14 wks post-infection
inflammation may be present in the lungs of BCG-infected mice. The term inflammation is being used here to convey both recognition of PAMPs by immune cells and inflammatory mediators produced as a result of that stimuli. Medzhitov, Sousa et. al.
have shown that recognition of PAMPs and not inflammatory cytokines is necessary for
activation of DCs to generate effector T-cell responses (Sporri and Reis e Sousa 2005)
(Iwasaki and Medzhitov 2004).
In Chapter 4 we postulated that the increased numbers of lung dendritic cells in
BCG-infected mice lead to enhanced migration of DCs to secondary lymphoid organs to
initiate naïve CD4+ T-cell responses. Migration of mature DCs to secondary lymphoid
organs is mediated by C-C chemokine receptor CCR7 on DCs, and the CCR7 ligands
CCL19 and CCL21. Therefore, the expression and functionality of CCR7 was examined
in lung DCs from uninfected mice and mice infected for 4 to 6 wks and 12 to 14 wks.
The later time point during infection was chosen to examine if bacterial burden
influenced migratory capacity of lung DCs. Functionality of CCR7 was determined
using transwell assays to measure chemotaxis of mature lung DCs to CCL21. This
obviated some difficulties in interpreting in vivo migration data where the specificity of the migration towards CCR7 ligands is not obvious without extensive controls. Surface expression of CCR7 was not different between myeloid DCs from infected and uninfected mice (Fig. 4.2B). Additionally quantitative RT-PCR showed that gene expression of CCR7 was only slightly increased in lung CD11c+ cells from infected mice
(Fig. 4.2A). However, upon BCG infection, transmigration of mature lung DCs towards
CCR7 ligand CCL21 was greater (Fig. 4.4A). But this could have been a result of greater
100 numbers of mature lung DCs present among the cells, from infected mice, used as input
for the transwell assay. When the transmigration data was expressed as a chemotaxis
index taking into consideration % of CD11chiMHChi cells used as input, mature lung DCs
from infected mice had unexpectedly lower chemotaxis index compared to uninfected
mice (Fig. 4.4B). The chemotaxis of mature lung DCs from mice infected for 12 to 14
wks returned back to the level of uninfected mice.
The reduction in APC function of mature (CD11chiMHChi) lung DCs from mice infected for 4 to 6 wks was at odds with the in vivo data indicating that during peak BCG infection naïve CD4+ T cell activation was enhanced in the lung and draining lymph node. One explanation for the disparity may be that apparent reduction in APC function of mature lung DCs is minor and in vivo inflammation may overcome this modulation to generate adequate CD4+ T-cell responses as observed in Chapter 2. Another possibility is
that our observations in Chapter 2 that there was enhanced naïve CD4+ T cell activation
in the lungs may in fact have represented enhanced naïve T cell activation in the lymph
node and greater recruitment to the inflamed lung.
When comparing the ability of lung CD11c+ cells to present peptide and activate
naïve CD4+ T cells, we found that mature lung DCs from mice infected for 4 to 6 wks
were apparently better at priming than DCs from uninfected mice (Fig. 4.5A). However,
when this data was expressed as the number of CD11chiMHChi cells placed into each well
as APCs instead of number of total CD11c+ cells, we found that mice infected for 4 to 6
wks had a reduced ability to activate naïve CD4+ T cells compared to uninfected mice
(Fig. 4.5B). Moreover, the ability of mature lung DCs from mice infected for 12 to 14 wks to activate CD4+ T cells was similar to that of lung DCs from uninfected mice. An
101 implication of these in vitro studies is that reduction in CCR7-mediated chemotaxis of
mature lung DCs during peak BCG infection may increase encounters between naïve
CD4+ T cells and mature DCs in the lungs. Naïve CD4+ T cells would be recruited to the
lungs due to MyD88-dependent expression of CCL19 in BCG-infected lungs (Fig. 4.3B).
Even though such encounters would result in sub-optimal activation of naïve CD4+ T cells (Fig. 4.5B), the inflammatory environment in vivo may overcome this BCG-induced modulation of mature lung DC function. Others have shown that effector CD4+ T cell
functions in the lungs of MTB infected mice with deficiencies in lymph node homing are
delayed but eventually catch up to that of wild-type infected mice (Schreiber, Ehlers et al.
2006). The mechanisms that cause BCG-induced modulation of mature lung DC
function and those that overcome it are yet unknown. Our observations so far have
helped us construct a model of pulmonary CD4+ T-cell responses during mycobacterial
infection (Fig. 5.1).
BCG-induced modulation of mature lung DC function correlated with bacterial
burden – more mycobacteria in the lungs led to greater reduction in migration of mature
lung DCs and their ability to activate naïve CD4+ T cells. Additional experiments need to
be performed to establish this relationship. One possible experiment will be to compare mice infected for 4 to 6 wks that are left untreated or treated with antibiotics to clear the
bacteria. Lung CD11c+ cells from the two groups of mice will be used in chemotaxis
and APC experiments to compare BCG-induced modulation of lung DC function. If
these experiments show that viability of bacteria is necessary for the modulation of lung
DC function, then that suggests recognition of bacteria or mycobacterial products could
also be necessary. Toll like receptors are one type of pattern recognition receptors that
102 A.
naïve CD4+ Lung MLN MΦ
CCL19 CCL21
Lung DC
B.
naïve naïve CD4+ CD4+ Lung MΦ
naïve CD4+ lung TLO
CCL19 lung DC CCL21
lung DC lung DC
Figure 5.1. Activation of naïve CD4+ T cells by lung dendritic cells. A. Naïve CD4+ T cells recirculate primarily between SLOs such as mediastinal lymph node (MLN) and the blood during homeostasis. Upon engagement of TLRs and other PAMP receptors on DCs in the periphery, DCs upregulate CCR7 expression and migrate to SLOs along with naïve T cells to initiate cell-mediated immune responses against invading pathogens. B. During mycobacterial infection, TLOs such as BALT begin to be organized in the lung tissue – the organization being aided by production of homeostatic chemokines: CCL19, CXCL13, and CCL21. Lung DCs become refractory to further CCR7-directed migration. Naïve CD4+ T cells use their CCR7 receptors to arrive towards TLOs producing CCR7 ligands. Stromal cells and B cells (not shown) are important organizers of the TLO. This setting allows for interaction between lung DCs and naïve CD4+ T cells that could result in naive T cell activation.
103 mediate innate recognition of pathogens. The in vitro experiments on chemotaxis and
APC function could be recapitulated using wild-type and MyD88-/- mice. MyD88-/- is a
common adaptor molecule used by many TLRs including TLR2 and TLR9, both of
which have been found to play a role during mycobacterial infection (Bafica, Scanga et
al. 2005).
The next step will be to reproduce the modulation of mature lung DC function in vitro so that additional studies could be done to derive a mechanism. To perform these studies primary lung DCs could be used although yield and purity of lung DCs would be much less than conventional bone-marrow derived DCs. Also, lung CD11c+ resident macrophages will need to be separated to specifically look at myeloid (CD11chiCD11bhi)
DCs. Recently, a report outlined generation of primary lung DCs (Wang, Peters et al.
2006) by cell culture, however it has met with criticism that resident lung macrophages
were not separated in the beginning to prevent contamination (von Garnier, Filgueira et
al. 2005). The published protocol may be modified then as follows: 1) lung cells are
separated by CD11c beads (Miltenyi Biotec.) to enrich for lung DCs, 2) lung CD11c+
cells are separated by CD11c and CD11b staining using BD Aria FACSorter, 3)
CD11bhiCD11chi lung cells are cultured in the presence of 20 ng/ml GM-CSF and
harvested 12 days later. However, the yield may be too small for protein extraction and analysis of cell signaling. A simpler protocol, which would contain contaminating lung
resident macrophages, will be to culture lung cells in 20 ng/ml GM-CSF for 12 days and then sorting the cells with CD11c beads.
Once a specific type of cultured DC is chosen, the design of the experiment will be to stimulate DCs with live mycobacteria, mycobacterial cell lysate, a well-
104 characterized mycobacterial PAMP such as LpqH (19-kDa lipoprotein), a commercial
TLR2 agonist Pam3Cys and a known inflammatory mediator with activity on DCs, TNF-
α. Dendritic cells will be washed and used in transwell assays to determine chemotaxis
towards CCL21. Upon establishing in vitro conditions pertaining to strength and
duration of stimulation necessary to elicit the reduction in chemotaxis seen ex vivo, cell
signaling studies can commence.
During peak BCG infection, chemotaxis of mature lung DCs to CCL21 and
activation of naïve CD4+ T cells by lung DCs are reduced. Surface expression of CCR7
and MHC-II are similar between mature lung DCs from all 3 groups of infected and
uninfected mice. Additionally, expression of costimulatory molecules, CD80 and CD86,
are slightly above isotype levels on lung DCs and are not different among the three
groups. The observed modulation in migration and APC function could then be the result
of intracellular modifications of cell signaling pathways induced by peak BCG infection.
Modulation of both DC functions may be independent of one another. But, there are many intracellular signaling pathways that impact on both. Recently, CCR7 signaling has been characterized as being modular to reflect the diverse functions governed by ligand binding to the receptor (Fig. 5.2) (Sanchez-Sanchez, Riol-Blanco et al. 2006).
Even though the modular nature is based on studies using pharmacologic inhibitors, the pathways are very much interdependent physiologically. One pathway is CCR7- mediated activation of mitogen-activated protein kinase (MAPK) family members p38,
Erk1/2, and JNK. Activation of all three MAPK family members is responsible for chemotaxis of DCs. A related but separated pathway controls migratory speed, random motion of the cell, that is regulated by Rho GTPases. In our transwell experiments, the
105
Figure 5.2. Chemokine receptor CCR7-signaling pathway (Sanchez-Sanchez, Riol-Blanco et al. 2006).
106 measured transmigration was comprised of both chemotaxis and migratory speed. Thus
both pathways could be modulated during peak BCG infection. The MAPK pathway is
downstream of many TLRs and has been shown to be important for mycobacterial
LpqH’s inhibitory activity in antigen processing and presentation by macrophages
(Pennini, Pai et al. 2006). Furthermore, Rho GTPases are important for cytoskeleton rearrangements and formation of dendrites that increase contact area between T cells and dendritic cells and facilitate T cell activation (Kobayashi, Azuma et al. 2001). In light of this, both modulation of mature lung DC migration and ability to activate naïve CD4+ T cells may be affected by MAPK and Rho GTPase pathways.
To examine the relevance of the MAPK pathway on our observation ex vivo, lung
CD11c+ cells from uninfected and infected mice (4 to 6 wks) and (12 to 14 wks) will be lysed and extracted by boiling in SDS-PAGE sample buffer and subsequently separated in SDS-PAGE and analyzed by Western blot. Antibodies against phospho-ERK1/2, phospho-p38, and phosphor-JNK along with antibodies against the unphosphorylated forms of these MAPK family members will be used to probe the blot. To measure RhoA activity lung CD11c+ cells from the 3 groups of mice will be dissolved in Rho lysis buffer and active GTP-Rho will be precipitated using Rho Activation Assay Kit (Upstate
Biotechnology). The bead pellets can then be boiled in SDS-PAGE sample buffer and separated by SDS-PAGE for Western blot using anti-RhoA Ab. Both protocols along with more detail has been published (Riol-Blanco, Sanchez-Sanchez et al. 2005; Shurin,
Tourkova et al. 2005). Aside from RhoA other Rho family of GTPases such as Cdc42 and Rac may play a role. These experiments will establish the ex vivo differences in the pathways involving MAPK and Rho GTPases during pulmonary BCG infection.
107 To determine how the patterns of phosphorylations found ex vivo may have come
about, the in vitro assay described earlier will be used with the same stimuli that
reproduce the reduction in chemotaxis observed. Only difference will be the DCs will be
stimulated in the absence and presence of CCL19 or CCL21 to engage CCR7 and initiate
downstream signaling. Importance of the MAPK and RhoA pathways can be
demonstrated using UO126, SB203580, SP600125, C3 exoenzyme, Y-27632 which
respectively inhibits ERK, p38, JNK, Rho, and Rho-associated kinase inhibitor. These studies will provide critical evidence to support the phenomena that peak BCG infection affects intracellular signaling machinery to influence both lung DC migration and APC function. More importantly, they will provide a framework addressing which mycobacterial components are involved in the modulation and what intracellular pathways are modulated. The studies can be complemented by examining direct interactions between lung CD11chiMHChi cells with naïve CD4+ T cells in vitro using confocal microscopy in the absence and presence of mycobacterial stimuli and CCR7 ligands along with appropriate inhibitors of MAPK and Rho pathways.
Due to the expression of CCL19, naïve CD4+ T cells are likely recruited to
infected lungs. It would be interesting to learn if in the infected milieu of the lungs naïve
CD4+ T cells become affected as do mature DCs with regards to loss of CCR7-guided
migration during peak bacterial burden in the lungs (4 to 6 wks). One would expect
MTB-specific naïve CD4+ T cells to become more affected than other T cells because
they are likely to undergo TCR mediated arrest in the lungs once they encounter lung
DCs presenting MTB-derived cognate peptides, while T cells with irrelevant antigen-
specificities “filter” through the lungs. The endeavor to generate the Ag85B TCR
108 transgenic mouse in Chapter 3 was undertaken to address just such questions as these.
Fortunately in the end the hurdles we encountered did not prevent us from obtaining a
MTB-specific TCR transgenic animal: P25 TCR transgenic mouse developed by Takatsu et. al. (Tamura, Ariga et al. 2004).
To determine if P25 TCR CD4+ T cells become refractory to CCR7-guided
signals akin to lung DCs, the experimental design would be to adoptively transfer naïve
P25 T cells with the allelic marker Thy 1.2 along with naïve OT-II OVA-specific CD4+ T cells (Thy 1.2) into Thy 1.1 recipient mice. Before adoptive transfer the recipient mice will be infected with mycobacteria such that at the time of adoptive transfer bacterial burden peaks in the lungs. Also, a day before adoptive transfer the recipient mice will be immunized in the footpad with LPS+OVA. For control, there should be recipient mice that are uninfected and unimmunized – these mice should have naïve T cells of both Thy
1.2 populations throughout the experiment. All recipient mice will be sacrificed 3 days after adoptive transfer and their spleen, lung, mediastinal lymph node, and popliteal lymph nodes harvested. Single cells from these tissues will be stained with KN7 (P25
TCR clonotypic mAb), anti-Thy 1.2 (KN7- Thy1.2+ are OT-II T cells), anti-CD44
(activation marker CD44hi), anti-CD62L and anti-CCR7 mAbs. Localization in the lungs
of activated T cells of both Thy 1.2 populations is not expected to be antigen-driven as
other studies have shown that activated T cells enter inflamed tissues without regard to
antigen presence (Topham, Castrucci et al. 2001). Analyzing the expression of CCR7 on
activated P25 T cells and OT-II T cells in the various tissues would shed light on the
hypothesis that MTB-specific T cells in the lungs have different CCR7 expression and
109 chemotactic response to CCR7 ligands compared to OT-II T cells activated elsewhere in the host.
The FACS analysis will need to be complemented with transwell assays demonstrating if in fact migration of MTB-specific T cells towards CCL21 or CCL19 is different from OT-II T cells. A technical challenge of purifying lung T cells may be overcome by using anti-Thy 1.2 microbeads (Miltenyi Biotec.) and adoptively transferring only 1 of the Thy 1.2 T cell populations for these latter studies. One caveat to this entire experiment is that P25 T cells will most likely be primed in the draining mediastinal lymph node and then enter the lungs. However, they should still be activated in the lungs by lung DCs presenting Ag85B peptides. To increase the signal to noise ratio, a less physiological model: plt/plt mice, which are deficient in naïve T cell homing to lymph nodes due to absence of CCL19 and CCL21-ser, could be used.
Examining P25 TCR tg (MTB-specific) T-cell responses in vivo Wolf et. al. have observed initiation of priming in the mediastinal lymph nodes (MLN) by day 12 post- infection using virulent Mycobacterium tuberculosis (unpublished data). This delay in priming of P25 T cells corresponded with lung bacterial burden reaching a plateau of 106 bacilli/lung on day 14 post-infection. In our experimental model using the attenuated strain M. bovis BCG, we notice bacterial burden in the lungs peaking at 4 to 6 wks.
Analogously, initiation of P25 T cell priming probably occurs at 3 wks post-infection in
MLN of mice infected with BCG. During this period of infection, we have found expression of CCL19 in lungs of infected mice. CCL19 may recruit naïve P25 T cells, some of which have not been primed in the MLN, into the lungs where they become sub- optimally activated by lung DCs. Peak BCG infection (4 to 6 wks) reduces the APC
110 function and CCR7-mediated migration of mature lung DCs. However, inflammation-
driven accumulation of mature (CD11chiMHChi) DCs in the lungs may overcome BCG-
induced modulation of lung DCs. This model would imply that initial trafficking of DCs
to MLN occurs earlier than 3 wks. We have not examined chemotaxis of lung DCs this early during BCG infection. It would be interesting to see if CCR7-mediated trafficking of mature lung DCs occurs early on in infection and then becomes refractory during peak bacterial burden only to return to steady-state levels once bacterial CFUs have declined.
Determining if MTB-specific T cells activated in the lungs in vivo are responsive to CCR7 ligands may be important in light of findings that CCR7 expression is necessary for effector-memory T cells in peripheral tissues to reenter lymph nodes (Bromley,
Thomas et al. 2005). Secondary lymphoid organs are necessary for the differentiation of effector CD4+ T cells to memory T cells (Obhrai, Oberbarnscheidt et al. 2006). It is unclear how these insights into CD4+ T-cell responses in vivo apply to a context where
naïve CD4+ T cell priming may also be occurring in BALT-like structures in the lungs
(Moyron-Quiroz, Rangel-Moreno et al. 2004). We hypothesize that MTB-specific T cells
that are primed or re-activated in the lung are activated less due to the modulation of lung
DC APC function and these T cells, like their DC counterparts, become refractory to
CCR7-mediated migration. Consequently these T cells don’t develop full potential
effector memory and central memory functions necessary to completely eradicate MTB.
The MTB-specific T cells remain at the infection foci having lost ability to reenter
afferent lymphatics using CCR7. Dissemination of bacteria or leakage of MTB-derived
antigens into the periphery by lymphatic drainage leads to efficient priming of MTB-
specific T cells in the periphery. These T cells primed in the periphery develop effector
111 T cell responses and cause delayed type hypersensitivity (DTH) to purified protein
derivative (PPD) of MTB. This proposition rests on the observations that T cells develop
homing properties based on the site where they are primed (Campbell and Butcher 2002),
and that mice deficient in fucosyltransferases have diminished DTH responses but
adequate pulmonary T-cell responses to MTB (Schreiber, Ehlers et al. 2006); thus,
indicating that the T cells mediating the two immune responses are not primed at the
same site.
A teleological model of naïve CD4+ T cell priming during mycobacterial infection
is that lymph nodes are the ‘preferred’ sites where naïve CD4+ T cells are activated.
When priming in the lymph node is compromised perhaps due to deficient homing of naïve T cells to lymph nodes then tertiary lymphoid structures such as BALT become important for priming T cells. Ordinarily, naïve CD4+ T cells arriving at the lymph node
don’t all arrive at once. Instead, there are early arrivers and late arrivers among naïve
CD4+ T cells (Catron, Rusch et al. 2006). We propose that due to limited number of
APCs in the lymph node, some late-arriving naïve CD4+ T cells are instead primed in
BALT, which develops during mycobacterial infection. These naïve CD4+ T cells are
recruited into the lungs by expression of CCL19 during mycobacterial infection and are
activated sub-optimally by mature lung DCs that have become refractory to CCR7-
mediated migration. The reduced migratory capacity of mature lung DCs could increase
the likelihood of naïve CD4+ T cells being activated by DCs in the lungs. BCG-induced
modulation of lung DC function could occur via mediator(s) that are found to be PAMPs,
which suppress immune responses in the setting of an established infection.
Inflammation induced by mycobacterial PAMPs also cause accumulation of mature lung
112 DCs in infected lungs, expression of CCL19, and as others have seen formation of
BALT. These changes induced by inflammation overcome the BCG-induced modulation of dendritic cell function and cause aequate CD4+ T-cell responses. Perhaps virulent
Mycobacterium tuberculosis causes a more substantial modulation of lung DC function that cannot be overcome by inflammation and thus leads to bacterial persistence.
113 Table 5.1 List of Experiments Corresponding to the Presented Figures
Fig. Experiment Fig. Experiment
2.1 MMA255 3.4 MMA112
2.2 MMA383 3.6 MMA134
2.3 MMA346 3.7 MMA188
2.4 MMA371 4.1 MMA472,479,506,528
2.5 MMA397 4.2 MMA481
2.6 A: MMA400, B: MMA403 4.3 A: MMA506, B: MMA504
2.7 A: MMA366, E: MMA379 4.4 MMA532
3.1 MMA4 4.5 MMA508
114
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